Full text of "soil analysis"

R. Margesin F.Schinner (Eds.)

Soil Biology

Series Editor: Ajit Varma

Volumes published in the series

Volume 1

A. Singh, O.P. Ward (Eds.)

Applied Bioremediation and Phytoremediation

2004

Volume 2

A. Singh, O.P. Ward (Eds.)

BlODEGRADATION AND BlOREMEDIATION

2004

Volume 3

E Buscot, A. Varma (Eds.)

Microorganisms in Soils: Roles in Genesis and Functions

2005

Volume 4

S. Declerck, D.-G. Strullu, J.A. Fortin (Eds.)

In Vitro Culture of Mycorrhizas

2005

Rosa Margesin
Franz Schinner (Eds.)

Manual for Soil Analysis
Monitoring and Assessing
Soil Bioremediation

With 3 1 Figures

4y Spri

ringer

Prof. Dr. Rosa Margesin

Leopold Franzens University

Institute of Microbiology

Technikerstr. 25

A-6020 Innsbruck

Austria

e-mail: Rosa.Margesin@uibk.ac.at

Prof. Dr. Franz Schinner

Leopold Franzens University

Institute of Microbiology

Technikerstr. 25

A-6020 Innsbruck

Austria

e-mail: Franz.Schinner@uibk.ac.at

Library of Congress Control Number: 2005926091

ISSN 1613-3382

ISBN-10 3-540-25346-7 Springer Berlin Heidelberg New York

ISBN-13 978-3-540-25346-4 Springer Berlin Heidelberg New York

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Preface

The increasing use of soil bioremediation technologies requires new con-
cepts and methods to assess the feasibility of a remediation technology
and to monitor the success of the treatment. The knowledge of the reac-
tion of the soil microflora to contamination facilitates the optimization of
biodegradation. Manual of Soil Analysis - Monitoring and Assessing Soil
Bioremediation differs from other books on soil analysis in that the moni-
toring and assessing of soil bioremediation are the central themes.

In this comprehensive laboratory manual, sampling, pretreatment and
storage of soil, feasibility studies for soil bioremediation, and the most
important methods to analyze physical, chemical, and biological soil pa-
rameters are presented. Chapters written by experts for those involved in
research, teaching, and routine analyses outline molecular and immunolog-
ical techniques, the use of conserved internal markers, radiorespirometry,
bioreporter technology, the interpretation of fatty acid profiles, soil mi-
crobial and enzymatic methods, and the assessment of ecotoxicity using
bioassays. Particular emphasis has been placed on the comprehensible and
complete description of the experimental procedures. The broad spectrum
of modern soil biological methods provides an excellent complementation
of traditional soil investigation and characterization. Our book, however,
does not claim to present all modern methods available, it rather contains
a selection of the most suitable methods for investigating contaminated
soil. More biological methods can be found in our volume Methods in Soil
Biology (Schinner, Ohlinger, Kandeler and Margesin 1996, Springer).

We are most grateful to the authors for their excellent contributions and
to Springer, especially to Dr. Jutta Lindenborn and Dr. Dieter Czeschlik, for
continuous support and cooperation. We also thank Dr. Ajit Varma for the
possibility to publish this book in the Soil Biology Series.

Innsbruck, Austria, Rosa Margesin

January 2005 and Franz Schinner

Contents

1 Soil Sampling and Storage 1

Andreas Paetz, Berndt-Michael Wilke

1.1 Objective of Soil Sampling 1

1.1.1 Principal Objectives 1

1.1.2 Specific Objectives 4

1.2 Selection of Sampling Technique 6

1.3 Sampling Strategy 7

1.3.1 General 7

1.3.2 Preliminary Investigation 7

1.3.3 Exploratory Investigation 9

1.3.4 Main Site Investigation 9

1.3.5 Samples and Sampling Points 10

1.4 Sampling Methods 25

1.4.1 General 25

1.4.2 Type of Sample 25

1.4.3 Undisturbed Samples 27

1.4.4 Cross-Contamination 34

1.4.5 Sampling Containers 34

1.5 Pretreatment 37

1.5.1 Chemical Analysis 37

1.5.2 Physical Analysis 40

1.5.3 Biological Analysis 40

1.6 Storage of Samples 41

1.6.1 General 41

1 .6.2 Specific Considerations for Biological Parameters 42

1.6.3 Preparing the Samples After Storage 44

References 44

2 Determination of Chemical and Physical Soil Properties 47

Berndt-Michael Wilke

2.1 Soil Dry Mass and Water Content 47

2.2 Water-Holding Capacity 50

VIII Contents

2.3 Bulk Density - Total Porosity 52

2.3.1 Core Method 52

2.3.2 Excavation Method 54

2.3.3 Clod Method 57

2.4 Water Retention Characteristics - Pore Size Distribution 59

2.4. 1 Determination of Soil Water Characteristics

Using Sand, Kaolin, and Ceramic Suction Tables 62

2.4.2 Determination of Soil Water Characteristics

by Pressure Plate Extractor 65

2.5 SoilpH 68

2.6 Soil Organic Matter - Soil Organic Carbon 71

2.6.1 Dry Combustion Method 72

2.6.2 Loss On Ignition Method (LOI) 74

2.7 Soil Nutrients: Total Nitrogen 76

2.7. 1 Dry Combustion Method ("Elemental Analysis") 77

2.7.2 Modified Kjeldahl Method 79

2.8 Soil Nutrients: Inorganic Nitrogen 82

2.8.1 Extraction 83

2.8.2 Quantification of Nitrate Nitrogen 84

2.8.3 Quantification of Ammonium Nitrogen 86

2.9 Soil Nutrients: Phosphorus 87

2.9.1 Extraction of Total Phosphorus 88

2.9.2 Extraction of Labile Phosphorus 90

2.9.3 Quantification of Phosphorus 91

References 93

3 Quantification of Soil Contamination 97

Kirsten S. j0rgensen, Olli Jarvinen, Pirjo Sainio, Jani Salminen,
Anna-Mari Suortti

3.1 General Introduction 97

3.2 Volatile Hydrocarbons 99

3.3 Hydrocarbons in the Range C 10 to C 40 103

3.4 Polyaromatic Hydrocarbons (PAHs) 109

3.5 Heavy Metals 115

References 118

4 Immunotechniques as a Tool for Detection of Hydrocarbons 121

Grazyna A. Plaza, Krzysztof Ulfig, Albert J. Tien

4.1 RaPID Assay Test System 121

4.2 EnviroGard Test System 126

References 130

Contents IX

5 Feasibility Studies for Microbial Remediation

of Hydrocarbon-Contaminated Soil 131

Ajay Singh, Owen P. Ward, Ramesh C. Kuhad

5.1 Introduction 131

5.2 Determination of Biodegradation Potential 132

5.2.1 Sampling and Soil Preparation 132

5.2.2 Selective Microbial Enrichment 134

5.2.3 Controls 135

5.2.4 Soil Microcosms 136

5.2.5 Slurry Bioreactors 137

5.2.6 Land Treatment 139

5.2.7 Composting 140

5.2.8 Scale-Up 141

5.3 Process Monitoring and Evaluation 142

5.4 Bioaugmentation 143

5.5 Effect of Surfactants 144

5.5. 1 Screening of Microbial Cultures

for Biosurfactant Production 145

5.5.2 Effect of Biosurfactants 146

5.5.3 Effect of Chemical Surfactants 146

5.6 Optimization of Environmental Conditions 147

5.7 Optimization of Nutritional Factors 148

5.8 Conclusions 150

References 151

6 Feasibility Studies for Microbial Remediation

of Metal-Contaminated Soil 155

Franz Schinner, Thomas Klauser

References 159

7 Feasibility Studies for Phytoremediation

of Metal-Contaminated Soil 161

Aleksandra Sas-Nowosielska, Rafal Kucharski, Eugeniusz Malkowski

7.1 Introduction 161

7.2 Phytoextraction 161

7.2.1 Treatability Study 162

7.2.2 Full-Scale Application 166

7.2.3 Conclusions 170

7.3 Phytostabilization Potential for Soils Highly Contaminated
with Lead, Cadmium and Zinc 171

7.3.1 Evaluation of Site Contaminants 171

7.3.2 Logistic Considerations 172

X Contents

7.3.3 Additives 172

7.3.4 Plants 173

7.3.5 Full-Scale Application 173

7.3.6 Effectiveness of Technology 174

7.3.7 Monitoring 174

7.3.8 Conclusions 175

References 176

8 Quantification of Hydrocarbon Biodegradation

Using Internal Markers 179

Roger C. Prince, Gregory S. Douglas

References 187

9 Assessment of Hydrocarbon Biodegradation Potential

Using Radiorespirometry 189

Jon E. Lindstrom, Joan F. Braddock

References 198

10 Molecular Techniques for Monitoring

and Assessing Soil Bioremediation 201

Lyle G. Whyte, Charles W. Greer

10.1 General Introduction 201

10.2 Extraction and Purification

of Nucleic Acids (DNA) from Soil 202

10.3 Amplification of Catabolic Genotypes

and 16SrDNA Genotypes by PCR 208

10.4 DGGE Analysis Soil Microbial Communities 218

10.5 Genomics in Environmental Microbiology 226

References 228

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 233

Steven Ripp

11.1 General Introduction 233

11.2 An Overview of Reporter Systems

for Soil Bioremediation Application 235

11.3 Single Point Measurements of Soil Contaminants 241

1 1.4 Continuous On-Line Vapor Phase Sensing

of Soil Contaminants 244

1 1.5 Quantification of Soil-Borne lux-Tagged Microbial Popula-
tions Using Most-Probable-Number (MPN) Analysis 247

References 249

Contents XI

12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 251

David B. Hedrick, Aaron Peacock, David C. White

12.1 Obtaining Fatty Acid Profiles from Soil Samples 251

12.2 Transforming Fatty Acid Peak Areas

to Total Microbial Biomass 252

12.3 Calculation and Interpretation of Community Structure 254

12.3.1 Standard Community Structure Method 254

12.3.2 Custom Community Structure Methods 255

12.3.3 Factor Analysis 255

12.4 Calculation and Interpretation

of Metabolic Stress Biomarkers 256

12.5 Naming of Fatty Acids 257

References 258

13 Enumeration of Soil Microorganisms 261

Julia Foght, Jackie Aislabie

13.1 Sample Preparation and Dilution 261

13.2 Direct (Microscopic) Enumeration 264

13.3 Enumeration by Culture in Liquid Medium

(Most Probable Number Technique) 268

13.4 Enumeration by Culture on Solid Medium

(Plate Count Technique) 272

References 279

14 Quantification of Soil Microbial Biomass

by Fumigation-Extraction 281

Rainer Georg Joergensen, Philip C. Brookes

14.1 General Introduction 281

14.2 Fumigation and Extraction 282

14.3 Biomass C 284

14.3.1 Biomass C by Dichromate Oxidation 284

14.3.2 Biomass C by UV-Persulfate Oxidation 286

14.3.3 Biomass C by Oven Oxidation 288

14.4 BiomassN 289

14.4.1 Ninhydrin-Reactive Nitrogen 289

14.4.2 Total Nitrogen 292

References 294

15 Determination of Adenylates and Adenylate Energy Charge 297

Rainer Georg Joergensen, Markus Raubuch

References 302

XII Contents

16 Determination of Aerobic N-Mineralization 303

Rainer Georg Joergensen

References 306

17 Determination of Enzyme Activities in Contaminated Soil 309

Rosa Margesin

17.1 General Introduction 309

17.2 Lipase-Esterase Activity 310

17.3 Fluorescein Diacetate Hydrolytic Activity 313

17.4 Dehydrogenase Activity 316

References 319

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 32 1

Adolf Eisentraeger, Kerstin Hund-Rinke, Joerg Roembke

18.1 General Introduction: Strategy 321

18.2 Sample Preparation 323

18.3 Water- Extractable Ecotoxicity 330

18.3.1 Vibrio fischeri Luminescence-Inhibition Assay 330

18.3.2 Desmodesmus subspicatus Growth- Inhibition Assay.... 331

18.4 Water-Extractable Genotoxicity 332

18.4.1 The umu Test 332

18.4.2 Salmonella/Microsome Assay (Ames Test) 333

18.5 Habitat Function:

Soil/Microorganisms, Soil/Soil Fauna, Soil/Higher Plants 334

18.5.1 Respiration Curve Test 334

18.5.2 Ammonium Oxidation Test 337

18.5.3 Combined Earthworm Mortality/Reproduction Test ... 340

18.5.4 Collembola Reproduction Test 342

18.5.5 Plant Growth Test 344

18.5.6 Test Performance for the Derivation

of Threshold Values 346

18.6 Combined Performance of Bioassays and Assessment of the
Results 348

18.6.1 Water-Extractable Ecotoxic Potential 348

18.6.2 Water-Extractable Genotoxicity 349

18.6.3 Assessment of the Habitat Function 350

18.6.4 Overall Assessment - Combined Strategy 353

References 355

Subject Index 361

Contributors

Aislabie, Jackie

Landcare Research, Private Bag 3127, Hamilton, New Zealand

Braddock, Joan R

College of Natural Science and Mathematics, University of Alaska Fair-
banks, Fairbanks, Alaska 99775, USA

Brookes, Philip C.

Agriculture and Environment Division, Rothamsted Research, Harpenden,

Herts., AL5 2JQ, UK

Douglas, Gregory S.

NewFields Environmental Forensic Practice LLC, Rockland, Massachusetts

02370, USA

Eisentraeger, Adolf

Institute of Hygiene and Environmental Medicine, Aachen University of

Technology, Pauwelsstr. 30, 52074 Aachen, Germany

Foght, Julia

Biological Sciences, University of Alberta, Edmonton AB, Canada T6G 2E9

Greer, Charles W.

Biotechnology Research Institute, National Research Council of Canada,

6100 Royalmount Ave., Montreal, Quebec, Canada H4P 2R2

Hedrick, David B.

Hedrick Services, Knoxville, TN 37932-2575; Center for Biomarker Anal-
ysis, University of Tennessee, 10515 Research Drive, Suite 300, Knoxville,
Tennessee 37932-2575, USA

Hund-Rinke, Kerstin

Fraunhofer Institute for Molecular Biology and Applied Ecology, P.O. Box

1260, 57377 Schmallenberg, Germany

XIV Contributors

Jarvinen, Olli

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland

Joergensen, Rainer Georg

Department of Soil Biology and Plant Nutrition, University of Kassel, Nord-

bahnhofstr. la, 37213 Witzenhausen, Germany

J0rgensen, Kirsten S.

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland

Klauser, Thomas

Institute of Microbiology, Leopold Franzens University, Technikerstrasse

25, 6020 Innsbruck, Austria

Kucharski, Rafal

Land Management Department, Institute for Ecology of Industrial Areas,

Kossutha 6 St, 40-833 Katowice, Poland

Kuhad, Ramesh C.

Department of Biotechnology, Kurukshetra University, Kurukshetra -

136 119, Haryana, India

Lindstrom, Jon E.

Shannon & Wilson, Inc., 2355 Hill Road, Fairbanks, Alaska 99709; Institute

of Arctic Biology, University of Alaska Fairbanks, Fairbanks, Alaska 99775,

USA

Malkowski, Eugeniusz

Department of Plant Physiology, Faculty of Biology and Environmental Pro-
tection, University of Silesia, Jagielloiiska 28 St, 40-032 Katowice, Poland

Margesin, Rosa

Institute of Microbiology, Leopold Franzens University, Technikerstrasse

25, 6020 Innsbruck, Austria

Paetz, Andreas

Deutsches Institut fur Normung (DIN), Normenausschuss Wasserwesen

(NAW), 10772 Berlin, Germany

Peacock, Aaron

Center for Biomarker Analysis, University of Tennessee, 10515 Research

Drive, Suite 300, Knoxville, Tennessee 37932-2575, USA

Contributors XV

Plaza, Grazyna A.

Institute for Ecology of Industrial Areas, 40-844 Katowice, 6 Kossutha,

Poland

Prince, Roger C.

ExxonMobil Research and Engineering Co., Annandale, New Jersey 08801,

USA

Raubuch, Markus

Department of Soil Biology and Plant Nutrition, University of Kassel, Nord-

bahnhofstr. la, 37213 Witzenhausen, Germany

Ripp, Steven

The University of Tennessee, Knoxville, Tennessee, 37996, USA

Roembke, Joerg

ECT Oekotoxikologie GmbH, Boettgerstr. 2-14, 65439 Floersheim, Ger-
many

Sainio, Pirjo

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland

Salminen, Jani

Finnish Environment Institute, P.O. Box 140, 00251 Helsinki, Finland

Sas-Nowosielska, Aleksandra

Land Management Department, Institute for Ecology of Industrial Areas,

Kossutha 6 St, 40-833 Katowice, Poland

Schinner, Franz

Institute of Microbiology, Leopold Franzens University, Technikerstrasse

25, 6020 Innsbruck, Austria

Singh, Ajay

Department of Biology, University of Waterloo, Waterloo, Ontario, Canada

N2L3G1

Suortti, Anna-Mari

SGS Inspection Services, Syvasatamantie 24, 49460 Hamina, Finland

Tien, Albert J.

Holcim Group Support Ltd Corporate Social Responsibility Occupational

Health and Safety, Im Schachen, 5113 Holderbank, Switzerland

XVI Contributors

Ulfig, Krzysztof

Institute for Ecology of Industrial Areas, 40-844 Katowice, 6 Kossutha,

Poland

Ward, Owen P.

Department of Biology, University of Waterloo, Waterloo, Ontario, Canada

N2L3G1

White, David C.

Center for Biomarker Analysis, University of Tennessee, 10515 Research

Drive, Suite 300, Knoxville, Tennessee 37932-2575, USA

Whyte, Lyle G.

Dept. of Natural Resource Sciences, McGill University, Macdonald Campus

21, 111 Lakeshore Road, St. Anne de Bellevue, Quebec, Canada H9X 3V9

Wilke, Berndt-Michael

Institute of Ecology, Berlin University of Technology, Franklinstrasse 29,

10587 Berlin, Germany

Soil Sampling and Storage

Andreas Paetz, Berndt-Michael Wilke

1.1

Objective of Soil Sampling
1.1.1

Principal Objectives

General

Samples are collected and examined primarily to determine their physical,
chemical, biological, and radiological properties. This section outlines the
more important factors which should be considered when devising a sam-
pling program for soil and related material. More detailed information is
given in subsequent sections.

Whenever a volume of soil is to be characterized, it is generally not pos-
sible to examine the whole and it is therefore necessary to take samples.
The samples collected should be as fully representative as possible, and all
precautions should be taken to ensure that, as far as possible, the samples
do not undergo any changes in the interval between sampling and exami-
nation. The sampling of multiphase systems, such as soils containing water
or other liquids, gases, biological material, radionuclides, or other solids
not naturally belonging to soil (e.g., waste materials), can present special
problems. In addition, the determination of some physical soil parameters
may require so-called undisturbed soil samples for correct execution of the
relevant measurement.

Before any sampling program is devised, it is important that the objec-
tives be first established since they are the major determining factors, e.g.,
the position and density of sampling points, time of sampling, sampling
procedures, subsequent treatment of samples and analytical requirements.
The details of a sampling program depend on whether the information
needed is the average value, the distribution, or the variability of given soil
parameters.

Andreas Paetz: Deutsches Institut fur Normung (DIN), Normenausschuss Wasserwesen
(NAW), 10772 Berlin, Germany

Berndt-Michael Wilke: Institute of Ecology, Berlin University of Technology, Franklinstrasse
29, 10587 Berlin, Germany, E-mail: bmwilke@tu-berlin.de

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

2 A. Paetz, B.-M. Wilke

Some consideration should be given to the degree of detail and precision
that will be required, and also to the manner in which the results are
to be expressed and presented, for example, concentrations of chemical
substances, maximum and minimum values, arithmetic means, median
values, etc. Additionally, a list of parameters of interest should be compiled
and the relevant analytical procedures consulted; these will usually give
guidance on precautions to be observed during sampling and subsequent
handling of soil samples.

It may often be necessary to carry out an exploratory sampling-and-
analysis program before the final objectives can be defined. It is important
to take into account all relevant data from previous programs at the same
or similar locations and other information on local conditions. Previous
personal experience can also be very valuable. Time and money allocated
to the design of a proper sampling program are usually well justified be-
cause they ensure that the required information is obtained efficiently and
economically.

It is emphasized that complete achievement of objectives of soil inves-
tigations depends mainly on the design and execution of an appropriate
sampling program. The four principal objectives of soil sampling may be
distinguished as follows and are discussed below:

• Sampling for determination of general soil quality

• Sampling for characterization purposes in preparation of soil maps

• Sampling to support legal or regulatory action

• Sampling as part of a hazard or risk assessmenthack

The utilization of the soil and site is of varying importance depending on
the primary objective of an investigation. For example, while consideration
of past, present, and future site use is particularly relevant to sampling for
risk assessment, it is less important for soil mapping where the focus is
on description rather than the evaluation of a soil. Objectives such as soil
quality assessment, land appraisal, and soil monitoring take utilization into
account to varying degrees.

The results obtained from sampling campaigns to assess soil quality for
mapping may indicate a need for further investigation. For example, if
contamination is detected, a need arises for identification and assessment
of potential hazards and risks.

Sampling for Determination of General Soil Quality

This is typically carried out at irregular time intervals to determine the
quality of the soil for a particular purpose, e.g., agriculture. As such, it will
tend to concentrate on factors such as nutrient status, pH, organic matter
content, trace element concentrations, and physical factors, which provide

1 Soil Sampling and Storage 3

a measure of current quality and which are amenable to manipulation.
Sampling is usually carried out within the main rooting zone and also
at greater depths but sometimes without exact distinction of horizons
or layers. The guidance given in ISO 10381-4 (2003) will be particularly
relevant.

Sampling for Preparation of Soil Maps

Soil maps maybe used in soil description, land appraisal (taxation), and for
soil monitoring sites to establish the basic information on the genesis and
distribution of naturally occurring or man-made soils, their chemical, min-
eralogical, biological composition, and their physical properties at selected
positions. The preparation of soil maps involves installation of trial pits
or core sampling with detailed consideration of soil layers and horizons.
Special strategies are required to preserve samples in their original physical
and chemical condition. Sampling is nearly always a once-off procedure.
The guidance given in ISO 10381-4 (2003) is particularly relevant.

Sampling to Support Legal or Regulatory Action

Sampling may be required to establish base-line conditions prior to an
activity that might affect the composition or quality of soil, or it may be re-
quired following an anthropogenic effect such as the input of an undesirable
material that may be from a point or a diffuse source. Sampling strategies
need to be developed on a site-specific basis. To adequately support legal
or regulatory action particular attention should be paid to all aspects of
quality assurance including, for example, £C chain-of-custody procedures."
The guidance given in ISO 10381-5 (1995) is particularly relevant; that in
ISO 10381-4 (2003) may also be relevant.

Sampling for Hazard and Risk Assessment

When land is contaminated with chemicals and other substances poten-
tially harmful to human health and safety or to the environment, it may
be necessary to carry out an investigation as a part of a hazard and/or risk
assessment i.e., to determine the nature and extent of contamination, to
identify hazards associated with the contamination, to identify potential
targets and routes of exposure, and to evaluate the risks relating to current
and future use of the site and neighboring land. A sampling program for
risk assessment (in this context: phase I, phase II, phase III, and phase IV
investigations) may have to comply with legal or regulatory requirements,
and careful attention to sample integrity is recommended. Sampling strate-
gies should be developed on a site-specific basis. The guidance given in
ISO 10381-5 (1995) is particularly relevant, and that in ISO 10381-4 (2003)
may also be relevant.

4 A. Paetz, B.-M. Wilke

1.1.2

Specific Objectives

General

Depending on the principal objective(s) it will usually be necessary to
determine for the body of soil or part thereof:

• The nature, concentrations, and distribution of naturally occurring sub-
stances

• The nature, concentrations, and distribution of contaminants (extrane-
ous substances)

• The physical properties and variations

• The presence and distribution of biological species of interest

It will often be necessary to take into account changes in the above
parameters with time, caused by migration, atmospheric conditions, and
land/soil use. Some detailed objectives are suggested in the clauses below.
The list is not exhaustive.

Sampling for the Determination of Chemical Soil Parameters

There are many reasons for chemical investigation of soil and related ma-
terial and only a few are mentioned here. It is important that each sampling
routine is tailored to fit the soil and the situation. Chemical investigations
are carried out

1. To identify immediate hazards to human health and safety and to the
environment

2. To determine the suitability of a soil for an intended use, e.g., agricultural
production, residential development

3. To study the effects of atmospheric pollutants including radioactive fall-
out on the quality of soil (which may also provide information on wa-
ter quality and indicate if problems are likely to arise in near-surface
aquifers)

4. To assess the effects of direct inputs to soil; there may be contributions
from:

- naturally occurring substances that exceed local background values,
e.g., certain mineral phases in metal deposits

- (un)expected contamination by application of agrochemicals

- (un)expected contamination due to industrial processes

1 Soil Sampling and Storage 5

5. To assess the effect of the accumulation and release of substances by soils
on other soil horizons or on other environmental compartments, e.g.,
the transfer of substances from a soil into a plant

6. To study the effect of waste disposal, including the disposal of sewage
sludge on a soil (which, apart from contributing to the pollution load,
may produce other chemical reactions such as the formation of persistent
compounds, metabolites, or the evolution of gases, such as methane)

7. To identify and quantify products released by industrial processes and by
accident (usually done by investigation of suspect sites or contaminated
sites)

8. To evaluate soil derived from construction works with view to possible
or further utilization of such soils or disposal as waste

Commonly, sampling strategies are employed that require samples to
be taken either from identifiable soil horizons or from specified depths
(below ground surface). It is best to avoid mixing the two approaches,
particularly when sampling natural strata, as this can make it difficult to
compare results. However, a coherent combination of the two approaches
can sometimes be useful on old industrial sites where there is variation in
both the nature of fill and the depth of penetration of mobile contaminants
into the ground, i.e., where there are two independent reasons for changes
in soil/fill properties.

Knowledge of the way in which particular chemical substances tend to be
distributed between the different compartments (air, soil, water, sediment,
and living organisms) is advantageous for the design of some sampling
programs. Similarly, knowledge of the behavior of living organisms affected
by chemical substances, or that influence the availability of substances due
to microbiological procedures, is also advantageous.

Sampling for the Determination of Physical Soil Parameters

The sampling of soil for the determination of some physical properties re-
quires special consideration since the accuracy and extrapolation of mea-
sured data rely on obtaining a sample that retains its in situ structural
characteristics. In many circumstances, it may be preferable to conduct
measurements in the field since the removal of even an undisturbed sample
can change the continuity and characteristics of soil physical properties and
lead to erroneous results. However, certain measurements are not possible
in the field. Others require specific field conditions, but the field situation
can only be controlled to a very limited extent; e.g., it may be possible to
modify the hydrological situation temporarily for measurement purposes
by irrigation. The time and expense necessary for field measurements may

6 A. Paetz, B.-M. Wilke

not be affordable. Laboratory measurements of physical properties are
therefore frequently necessary.

Differences and changes in soil structure affect the choice of size of
sample. Hence, a representative volume or minimum number of replicates
must be determined for each soil type to be studied. The moisture status of
the soil at sampling can influence physical measurements, e.g., hysteresis
on rewetting can occur. Many physical properties have vertical and lat-
eral components, and this should be considered prior to sampling. Where
small undisturbed soil samples are required, manual excavation of cores,
clods, or soil aggregates can be applied. Sampling equipment should be de-
signed such that minimal physical disturbance to the soil occurs. For larger
samples, the use of hydraulic sampling equipment and cutting devices is
preferable in order to obtain a sample with minimal disturbance. Care
should be taken in both equipment design and manufacture to ensure that
internal compression or compaction of the sample does not occur. Where it
is difficult to obtain an undisturbed sample for laboratory measurements,
e.g., in stony or iron pan soils, then in situ measurements may be the most
appropriate method.

Sampling for the Assessment of Biological Soil Parameters

Biological soil investigations address a number of different questions re-
lated to what is happening to or caused by life forms in and on the soil,
including both fauna and flora in the micro and macro range. Ecotoxico-
logical questions are usually given first priority. For example, tests should
be made to verify the effects of chemicals added to the soil on life-forms
and also the possible effects of life-forms in the soil on plants (e.g., high-
value crops) and on the environment, especially on human health. In some
cases, biological soil test procedures operate with fully artificial soils, but
normally the major task of sampling is to choose a reliable soil or site to
carry out the tests. Sampling for the assessment of aerobic microbial pro-
cesses is covered in ISO 10381-6 (1993). The sampling for the assessment
of anaerobic processes is described in ISO 15473 (2002).

1.2

Selection of Sampling Technique

The selection of appropriate sampling equipment depends on the objective
of sampling and should be done after consideration by the analyst or
scientist responsible for subsequent determination. ISO 10381-2 (2002)
gives guidance on commonly used equipment for sampling soil and related
material. Parts 4, 5, and 6 of ISO 10381 describe needs for specific purposes
within their scopes.

1 Soil Sampling and Storage 7

1.3

Sampling Strategy

1.3.1
General

The strategy for the site investigation (whether preliminary, exploratory,
or main) will be determined by the objectives. For example, the different
requirements of site investigations for the purpose of selling, determining
whether contamination is present as suspected, or redevelopment will influ-
ence the spacing of sample locations and the number of samples analyzed,
and hence the cost of the investigation.

Before embarking on any phase or stage of investigation it is important
to set data quality objectives in terms of the type, quantity, and quality (e.g.,
analytical quality) of the data and other information to be collected. These
data quality objectives will depend in part on the nature of the decisions
to be made on the basis of the investigation and the confidence required in
those decisions. Failure to set data quality objectives at the outset can lead
to considerable waste of money if, for example, the data collected are not
suitable or sufficient for a reliable hazard assessment, or leave too many
uncertainties about the "conceptual model" developed for the site.

1.3.2

Preliminary Investigation

General

This is an investigation comprising a desk study (see below) and site recon-
naissance (walk-over survey, site inspection). It is carried out using histori-
cal records and other sources to obtain information on the past and present
usage of the site together with information about local soil properties, ge-
ology, hydrogeology, and environmental setting. From this investigation,
the possibility of contamination can be deduced, and hypotheses can be
formulated on the nature, location, and distribution of the contamination.
These hypotheses form part of the overall conceptual model of the site
that should be developed, encompassing not only the contamination as-
pects but also the geology, hydrogeology, geotechnical properties, and en-
vironmental setting. The current and planned site uses are also important
aspects of the conceptual model. The preliminary investigation should
provide sufficient information:

• For initial conclusions about potential hazards and hazards to actual or
potential human and other receptors, and

• For determination as to need for further action.

8 A. Paetz, B.-M. Wilke

The amount and type of information required will depend on the objec-
tives of the investigation and the ease with which the information can be
obtained, i.e., the amount of work required will vary with the age of the
site, the complexity of its historic usage, the complexity of the underlying
geology, etc.

It shall be remembered that the contamination on a site may be more
complex than initially indicated (for example by current usage) and ade-
quate information on site history should always be obtained in the prelim-
inary investigation.

Desk Study

This includes collection of relevant information on the site, e.g., location,
infrastructure, utilization, history. Possible sources of this information are
publications, maps (check accuracy of map used), aerial photographs, and
satellite imagery from, e.g., land surveyor's offices, geological surveys, water
management boards, industrial inspection boards, mining boards, mining
companies, geotechnical institutions, regional and local (city) archives,
agricultural and forestry authorities, and building supervisory boards.
Particularly important is information on the physical and chemical prop-
erties and the possible spatial distribution of the soil parameter under
investigation; special attention must be paid to geological features such as
stratigraphy and hydrogeology.

Site Reconnaissance

A visit of the site should be part of the preliminary investigation, prefer-
ably in conjunction with the desk study, although it may be independent.
Depending on the local variability of the site and the technical difficulty
of the planned investigation, an experienced person should be chosen for
this task. Such a visit gives a first impression about the correlation of ex-
isting maps with reality, and it will provide much additional information
in a comparatively short time. In some cases, it may be necessary to draw
a first or additional map at this stage.

Samples are not often taken during preliminary investigations; if they
are, they are usually needed to obtain an overview of the kind of soil in
order to chose the right equipment for later activities. Parts 4, 5, and 6 of
ISO 10381 specify the range of preliminary investigations used within their
scopes.

Output from Preliminary Investigation

A report should be prepared summarizing the findings of the preliminary
investigations and stating the conclusions (or hypotheses) drawn concern-

1 Soil Sampling and Storage 9

ing the anticipated site conditions (e.g., geology, hydrology, possible con-
tamination) relevant to the design of the sampling program. This should
enable the appropriateness of the sampling strategy adopted to be assessed
at a later date.

1.3.3

Exploratory Investigation

This involves on-site investigation including collecting samples of soil or
fill, surface water, groundwater, and soil gas, where appropriate, to be
analyzed or tested. The data and information produced are then assessed to
determine if the hypotheses from the preliminary investigation are correct
and, where appropriate, to test other aspects of the conceptual model. It is
therefore mainly a qualitative investigation rather than quantitative.

In some cases, where the hypotheses are found to be correct, no further
investigation may need to be carried out. However, it may become apparent
as a result of the exploratory site investigation, for example, that the con-
tamination pattern is more complex or concentrations of contamination
are higher than anticipated and may have already caused or in the future
may cause a hazard. In this situation the information obtained may be
inadequate to make decisions with a satisfactory degree of confidence. It
will be necessary to carry out a main site investigation to produce sufficient
information to make a full hazard assessment, to determine the need for
protective or remedial measures, and in due course (and possibly following
further stages of investigation), to select, design, and apply protective or
remedial measures.

1.3.4

Main Site Investigation

The main site investigation quantitatively determines the amount and spa-
tial distribution of contaminants, their mobile and mobilizable fractions,
and the possibilities of spread into the environment. Also included is the
possible future development of the contamination situation. This will in-
volve the collection and analysis of soil or fill, surface water, ground water,
and soil gas samples in order to obtain the information necessary to enable
a full assessment of the hazards presented by the contamination to humans
and other potential receptors and also to enable appropriate containment
or remediation actions to be identified (sometimes) together with an initial
estimate of costs. The analysis of samples can be supported by model cal-
culations and investigation techniques that do not make use of sampling.

10 A. Paetz, B.-M. Wilke

Detailed design of protective or remedial works may require further stages
of investigation.

The amount and nature of the information required from the main site
investigation (or any particular stage of it) will vary depending on the
nature of the site and the objectives of the investigation. The implications
of the decisions on what actions should be implemented on a site will vary
from site to site. Additionally, the amount and quality of the information
required will also vary according to the requirements of the decision-
making processes (e.g., the risk assessment, decisions regarding the need
for and type of remedial actions). All parties involved in the decision-
making process should be kept fully informed as information is produced
to ensure that the information is sufficient for the purpose intended.

After completion of the interpretation of the information generated,
including any risk assessment, it should be possible to determine whether
protective or remedial measures are required and to make generalizations
about the type of measures that might be appropriate.

1.3.5

Samples and Sampling Points

General

The selection, location and preparation of the sampling points depend on:

• The objectives of the investigation

• The preliminary information available

• The on-site conditions

The nature of samples to be obtained shall be appropriate to the aim of
the investigation and shall be specified in the program before fieldwork
begins.

Sampling Patterns

Sampling patterns are based on the estimation of the distribution of soil
constituents (in most cases chemical substances) in an area or, when appro-
priate, on the type of substance input. Four major fixed sampling patterns
can be identified as being based on:

• No specific estimate of substance distribution

• Local substance distribution and known as a "hot spot"

• Distributions along a line

• Strip-like distributions

1 Soil Sampling and Storage 1 1

Along with these, several other patterns exist (e.g., based on deposition
of substances from the air, input due to flooding). All fixed patterns have
to be adjusted to local conditions and are subject to modification.

In agricultural sampling a small number of convenient sampling patterns
are established in order to obtain information on, e.g., nutrient demand
or pesticide residues of rather large areas. For additional information refer
to ISO 10381-4 (2003). However, it must be emphasized that most grid
sampling patterns are not very efficient during the growing season, and are
rarely applicable. The investigation of contaminated sites which may have
profound health and economic consequences usually requires a much more
detailed selection and application of sampling patterns, to give calculated,
estimated, or randomly chosen sampling points on a one-, two- or three-
dimensional figure. The choice of pattern should be the result of preliminary
investigation of a site rather than of an ad hoc decision taken in the field.

Some investigations are carried out without predetermined pattern
plans. This should not be confused with the application of random dis-
tribution of sampling points, because a person usually cannot distribute
sampling points randomly without preparation, i.e., without ensuring that
at every point in the area, despite the position of the other sampling points,
a sample will be obtained with equal probability. Where sampling is to be
carried out without a predetermined pattern (ad hoc sampling) care shall
be taken that sampling is carried out by an appropriately experienced in-
vestigator. It also should not be confused with the application of sampling
plans to verify special hypotheses which, with regard to the problem, will
be developed and justified by the investigator (judgmental sampling).

In the following are examples of a number of commonly applied sampling
patterns which meet different statistical requirements (Figs. 1.1-1.5). Expe-
rience (and theoretical considerations) show that in many cases systematic
sampling on a regular grid is both practical and sufficiently productive
to allow the creation of a detailed picture of variations in soil properties.
The number of sampling points can be easily increased (e.g., in areas mer-
iting more detailed investigation), the grid is easy to mark out on site,
and sampling points are usually easily relocated. Systematic sampling can
be supplemented by judgmental sampling when appropriate. ISO 10381-5
(1995) provides examples of pattern application for sampling contaminated
sites. For selection of sampling patterns see Fig. 1.5.

Most natural properties of the soil vary continuously in space and, as
a consequence, the values at sites that are close together are more similar
than those further apart. They depend upon one another in a statistical
sense. This property is known as spatial dependence and its implications
for sampling are covered by methods of geostatistics, i.e., spatial statis-
tics. Viewed mathematically, the value of a soil property at any place is
a function of its position. The only practicable approach is to regard such

12 A. Paetz, B.-M. Wilke

a property as a random variable and to treat its variation in space statisti-
cally. Such properties are known as regionalized variables. The application
of regionalized variable theory by developing variograms is a common tool
in geostatistics.

Another geostatistical approach is multi-stage or nested sampling and
analysis which also can be linked with a regionalized variable theory.

The applicability of geostatistical methods does not depend on the ob-
served values at those sites, but on the configuration of the sampling points
in relation to the area (or block if three dimensions are considered) to be
estimated. A general criterion for the usefulness of a sampling pattern lies
in the smallness of the largest parcel not being sampled. In terms of sta-
tistically efficient sampling, a regular equilateral triangular grid provides
the best selection of sampling points. For a grid with one node per unit
area, the sampling points are 1.0746 units of distance apart, and no other
point is more then 0.6204 units of distance away from a sampling point. For
practical purposes, sampling patterns are based on rectangular grids. For
such a grid with one node per unit area, no point is more than 0.7071 units
of distance from a sampling point, i.e., the greater ease of use of the square
grid is offset by the slightly greater area of unsampled site.

Example 1: Non-Systematic Patterns (Irregular Sampling)

Widely used in agricultural/horticultural land investigations are the "N",
"S", " W", and "X" patterns of sampling (Fig. 1.1). The general premise is that
the distribution of soil constituents is relatively homogeneous. The patterns
used are simplifications of the stratified random sampling method. Along
the outline of such a pattern, a number of samples are taken and then may
be bulked and mixed to provide one sample for analysis. The distribution
of sampling points is likely to be inadequate to provide the location of point
pollution, and in any event high contaminant levels will be lost in mixing
of these samples. Thus, in most contaminated-land investigations, these
patterns are unlikely to be useful because they obscure high levels of point
contamination. Wherever there are likely to be differences in soil type or
conditions, crop growth, plant species, previous cultivation, etc., the site
should be subdivided according to these differences and a separate sample
taken from each area.

Sampling along a single diagonal of a field or a unit is only recom-
mended in case of strip-like distribution of contaminants on agricultural
areas due to application of fertilizers. Applying a diagonal for sampling
avoids systematic bias by simple and effective means, which would arise
with strip-parallel sampling. However, the more diagonals, the better. Two
diagonals (X-shape) introduce a serious bias to the central area of the field
(Fig. 1.1). This should be considered for the evaluation of the results of the

1 Soil Sampling and Storage

Fig. 1.1. Non- systematic patterns

determinations. Application of diagonal patterns should be based on the
following:

• Estimation that substances are distributed uniformly

• Recognition of usefulness only for uniformly developed areas, and of the
need to sample deviating parts separately

• Application of more than one diagonal if possible (e.g., parallel or X-
shape)

• Equidistant placing of sampling points for all diagonals, i.e. shorter
diagonals have fewer sampling points

• Selection of sampling point independent of local characteristics, points
being fixed (preferably by pacing)

Traversing the area in a zigzag manner similar to that shown in Fig. 1.2
is another way of applying a non-systematic pattern.

Fig. 1.2. Zigzag traverse
sampling pattern

A. Paetz, B.-M. Wilke

A general exception to the "biased diagonals pattern" was developed for
permanently monitored areas within selected sites to achieve information
about long-term changes due to human influence. The aim is to make sam-
ples available from an area representative of the surrounding environment
for a defined number of examinations to be carried out over a period of
some years. The following procedure is recommended (Fig. 1.3):

1. Select a representative area of approx. 1,000 m 2 .

2. Divide this area into four squares, each of 250 m 2 .

3. Within each square draw two diagonals, along each of which nine samples
are obtained (Fig. 1.3).

4. Take samples according to the specified requirements.

5. Prepare composite samples 1, 2, and 3 by:

- Mixing single samples of positions 1, 4, 7, 10, 13, and 16 to give
composite sample 1

- Mixing single samples of positions 2, 5, 8, 11, 14, and 17 to give
composite sample 2

- Mixing single samples of positions 3, 6, 9, 12, 15, and 18 to give
composite sample 3

Fig. 1.3. Rotating diagonals
pattern for permanently
monitored areas

1 Soil Sampling and Storage

6. Rotational sampling of the area may be conducted by:

- Taking samples in the intersections of the sampling points (positions
1-18 in Fig. 1.3)

- Rotating the diagonals clockwise around the center of the square in
steps of 22. 5° so that, all in all, four series of samplings can be carried
out at undisturbed positions

An area selected and sampled according to the above-mentioned scheme
serves for eight sampling series. After the final series, the area may be
considered unsuitable for further sampling. Extensions or reductions in
the dimensions of the test area may impose changes in the total number of
samples, and thus also affect composite samples.

Example 2: Circular Grids

Circular grids are useful for delineating local contaminations, such as from
storage tanks, but also for indicating influence around a regional emitting
source, e.g., precipitation from an industrial plant. Sampling is carried out
at the intersection of concentric circles (the radii of which will depend on
the suspected area of contamination) and the radial lines of the main eight
points of the compass (Fig. 1.4). Sampling based on circular grids may lead
to information on:

• Substance concentrations at the grid center (maximum values)

• Distribution of contamination (size of particular area with increased
contamination)

• Shape of distribution of contamination

Fig. 1.4. Circular grid

16 A. Paetz, B.-M. Wilke

Disadvantages of circular grids are:

• Star-shaped (radial) location of sampling points is practicable but not
optimal. Rotation of concentric circles by 22. 5° leads to a higher quality
pattern (Fig. 1.3).

• Relationship of sampling-point densities of the (usually) eight samples
close to the center and those (usually) eight samples at greater distance
might not to be optimal in every case. If, for example, borders of distri-
bution of a contaminated area are looked for, fewer central points should
be sampled and more toward the margins of the grid.

• Circular grids might imply a uniform extension of contamination in
all directions. This is usually not the case. Preferred directions, e.g.,
due to main wind direction in case of airborne contaminants, should be
considered in modifying of the circular grid, e.g., an increased number of
sampling points in critical directions, an extended distance of sampling
in critical directions.

• Circular grids generally do not serve for taking composite samples be-
cause the values thus measured give information neither on the average
nor on the maximum concentration in the area sampled.

Example 3: Systematic Sampling (Regular Grids)

In many cases a regular grid is selected (Fig. 1.5). Because there is a direct
relationship between optimal sampling point distance and the (estimated)
dimension of the contamination, spacing between sampling points should
not exceed the greatest (estimated) extent of the contamination. Grid di-
mensions will depend on how much detail is required. The assigned spacing
will differ according to the objective of sampling, e.g., to collect samples of
average degree of contamination, to locate isolated sources of contamina-
tion, or to establish the extent of contaminated zones (horizontal and verti-
cal). The latter is of particular importance in cases where a contamination
is already located and a follow-up sampling program becomes necessary.
Although more frequently used for the investigation of soil contamination,
regular grids are also suitable for soil fertility investigations, general soil
monitoring programs, etc. An advantage of a regular grid is that it may
be set up easily and grid dimensions may be readily varied. Interpolation
between sampling points and return to the grid to carry out a more in-
tensive sampling in localized areas to further delineate point sources of
contamination is easy. It is also possible to fix the sampling points at the
intersections of the grid lines.

1 Soil Sampling and Storage

•

/• JM

# A

•

• /

/V y I^v.

•

y / /2^>L

/*%//

/ •

•

Fig. 1.5. Regular distribution of sampling points on a regular grid hatched areas indicate
contamination

Example 4: Random Sampling

In cases of presumed irregular occurrences of contaminated zones, random
sampling may be applied. Sampling points within the area are selected by
using random numbers, which can be found in tables included in manuals
on statistics or which may be generated by computer programs. This tech-
nique has the disadvantage of irregular coverage and makes interpolation
between sampling points difficult (Fig. 1.6). In general, random sampling
can also be applied for soil fertility investigations, etc. In practice, random
sampling (in its purest form) is rarely used in soil surveys.

Example 5: Stratified Random Sampling

This method avoids some of the disadvantages of strictly random sam-
pling. The site is divided into a number of grid cells, and a given number
of randomly distributed sampling points is chosen in each cell (Fig. 1.7).
In general, stratified random sampling can also be applied for soil fertility
investigations, etc. The method has disadvantages in terms of interpola-
tion between the sampling points. Further sampling of the site to identify
local areas of contamination based on the original sampling locations is
difficult.

A. Paetz, B.-M. Wilke

Fig. 1.6. Random sampling without grid
x

•

W^W./

•
# ^

•

/•

w/%\ /

fyf //

^^ •

• \

I / n^>L

J* jMz 1

/wfyfrJ^

/ •

•

•^

^<4m iffi

•

Fig. 1.7. Stratified random sampling

1 Soil Sampling and Storage

Example 6: Unaligned Random Sampling

The term "unaligned" means "irregular" in the sense of "not-in-a-line."
The method is similar to stratified random sampling but in this case only
one of two coordinates is chosen at random. The procedure is as follows.
For example: given a grid with 24 cells (squares), arranged in 4 lines and
6 columns (Fig. 1.8):

1. For the first cell (line 1, column 1), x- and y-coordinates are chosen at
random.

2. For cells 2, 3, 4, 5, and 6 only the y-coordinates are chosen at random.

3. For cells 7, 13, and 19, only the x-coordinates are chosen at random.

4. All sampling points are now located on the grid: For all sampling points
in the columns, the y-coordinates of cells 2, 3, 4, 5, and 6 are valid, and
for all sampling points in the lines the x-coordinates of cells 7, 13, and
19 are valid.

The method has disadvantages in terms of interpolation between the
sampling points. Further sampling of the site to identify local areas of
contamination based on the original sampling location is difficult.

Fig. 1.8. Unaligned random sampling on a regular grid

A. Paetz, B.-M. Wilke

Example 7: Systematic Sampling on a Non-Rectangular Grid

In case of an equilateral triangular grid (Fig. 1.9), each grid point is neigh-
bored by three grid points at the unique distance d x . No other adjacent
points exist. The free, unsampled distance between the related adjacent
points has a radius of

(1.1)

The circular area (A) not being sampled therefore is

d 2
A = n • r ' = n • —

(1.2)

Example: given an area of 10 x 10 m and using 99 sampling points arranged
in 11 rows with 9 sampling points each (distance between rows = 1.11 m)
the area not being sampled is 1.29 m 2 . This unsampled area is thus smaller
than for example a rectangular grid of the same size and using 100 sampling
points arranged at a distance of 1 m one from another, where the area not
being sampled is 1.57m 2 . Any circular contamination with r > 0.64m is
certain to be detected. Thus, just by changing the pattern (and with one
sample less) the size of the unsampled area decreases to approx. 18%.

Sampling points at the site are fixed at a distance of d x in parallel rows
spaced at a distance

d y — — • V3

(1.3)

That is, approx. 0. 87 x d x . The sampling points on the parallel rows are
staggered by

(1.4)

Fig. 1.9. Triangular grid

1 Soil Sampling and Storage

•

W
/

■

•

Fig. 1.10. Sampling along
a linear source

Example 8: Sampling Along a Linear Source

In case of contamination following a line, e.g., caused by leaking pipelines,
sampling points can be arranged in the covering soil directly above the
pipeline or, if not practicable for certain reasons, close to the pipeline. If
the distribution of contaminants caused by a line-like structure is also of
interest, it is recommended to take samples at a distance x one from another
above the line and further samples at increasing distances (e.g., 2x) parallel
to the line (Fig. 1.10).

Identifying the Sampling Location

Identification of sampling points is not usually necessary when taking
composite samples for agricultural purposes. Where samples are taken at
pre-defined points, their accurate location and identification is important
for three principal reasons:

1. To enable actual sampling locations to be revisited if necessary

2. To enable accurate plotting of data in relation to site features so that any
needed treatment (e.g., additions of nutrients or removal of contamina-
tion) can be properly planned

3. To enable the data to be stored and processed by computers (e.g., for
modeling studies, preparation of maps, input into geographic informa-
tion systems)

Moreover, it is recommended that a sketch map be prepared present-
ing all relevant information on the sampling location. Both maps and
photographs should include a scale and a direction marker. It is impor-
tant for the interpretation of data, particularly on abandoned industrial
sites to have detailed information on surface levels at sampling locations.

22 A. Paetz, B.-M. Wilke

Sampling locations should be determined with an appropriate degree of
accuracy. Because it may be necessary to vary the actual location away from
the predetermined location because of the presence of obstructions, it may
be preferable to do the accurate surveying of sampling locations once the
sampling exercise is completed or as it progresses. Surface levels can be
determined at the same time.

When investigating abandoned industrial, waste disposal, or other po-
tentially contaminated sites, the horizontal and vertical location of sam-
pling points or probing points should be recorded. The location of sampling
points should be marked before sampling begins using poles/markers with
color sprays. Color sprays should not be used if soil air has to be sampled.

Preparation of the Sampling Site

Depending on the objective of the investigation, a sampling pattern is
chosen at the design stage and is then applied in the field. Within the
range of patterns are some very complex ones developed with the help of
computer-aided statistics. Preparation for sampling with the use of such
patterns, e.g., location of desired sampling points on the ground, can be
very time-consuming, especially when samples are to be obtained by bor-
ing/drilling techniques or from trial pits. Preparation of the site includes,
for example, removal of superficial deposits (e.g., uncontrolled deposition
of urban wastes), establishment of safety measures, installation of mea-
surement devices (if field tests are carried out together with sampling), as
well as exactly locating the sampling points. In many cases, preparation
of the site takes longer than the actual sampling procedures. Both during
and on completion of sampling all necessary measures must be taken to
avoid hazards to the health and safety of anyone entering the site, and to
the environment.

Barriers to Sampling

It may not be possible to sample at a planned location due to a variety
of reasons (e.g., trees, large rocks, buildings, buried foundations or public
utility services, difficulties of access) and contingency plans for dealing with
such situations should be made in advance. The action to take will depend
on the circumstances. The investigator may ignore the unavailable point
or follow predetermined rules for choosing a nearby substitute location
(e.g., alternative position within 10% of grid spacing or paired sampling
along grid lines on either side the obstruction). Ad hoc decisions made in
the field can lead to bias. An attempt should be made when mapping out
the site to identify such obstructions in advance of actual field work. In all
cases when a sampling point has to be relocated, this fact, and the reasons
for relocation, should be clearly indicated in the report.

1 Soil Sampling and Storage 23

Preliminary investigations as described in Sect. 1.3.2 should provide as
much detail as possible about conditions expected to exist on the site and
should therefore guide the design and execution of the sampling program.
However, such investigations cannot totally prevent the danger of misin-
terpretation of the results of borings, and the selection of sampling points
should take this into account.

Depth of Sampling

No general recommendation can be given on the depths at which samples
should be taken or on the final depths to which trial pits or boring/ drilling
should extend. This depends on the objectives and might be subject to
change during a running program. Investigation of soil for chemical char-
acteristics can be divided into two general types:

1. The investigation of agricultural and similar near-natural sites, where
information is required mostly on the topsoil or plowed horizon or arable
zone but often over an extended area.

2. The investigation of sites which are known or suspected to be contami-
nated, where information is required from deeper layers, sometimes to
a depth of several tens of meters, the extent of the area usually being
rather small compared to agricultural sites.

A mixture of both cases is realized in so-called "soil-monitoring sites,"
which represent larger areas of homogeneous soil development and in most
cases are established to monitor environmental effects to the complete
profile over a long-term scale. A precise description should be made of
all soil horizons or layers encountered during the sampling exercise and
included in the report.

If a profile is to be sampled, care should be taken that every horizon/layer
of interest is sampled and that different horizons/layers are not mixed. In
general, contaminated sites should be sampled horizon by horizon unless
stated otherwise by the client. Care should be taken in a site investigation
to ensure that pathways for migration of contamination are not created,
particularly where impermeable strata maybe penetrated.

When trial pits are used it may be appropriate to sample from more
than one site. A depth-related sampling program is based on a number
of conventions, depending on the project. It is not as representative with
regard to the soil as a horizon-related sampling program can be. The
mode of sampling from each depth should be carefully specified; e.g., the
maximum depth range (usually not more than 0.1 m) and how horizontal
variations are to be dealt with.

The total depth reached, the thickness of the horizons/layers penetrated,
and the depth from which the samples are obtained should be recorded. All

24 A. Paetz, B.-M. Wilke

data should be recorded in meters below surface. The soil depth should be
measured from the ground surface with the thickness of the humus litter
layer recorded separately.

Mountain regions or hilly areas with pronounced slopes require special
consideration. For slopes of 10° and greater, vertical drilling lengths should
be extended according to the cosine rule in order to maintain constant
slope-parallel thicknesses of soil layers. The extension factor is 1/cos of
slope. Without correction, for example, the error will be 2% at a slope of
11.5°.

Timing of Investigation

In some circumstances, it may be necessary to restrict sampling to specific
periods of the year. For example, if the characteristic or substance to be
determined is likely to be affected by seasonal factors or human activities
(weather, soil conditioning/fertilization, use of plant protecting agents), this
should be taken into account in the design of the sampling program. This is
particularly important where monitoring continues for several months or
years or is repeated periodically, and therefore requires similar conditions
every time sampling is carried out.

Sample Quantity

At least 1,000 g of fine soil should be obtained for chemical analysis. This
figure applies both to single samples and composite samples, in the latter
case after sufficient homogenization. Samples obtained to serve as reference
material or to be stored in a soil specimen bank should be of larger size,
usually larger than 2,000 g.

Where the sampling of soil involves the separation of oversized material
(i.e., mineral grains, sand, pebbles, and all other materials) due to very
coarse-grained or heterogeneous soil conditions, the material removed
shall be weighed or estimated and recorded and described to enable the
analytical results to be given with reference to the composition of the
original sample. These procedures should be carried out in accordance
with ISO 11464(1994).

Details on the amount of sample materials needed for determination
of specific physical soil parameters are given in the respective methods
(Chap. 2). In particular, the determination of the particle size distribution
may need a very large mass of soil material. The actual mass required will
usually depend on the largest grain size to be determined (see ISO 11277
1998). The quantity of soil sample needed for biological or ecotoxicological
investigations is highly dependent on the aim of the investigation and the
related soil organisms.

1 Soil Sampling and Storage 25

Single Samples vs. Composite Samples

Composite samples are usually required in cases where the average con-
centration of a substance in a defined horizon/layer is to be determined.
Single samples are required in cases in which the distribution of a sub-
stance over a defined area and/or depth is sought. In most guidelines on
sampling for agricultural or similar investigations, it is recommended that
composite samples be collected by taking a number of increments (accord-
ing to ISO 10381-4 (2003) at least 25 increments should be obtained) and
combining them to form a composite sample. When preparing composite
samples regard should be paid to analytical requirements. For example,
composite samples should never be used if volatile compounds are to be
determined.

1.4

Sampling Methods

1.4.1
General

The most commonly used methods of sampling and forming holes in the
ground to collect samples are covered in this text. This does not preclude
the use of other techniques that are suited to the problems of a partic-
ular location, e.g., areas of permafrost, nor does it preclude the use of
other methods that have been developed. Whatever technique is used, the
principles of sample collection and the approach to sampling to obtain an
appropriately representative sample should be adhered to. This will include
the minimization of contamination of the sample and the protection of the
samplers and other personnel involved. The choice of sampling method
will be determined by taking into account all the needs of the investigation,
including distribution of sampling locations, size and type of sample, and
the nature of the site, including any problems the site poses in carrying out
the investigation.

1.4.2

Type of Sample

There are three basic approaches to taking samples from the ground for
the purpose of investigating soil and ground conditions. A sample maybe:

Type 1 Material collected from a single point (disturbed or undisturbed
sample).

26 A. Paetz, B.-M. Wilke

Type 2 A composite of small incremental point samples taken close to-
gether [disturbed sample; perhaps not suitable for certain tests,
e.g., the determination of volatile organic compounds; (VOCs)].

Type 3 A composite of small incremental point samples taken over an area
(such as a field; disturbed sample).

Samples taken to identify the distribution and concentration of particu-
lar elements or compounds will normally be samples of type 1 or perhaps
type 2 within the area being examined. Such samples would be appropriate
for geological or contamination investigations and any other investigation
involving disturbed samples. Samples taken to assess the overall quality or
nature of the ground in an area would be type 3. Such samples would be
taken for agricultural purposes.

Disturbed samples may be taken by any of the three basic methods since
these samples do not require the maintenance of the original ground struc-
ture. Undisturbed samples will always require type 1 sampling because the
original ground structure needs to be retained in the sample. Undisturbed
samples can be taken using a coring tool or cylinder or with a sampling
frame. Whichever of these sampling devices is used, the mode of operation
is the same. The sampling device is pushed into the ground to be sam-
pled and then subsequently removed complete with the sample so that the
ground is collected in its original physical form.

Type 1 samples can be readily collected using hand augers and other
similar sampling techniques. Any of the following tools (as well as others)
may be used as appropriate:

• Cutting cylinders of different size, cutting frame

• Special hand augers [gauge auger (shallow-profile sampler), bucket auger
to bring down borings for cutting cylinder application];

• Protective cap, hydraulic or handpowered supporting ring

Special bags should be used for storage and transport of "sample rings"
(actually sample cylinders of limited height) to prevent disturbance and
drying out. Where undisturbed samples are required, special equipment
(see above) will be necessary in order to collect the sample while maintain-
ing the original ground structure.

Type 2 samples will be appropriate when using machines for excavating
ground to obtain samples. In these circumstances the samples should be
formed by taking portions from locations within the bucket of excavated
material (e.g., nine-point sample, according to Fig. 1.4).

Type 3 samples can be collected using hand or powered augers, but care
needs to be taken to ensure the auger repetitively collects the same amount
of sample.

1 Soil Sampling and Storage 27

Disturbed samples are suitable for most purposes except for some physi-
cal measurements, profiles, and microbiological examinations when undis-
turbed samples maybe required. Undisturbed samples should be collected
where it is intended to determine the presence and concentration of VOCs,
since disturbance will result in loss of these compounds to the atmosphere.

Choices of sampling method include the use of machinery or manual
methods. The sampling may be carried out near the ground surface, at
some depth below ground level, or from locations deep below the ground
surface. Methods of achieving the desired depth for sampling are either by
excavating (e.g., trial pits), driving probes, or drilling (e.g., boreholes).

Sampling during borehole creation allows the required integrity for the
chemical, physical, and biological investigation of selected soil horizons.
Gas and water sampling may also be undertaken for specific purposes re-
lating to the need to acquire information rapidly, for example monitoring
a borehole for methane and carbon dioxide or VOCs on occasions when the
rapid identification of chemical constituents in groundwater is required. It
is recommended that monitoring groundwater horizons over time for hy-
drogeological and chemical parameters, as well as ground composition, be
undertaken from cased wells or standpipes installed in boreholes. The re-
quirements of the sampling strategy should identify the nature of borehole
construction so that the appropriate monitoring design can be specified.

1.4.3

Undisturbed Samples

If undisturbed samples are required for soil sampling, these can easily
be taken, for example, using a Kubiena box, a coring tool, or cylinder. In
each case the sampling device is pushed into the soil and subsequently
removed with the sample so that the soil is collected in its original physical
form. Beside these simple techniques, many others exist, some of which are
described later.

Hand-Operated Auger Techniques

There are many designs of hand auger samplers available. The designs
have been developed over many years to deal with different soil types
and conditions. Ease of use depends upon the nature of the ground to be
sampled. In general, handaugers are easier to use in a sandy soil than in
other soils, particularly where obstructions such as stones are encountered.
In sandy soils, hand augers can be used to sample to a depth of about
5 m. Hand augers are usually used for sampling homogeneous soils, e.g.,
agricultural soils. When using hand augers, care should be taken to ensure
that the soil is not contaminated by material dropping into the sample from

28 A. Paetz, B.-M. Wilke

higher up the bore either during augering or during withdrawal of the
samples. Lining the borehole carefully with a plastic tube can prevent this
cross contamination.

Preferred forms of hand augers to be used for collection of soil samples
are those which take a core sample. Other types of auger may be used to
facilitate drilling to the requisite depth for sampling providing it is possible
to clean the bore to prevent cross contamination.

Sampling by hand augers allows observation of the ground profile and
the collection of samples at preselected depths. Particular care should
be taken to obtain representative samples if localized contamination is
penetrated. When a hand auger is to be used to take samples for testing
soil for agricultural purposes, and the samples are to be composited, it
is essential that the auger should be capable of consistently collecting the
same sample volume. Such sampling of the near-surface soil is normally
done at approx. 150-250 mm depth.

Power-Operated Auger Techniques

It is possible to obtain augers powered by small motors to reduce the labor
required to carry out the sampling. The need to avoid cross contamination
within the bore applies equally to augering with power-operated augers
as with hand augers. Powered augers mounted on rough-terrain vehicles
are available for repetitive sampling for agricultural purposes. Care should
be exercised when using fuel-driven motors to avoid contamination of the
sample by the fuel, the motor lubricant, and the exhaust fumes. Augers
powered by electric motors that minimize the risk of such contamination
are available.

Light Cable Percussion Boring

Light cable percussion boring general uses a mobile rig with winch of 1-2 1
capacity driven by a diesel engine and a tripod derrick of about 6 m height.
With many types the derrick folds down so that the rig can be towed by
a small vehicle (frequently four-wheel drive). The light cable percussion
technique is commonly used for geotechnical purposes, and boreholes
over 20 m deep can be created. This technique can be of particular use in
investigating deep sites such as refuse tips and other unstable ground. The
ground is penetrated using different tools, depending on the strata. A clay
cutter is used for cohesive soils and a shell (or bailer) for cohesionless
soils. Chisels may be used to penetrate very hard ground and obstructions.
The borehole formed by these tools is supported by a steel casing that is
advanced as the borehole proceeds.

Depending upon the nature of the ground, the tool may form the borehole
in advance of the steel casing being pushed down the hole, e.g., in clay

1 Soil Sampling and Storage 29

strata. This often results in material from the side of the borehole being
dislodged as the casing is pushed down the borehole, and can result in
cross-contamination. If the borehole is being formed in sands or gravels,
particularly in the saturated zone, the steel casing may be pushed into place
to support the borehole sides before the material is removed with the shell.
This can disturb the ground and make sampling difficult.

In some strata it may be necessary to add water to the borehole to
provide lubrication. In this situation tap water may be used, if available,
and care should be taken with respect to the effects on both soil and
water samples. The addition of water should be recorded on the borehole
log and, if appropriate, on the sample details. The clay cutter and the
shell bring up disturbed material from the borehole which is generally
sufficiently representative to permit recording of the strata, but care has
to be taken to avoid misinterpretation due to ground being pushed down
within the borehole - for example, when the casing is moved. The casing
avoids most of the problems of cross contamination, but the borehole
should be cleaned out each time the supporting casing is driven further
into the borehole, before taking a sample. Samples may be collected from
both the clay cutter and the shell. The resultant sample size, although larger
than obtained by hand-augering techniques, is still restricted. Undisturbed
samples may be collected in cohesive strata and in weak rock (e.g., chalk)
by driving a hollow tube (100 mm open-tube sampler) into the ground and
withdrawing the resultant core for examination and analysis. Use of such
undisturbed sampling equipment may be preferred in order to minimize
cross-contamination of samples collected for testing purposes.

Water samples may be obtained as drilling proceeds and, because the
casing of the borehole seals the borehole from the surrounding ground as
the borehole advances, it is possible to sample water horizons at different
depths with minimal risk of cross contamination. However water samples
that are truly representative of the ground water necessitate the installation
of an appropriately designed monitoring well. The borehole atmosphere
can be monitored for gas concentrations as the borehole proceeds, or gas
samples maybe taken so that the profile of the ground gas composition can
be determined.

Rotary Drilling

Powered rotary cutting tools use a shaft fitted with a cutter head that
is driven into the ground as it rotates. The system requires some form
of lubrication (air, water, or drilling mud) to keep the cutting head cool
and remove the soil and other material that has been cut through. The
lubricant lifts the debris from the cutting head up the borehole formed
and ejects the material at ground level. This results in the potential for

30 A. Paetz, B.-M. Wilke

cross contamination due to contact with the ground forming the sides of
the hole. This technique is particularly useful for digging a hole quickly
in order to form a deep observation well or for obtaining samples using
a technique appropriate at greater depths only. The uncontrolled ejection
of material that can occur with this technique (for instance where air or
water is used for lubrication) can lead to extensive surface contamination
when drilling through contaminated ground. This may be hazardous, both
to the investigation team and the environment.

There are two basic types of rotary drilling, (1) open hole (or full hole)
drilling in which the drill cuts all the material within the diameter of the
borehole, and (2) core drilling where an annular bit fixed to the bottom
of the outer rotating tube of the core barrel assembly cuts a core that is
recovered within the inner most tube of the core barrel assembly and is
brought to the surface for examination and testing. Rotary drilling requires
well-maintained equipment operated by a specialist driller with adequate
training and considerable experience.

Driven Auger

The driven auger is powered by machine, so that great force can be exerted
downwards. The cutter head consists of one or more 360° spirals, usually
with a shallow pitch to prevent ground falling off when withdrawn from
the borehole. The method of forming the borehole is to advance the cutter
head approx. 1 m into the ground, withdraw the head from the hole and spin
off the spoil. This process is repeated until the required depth is reached.
This method is not very satisfactory for sampling, because of the potential
for cross contamination, nor is it suitable for strata logging. The method
does enable the formation of a large diameter hole (up to 25 cm) into the
ground relatively quickly. Lubrication of the auger is not required, but some
dispersal of contaminated material may occur as the spoil is spun from the
cutter head.

Continuous Flight Auger

A similar system is the continuous flight auger, which consists of a contin-
uous helix welded to the center shaft. Downward force is again provided by
the machine and continuous rotation lifts the ground to the surface from
the base of the hole. This technique is only of use in site investigations in
forming a hole rapidly to give depth in the ground and cannot be used for
sampling or strata logging. Lubrication of the auger is not required.

Hollow Stem Auger

Hollow stem augers are a form of continuous flight auger in which the
continuous helix is attached to a hollow central shaft. The drill head is

1 Soil Sampling and Storage 31

formed of two pieces, a circular outer head and an inner pilot or center bit
that is fixed on a plug on the hollow shaft that can be withdrawn through
the center of the auger up to the surface. This ability to withdraw the center
bit and plug whilst leaving the auger in place is the principal advantage of
the hollow stem auger. Withdrawing the plug provides an open cored hole
into which samplers, undisturbed samplers, instruments, borehole casing,
and numerous other items can be inserted to the depth achieved.

Removal of any such equipment and replacing the center plug and bit
enables the continuation of the borehole. The technique provides a fully
cased hole and can avoid some of the potential cross-contamination prob-
lems of percussion boring. Ground samples are collected by open drive
samplers or core barrels inserted down the hollow stem. The method has
been successful on some landfill sites and can be used for the installation
of groundwater monitoring wells and gas standpipes. Some versions of the
hollow stem auger allow continuous access to the bottom of the borehole
and will permit percussion drilling or driven sampling through the center,
while the hollow stem auger is actually forming the hole. The technique will
allow collection of samples, particularly undisturbed samples, in addition
to other down-hole testing, and also enables strata logs to be produced.
Lubrication of the auger is not required.

Percussive window sampling involves driving cylindrical steel tubes into
the ground using a high frequency percussive hammer. Usually, the hammer
is driven by a hydraulic power pack, but electric and pneumatic hammers
are also available to suit particular site conditions. Sample tubes are 1 or 2 m
long and have a broad slot or window cut down one side. The soil material
passes into the sample tube, through a cutting shoe at the end, as it is driven
into the ground. Drill rods are used to drive the sample tubes to greater
depths. On reaching the required depth for sampling, the sample tube and
any drill rods are withdrawn using a mechanical jack. After removal from
the probe hole, the soil material can then be inspected and the strata logged
and sampled from the window.

Soil samples may also be obtained using split tubes or split spoon sam-
plers. These are effectively tubes linearly split in half but held together by
securing rings during sampling. Such devices are often used in conjunc-
tion with driven bar probes, and they allow ready retrieval of the core. Soil
samples may also be obtained using a tube combined with an inert liner
to enable ease of removal of the core from the sampler. The system can be
used to collect samples at different depths, to rapidly penetrate to the depth
at which the sample is to be taken, or to provide a continuous core.

Sample tubes of various diameters are available (35-80 mm) and se-
lected according to the ground conditions. Tubes are normally selected in
a sequence of reducing diameters to penetrate to depth. The depth that
can be achieved depends on the soil type and particularly on the presence

32 A. Paetz, B.-M. Wilke

(or absence) of obstructions. Depths of 10-12 m can be achieved where
the probe hole remains open without support. Piezometers and ground gas
monitoring pipes can be installed in the resultant probe holes where the
ground is sufficiently stable. Systems are available to allow a probe head,
with a sampling device, to be inserted into the previously formed hole to the
desired sampling depth. The probe head is then unscrewed and withdrawn
up the inside of the shaft, and the exposed sampling device is pushed into
the ground to collect the sample. The sampling head is then withdrawn and
removed for analysis. This system also enables undisturbed samples to be
collected.

Continuous Samplers

Continuous soil samplers can produce core samples up to 30 m length in
ground such as fine alluvial deposits. This may be of particular value and
is considered to yield superior samples to those obtained by consecutive
drive-in sampling. The samplers normally are made in sizes between 30
and 70 mm diameter and consist of an outer driven tube with an internal
system providing a sheath to the core as the sampler is driven into the
ground. Extension tubes of 1 m length are added to the sampler as the
ground is penetrated. On removal from the ground, the continuous core is
cut to suitable lengths, frequently 1 m, and placed in purpose-made sample
cases for storage. Samples may be removed from the core for testing and
the core itself observed and recorded.

Driven Probes

Driven probes maybe used to make continuous geophysical measurements,
for example, resistance to penetration, or may be fitted with instruments
for gathering other data. Care should be taken to avoid cross contamination
from the sides of the probe hole and from the base of the probe hole. This
system can be used to either monitor ground water parameters (such pH,
electrical conductivity, temperature, etc.) using monitors in the probe, or to
access groundwater so that a representative sample can be taken without the
need for purging as associated with conventional monitoring wells. Ground
gases can be similarly accessed and sampled. Driven probes have the usual
disadvantage of difficulty in penetrating ground with obstructions, and
cannot be used for logging the ground strata unless continuous soil samples
are taken. Driven probes are, however, considerably faster than traditional
boreholing techniques.

Excavations (Trial Pits)

This is a widely used technique for collecting samples for site investigations
related to contamination. The advantages of the method are the applicability

1 Soil Sampling and Storage 33

over a wide range of ground conditions, the opportunity for close visual
examination of the strata, and the speed with which the work can be
carried out. Trial pits can be dug where the ground will stand temporarily
unsupported and permit the observation of the in-situ condition of the
ground both vertically and laterally. Where there is water present in the
excavation, problems are presented due to instability of the sides and the
difficulty of obtaining representative samples of the ground (finer material
tends to wash out with the water as the sample is collected). In this situation
the trial pit may be dewatered by pumping, providing there is a safe and
suitable means of disposal of the water - or an alternative technique of
sampling should be used. In deeper trial pits formed by machines, samples
of the ground can be collected by careful use of the machine bucket, thereby
avoiding any need to enter the pit. In carrying out excavations, whatever
technique is used to form a trial pit, the excavated material should be
placed on the adjacent ground (this should be protected as necessary from
contamination) in a way that ensures it will not fall back into the excavation
causing cross contamination.

The surface soil layer should be kept separate so that it can be replaced
on the surface after the trial pit is backfilled. It may be necessary to separate
other material as it is excavated so that any deep lying contamination is
replaced at the same depth when back filling and not mixed with other
material or replaced near the surface. For environmental reasons and due
to legislation, it may be necessary to dispose of excavated material off-site
and to complete the backfilling of the trial pit and restoration of the site
using clean imported material.

Entry of the excavation by personnel should be avoided where possible
since the unsupported sides of a trial pit may readily collapse. If it is
essential that an excavation is to be entered for sampling purposes, e.g.,
the collection of undisturbed samples, then shoring should be used and
reference should be made to the guidance given in ISO 10381-3 (2001). In
unstable ground the trial pit may collapse and extra care should be taken
when observing the excavation and collecting samples. If necessary, the
sides should be supported or made to slope to improve stability. For all
ground conditions, if the depth of excavation is greater than 1-1.2 m and
the excavation is to be entered by personnel, the sides should be adequately
shored to prevent collapse.

Manual

Shovel, pick, and fork may be used to excavate trial pits down to about
2 m and, if only a small number of such excavations are required, this may
be the easiest technique for collecting soil samples. The trial pit should
have a plan area of approx. 1 x 1 m to enable easy collection of samples

34 A. Paetz, B.-M. Wilke

and recording of the soil profile. Hand excavation is necessary particularly
in urban areas if services (water, gas, electricity, etc.) are known to exist
in the vicinity, and particularly if their location is uncertain. Once the
base of the excavation is below the depth at which any services may exist,
then the excavation or boreholing may be continued using the appropriate
machinery.

1.4.4
Cross-Contamination

Whatever method is used, it is important that nothing connected with the
sampling system itself contaminates the sample. This includes avoiding
contamination by contact with the sampling equipment or containers and
also avoiding the loss of contaminants from the sample by adsorption or
volatilization. The sampling equipment should be kept clean so that parts
of a previous sample are not transmitted to a subsequent sample causing
cross contamination. For agricultural purposes, even with repetitive sam-
pling across a field to form a composite sample, the sampling device should
at least be brushed clean between each location. For geological and con-
tamination investigations, all sampling equipment should be thoroughly
cleaned between each sample. Contamination of samples due to lubrica-
tion used to ease sample collection, or contamination due to equipment
lubricants, oils, greases, or fuels should be avoided. Where it is necessary
to use lubrication, e.g., water, to ease forming a borehole to enable sample
collection, only lubrication that will not conflict with nor confound the
analysis of the samples (in the sense of matrix effects or contribution to the
contamination) should be used.

A hand trowel of stainless steel should be used to place samples into
sample containers. The quality of the stainless steel should, however, first
be verified to ensure that cross contamination of the samples will not occur
or interfere with the quality of the analytical data. The most commonly
used methods of drilling, excavating, and sampling of the ground produce
disturbed samples. If undisturbed samples are required, special sampling
equipment is required and extra care should be taken in collection.

1.4.5

Sampling Containers

General Considerations

Samples of soils and related materials are liable to change to differing
extents as a result of physical, chemical or biological reactions that may take
place between the time of sampling and the analysis. This is especially true

1 Soil Sampling and Storage 35

of soils contaminated with volatile constituents. The causes of variations
are numerous and may include:

• Changes of certain constituents due to the activities of living organisms
in the soil

• Oxidation of certain compounds by atmospheric oxygen

• Changes in the chemical nature of certain substances due to changes of
temperature, pressure, and hygroscopicity (e.g., loss to the vapor phase);

• Modification of pH, conductivity, carbon dioxide content, etc., by the
absorption of carbon dioxide from the air

• Irreversible adsorption on the surface of containers by metals in solution
or in a colloidal state, or by certain organic compounds

• Polymerization or depolymerization

The extent of these reactions is a function of the chemical and biological
nature of the sample, its temperature, its exposure to light, the nature
of the container in which it is placed, the time between sampling and
analysis, conditions such as rest or agitation during transport, seasonal
conditions, etc. It must be emphasized, moreover, that these variations are
often sufficiently rapid so as to modify the sample considerably within
several hours. It is therefore essential in all cases to take the necessary
precautions to minimize these reactions, and in the case of many parameters
to analyze the sample with a minimum of delay. Any of the procedures
should be mentioned in the sampling report if applied during sampling.

Preservation

The addition of chemical preservatives or stabilizing agents is not a com-
mon practice for soil sampling. This is because a single soil sample is usually
used for a large number of different determinations, and moreover has to
undergo preparation (drying, milling, etc.) during which unwanted and
unquantifiable reactions of the preservatives may occur. If, in special cases,
it is necessary to preserve samples a method that does not introduce un-
acceptable contamination should be chosen. Generally, stability of samples
can be considered in three classes:

1. Samples in which the contaminant(s) is/are stable

2. Samples in which the contaminant(s) is/are unstable but stability can be
achieved by a preservation method

3. Samples in which the contaminant(s) is/are unstable and cannot be
readily stabilized

36 A. Paetz, B.-M. Wilke

For those contaminants that are unstable, loss or change (chemical or
biological) of the contaminant should be minimized by either preserving
the contaminant (e.g., freezing or adding a stabilizing agent) or by arrang-
ing for analysis to be undertaken immediately or soon after sampling. The
use of liquid nitrogen for immediate deep freezing of soil samples in vapor
phase is effective, and containers made of stainless steel (not chromium or
nickel plated) are recommended. Some contaminants are not easily stabi-
lized in a manner compatible with subsequent analysis. Volatile solvents
fall into this category and some of them may begin to volatilize as soon as
the soil is exposed by sampling. A special sampling procedure is needed
to minimize such loss. In spite of numerous investigations carried out in
search of methods that will enable soil samples to be stored without modifi-
cation of their composition, it is impossible to give absolute rules that cover
all cases and all situations and that do not have exceptions. In every case,
the method of storage must be compatible with the analytical techniques
to be used and should be discussed with the analytical laboratory.

Use of Appropriate Containers

The choice and the preparation of containers can be of major importance.
The most frequently encountered problems are:

• Adsorption onto the walls of the containers

• Improper cleaning resulting in contamination of the container prior to
sampling

• Contamination of the sample by the material of which the container is
made

• Reaction between constituents of the sample and the container

The purpose of the container is to protect the sample from losses due to
adsorption or volatilization, or from contamination by foreign substances.
Other factors to be considered in selection of the sample container used to
collect and store the sample include:

• Resistance to temperature extremes

• Resistance to breakage

• Water and gas tightness

• Ease of reopening

• Size, shape, and mass

• Availability

• Potential for cleaning and re-use

1 Soil Sampling and Storage 37

Cleaning of the sample container is a very important part of any sam-
pling/analysis program. Two basic situations can be distinguished: ( 1 ) clean-
ing of new containers to remove dust and packing material; (2) cleaning of
used containers prior to re-use. The type of cleaners used depends on the
kind of container material and on the material to be analyzed. The selection
of acids or other cleaning agents should ensure that no contamination of
the containers results with regard to the constituents to be analyzed and,
moreover, that there is no harm to the environment or human health.

Containers already used for investigations of contaminated sites should
not be used again because cleaning containers of soils containing unknown
substances may cause risks to health. The determination of organic con-
stituents may require drying or cooling procedures under carefully con-
trolled conditions to avoid microbial contamination. Sterilization is re-
quired whenever biological or microbiological determinations are to be
carried out.

1.5
Pretreatment

1.5.1

Chemical Analysis

Inorganic Parameters and Soil Characteristics

Soil samples are dried in the air or in an oven at temperature not exceeding
40 °C, or are freeze-dried. If necessary, the soil sample is crushed while still
damp and friable and again after drying. The soil is sieved and the fraction
smaller than 2 mm is divided into portions mechanically or by hand, to
enable representative subsampling for analysis. If small subsamples (< 2 g)
are required for analysis, the size of the particles of the fraction smaller
than 2 mm is further decreased.

• A drying temperature of 40 °C in an oven is preferable to air drying
at room temperature because the increased speed of the drying limits
changes due to microbial activity.

• It should be noted that every type of pretreatment will have an influence
on several soil properties.

• The sieve aperture size of 2 mm is generally used. However, before the
pretreatment is started, check should be made to see if any of the ana-
lytical methods to be applied require other sieve sizes.

• Storing soil samples, including samples that are air dried, refrigerated
or stored in the absence of light, for a long time may have an influence

38 A. Paetz, B.-M. Wilke

on a number of soil parameters, especially solubilities of both inorganic
and organic fractions.

• Special measures should usually be taken for samples from contaminated
soils. It is important to avoid contact with the skin, and special measures
should be taken when drying such samples (ventilation, air removal,
etc.). Samples may be hazardous because of the presence of chemical
contaminants, fungal spores, or pathogens such as leptospirosis, and
appropriate safety precautions should be taken.

• According to the international standard, it is generally assumed that at
least 500 g of fresh soil shall be available.

• Keeping an archive sample is optional and should be clearly stated in the
overall description of the investigation program.

Organic Contaminants

The properties of organic micro-pollutants may differ greatly according to
chemical species:

• They can range from non volatile to very volatile compounds (low to
high vapor pressure).

• They may be labile or reactive at ambient or elevated temperatures.

• They may be biodegradable or UV degradable.

• They may have considerably different solubilities in water.

• They require different analytical procedures.

Because of these differences a general pretreatment procedure cannot be
proposed. The goal of a pretreatment procedure is to prepare a test sample
in which the concentration of the contaminant is equal to the concentration
in the original soil, provided, however, that this procedure does not alter the
chemical species to be analyzed. If the sample contains only small particles
and the contaminant is homogeneously distributed it is, for instance, not
necessary to grind the sample. According to the international standard the
size 2 mm is used to distinguish between small and large soil particles. Care
should be taken to ensure consistency among the following aspects:

• Soil diversity

• The aim and accuracy of the analysis

• The nature of the chemical species to be analyzed

Important to pretreatment is the particle size distribution of the sample
in relation to the mass of sample taken for analysis. For the analysis of

1 Soil Sampling and Storage 39

organic contaminants, the mass taken in most cases is about 20 g. With
such a sample mass, and provided that the contaminant is homogeneously
distributed and the particles in the sample are smaller than about 2 mm,
further grinding of the sample is not necessary. If the sample contains large
particles or if the contaminant is heterogeneously distributed (for instance,
tar particles), it is not possible to take a representative test sample of about
20 g without grinding the sample. To improve the homogeneity, samples
are grinded to a size smaller than 1 mm. Prior to analysis very often no
information about the distribution of the contaminant in the soil is known.
Some analytical procedures start with a field-moist sample. Drying of the
sample will give lower extraction results, But because the sample is not dry,
grinding is not possible. In a situation in which accurate results are needed,
the best available pretreatment procedure should be used. If it is necessary
to establish whether the concentration is above a certain limit, and it is
already known that the soil is heavily polluted, the simplest pretreatment
procedure will perhaps meet the needs despite drawbacks. In that case,
however, the result may have to be presented as not representative of the
whole sample.

Three methods for the pretreatment of soil samples in the laboratory
prior to the determination of organic contaminants are applied in routine
analysis:

1. A method for pretreatment if VOCs are to be measured. Core test sam-
ples are taken from the sample and extracted according to the specific
analytical procedure. If composite samples are required, extracts of indi-
vidual samples are mixed. It is usually not possible to obtain composite
samples without severe losses of volatiles.

2. A method for pretreatment of moderately volatile to non-volatile organic
compounds where the result of analysis must be accurate and repro-
ducible. The sample contains particles larger than 2 mm and/or the con-
taminant is heterogeneously distributed: Samples are chemically dried
at a low temperature (-196 °C, liquid nitrogen). The freeze-dried sam-
ples are ground with a cross beater mill with a sieve of 1 mm (cryogenic
crushing). After grinding suitable test portions are processed according
to the specific analytical procedures. Composite samples can be prepared
by mixing of the ground samples. If the extraction procedure prescribes
a field-moist sample, drying and grinding is not possible. If the original
samples only contain a small fraction of particles greater than 2 mm and
the distribution of contaminants is likely to be homogeneous, grinding
may be omitted. In these two cases suitable test portions are directly
taken after mixing of the sample. To distinguish more volatile from less
volatile organic compounds, boiling points are used instead of vapor
pressure at ambient temperature. For some specific components in the

40 A. Paetz, B.-M. Wilke

group of moderately volatile compounds, freeze drying may give good
results. (In the International Standard freeze drying is not described.)

3. A method for pretreatment if non volatile organic compounds are to be
measured and the extraction procedure prescribes a field-moist sample,
or if the largest particles of the sample are smaller than 2 mm and the
contaminant is homogeneously distributed, mixing by hand is the only
pretreatment that need be applied. This procedure may also be used if
reduced accuracy and repeatability are acceptable.

The choice depends above all on the volatility of the organic compounds
under analysis. It also depends on the soil particle size distribution, the
heterogeneity of the sample, and the analytical procedure that is to follow.

1.5.2

Physical Analysis

Usually, the determination of soil physical parameters requires undisturbed
soil samples. Thus, pretreatment plays only a minor role. Exceptions are:

• Determination of the water content, which can be carried out to support
calculation of the analytical result, i.e., to standardize the result on dry
soil mass. In this case, the analysis can be performed in the laboratory
based on disturbed soil sample material. On the other hand, if the soil
water content needs to be determined on a volume basis, an undisturbed
sample must be taken and no further treatment applied before testing.

• The determination of the particle size distribution. Depending on the
range of particle sizes, the nature (chemistry) of the soil material, and the
objective of the investigation, a suite of different pretreatment procedures
maybe applied, including drying, slightly breaking aggregates, removing
specific kinds of materials, chemically breaking aggregates down, etc.
The matter is very complex and needs specialists' advice in most cases.

1.5.3

Biological Analysis

As a rule, soil for microbiological analyses under laboratory conditions
should be sampled in the field with a water content that facilitates sieving.
In the laboratory the soil should be processed (sieving) as soon as possible
after sampling. Soil fauna and plant tests can be also carried out on air-dried
samples. In this case the samples must be pretreated in order to achieve
optimum conditions for the species present.

1 Soil Sampling and Storage 41

1.6

Storage of Samples

1.6.1
General

• Soils samples for laboratory determinations are collected in many stud-
ies. In general the samples are taken at the site, mixed, or otherwise
treated at the site, packed in containers, and then transported to the lab-
oratory. Upon arrival at the laboratory the samples may again be treated
before being sent for analysis. Some samples may be stored directly for
later analysis. After analysis the samples may be discarded or stored.
The samples are stored when there is a need for further analysis, either
because parameters already determined require rechecking or a need
exists for making additional determinations in the future.

• The conditions for storage should be selected carefully at all stages from
the point of taking the sample. Examples of storage conditions are light,
temperature, humidity, accessibility, duration of storage, type of con-
tainers, and amount of storage. The documentation is also important.
Risk and security problems should be considered. Well-designed storage
conditions, such as provisions for monitoring, are particularly impor-
tant in large-scale studies where the number of samples may become
quite large over the years. Incorrectly chosen storage conditions may
lead to high costs and may render the samples unfit for future use.

• The effect of storage on biodiversity is not discussed because of the
difficulty to define this parameter.

• Radioactivity decay is generally not affected by storage and is not treated
in this standard. Radioactive change caused by loss or gain of matter
should be considered in connection with the appropriate compounds.

• Containers holding samples should be protected and sealed in such
a way that the samples do not deteriorate or lose any part of their content
during transport. Packaging should protect the containers from possible
external contamination, particularly near the opening, and should not
itself be a source of contamination. Most of the analytical procedures
used in chemical soil analysis recommend that soil samples be taken to
the laboratory immediately after sampling, but in some cases a range of
time is given during which the sample should arrive in the laboratory.

• Soil samples should be kept cool and dark during transportation and
storage.

42 A. Paetz, B.-M. Wilke

• Cooling or freezing procedures can be applied to increase the period
available for transport and storage. A cooling temperature of 4 ± 2°C
has been found suitable for many applications. But cooling and freezing
procedures should only be used in consultation with the analytical lab-
oratory. Freezing especially requires detailed control of the freezing and
thawing process in order to return the sample to its initial equilibrium
after thawing.

• Light-sensitive soil constituents require storage in darkness or, at least,
in light-absorbent containers.

• Undisturbed samples should be transported in the absence vibration or
other physical disorder in order to maintain the original structure.

• Disturbed samples, and especially non-cohesive, very dry soils, tend to
separate into different particle fractions during transportation. In such
cases the soil material should be re-homogenized before pretreatment
and analysis.

• Any national regulations regarding the packaging and transport of haz-
ardous materials should be observed.

1.6.2

Specific Considerations for Biological Parameters

Biological tests can be separated into soil microbiological, fauna, plant, and
biodegradation tests, and tests for the ecotoxicological characterization of
soils and soil materials. Storage conditions for soils used for these tests vary
over a wide range and depend on the organism or parameter to be tested.

Microbiological Tests

Samples should be stored in the dark at 4 ± 2 °C with free access of air. It
is preferable to use soils as soon as possible after sampling. If storage is
unavoidable, this should not exceed 3 months unless evidence showing con-
tinued microbial activity is provided. The active soil microflora decreases
with storage time, even at low temperatures, and the rate of decrease de-
pends on the composition of the soil and the microflora involved (see also
ISO 10381-6 1993).

If soil samples have to be stored for longer periods than 3 months,
freezing of samples at -20, -80, or -150 °C may be appropriate, although
not generally recommended. It has been shown for a number of soils from
temperate climates that storage at -20 °C for up to 12 months does not
inhibit microbial activity (e.g., ammonium oxidation). Soil samples for
phospholipid fatty acid (PLFA) and DNA analyses can be stored at -20 °C

1 Soil Sampling and Storage 43

for 1-2 years. Samples for rRNA analyses can be stored at -80 °C for the
same period. In the latter case the samples should be frozen immediately
at -180 °C (shock freezing with liquid nitrogen).

Longer storage periods are mainly needed if the influence of added
pollutants on soil microbes and microbial processes has to be tested with
the same soil material, or if the community structure (structural diversity;
PLFA, DNA, RNA) of soils has to be evaluated at a distinct point of time
during the year. In these cases the time needed for analyses can easily
exceed 3 months (chemical, pollutant testing). For structural analyses of
the microflora, storage at -4 °C is not suitable.

If longer storage of samples at temperatures below -20 °C is used, special
attention has to be given to the thawing of samples. Freeze-thaw cycles can
increase the availability of organic matter to micro-organisms (Haynes and
Beare 1996). For analyses of microbial activity (e.g., soil respiration) a thaw-
ing period of 1 day at 4°C and another 3 days at 20 °C is recommended.
Generally drying of soils is not recommended although air drying and
rewetting is a common physiological stress for the microbial communities
in surface soils. It has been shown that drying- rewetting events can induce
significant changes in microbial carbon and nitrogen dynamics that can
last for more than a month after the last stress (Fierer and Schimel 2002).
Rewetting after drying causes bursts of respiration and growth of distinct
populations of bacteria (Lund and Goks0r 1980).

Investigations indicate that fast thawing (1 day at 20 °C in an incubator)
results in smaller variations in microbiological parameters (e.g., microbial
activity) than the recommended slow thawing when compared to a frozen
control (Weinfurtner et al. 2002).

Biodegradation Tests

For testing the biodegradation of organic chemicals in soils (ISO 11266
1994; ISO 15473 2002), storage of soils should be avoided if possible because
activity of soil microorganisms will decrease in the course of time. Storage
at 4 °C up to 3 months is permissible. For the assessment of degradation
of chemicals in anaerobic soils under anaerobic conditions, the access of
oxygen should be avoided during storage.

Tests Involving Soil Fauna and Higher Plants

There are no specific recommendations for soil storage with respect to
soil fauna and higher plant tests in ISO standards. It is recommended to
store the soil samples under the same conditions as for testing of microbes
and microbial processes. The reason for this is that the availability and
effectiveness of pollutants is essentially governed by microbial activity.
The same is also true for plant testing. Additionally, the nutrient supply of

44 A. Paetz, B.-M. Wilke

test soils should be considered, especially if unknown contaminated soils
are tested, to avoid false negative results.

Ecotoxicological Testing

Generally, sieved samples should be stored in darkness. For microbial
analyses, soils and soil materials should be handled as described above.
For terrestrial analyses (e.g., plant tests, earthworm tests) samples can be
stored at 4 ± 2 °C for 3 months. For testing the leaching potential/ retention
function of soils and soil materials, water extracts for aquatic tests should
be prepared immediately after sieving. If the tests cannot be performed
within 7 days (storage of the extracts at 4 ± 2°C in the dark), extracts
should be stored at -20 °C.

1.6.3

Preparing the Samples After Storage

The procedures for preparing the samples after storage will depend on the
storage conditions and the analyses. It is not possible to give a general spec-
ification. Existing standards (e.g., ISO 11464 1994) have to be considered.
When a soil sample is stored for a long period of time, a vertical redistri-
bution may occur. A new mixing in a suitable mixer is advisable. For large
samples, this may not be sufficient. It is recommended that the sample
be spread in a thin layer on a plastic foil, and then the layer repeatedly
folded and spread it out again. Especially, the conditions of thawing have
to be defined because this can influence the determination of biological,
microbiological, and organic parameters. The soil samples stored below
°C must be unfrozen in original bags or containers.

References

Fierer N, Schimel P (2002) Effects of drying- rewetting frequency on soil carbon and nitrogen
transformations. Soil Biol Biochem 34:777-787

Haynes RJ, Beare MH (1996) Aggregation and organic matter storage in meso-thermal,
humid soils. In: Carter MR, Steward BA (eds) Soil structure and organic matter storage.
CRC/Lewis, Boca Raton, pp 213-262

ISO 10381-1 (2002) Soil quality - Sampling - Part 1: Guidance on the design of sampling
programmes

ISO 10381-2 (2002) Soil quality - Sampling - Part 2: Guidance on sampling techniques

ISO 10381-3 (2001) Soil quality - Sampling - Part 3: Guidance on safety

ISO 10381-4 (2003) Soil quality - Sampling - Part 4: Guidance on the procedure for investi-
gation of natural, near- natural and cultivated sites

ISO 10381-5 (1995) Soil quality - Sampling

ISO 10381-6 (1993) Soil quality - Sampling - Part 6: Guidance on the collection, handling
and storage of soil for the assessment of aerobic microbial processes in the laboratory

1 Soil Sampling and Storage 45

ISO 11266 (1994) Soil quality- Guidance on laboratory testing for biodegradation of organic

chemicals in soil under aerobic conditions
ISO 11277 (1998) Soil quality - Determination of particle size distribution in mineral soil

material - Method by sieving and sedimentation
ISO 11464 (1994) Soil quality - Pretreatment of samples for physico-chemical analysis
ISO 1 5473 (2002) Soil quality - Guidance on laboratory testing for biodegradation of organic

chemicals in soil under anaerobic conditions
ISO 15799 (2003) Soil quality - Guidance on the ecotoxicological characterization of soils

and soil materials
Lund V, Goks0r J (1980) Effects of water fluctuations on microbial mass and activity in soil.

Microbial Ecol 6:115-123
Weinfurtner K, Koerdel W, Schlueter C (2002) Probenahmerichtlinie fur eine Kryo-

lagerung von Bodenproben. Jahrestagungen GDCh-Fachgruppe Umweltchemie und

Okotoxikologie / SETAC-GLB, Braunschweig 2002

Determination of Chemical
and Physical Soil Properties

Berndt-Michael Wilke

2.1

Soil Dry Mass and Water Content

■ Introduction

Objectives. Measures of soil water content and dry mass are needed in prac-
tically all types of soil studies, e.g., determination of water holding capacity,
plant available water, infiltration, pore size distribution, permeability. With
respect to soil microbial processes and biological soil remediation, determi-
nation of optimum water content for measurement of microbial parameters
and activity, as well as determination of soil permeability for estimation of
the success of in situ remediation, is of essential importance.

Principle. Soil samples are dried at 105 ± 5°C until mass constancy is
reached. The differences in masses before and after drying are a measure
for the water content of soils. The water content is calculated on gravimetric
(gwater/^soii) o r o n volumetric basis (cm 3 water /cm 3 SO ii). The method described
below can be used for disturbed and undisturbed (sampling of soil using
coring sieves) soil samples. It is a direct laboratory measurement. The
procedure described can be used for the determination of dry mass on
a mass basis (ISO 11465 1993).

Theory. Under natural conditions all soils contain water. The amount of
water can be very low in air-dried soils. As a convention the total water
content and dry mass of soils are measured after drying at 105 °C (ISO
1 1465 1993). Thus, the water content of a soil is given as percent by weight
or volume of oven-dried soil. Water which is removed at higher tempera-
tures is not included in the definition of soil water. The soil water content
can be determined with direct and indirect methods. Direct methods are
more precise but time consuming. Indirect methods are mainly used for
continuous determination of water contents in the field. The most appro-
priate indirect method is the time domain reflectometry (TRD) method
(Topp et al. 2000). The optimum water content for microbial processes is in

Berndt-Michael Wilke: Institute of Ecology, Berlin University of Technology, Franklinstrafte
29, 10587 Berlin, Germany, E-mail: bmwilke@tu-berlin.de

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

48 B.-M. Wilke

the range of 40-60% of maximum water-holding capacity (WHC, Sect. 2.2),
or corresponds to the water content that is held in soil at suction pressures
of -0.01 to -0.031 MPa.

■ Equipment

• Drying oven, thermostatically controlled with forced air ventilation and
capable of maintaining a temperature of 105 ± 5 °C

• Desiccator with an active drying agent

• Analytical balance, accuracy 1 mg

• Container (moisture box, 25-100 mL) with lid, made of waterproof ma-
terial that does not adsorb moisture, capacity 25-100 mL for air-dried
soil samples and at least 100 mL for field-moist soil samples

• Spoon

■ Procedure

Air-Dried Soil Samples

1. Dry container with lid at 105 ± 5 °C and then cool it, with the lid closed,
in a desiccator for at least 45 min. Determine the mass (m ) of the closed
container with an accuracy of ± 1 mg.

2. Transfer 10-15 g of air-dried soil to this container using a spoon.

3. Determine the mass (mi) of the closed container and soil with an accu-
racy of ±1 mg.

4. Dry the container and soil in an oven at 105 °C until constant mass is
achieved. Dry the lid at the same time.

5. Cool the container with the lid closed in a desiccator for at least 45 min.

6. Remove the container from the desiccator and immediately determine
the mass (m 2 ) of the closed container containing the oven-dried soil with
an accuracy of ± 1 mg.

Field-Moist Soil Samples

1. Place the soil on a clean surface that does not absorb moisture (e.g.,
a glass plate) and mix well. Remove particles with a diameter > 2 mm.

2. Dry container with lid at 105 ± 5 °C and then cool it, with the lid closed,
in desiccator for at least 45 min. Determine the mass (m ) of the closed
container with an accuracy of ± 1 mg.

3. Transfer 30-40 g of soil to this container using a spoon.

2 Determination of Chemical and Physical Soil Properties 49

4. Determine the mass (mi) of the closed container and soil with an accu-
racy of ±10 mg.

5. Dry the container and soil in an oven at 105 °C until constant mass is
achieved. Dry the lid at the same time.

6. Cool the container with the lid closed in a desiccator for at least 45 min.

7. Remove the container from the desiccator and immediately determine
the mass (m 2 ) of the closed container containing the oven-dried soil with
an accuracy of ± 10 mg.

■ Calculation

Calculate the dry mass content ( w dm ) or water content ( w H2 o ) on a dry mass
basis expressed as percentages by mass to an accuracy of 0.1% (m/m) using
the following equations:

m 2 - m /

w dm = — x 100 (2.1)

mi - m

mi - m 2 ,

w H0 = — x 100 (2.2)

m 2 -m

m mass of the empty container with lid (g)

mi mass of the container with air-dried soil or field-moist soil (g)

ra 2 mass of the container plus oven-dried soil (g)

■ Notes and Points to Watch

• With contaminated soil samples, special measures must be taken. Avoid
any contact with the skin. Special measures must be taken during the
drying process in order to prevent contamination of the laboratory at-
mosphere. The procedures must be performed as quickly as possible to
prevent evaporation.

• In general decomposition of organic material can be neglected at tem-
peratures up to 105 °C. However, for soil samples with a high organic
matter content (> 10% m/m) the method of drying should be adapted
by drying to a constant mass at 50 °C.

• Some minerals similar to gypsum lose chemically combined water at
a temperature of 105 °C.

• If volatile organic substances are present, the method will not give a re-
liable determination of the water content.

50 B.-M. Wilke

2.2

Water-Holding Capacity

■ Introduction

Objectives. Microbiological laboratory tests (e.g., respiration measure-
ments ISO 16072 2002; nitrogen mineralization ISO 14238 1997; biodegra-
dation ISO 11266 1994) are carried out under optimum water conditions.
These range from 40-60% of soil WHC. In order to adjust the optimum
water content of a given soil the maximum WHC has to be determined.

Principle. A cylinder with a perforated base is filled with soil, capped and
immersed in water and drained. The quantity of water taken up by the
soil is determined by weighing, drying to constant mass at 105 °C, and
reweighing.

Theory. Microbial transformations in soils are moisture dependent. Mois-
ture must be adequate for decomposition to proceed. High moisture levels
reduce activities of aerobic microorganisms due to a deficiency of oxygen.
Therefore, the soil moisture content is adjusted to optimum conditions
in microbial laboratory experiments. For most aerobic processes it ranges
from 40 to 60% of WHC. Alternatively, the water content can be also ad-
justed by means of pore water pressure. Water contents of 40-60% of WHC
equal 0.01-0.03 MPa.

■ Equipment

• Cylinder (glass, plastic, metal; with cap) of known volume, of about
50-150 mm length and 50-100 mm in diameter with a perforated base

• Water bath at room temperature

• Tray with a drainage hole, containing wet, fine, quartz sand (20-50 mm)

• Oven, capable of maintaining a temperature of 105 ± 2 °C

• Beaker, 250 mL

• Desiccator

• Balance, accurate to ±0.01 g.

■ Sample Preparation

As rule fresh soil samples screened through a 2 mm sieve are used.

2 Determination of Chemical and Physical Soil Properties 51

■ Procedure

Determination of Maximum WHC

1 . Cover the perforated base of the cylinder with a filter paper and fill it with
field-moist soil (three parallels per sample). Fill the soil in small portions
and provide homogeneous spreading by gentle tapping of the cylinder.

2. Submerge the cylinder in the water bath at room temperature with the
water level lower than the soil surface. When the soil is moistened to the
surface, lower the cylinder to the soil surface and leave it in this position
overnight.

3. Remove the cylinder from the water and place the capped cylinder on
the tray of sand and allow to drain. Capping of the cylinder is crucial to
avoid evaporation of water.

4. Weigh the cylinder hourly beginning after 3 h until constant weight is
achieved. Remove the soil from the cylinder into a 250 mL beaker and
dry it at 105 °C in an oven for 24 h (minimum). Cool the samples in
a desiccator and weigh again.

Adjustment of a Defined Water Content

If the actual water content is lower than wanted, the soil is spread as a thin
layer and the needed amount of water is evenly sprayed in small portions on
the surface. The soil should be mixed thoroughly after each water addition.
Reduction of soil volume (formation of aggregates) should be avoided
during addition of water.

In case of a higher actual water content the soil is dried at room temper-
ature until the wanted moisture is reached. Drying of the soil surface has
to be avoided by periodic mixing.

The water content can also be adjusted by using a device (e.g., porous
funnel apparatus) whereby the saturated soil can be drained stepwise to
a known soil water (matric) potential. First, the field-moist soil is saturated
on a ceramic plate. Subsequently the surplus of water is drained until the
wanted water content is reached using a vacuum pump.

■ Calculation

Calculate the WHC using the following equation:

WHC max (% dry mass) = — x 100 (2.3)

m t - rab

m s mass of beaker containing water saturated soil (g)
m t mass of beaker containing oven-dried soil (g)
mb mass of beaker (g)

52 B.-M. Wilke

2.3

Bulk Density - Total Porosity

■ Introduction

Objectives. Determination of bulk density is a widely used soil parameter.
Bulk density is needed for converting water percentage by weight to content
by volume, calculating the porosity and void ratio when the particle density
is known (Blake and Hartge 1986). It can by used to estimate the weight
of a volume of soil too large to weigh and to calculate the total mass of
a pollutant in a given soil volume.

The bulk density gives a rough estimation of the aeration and permeabil-
ity of a soil. The lower the bulk density, the higher is the permeability. Bulk
density varies with structural conditions of the soil. Therefore, it is related
to packing and often used as a measure for soil structure. In swelling soils
(e.g., clay soils) it varies with soil water content (Hartge 1968). In these
soils the bulk density obtained should be compared with the soil water
content at the sampling time. There are three methods available for the
determination of soil bulk density: core method, excavation method, and
clod method. All methods are standardized (ISO DIS 11272 1998).

Theory. Soil is a porous three-phase system composed of air, water, and
solids. The relative distribution of these three components is important
to understand the hydraulic properties of the soil. The dry bulk density
is the ratio of oven-dried solids to volume of soil. It is expressed in SI
units, e.g., g/cm 3 , kg/m 3 , or Mg/m 3 . It reflects the structural condition of
the soil at given depth. Bulk densities of mineral soils may range from
< 0. 8 to > 1.75 g/cm 3 (Schlichting et al. 1995). The total porosity (S t ) can be
calculated if the particle density (p p ) and the bulk density (f>b) are known,
according to the following equation:

St = 1 - (pi/ft) (2.4)

(Danielson and Sutherland 1986). As a rule of thumb the density of quartz
(p p = 2.65 g/cm 3 ) is used as particle density (p p ) of mineral soils.

2.3.1

Core Method

Principle. This method is only applicable to stoneless and slightly stony
soils. A cylindrical metal sampler is pressed or driven into the soil to the
desired depth. It is carefully removed to preserve a known volume of sample
as it existed in situ. The sample is dried in an oven at 105 °C and weighed.

2 Determination of Chemical and Physical Soil Properties 53

■ Equipment

• Core sampler holder, thin walled metal cylinders with a volume of
100-400 cm 3 , a steel cap for driving into the soil, and a driver

• Oven, heated and ventilated, capable of maintaining a temperature of
105 °C

• Desiccator

• Laboratory balance, capable of weighing to an accuracy of 1/1,000 of the
measured value

■ Procedure

1. Drive or press the core sampler into either a vertical or a horizontal soil
surface enough to fill the sampler but not so firmly as to compress the
soil in the confined space of the sampler.

2. Carefully remove the sampler and its contents to preserve the natural
structure, and trim the soil extending beyond each end of the sample
holder with a straight-edged knife or a sharp spatula. The soil sample
volume thus established is the same as the volume of the sample holder.

3. Take at least six core samples from each soil horizon.

4. Place the holders containing the sample in an oven at 105 °C until con-
stant mass is achieved (minimum 48 h).

5. Remove the samples from the oven and allow them to cool in the desic-
cator.

6. Weigh the samples immediately after removal from the desiccator (m t ).
Control mass is reached when the differences in successive weighing of
the cooled sample, at intervals of 4 h, do not exceed 0.01% of the original
mass of the sample.

■ Calculation

The dry bulk density is calculated using the following equations:

Qh = — (2.5)

m& = m t — m s (2.6)

£>b bulk density (g/cm 3 )

ma mass of the core sample dried at 105 °C minus mass of the core sample
holder (g)

54 B.-M. Wilke

V volume of the core sample holder (cm 3 )

m t mass of the sample holder plus soil sample dried at 105 °C (g)

m s mass of the empty core sample holder (g)

■ Notes and Points to Watch

• Swell/shrink soils (e.g., clays, muds, peats) change their bulk density
with changing water content. Such soils should be sampled in a moist
state (e.g., field capacity). In addition they should be sampled in a wet
state (water saturation) and a dry state.

• If bulk density and water content (Sect. 2.1) are the only parameters of
interest, it is not necessary to keep the samples in their holders. A single
core sample holder can be reused if each sample is transferred to another
container.

• The undisturbed samples in the core sample holder can be also used for
other measurements such as pore-size distribution (Sect. 2.5), conduc-
tivity, or water retention.

• It is normally worthwhile to combine a measurement of the water content
with a measurement of bulk density. In this case, it is necessary to
transport the samples without allowing loss of water by evaporation and
to begin the laboratory operations by weighing the fresh sample.

2.3.2

Excavation Method

Principle. Dry bulk density is determined by excavating a quantity of soil,
drying and weighing it, and determining the volume of the excavation by
filling it with sand. The method is applicable to soils containing gravel
and/or stones.

■ Equipment

• Earth-digging equipment, e.g., spade with sharp-edged blade

• Sampling equipment (flat blade spade, knife, pick, spade chisel, hammer)

• Equipment for collecting and cleaning (plastic sheet, brush, heat-resistant
plastic bags or containers)

• Plastic film, thin, flexible but stable

2 Determination of Chemical and Physical Soil Properties 55

• Equipment for spreading sand, including funnel with a gauging rod
(falling height beneath the funnel mouth should be 5 cm), graduated
cylinder of 1 dm 3 capacity

• Dry, graded sand of known volume, particle diameter 500-700 yam

• Balance capable of weighing 1 g

• Oven, heated and ventilated, capable of maintaining a temperature of
105 °C

• Vacuum desiccator with self indicating desiccant

• Sieve with 2-mm mesh size

■ Procedure

Field

1. Level off the soil surface with the straight metal blade (Fig. 2.1a).

2. Dig a hole in the leveled soil having a representative content of larger
gravel and stones (volume 20 dm 3 containing 30% stones) avoiding com-
paction of sides (Fig. 2.1b).

3. Put the excavated soil in bags or containers for laboratory analysis. (Large
nonporous stones such as granite can be separated in the field, cleaned
with a stiff brush, and weighed on a field balance).

4. Line the hole with a plastic film.

5. Fill the hole to excess with a known volume of sand from a height of 5 cm
using the funnel (Fig. 2.1c). Level the surface with the blade without
packing down.

6. Pour the excess sand into the graduated measuring cylinder and read
the volume (Fig. 2. Id). The difference from the initial volume of sand is
the volume V in the hole.

Laboratory

1. Determine the mass of the moist excavated soil (in g) with a balance

2. Separate the stones and gravel from the fine soil with a 2-mm sieve and
weigh them on a balance (m xw ).

3. Dry the stones and the gravel in the oven at 105 °C and weigh them after
cooling on the laboratory balance (m x ).

4. Determine the water content of the fine soil (< 2 mm) by drying a rep-
resentative sample (5-10 g) of known mass in the oven (105 °C) until

B.-M. Wilke

Fig. 2.1. Excavation method, field procedure (adapted from ISO 11272 1998). a Level off soil
surface; b dig a hole; c fill with sand; d remove excess sand and measure its volume

constant mass is reached. Remove the sample from the oven and cool it
in the desiccator. Weigh the sample on a laboratory balance. Calculate
the water content (w) as a mass ratio of the moist sample.

Qh =

m x - m tp

■ Calculation

The bulk density of the soil layer is calculated using the following equations:

(2.7)

(2.8)
m w = m pw x m tw (2.9)

(2.10)

™tp ~ wip w ■ ■ trixw ' ' wi w

TH-Ua; — ffltiAA; ' ' ffl

■tw

l xw

£>b bulk density (g/cm 3 )

m x mass of stones and dry gravel (g)

m tp mass of the dry fine soil (g)

2 Determination of Chemical and Physical Soil Properties 57

V volume of the hole (cm 3 )

m pw mass of the excavated moist soil (g)

m w mass of the water from excavated fine soil (g)

w water content of the excavated moist fine soil (g/g oven-dried soil)

m tw mass of the moist fine soil (g)

m xw mass of moist gravel and stones (g)

■ Notes and Points to Watch

• Holes should have smooth, rounded walls.

• Protruding stones should be included in the sample

• A heavy pair of scissors can be used to cut roots at the wall surface.

2.3.3

Clod Method

Principle. The bulk density of clods, or coarse peds, is calculated from
their mass and volume. The volume is determined by coating the clod
with a water-repellent substance and by weighing it first in air, then again
while immersed in a liquid of known density, making use of Archimedes'
principle.

■ Equipment

• Earth digging equipment (flat shovel, spade, pick)

• Sampling equipment (small flat-bladed spade, knife, chisel, hammer)

• Container of molybdenum sulfide (MoS 2 ) in heavy oil

• Laboratory balance, capable of weighing suspended samples (Fig. 2.2)

• Thermometer

■ Procedure

1. Separate clods or peds of about 50-200 cm 3 , trim off protrusions and
cut off roots with scissors.

2. Weigh soil clods or peds with a laboratory balance and coat them in oil.

B.-M. Wilke

1
2

3
4

Fig. 2.2. Laboratory balance to determine the volume of clods by weighing in air and water.
1 Compensating weights; 2 thin wire; 3 small container; 4 large container filled with water;
5 balance

3. Weigh the coated clod again once in air and once immersed in water.

4. Measure the temperature of the water and determine its density

5. To obtain a correction for the water content of the soil, break the clod,
remove an aliquot of soil, and weigh it before and after drying in an oven
atl05±2°C.

■ Calculation

1. The oven dry mass of the soil clods is calculated using the equation

m& = ra/(l + w)

(2.11)

ma net mass of the oven-dried clod (g)
m net mass of the moist clod in air (g)
w water content of the subsample (g of water/g of oven-dried soil)

2. Calculate the bulk density of the dried clod using the equation

mass

Pb =

pw x m d

volume m- m w + m (p - £>w)

(2.12)

05 bulk density of oven-dried sample (g/cm 3 )

(? density of the coating oil (g/cm 3 )

p w density of water at temperature of determination (g/cm 3 )

ma oven-dried mass of soil sample, i.e., clod or ped (g)

2 Determination of Chemical and Physical Soil Properties 59

m mass of soil sample in air (g)

m w mass of soil sample plus coating in water (g)

m mass of coating in air (g)

■ Notes and Points to Watch

• The clod method gives usually higher bulk density values than the other
methods as it does not take interclod space into account.

• Clods on or near the soil surface are likely to be unrepresentative as these
are often formed by packing or plowing.

• Several other substances have been used to seal the clods against water
including Saran solution (Dow Chemical, Rolling Meadows, IL, USA),
paraffin, and wax mixtures.

• See also Blake and Hartge (1986).

2.4

Water Retention Characteristics - Pore Size Distribution

■ Introduction

Objectives. The spaces between soil particles are known as the soil pores.
They are filled either with soil-air or water (soil solution) depending on the
pore size and the water saturation of the soil. With respect to their equiv-
alent diameter, soil pores can be divided into wide coarse (> 50 yam), tight
coarse (10-50 yam), medium (0.2-10 yam) and fine (< 0.2 yam). Pore sizes
were assigned in accordance with adaptation to the water content at char-
acteristic matric pressures. Equivalent diameters of 50 and 10 yam comply
with the water content of soils at field capacity (6 and 30 kPa), 0.2 yim with
the water content at the permanent wilting point (1,500 kPa). The range of
water available to plants and microorganisms is between field capacity and
the permanent wilting point. Water stored at matric pressures > 1,500 kPa
is neither accessible to fine plant roots nor to microorganisms. The pore size
distribution of a given soil depends on its density and texture. Thus, it in-
fluences its aeration, permeability, transport of chemicals dissolved in soil
water, and the water- retention characteristics of a soil. Direct evaluation of
the size, configuration, and distribution of soil pores is impossible due to
their extremely complicated nature. However, the size distribution can be
measured by determination of water content at different matric pressures.
Besides providing an assessment of the equivalent pore size distribution
(e.g., identification of coarse, medium, and fine pores), the results using
the methods to be described can be used for other purposes, for example:

60 B.-M. Wilke

• For assessment of the water retention characteristics

• To determine water content at specific matric pressures (e.g., for micro-
bial degradation studies)

• To ascertain the relationship between the negative matric pressures and
other soil physical properties (e.g., hydraulic conductivity, thermal con-
ductivity)

• To determine the drainable pore space (e.g., pollution risk assessment)

• To determine indices for plant- available water in the soil (e.g., for irri-
gation purposes)

Principle. Undisturbed soil samples (soil cores) are used for the measure-
ment at the high matric pressure range 0-100 kPa. The samples are satu-
rated with de-aerated water or calcium sulfate solution (0.005 mol/L) and
subsequently drained using sand, kaolin, or ceramic suction tables (for
pressures from to 20 kPa) and pressure plate extractors (for determina-
tion of pressures from -5 to -1,500 kPa). At equilibrium status, soil samples
are weighed, oven dried and reweighed to determine the water content. The
results are given either as volume fraction or mass ratio. The differences in
volume fractions at different suction pressures give the pore volume (e.g.,
medium pores in vol%), the differences in mass fractions give the water
content retained in these pores. Two standardized (ISO 11274 1998) meth-
ods are described, namely use of sand, kaolin, or ceramic suction tables
for determination of water contents at pressures of to -50 kPa, and use of
pressure plates for determination of pressures from -5 to -1,500 kPa.

Theory. Soil water content and matric pressure are related to each other. At
zero matric pressure the soil is saturated and all pores are filled with water.
As the soil dries matric pressure decreases and pores will empty according
to their equivalent diameter. Large coarse pores (> 50 p.m) will drain at
a matric pressure of > -6kPa, tight coarse pores (10-50 }im) at -6 to
30 kPa, medium pores at -30 to -1,500 kPa, and fine pores at < -1,500 kPa.

■ Sampling

1. It is essential that undisturbed soil samples be used for measurement
at the matric pressure range to -lOOkPa, since soil structure has
a strong influence on water- retention properties. Use either undisturbed
cores or, if appropriate, individual peds for low matric pressure methods
(< -100 kPa). Soil cores shall be taken in a metal or plastic cylinder of
a height and diameter such that they are representative of the natural
soil variability and structure. The dimensions of samples taken in the
field are dependent on the texture and structure of the soil and the test

2 Determination of Chemical and Physical Soil Properties 61

Table 2.1. Recommended sample sizes (height x diameter) for the different test methods

Test method

Structure

Coarse

Medium

Fine

Suction table
Pressure plate

50 x 100 mm

40 x 76 mm
10 x 76 mm

24 x 50 mm
10 x 50 mm

method which is to be used. Table 2.1 gives guidance on suitable sample
sizes for the different methods and soil structure.

2. To ensure minimal compaction and disturbance to structure, take soil
cores carefully, either by hand pressure in suitable material or by using
a suitable soil corer. Take a minimum of three representative replicates for
each freshly exposed soil horizon or layer; more replicates are required
in stony soils. Dig out the cylinder carefully with a trowel, roughly trim
the two faces of the cylinder with a knife. If necessary adjust the sample
within the cylinder before fitting lids to each end, and label the top clearly
with the sample grid reference, the direction of the sampling (horizontal
or vertical), the horizon number, and the sample depth.

3. Wrap the samples (e.g., in plastic bags) to prevent drying. Wrap ag-
gregates (e.g., in aluminum foil or plastic film) to retain structure and
prevent drying. Alternatively, excavate undisturbed soil blocks measur-
ing approx. 30 cm 3 in the field, wrap in metal foil, wax (to retain structure
and prevent drying), and take to the laboratory for subdivision. Store the
samples at 1-2 °C to reduce water loss and suppress biological activity
until they can be analyzed. Treat samples having obvious macrofaunal
activity with a suitable biocide, e.g., 0.05% copper sulfate solution.

■ Sample Preparation

1. To prepare samples for water- retention measurements at pressures great-
er than -50kPa, trim undisturbed cores flush with the ends of the con-
tainer and replace one lid with a circle of polyamide (nylon) mesh (or
similar close-weave material or paper if the water- retention character-
istic is known) secured with an elastic band. The mesh will retain the
soil sample in the cylinder and enable direct contact with the soil and
the porous contact medium. Avoid smearing the surface of clayey soils.
Remove any small projecting stones to ensure maximum contact and
correct the soil volume if necessary. Replace the other lid to prevent
drying of the sample by evaporation. Prepare soil aggregates for high
matric pressure measurements by leveling one face and wrapping other

62 B.-M. Wilke

faces in aluminum foil to minimize water loss. Disturbed soils should be
packed into a cylinder with a mesh attached. Firm the soil by tapping
and gentle pressure to obtain a specified bulk density.

2. Weigh the prepared samples. Ensure that the samples are brought to
a pressure of less than the first equilibration point by wetting them, if
necessary, by capillary rise, mesh side or leveled face down on a sheet of
foam rubber saturated with de-aerated tap water or 0.005 mol/L calcium
sulfate solution. Weigh the wet sample when a thin film of water is seen
on the surface. The time required for wetting varies with initial soil
water content and texture. Soils are ideally field moist when the wetting
is commenced. General guidelines for wetting times are:

sand: 1-5 days

loam: 5-10 days

clay: 5-14 days or longer

peat: 5-20 days.

Very coarse pores are not water filled when the soil sample is saturated
by capillary rise.

2.4.1

Determination of Soil Water Characteristics

Using Sand, Kaolin, and Ceramic Suction Tables

Principle. Suction tables are suitable for measurement of water contents at
matric pressure from to -50 kPa. A negative matric pressure is applied to
coarse silt or very fine sand held in a rigid watertight non-rusting container
(a ceramic sink is particularly suitable). Soil samples placed in contact
with the surface of the table lose pore water until their matric pressure is
equivalent to that of the suction table. Equilibrium status is determined by
weighing samples on a regular basis, and soil water content by weighing,
oven drying, and reweighing.

■ Equipment

• Large ceramic sink or other watertight, rigid, non-rusting container with
outlet in base (dimensions about (50 x 70 x 25 cm) and close-fitting cover

• Tubing and connecting pieces to construct a draining system for the
suction table

• Sand, silt, or kaolin, as packing material for the suction table (Com-
mercially available graded and washed industrial sands with a narrow

2 Determination of Chemical and Physical Soil Properties

particle size distribution are most suitable. The particle size distribu-
tions of some suitable sand grades and the approximate suctions they
can attain are given in Table 2.2. It is permissible to use other packing
materials, such as fine glass beards or aluminum oxide powder, if they
can achieve the required air entry values. Alternatively to sand, silt, or
kaolin suction tables, ceramic plates can be used.

Leveling bottle, stopcock, and 5-L aspirator bottle

Tensiometer system (optional)

Drying oven, capable of maintaining a temperature of 105 ± 2 °C

Balance capable of weighing with an accuracy of 0.1% of the measured
value

■ Procedure

1 . Prepare suction tables using packing material that can attain the required
air entry values (Table 2.2).

2. Prepare soil cores as described (see above).

3. Weigh the cores and then place them on a suction table at the desired
matric pressure.

4. Leave the cores for 7 days. The sample is than weighed, and thereafter
weighed as frequently as needed to verify that the daily change in mass of
the core is less than 0.02%. The sample is than regarded as equilibrated.

Table 2.2. Examples of sands and silica flour suitable for suction tables

Type

Coarse sand

Medium

Fine sand

Silica flour

Use

Base of

suction

tables

Surface of Surface of Surface of

suction tables suction tables suction tables

(5 kPa matric (11 kPa matric (21 kPa matric

pressure) pressure) pressure)

Typical particle
size distribution

Percent content

> 600 um

200-600 um

100-200 jim

63-100 um

20-63 um

<20um

64 B.-M. Wilke

5. Move the equilibrated sample to a suction table of a lower pressure or
dry it in an oven at 105 ± 5 C.

6. Samples which have not attained equilibrium should be replaced firmly
onto the suction table and the table cover replaced to minimize evapo-
ration from the table.

■ Calculation

Soils Containing <20% Stones (>2mm)

1. Calculate the water content mass ratio at a matric pressure p m using the
formula:

w(p m ) = -^ (2.13)

m d

w(p m ) water content mass ratio at a matric pressure p m (g)
m(p m ) mass of the soil sample at a matric pressure p m (g)
ma mass of the oven-dried soil sample (g)

2. Calculate the water content on a volume basis at matric pressure p m using
the formula:

/ x m (pm) - md

e p m = -^r 1 — - (2.14)

v y V x p w

0(pm) water content mass ratio at a matric pressure p m
(cm 3 water/ cm 3 soil)

m(p m ) mass of the soil sample at a matric pressure p m (g)

ma mass of the oven-dried soil sample (g)

V volume of the soil sample (cm 3 )

p w density of water (g/cm 3 )

Conversion of Results to a Fine Earth Basis

The stone content of a laboratory sample may not accurately represent the
field situation. Therefore, conversion of data to a fine earth basis may be
required. Conversion of results derived from suction methods to a fine
earth basis (f) is required for soils containing stones (> 2 mm) according
to the following equation:

"• ■ Ay <2 - i5)

2 Determination of Chemical and Physical Soil Properties 65

0f water content of the fine earth expressed as a volume fraction
6 S volume of stones, expressed as a fraction of total core volume
6 t water content of the total soil, expressed as a volume fraction

2.4.2

Determination of Soil Water Characteristics

by Pressure Plate Extractor

Principle. Pressure plate extractors are suitable for measurement of water
contents at matric pressure -5 to -1,500 kPa. Several small soil cores are
placed in contact with a porous ceramic plate contained within a pressure
chamber. A gas pressure is applied to the air space above the samples and
soil water moves through the plate to be collected in a burette/measuring
cylinder or similar collecting device. At equilibrium status, soil samples
are weighed, oven-dried, and reweighed to determine the water content at
the predetermined pressures.

■ Equipment

• Pressure chamber with porous ceramic plate

• Sample retaining rings/soil cores with plastic discs or lids

• Graduated burette

• Air compressor (1.700 kPa), nitrogen cylinder, or other suitable pressur-
ized gas

• Pressure regulator and test gauge

• Drying oven capable of maintaining a temperature of 105 ± 2.0 °C

• Balance capable of weighing to ±0.01 g

■ Procedure

1. Take small soil cores of approx. 5 cm diameter and 5-10 mm in height
in situ or from larger undisturbed cores.

2. Place at least three replicates on a pre-saturated plate of appropriate
bubbling pressure.

3. Wet the samples by immersing the plate and the samples to a level just
above the base of the core and waiting until a thin film of water can be
seen on the surface of the sample.

66 B.-M. Wilke

4. Cover the bottom of the extractor with water to create a saturated
atmosphere.

5. Place a plastic disc lightly on top of each sample to prevent evaporation.

6. To apply the desired pressure, remove excess water from the porous
plate and connect the outflow tube to the burette via the connector in
the chamber wall. The pressure is supplied via regulators and gauges
from a nitrogen cylinder or by a mechanical air compressor.

7. The pressure (from whatever source) should slightly exceed the lowest
matric pressure required.

8. Apply the desired gas pressure p, check for any gas leaks, and allow the
samples to come to equilibrium by recording on a daily basis the volume
increase in the burette. When this remains static, the samples have come
to equilibrium; the matric pressure p m of the samples equals -p.

9. To remove the samples, clamp the outflow tube to prevent a backflow
of water, and release the air pressure.

10. Weigh the samples plus sleeve immediately.

11. Carry out sequential equilibration of the core at different pressures
by removing and weighing the core at equilibrium, reinserting it, and
resetting the pressure.

12. Moisten the ceramic plate with a fine spray of water to re-establish
hydraulic contact.

13. When the last equilibrium has taken place, dry at 105 °C and determine
the oven-dried mass of the soil plus sleeve.

■ Calculation

Stoneless Soils

Calculate the water content volume fraction (0) using the formula:

0{p m ) = ^l 1 ( 2 .1 6 )

Q(pm) water content mass ratio at a matric pressure p m (cm 3 water/cm 3
soil)

m(p m ) mass of the soil sample at a matric pressure p m (g)

ma mass of the oven-dried soil sample (g)

2 Determination of Chemical and Physical Soil Properties 67

V volume of the soil sample (cm 3 )

p w density of water (g/cm 3 )

Stony Soils

Samples containing any stones (> 2 mm) shall not form part of the pressure
chamber or membrane sample since the sample volume is very small. After
oven-drying, determine the volume of stones in the original soil core from
a field measurement and make a correction to convert 0f values to total
soil (0 t ).

e t = e f (i-e s ) (2.17)

0f water content of the fine earth in the pressure vessel at equilibrium
expressed as volume fraction

6 S volume of stones, expressed as a fraction of total core volume

6 t water content of the total soil, expressed as a volume fraction

For a soil containing a volume fraction of non porous stones of 0.05 the
water content is:

t = f x 0.95 (2.18)

Evaluation of Results: Pore Size Distribution

Pore volumes of coarse, medium, and tight pores in vol% of total soil
volume can be calculated as follows:

Large Coarse Pores (Equivalent Diameter > SOjjm)

V\c V = (0pmO-0pm-6) * 100 (2.19)

Vi C p volume of large coarse pores (% of total soil volume)
0pmo volumetric water content at water saturation (p m = kPa)
0pm-6 volumetric water content at a matric pressure of p m = -6 kPa

Tight Coarse Pores (Equivalent Diameter 10-50 pm)

V tcv = (0 pm -6 " 0pm-3o) x 100 (2.20)

V t cp volume of large coarse pores (% of total soil volume)
0pm-6 volumetric water content at water saturation (p m = -6 kPa)
0pm-3o volumetric water content at a matric pressure of p m = -30 kPa

68 B.-M. Wilke

Medium Pores (Equivalent Diameter 0.2-10 pm)

V mV = {0-30 - 0pm-15Oo) X 100 ( 2 - 21 )

V mp volume of large coarse pores (percent of total soil volume)
0pm-3o volumetric water content at water saturation (p m = -30 kPa)
0pm-i5oo volumetric water content at a matric pressure of p m = -1,500 kPa

Fine Pores (Equivalent Diameter < 0.2 pm)

Vfp = 0pm-15OO x 100 (2.22)

Vfp volume of fine pores (% of total soil volume)

0pm-i5oo volumetric water content at a matric pressure of p m = -1,500 kPa

■ Notes and Points to Watch

• If a containing sleeve is used, it should be weighed and the mass deducted
from the total mass of the soil core to give m(p m ).

• If stones are porous, carry out separate water retention measurements
and correct fine earth values according to their volume.

2.5
Soil pH

■ Introduction

Objectives. Soil pH is one of the most indicative measurements of the soil
chemical properties. All (bio)chemical reactions in soils are influenced by
proton (H + ) activity, which is measured by soil pH. Values of pH of most
natural soils (measured in 0.01 M CaCl 2 ) vary between < 3. 00 (extremely
acid) and 8.00 (weakly alkaline). Solubility of various compounds in soils
is influenced by soil pH (e.g., heavy metals) as well as by microbial activ-
ity and microbial degradation of pollutants. The optimum pH values for
pollutant-degrading microorganisms range from 6.5 to 7.5 (Kastner 2001).
Determination of soil pH is standardized in ISO DIS 10390 (2002).

2 Determination of Chemical and Physical Soil Properties 69

Principle. A pH measurement is normally made by either a colorimetric or
an electrometric method. The former involves suitable dyes or acid-base
indicators. Indicator strips can be used for rough estimation of soil pH.
Normally, pH values of soils are measured by means of a glass electrode
in a soil solution slurry that contains a fivefold volume of water containing
lMKClor0.01MCaCl 2 .

Theory Soil pH is a measure of the activity of ionized H (H + , H 3 + ) and
defined as the negative logarithm of the H + /H 3 + ion activity in mol/L.
Soil acidity results from soluble acids in the soil solution, e.g., organic acids
and carbonic acid. Further acidic cations in the soil solution are Al 3+ and
Fe 3+ . Al 3+ ions exists in water as an [A1(H 2 0) 6 ] 3+ complex which dissociates
intoH 3 + ions according to [A1(H 2 0) 6 ] 3+ + H 2 ^ [A1(H 2 0) 5 ] 2+ + H 3 +
(pK a = 5. 0). A stronger cationic acid producer is Fe 3+ (pK a = 2. 2), which
due to the low solubility of iron oxides only exists below pH 3.

Soil pH is influenced by various factors, namely, the nature and type of
inorganic and organic constituents (that contribute to soil acidity), the
soil/solution ratio, the salt or electrolyte content, and the C0 2 partial
pressure. A pH measurement in water includes easily dissociated pro-
tons while 0.01 M CaCl 2 and 1 M KC1 solutions also mobilize exchangeable
H + . They are used to simulate soil solutions of arable soils (CaCl 2 ) and
forest soils (KC1) in temperate humid climates. Values of pH measured at
constant salt concentrations reflect seasonal variations to a lower degree
(Page et al. 1982); and those measured in 0.01 M CaCl 2 are 0. 6 ± 0. 2 units
lower than pH H2 o values, because H + and Al 3+ ions are partly exchanged
byCa 2+ .

■ Equipment

• Shaking or mixing machine

• pH meter with slope adjustment and temperature control (in case of pH
values > 10, an electrode specifically designed for that range is to be
used)

• Glass electrode and a reference electrode or a combined electrode of
equivalent performance

• Thermometer capable of measuring to the nearest 1 °C

• Sample bottle (50 mL) made of borosilicate glass or polyethylene with
a tightly fitting cap

• Spoon of known capacity (at least 5.0 mL)

70 B.-M. Wilke

■ Reagents

• Water with a specific conductivity not higher than 0.2 mS/m at 25 °C and
apH> 5.6

• Potassium chloride solution (KC1 1 mol/L)

• Calcium chloride solution (CaCl 2 0.01 mol/L)

• Solution for the calibration of the pH meter

• Buffer solution, pH 4.00 at 20 °C: dissolve 10.21 g of potassium hydrogen
phthalate (C6H5O4K, dried at 1 10-120 °C for 2h before use) in water
and dilute to 1,000 mL at 20 °C.

• Buffer solution, pH 6.88 at 20 °C: dissolve 3.39 g of KH 2 P0 4 and 3.53 g of
Na 2 HP0 4 in water and dilute to 1,000 mL at 20 °C.

• Buffer solution, pH 9.22 at 20 °C: dissolve 3.80 g of Na 2 B 4 7 x 10H 2 Oin
water and dilute to 1,000 mL at 20 °C. The buffer solutions are stable for
1 month when stored in polyethylene bottles. Alternatively, commercially
available buffer solutions may be used.

■ Sample Preparation

Use the fraction of particles of air-dried soil or soil dried at temperatures
< 40 °C and passed through a square-hole sieve with 2-mm mesh size.
Alternatively, field-moist soil passed through a 2-mm sieve can be used.

■ Procedure

1. Take a representative test portion of at least 5 mL from the soil sample
using the spoon.

2. Place the test portion in the sample bottle and add five volumes of water,
potassium chloride solution, or calcium chloride solution.

3. Shake or mix the suspension for 60 ± 10 min using a mechanical shaker
(never longer than 3h). The stirring should be at a rate that achieves
a homogenous soil suspension. Entrainment of air should be avoided.

4. Calibrate the pH meter as prescribed in the manufacturer's manual using
the buffer solutions.

5. Adjust the pH meter as indicated in the manufacturer's manual. Measure
the temperature of the suspension and take care that the temperature of
the buffer and the soil solution does differ more than 1 °C. Measure the
pH in the suspension while or immediately after being stirred. Read the
pH after stabilization is reached. Record the pH values to two decimals.

2 Determination of Chemical and Physical Soil Properties 71

■ Notes and Points to Watch

• Drying may influence the pH of soils, especially those containing sulfides.
In such soils drying will lower the pH substantially.

• In calcareous soil samples the pH depends on calcium ion activity and
C0 2 partial pressure (pC0 2 ), and also on the quality of the laboratory
air (Schlichting et al. 1995).

• If a swinging-needle pH meter is used, the second decimal place should
be estimated (ISO DIS 10390 2002).

• In samples with a high content of organic material (e.g., peat soils,
pot soils) the suspension effect can play a role. In calcareous soils it is
possible for carbon dioxide to be adsorbed by the suspension. Under
these circumstances it is difficult to reach equilibrium pH values (ISO
DIS 10390 2002).

• Magnetic stirring of the suspension is not suitable since this can affect
the reading of pH.

• pH indicator strips may be used for rough estimations.

2.6

Soil Organic Matter - Soil Organic Carbon

■ Introduction

Objectives. Soil organic matter (SOM) is one of the most important indi-
cators of soil quality. It influences many soil properties including nutrient
supply (mainly N, P, S), cation exchange capacity, adsorption of pollu-
tants, infiltration and retention of water, soil structure, and soil color, most
of which in turn affect soil temperature. SOM consists of microbial cells,
plant and animal residues at various stages of decomposition, stable humus
(humic acids, humins) synthesized from residues by microorganisms, and
highly carbonized compounds (e.g., charcoal, graphite, coal; Nelson and
Sommers 1996). The term humus is used synonymously with SOM; that is,
it denotes all organic material in the soil. Organic material is essential as
a nutrient source for all heterotrophic soil organisms, which in turn hold
a key position in the processes of humification and mineralization of humic
substrates that lead to the production of stable humus, degradable organic
compounds, and carbon dioxide (Forster 1995a). There is often a direct
relationship between the organic carbon contents of soils and microbial
biomass and activity. Several methods are available for the determination
of SOM in soils. Most often SOM content of soils is determined by carbon

72 B.-M. Wilke

analysis. Two methods are described in this Section, namely dry combus-
tion and loss on ignition (LOI).

Theory. Carbon is the chief element (48-58%) in SOM. Therefore, organic
C determination is used as a basis of SOM estimates in soils. Based on
the assumption that SOM contains 58% organic C, a conversion factor of
1.724 has been proposed for the conversion of organic C content to SOM
(humus content) of soils (Nelson and Sommers 1996). C content of soil
can be determined by wet and dry combustion techniques. If inorganic C
is also extracted, corrections have to be made for the inorganic portion.
This can be done either by destruction of inorganic C prior to C analysis
or by separate measurement and subtraction of inorganic C from total
C content. Wet digestion procedures are based on oxidation of organic C
compounds by Q^Oy". Because of the high toxicity of Cr(VI) compounds,
this method should not be used. Dry combustion techniques are based on
heating the soil gradually up to > 900 °C and subsequent measurement of
evolved C0 2 trapped in a suitable reagent and determined titrimetrically
or gravimetrically. There are also other measuring devices in use (see below
and ISO 10694 1995). A simple technique for the estimation of SOM is the
LOI method that was standardized in Germany under DIN 19684-3 (1977).

2.6.1

Dry Combustion Method

Principle. The soil sample is gradually heated in a stream of purified oxy-
gen to > 900 °C. Organic and inorganic soil carbon is converted to C0 2 .
The C0 2 evolved is measured by titrimetry, gas chromatography, infrared
spectrometry, or gravimetry. In the presence of carbonates, the samples
are pretreated with HC1. If the carbonate content is known (determination
according to ISO 10693 1995), the organic carbon can be calculated. Soils
with pH(CaCl 2 ) < 6. 5 are unlikely to contain carbonates!

■ Equipment

• Analytical balance, accuracy 0.1 mg, or microbalance, accuracy 0.01 mg.

• Apparatus for determination of total organic carbon by dry combustion
at a temperature of > 900 °C equipped with an appropriate C0 2 de-
tector. The following detection devices are currently available: titrime-
try, gravimetry, gas chromatography, conductometry, and infrared spec-
troscopy. Some of the devices are able to measure separately inorganic
and organic carbon, others also measure total C and N contents (CN
analyzer)

2 Determination of Chemical and Physical Soil Properties 73

• Crucibles made of porcelain, quartz, silver, tin, or nickel of different size;
crucibles made of tin and nickel are not acid resistant.

■ Reagents

• Distilled or demineralized water with an electric conductivity of <
0.2mS/mat25°C

• Reagents for calibration, e.g., acetanilide (C 8 H 9 NO); atropine
(Q7H23NO3); calcium carbonate (CaC0 3 ); graphite powder for spec-
troscopy (C); sodium hydrogen phthalate (C 8 H 5 K0 4 )

• HC1 (4 mol/L)

■ Sample Preparation

Use air-dried, sieved (< 2 mm) soil.

■ Procedure

1. Weigh out mi g of the air-dried sample or subsample into a crucible. The
amount for analysis depends on carbon content and on the apparatus
used!

2. Carry out the analysis according to the manufacturer's manual.

3. Soils containing carbonates should be pretreated as follows: add an excess
of HC1 to the crucible containing a weighed quantity of air-dried soil and
mix. Wait 4 h and dry the crucible for 16 h at a temperature of 60-70 °C.
Then carry out the analysis in accordance to the manufacturer's manual.
The quantity of HC1 depends on the weight of the subsample and its
carbonate content. In all cases an excess of acid should be added!

■ Calculation

Organic Carbon Content

The total carbon content is calculated according to the following equation:

m 2 100 + w H? o

w ct = 1000 x — x 0.2727 x — (2.23)

mi 100

wet total carbon content on the basis of oven-dried soil (g/kg)

mi mass of the test portion (g)

m 2 mass of carbon dioxide released by the soil sample (g)

74 B.-M. Wilke

0.2727 conversion factor for C0 2 to C

Wh 2 o water content expressed as a percentage by mass on a dry mass
basis (Sect. 2.1)

Organic Matter Content

The organic matter content of the soil sample can be calculated using the
following equation:

Wom = / X Wcorg (2.24)

w om organic matter content of the soil on the basis of oven-dried soil
(g/kg)

wcorg organic carbon content of the soil on the basis of oven-dried soil
(g/kg)

/ conversion factor

2.6.2

Loss On Ignition Method (LOI)

Principle. The LOI method is based on ignition (550 ± 25 °C) of a dried
(105 °C) soil sample until mass constancy is achieved. The SOM content is
calculated from the mass difference before and after heating.

■ Equipment

• Sieves, 2- or 5-mm mesh size

• Drying oven, capable of maintaining a temperature of 105 ± 2 °C

• Muffle furnace, capable of maintaining a temperature of 550 ± 25 °C
installed under a fume hood

• Analytical balance, accuracy 0.01 g

• Porcelain crucibles or bowls

• Desiccator with an active drying agent

■ Sample Preparation

Use field-moist, sieved (< 5 mm) soil or air-dried, sieved (< 2 mm) soil.
Dry the soil to 105 °C prior to organic matter determination.

2 Determination of Chemical and Physical Soil Properties 75

■ Procedure

1. Determine the dry mass (m s ) of the soil according to Sect. 2.1.

2. Heat crucibles or bowls in the muffle furnace at 550 ± 25 °C for 20 min,
cool in a desiccator and determine tare mass (m t ) to 0.1 g.

3. Weigh 5-20 g (accuracy 0.01 g) of oven-dried (105 °C) soil (see step 1)
depending on its organic matter content in crucibles or bowls, and place
them in the cold muffle furnace.

4. Heat the muffle furnace gradually to 550 ± 25 °C for 2-4 h until mass
constancy is achieved.

5. Open the door and cool the muffle furnace down to 100 °C.

6. Place the crucibles/bowls in the desiccator and cool them to room tem-
perature (approx. 1 h).

7. Measure the mass of the filled crucibles/bowls (m c + m t ) twice. The
difference of each individual measurements from the mean should not
exceed 5% of the mean.

■ Calculation

1. Calculate the loss of mass (Am; g) after ignition at 550 °C using the
following equation:

Am = (m s + m t ) - (m c + m t ) = m s - m c (2.25)

2. The LOI corresponds to the SOM content and can be calculated using
the following equation:

Am
LOI (%) = x 100 (2.26)

m s

Am loss of mass of the soil after ignition at 550 °C (g)

m s mass of the soil dried at 105 °C (g)

m t mass of the crucibles/bowls ignited to 550 °C (g)

m c mass of the soil ignited to 550 °C (g)

■ Notes and Points to Watch

• Humus-rich samples should be weighed in the crucibles/bowls in a field-
moist state and dried and heated in the same crucible. In order to avoid
dusting the organic samples, the crucibles/bowls should be covered with
a porcelain lid or a metal mesh.

76 B.-M. Wilke

• The incineration of the samples should be controlled. The process is
complete if black particles cannot be found in the sample or if it has
a light gray to reddish color.

• Samples which do not show complete incineration should be treated
with a few drops of saturated ammonium nitrate solution or hydrogen
peroxide and heated again to 550 °C for 1 h.

• The LOI is assumed to be equal in most surface soils. Losses of crystalline
water of clay minerals and gypsum may result in an overestimation
of SOM contents. The same is true for carbonate-rich soils, because
decomposition of CaC0 3 , which starts at temperatures of approx. 500 °C.
Therefore, the method is mainly recommended for sandy and carbonate-
free soils and peats. Nevertheless, results for clayey soils and soils rich in
gypsum can be corrected by subtraction of 0.1% SOM per 1% of clayey
soil and 0.26% SOM per 1% of gypsum-rich soil.

• The error caused by the destruction of clay minerals may be avoided by
pre-heating at 430 °C in an N 2 atmosphere.

• For peat soils the LOI method is advantageous over the carbon determi-
nation procedures because the carbon content of these materials varies
between 40 and 100 mass%.

2.7

Soil Nutrients: Total Nitrogen

■ Introduction

Objectives. Analysis of total N, the C/N ratio, and inorganic N (ammo-
nium, nitrate) provides an insight into the nitrogen supply to soil mi-
croflora and plants. The total N content ranges from < 0. 02% (subsoils)
to > 2. 5% (peats). A-horizons of mineral soils contain 0.06-0.5% N. Ni-
trogen, phosphorous, and/or potassium deficiency may limit the microbial
decomposition (mainly cometabolic) of pollutants in soil. Optimum con-
ditions are achieved at C:N:P ratios of 100:10:2 (Kastner 2001). Therefore,
the concentrations of these nutrients have to be analyzed and adjusted if
necessary. Two methods have gained general acceptance for the determi-
nation of total N in soils, namely the Kjeldahl and the Dumas methods
(Bremner 1996). The Kjeldahl method is a wet oxidation procedure, the
Dumas method a dry oxidation (combustion) method. Both methods have
been standardized (ISO 11261 1995; ISO 13878 1998).

2 Determination of Chemical and Physical Soil Properties 11

2.7.1

Dry Combustion Method ("Elemental Analysis")

Principle and Theory. The soil is heated in a purified oxygen stream to a tem-
perature of > 900 °C. Mineral and organic N species are oxidized and/or
volatilized. Products are oxides of N (NO x ) and molecular N (N 2 ) mainly.
After transforming into N 2 by reduction on surfaces of metallic copper,
the N content is measured by means of thermal conductivity detection
(method adapted from ISO 13878 1998).

■ Equipment

• Balance, capable of weighing accurately to 0.1 mg, or microbalance, ca-
pable of weighing accurately to 0.01 mg

• Combustion apparatus to determine total N at a temperature > 900 °C,
including a detector for measuring the nitrogen gas formed

• Crucibles of various sizes, e.g. 10 or 20 mL, special requirements being
given in the manual of the apparatus used

■ Reagents

• Combustion gas (oxygen), special requirements being given in the in-
struction manual of the apparatus used

• Chemicals and/or catalysts for reduction, oxidation, and/or fixing of
combustion gases that interfere with the analysis

• Calibration substances, for example acetanilide (C 8 H 9 NO), amino acids
of known composition, or soil samples with certified N contents, the N
content of the calibration substance being as similar to the suspected soil
N content as possible

■ Sample Preparation

Soil samples dried in the air, dried in an oven at a temperature not exceeding
40 °C, or freeze dried (see Chapt. 1) are sieved (2 mm); if a soil mass < 2 g is
required for the analysis, mill a representative subsample to 0.1-0.15 mm.

■ Procedure

1. Calibrate the apparatus as described in the manufacturer's manual.

2. Weigh out m x g of the air-dried sample or subsample into a crucible. The
amount for analysis depends on N contents and on the apparatus used.

78 B.-M. Wilke

3. Carry out the analysis according to the manufacturer's manual.

4. Determine the percentage of water content (mass fraction) according to
the method described in Sect. 2.1.

■ Calculation

1. Normally, the primary results (from the apparatus) are given in mg N
(Xi) or a mass fraction of N (X 2 ), expressed as a percentage of the air-
dried soil used (mi ). Calculate total N content (w Nt ), in mg/g, on the basis
the oven-dried soil according to following equations:

For primary results given in mg of N:

X x (100 + w) _

w Nt = — x — — (2.27)

m\ 100

For primary results, given as percent mass fraction of N:

(100 + w)

w Nt = X 2 x 10 x (2.28)

100

w N t content of N (mg/g oven-dried soil)

Xi primary result in N (mg)

X 2 primary result in percentage N (mass fraction)

mi mass of air-dried soil for analysis (g)

w percentage of water content (mass fraction) on the basis of oven-
dried soil (Sect. 2.1)

2. If oven-dried samples are used, the N content is calculated as follows:
For primary results given in mg of N:

WNt - — (2.29)

For primary results given as percent of mass fraction of N:

w Nt = X 2 x 10 (2.30)

■ Notes and Points to Watch

• Today, several automated elemental analyzers for the determination of
total N are in use (see Bremner 1996). Most of them can be used for
determination of total N and total C, others also for hydrogen or sulfur.

2 Determination of Chemical and Physical Soil Properties 79

• One of the problems linked with the use of elemental analyzers is sample-
size limitation. This makes it essential to grind soil samples very finely
in order to get representative subsamples.

• Soil pores are filled with air, which contains up to 80% N 2 . Nitrogen gas
can also enter the combustion cell when it is opened for sample exchange.
Both facts may lead to overestimation of the soil N content. Therefore,
sufficient purging should be carried out by oxygen gas flow before the
combustion step.

2.7.2

Modified Kjeldahl Method

Principle. The method (as in ISO 11261 1995) is based on the Kjeldahl
digestion. Additional reagents are salicylic acid to avoid loss of nitrates,
and thiosulfate to detect azo-, nitroso- and, nitrocompounds. Instead of
selenium, titanium dioxide is used as catalyst.

Theory. The Kjeldahl method generally employed for determination of
total N involve two steps: (1) digestion of the sample to convert organic N
into NHj-N and (2) determination of NHj-N in the digest. The digestion is
usually performed by heating the sample with H 2 S0 4 containing substances
that promote the oxidation of organic matter and conversion of organic N
into NHJ-N. For increasing the temperature K 2 S0 4 or Na 2 S0 4 are used.
Catalysts such as Hg, Cu, Se, or Ti0 2 increase the rate of oxidation of
organic matter by H 2 S0 4 . A general equation for the digestion process is
given below:

Organic-N + H 2 S0 4 -^(NH 4 ) 2 S0 4 + H 2 + C0 2

+ other matrix by-products.

For the determination of NHJ -N several methods are possible, namely ion-
sensitive electrodes, colorimetric analyses, or analyses by steam distillation
and titration (Forster 1995b). In the distillation method employed by most
workers, NHJ -N in digests is converted under excess alkali into NH 3 , which
is collected in boric acid. By this procedure ammonium borate is formed,
which is titrated back to boric acid with hydrochloric acid according to the
following equations:

2NH 3 + H3BO3 -> (NH 4 ) 2 HB0 3
(NH 4 ) 2 HB0 3 + 2HC1 -> 2NH 4 C1 + H3BO3

80 B.-M. Wilke

■ Equipment

• Digestion flasks or tubes, of nominal volume 50 mL, suitable for the
digestion stand

• Digestion stand, glass tubes placed in holes drilled into an aluminum
block also being suitable

• Distillation apparatus, preferably of the Parnass- Wagner type

• Burette, graduated in intervals of 0.05 mL or smaller

■ Reagents

• Salicylic acid/sulfuric acid: dissolve 25 g of salicylic acid in 1 L of cone,
sulfuric acid (p = 1.84 g/cm 3 ).

• Potassium sulfate catalyst mixture: grind and thoroughly mix 200 g of
potassium sulfate, 6 g of copper(II) sulfate pentahydrate, and 6 g of tita-
nium dioxide with the crystal structure of anatase.

• Sodium thiosulfate pentahydrate: crush the crystals to form a powder
that passes through a sieve with 0.25-mm mesh size.

• NaOH solution ( 10 mol/L).

• Boric acid solution (H3BO3, 20 g/L).

• Mixed indicator: dissolve 0.1 g of bromocresol green and 0.02 g of methyl
red in 100 mL of ethanol.

• Sulfuric acid (0.01 mol/L).

■ Sample Preparation

Soil samples dried in the air, dried in an oven at temperature not exceeding
40 °C, or freeze-dried (see Chapt. 1) are sieved (2 mm) and ground to
0.1-0.15 mm.

■ Procedure

1. Place a test portion of the air-dried soil sample of about 0.2 g (expected
N content ca. 0.5%) to 1 g (expected N content ca. 0.1%) in the digestion
flask.

2 Determination of Chemical and Physical Soil Properties 81

2. Add 4 mL of salicylic/sulfuric acid and swirl the flask until the acid is
thoroughly mixed with the soil.

3. Allow the mixture to stand for at least several hours (or overnight).

4. Add 0.5 g of sodium thiosulfate through a dry funnel with a long stem
that reaches down into the bulb of the digestion flask, and heat the
mixture cautiously on the digestion stand until frothing has ceased.

5. Cool the flask.

6. Add 1.1 g of the catalyst mixture and heat until the digestion mixture
becomes clear.

7. Boil the mixture gently for up to 5 h so that the sulfuric acid condenses
about 1/3 of the way up to the neck of the flask. Ensure that the tem-
perature of the solution does not exceed 400 °C. In most cases a boiling
period of 2 h is sufficient.

8. After completion of the digestion step, allow the flask to cool, and add
about 20 mL of water slowly while shaking.

9. Swirl the flask to bring any insoluble material into suspension and
transfer the contents to the distillation apparatus.

10. Rinse three times with water to complete the transfer.

11. Add 5 mL of boric acid solution to a 100 mL conical flask and place the
flask under the condenser of the distillation apparatus in such a way
that the end of the condenser dips into the solution.

12. Add 20 mL of sodium hydroxide solution to the funnel of the apparatus
and run the alkali slowly into the distillation chamber.

13. Distill about 40 mL of condensate (the amount depends on the dimen-
sions of the conical flask).

14. Rinse the end of the condenser.

15. Add a few drops of indicator to the distillate.

16. Titrate with sulfuric acid to a violet endpoint.

■ Calculation

Calculate total N content (w Nt ) using the following equation and round the
result to two significant figures:

< xt/ -n (^i " ^o) x c(H + ) x M N 100 + w H2 o „ al ,

w Nt (mg N/g soil) = x — (2.31)

m 100

82 B.-M. Wilke

V\ volume of the sulfuric acid (mL)

Vq volume of the sulfuric acid used in the blank test (mL)

c(H + ) concentration of H + in the sulfuric acid (mol/L; e.g., if O.Olmol/L
sulfuric acid is used, c(H + ) is 0.02 mol/L

M N molar mass of nitrogen (g/mol; 14)

m mass of the air-dried soil subsample used (g)

Wh 2 o water content expressed as a percentage by mass on the basis of
oven-dried soil (Sect. 2.1)

■ Notes and Points to Watch

• The modified Kjeldahl procedure is satisfying for the analyses of most N
compounds in soils, but it detects compounds containing N-N and N-O
linkages and some heterocyclics (e.g., pyridine) only partially (Bremner
1996; ISO 11261 1995).

• Losses of nitrogen can occur with samples of high NH 4 -N and NO3-N
content. Therefore, excessive drying prior to analysis should be avoided.

• A potentiometric titration is also possible. The endpoint of the titration
is pH 5.0.

• If a steam distillation is used, a distillation rate up to 25 mL/min is
applicable. Stop the distillation when about 100 mL of distillate have
been collected.

2.8

Soil Nutrients: Inorand place it immediately on the shaker.

2 Determination of Chemical and Physical Soil Properties 91

4. Shake for exactly 30 min at 20 ± 1 °C.

5. Filter immediately (within 1 min) into a dry vessel using a P free-fluted
filter paper.

6. Prepare a blank by following the above procedure without soil.

7. Measure the P concentration of filtrates as given below.

■ Calculation

The content of phosphorous soluble in sodium hydrogen carbonate (w p ;
mg/kg of oven-dried soil) is calculated using the following equation:

100 + w w ,

Wp = Pp x 20 x (2.34)

Y 100

p P concentration of P (mg/L) as measured in the extract according to the
method described in Sect. 2.9.3

20 quotient of the volume of extracting solution (100 mL) and the mass
of air-dried soil (5 g)

w w percentage of water content (mass fraction) on the basis of oven-dried
soil (Sect. 2.1)

■ Notes and Points to Watch

• The extracting solution must be used within 4 h after preparation.

• A NaHC0 3 test level of 10 mg P/kg soil is considered to be in the "high"
category (see Kuo 1996).

2.9.3

Quantification of Phosphorus

■ Equipment

• Analytical balance, with a readability of ±0.001 g

• Spectrophotometer, capable of measuring the absorbance in wavelength
up to 900 nm and accepting cells of path 10 mm (readability 0.001 units
of absorbance)

• Optical cells, of path length 10 mm

• Volumetric flasks, 50 mL

92 B.-M. Wilke

■ Reagents

• Sulfuric acid, (H 2 S0 4 2.5 mol/L)

• Ammonium molybdate solution: dissolve 20 g of ammonium hepta-
molybdate tetrahydrate ([(NH 4 ) 6 Mo 7 2 4 4H 2 0]) in 500 mL of deionized
water. Store the solution in a glass-stoppered bottle.

• Antimony potassium tartrate solution (1 mg Sb/mL): dissolve 0.2728 g of
K(SbO) C 4 H 4 6 x 1/2H 2 in 100 mL of deionized water.

• Ascorbic acid solution (0.1 mol/L): dissolve 1.76 g of C6H 8 06 in 100 mL
of deionized water. Prepare the solution fresh daily.

• Mixed reagent: thoroughly mix 50 mL of H 2 S0 4 (2.5 mol/L), 15 mL of
ammonium molybdate solution, 30 mL of ascorbic acid solution, and
5 mL of antimony potassium tartrate solution. Prepare fresh daily.

• Phosphate stock solution (50mgP/L): dissolve 0.2197 g of oven-dried
(40 °C) KH 2 P0 4 in deionized water. Add 25 mL of H 2 S0 4 (2.5 mol/L) and
dilute to 1 L with deionized water.

• Working phosphate standard solution (5mgP/L): dilute 10 mL stock
solution to 100 mL with deionized water.

■ Sample Preparation

Soil samples are extracted as described in Sects. 2.9.1 and 2.9.2.

■ Procedure

1. Transfer into 50 mL volumetric flasks:

- A standard series, ranging from 0.5 mL up to 8mL of the working
phosphate standard solution, corresponding to P concentrations in
the measuring solution between 0.05 and 0.8 mg/L.

- An aliquot of lOmL of the soil extract containing total phosphorus
if the expected concentration of P in the soil is less than 400 mg/kg;
otherwise use 5 mL or less.

- An aliquot of 25 mL of the soil extract containing labile phosphorus.

2. Dilute the aliquots with deionized water to about 25 mL (if necessary),
and add 8 mL of mixed reagent.

3. Dilute the solution to volume and mix well.

4. Measure the absorbance at 880 nm after 10 min in a spectrophotometer.

5. Prepare a blank that contains all reagents except the P solution.

2 Determination of Chemical and Physical Soil Properties 93

■ Calculation

For the evaluation of the spectrophotometric measurements prepare a cal-
ibration graph by plotting absorbance units versus the P concentrations of
standard solutions (mg P/L). The correlation between both parameters is
linear in the relevant range up to 0.8 mg P/L.

The phosphorous concentration of the measuring solution (p P ; mg/L)
can be calculated using the following equation:

(A ES - A B ) x 50
Pp = 7 77 ( 2 - 35 )

/x y s

A E s absorbance of the soil extract

A B absorbance of the blank

/ slope of the regression line (absorbance per mg P/L)

V s volume of the aliquot (mL)

50 volume of the volumetric flasks (mL)

References

Blake JR, Hartge KH (1986) Bulk density. In: Klute A (ed) Methods in soil analysis, part 1.

Physical and mineralogical methods, 2nd ed, Am Soc Agron, Madison, WI, pp 363-376
Bowman RA (1988) A rapid method to determine phosphorus in soils. Soil Sci Soc Am J

52:1301-1304
Bremner JM (1996) Nitrogen - total. In: Bigham JM (ed) Methods of soil analysis, part 3,

chemical methods. Soil Sci Soc Am Am Soc Agron, SSSA Book, Series no 5, Madison,

WI,pp 1085-1121
Danielson RE, Sutherland PL (1986) Porosity. In: Klute A (ed) Methods in soil analysis,

part 1, physical and mineralogical methods, 2nd edn. Soil Sci Soc Am Am Soc Agron,

Madison WI,pp 443-461
DIN 19684-3 (1977) Bodenuntersuchungsverfahren fur den Landwirschaftlichen Wasser-

bau - Chemische Laboruntersuchungen - Teil 3: Bestimmung des Gliihverlustes und

des Gliihriickstandes
Forster JC (1995a) Soil sampling, handling, storage and analysis - organic carbon - In:

Alef K, Nannipieri P (eds) Methods in applied soil microbiology and biochemistry.

Academic Press, pp 59-65
Forster JC (1995b) Soil sampling, handling, storage and analysis - soil nitrogen. In: Alef K,

Nannipieri P (eds) Methods in applied soil microbiology and biochemistry. Academic

Press, pp 79-87
Forster JC (1995c) Soil sampling, handling, storage and analysis - soil phosphorus. In:

Alef K, Nannipieri P (eds) Methods in applied soil microbiology and biochemistry.

Academic Press, pp 88-93
Hartge KH (1968) Heterogenitat des Bodens oder Quellung? Trans Int Congr Soil Sci 3:591-

597

94 B.-M. Wilke

ISO 10693 (1995) Soil Quality - Determination of carbonate content - volumetric method

ISO 10694 (1995) Soil Quality - Determination of organic and total carbon after dry com-
bustion (elementary analysis)

ISO 11261 (1995) Soil Quality- Determination of total nitrogen- Modified Kjeldahl method

ISO 11263 (1994) Soil Quality - Determination of phosphorous - spectrometric determina-
tion of phosphorous soluble in sodium hydrogen carbonate solution

ISO 1 1266 ( 1 994) Soil Quality - Guidance on laboratory testing for biodegradation of organic
chemicals in soil under aerobic conditions

ISO 11272 (1998) Soil Quality - Determination of dry bulk density

ISO 11274 (1998) Soil Quality - Determination of the water- retention characteristic - Lab-
oratory methods

ISO 11465 (1993) Soil Quality - Determination of dry matter and water content on a mass
basis - Gravimetric method

ISO 13878 (1998) Soil Quality - Determination of total nitrogen content by dry combustion
("elemental analysis")

ISO 14238 (1997) Soil quality - Biological methods - Determination of nitrogen mineral-
ization and nitrification in soils and the influence of chemicals on these processes

ISO 14255 (1998) Soil Quality - Determination of nitrate nitrogen, ammonium nitrogen
and total soluble nitrogen in air-dry soils using calcium chloride solution as extractant

ISO 16072 (2002) Soil quality - Laboratory methods for determination of microbial soil
respiration

ISO 3696 (1987) Water for analytical laboratory use - Specifications and test methods

ISO DIS 10390 (2002) Soil Quality - Determination of pH

ISO/TS 14256-1 (2003) Soil Quality - Determination of nitrate, nitrite and ammonium in
field moist soils by extraction with potassium chloride solution - Part 1 : Manual method

Kastner M (2001) Parameter und Methoden zur Beurteilung der biologischen Sanierbarkeit
von Boden. In: Michels J, Track Th, Gehrke U, Sell D (eds) Leitfaden, Biologische
Verfahren zur Bodensanierung. DECHEMA Gesellschaft fur Chemische Technik und
Biotechnologie eV, Bericht im Auftrag des BMBF Projekttrager Abfallwirtschaft und
Altlastensanierung im Umweltbundesamt Berlin. Forderkennzeichen 1491064, pp 191 —
234

Kuo S (1996) Phosphorous. In: Bigham JM (ed) Methods of soil analysis, part 3, chemical
methods. Soil Sci Soc Am Am Soc Agron, SSSA Book, Series no 5, Madison WI, pp 869-
919

In: Bigham JM (ed) Methods of soil analysis, part 3, chemical methods. Soil Sci Soc Am Am
Soc Agron, SSSA Book, Series no 5, Madison WI, pp 1123-1184

Li C, Fan X, Mengel K (1990) Turnover of interlayer ammonium in loess-derived soil grown
with winter wheat in the Shaanxi Province of China. Biol Fert Soils 9:21 1-214

Mulvaney RL (1996) Nitrogen - inorganic forms. In: Bigham JM (ed) Methods of soil
analysis, part 3, chemical methods. Soil Sci Soc Am Am Soc Agron, SSSA Book, Series
no 5, Madison WI, pp 1123-1184

Murphy J, Riley HP (1962) A modified single solution method for the determination of
phosphate in natural waters. Anal Chim Acta 27:31-36

Nelson, DW, Sommer LE (1996) Total carbon, organic carbon, and organic matter. In:
Bigham JM (ed) Methods of soil analysis, part 3, chemical methods. Soil Sci Soc Am
Am Soc Agron, SSSA Book, Series no 5, Madison WI, pp 961-1010

Page AL, Miller RH, Keeney DR (eds, 1982) Methods in soil analysis, part 2, chemical and
microbiological properties. Soil Sci Soc Am Am Soc Agron, SSSA Book, Madison WI

Sahrawat KL ( 1 995) Fixed ammonium and carbon-nitrogen ratios of some semi-arid tropical
Indian soils. Geoderma 68:219-224

2 Determination of Chemical and Physical Soil Properties 95

Scheffer F (2002) Lehrbuch der Bodenkunde, Scheffer/Schachtschabel. 15. Auflage, neubear-
beitet und erweitert von Blume HP et al. Akademischer Verlag GmbH, Heidelberg

Schlichting E, Blume H-P, Stahr K (1995) Bodenkundliches Praktikum, 2nd ed, Blackwell,
Berlin Wien

Topp GC, Dow B, Edwards M, Gregorich EG, Curnoe WE, Cook EJ (2000) Oxygen measure-
ments in the root zone facilitated by TRD. Can J Soil Sci 80:33-41

VDLUFA (2000) Methodenbuch 1 3. Teilliefg. 2000, Nmin-Methode Labor A6. 1.4.1, VDLUFA-
Verlag Darmstadt

Quantification of Soil Contamination

Kirsten S. j0rgensen, Olli Jarvinen, Pirjo Sainio,
Jani Salminen, Anna-Mari Suortti

3.1

General Introduction

Even though better toxicity tests have become available and their use is
increasing in risk assessment of contaminated sites and of the reuse of
bioremediated soil, chemical data is still today the main information used
in decision-making. The awareness of soil contamination increased in the
1980s and large-scale bioremediation became frequent during the 1990s.
However, the development and standardization of reproducible chemical
methods for the determination of specific organic contaminants in soils
have been very slow (Karstensen et al. 1998). The methods for the de-
termination of heavy metals have been in use longer, metal examination
of soil samples having been performed for geological purposes, e.g., in
the search for ore by the mining industry. The older methods for deter-
mination of oil and polyaromatic hydrocarbons (PAHs) were often based
on extraction of dried and sieved samples (resembling the procedures for
heavy metal pretreatment) followed by an extraction with non-polar sol-
vents. The use of halogenated solvents such as CC1 4 and Freon (ISO/TR
11046 1994) were common, but due to occupational health and environ-
mental aspects these are being phased out. Methods that are based on the
extraction of field-moist samples with a mixture of polar (e.g., acetone,
methanol) and non-polar (hexane, pentane, heptane) solvents have been
proven to give sufficient yield and to be reproducible. Methods that are fea-
sible and reproducible in the laboratory are currently being standardized
by the International Standardization Organization (ISO).

The objectives of determining contaminant concentrations in soil may
be to assess the appearance of particular contaminants at a site or to
monitor the progress of a bioremediation action either in the field or in
laboratory feasibility studies. The monitoring of bioremediation involves
repeated measurements of contaminant concentrations over time. Based on
a time series with, e.g., five points, a biodegradation rate can be obtained.

Kirsten j0rgensen, Olli Jarvinen, Pirjo Sainio, Jani Salminen: Finnish Environment Institute,
P.O. Box 140, 00251 Helsinki, Finland, E-mail: Kirsten.Jorgensen@ymparisto.fi

Anna-Mari Suortti: SGS Inspection Services, Syvasatamantie 24, 49460 Hamina, Finland

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

98 K.S. j0rgensen et al.

The biodegradation rate can be linear or represent first order decay depen-
dent on, e.g., the contaminant concentration and bioavailability. Field-scale
bioremediation can be classified as either in situ methods, where the treat-
ment takes place without excavating the soil, and ex situ methods, where ex-
cavated soil is treated typically in piles. When monitoring a site undergoing
in situ treatment by drilling for subsurface samples, it is essential to remem-
ber that true replicate samples cannot be obtained and a large variation is to
be expected. When sampling stock piles or biopiles, a combination sample
consisting of subsamples from different places in the piles typically will be
assembled, and parallel combination samples can be made ( j0rgensen et al.
2000). When monitoring biodegradation by laboratory microcosms, it is of
great importance that all the sample material representing a certain depth,
treatment, etc., is homogenous. This is best ensured by homogenizing and
sieving a larger batch from the field and by distributing this into separate
parallel bottles or other containers for laboratory incubations. A mesh size
of 8 mm has proven to be a good size for sieving field-moist soil (Laine
and j0rgensen 1997; Salminen et al. 2004). However, the measurement of
contaminant disappearance only shows that the parent compound has been
transformed; it does not reveal whether the degradation is complete to C0 2
or CH 4 or if other degradation products are produced.

Contamination with petroleum hydrocarbon products is one of the most
frequent types of soil contamination. Refineries, surface and underground
storage tanks, petrol service stations, etc., are the most common sites for
such contamination. Most petroleum products also contain minor amounts
of PAHs. No single method is reliable for the determination of all petroleum
hydrocarbons, and we therefore describe three methods for the determi-
nation of different fractions of hydrocarbons in soil samples.

Volatile hydrocarbons (Sect. 3.2) should be determined at sites where
gasoline and jet fuel are the sources of contamination. The pertinent
method here quantitatively determines these separate compounds: ben-
zene, toluene, ethylbenzene and xylenes (BTEX compounds), naphthalene,
and gasoline additives such as MTBE (methyl tert-butyl ether) and TAME
(tert-amyl methyl ether). This method can also be used to determine halo-
genated volatiles, which may often be found together with fuel products
because such solvents often are used, e.g., for cleaning engines.

Contamination with oil products such as heating oil, diesel or lubricating
oil is best determined using the method (Sect. 3.3) for hydrocarbons in
the range C 10 to C 40 . The result is a sum parameter, which does not give
concentrations of specific compounds. But still the sum of the hundreds of
compounds in this range is very useful for quantifying contamination with
them and for monitoring bioremediation. Based on the chromatogram,
a qualitative estimation of the type of contamination can be obtained.
This Qo-40 parameter is often referred to as mineral oil or total petroleum

3 Quantification of Soil Contamination 99

hydrocarbons (TPH), but these terms are somewhat unspecific. Crude oil is
often determined with this method, but it also includes volatiles and PAHs
that should be determined separately with the methods for volatiles and
PAHs, respectively.

Contamination with PAHs is commonly found at gas works and at sites
where coal tar and oil shale are handled. Oil containing heavy fractions
or waste oil may also contain significant amounts of PAHs. The method
described here (Sect. 3.4) allows for a single determination of 16 different
PAH compounds. In the literature the sum of PAHs is often reported, but
the fact that different countries and different laboratories analyze different
number of compounds has made this term very unspecific. Guideline values
for clean-up needs also differ between countries, so it is important to
check which compounds require reports. Since the toxicities of the PAH
compounds differ, there may not be any guideline value set out for all
compounds.

Contamination with heavy metals is difficult to assess because clean
soil itself may contain many heavy metals, depending on the geological
structure. Furthermore, many metals are not necessarily bioavailable in
soil, and for that reason different types of less exhaustive extractions are
being developed to determine the bioavailable fractions. The background
contents of metals in soil are in many countries known and they are taken
into account when guideline values for clean-up are determined. Still today
most guideline values are based on the total or near- total content of metals.
The method described here (Sect. 3.5) reveals the near-total content and is
aiming at determining the anthropogenic contamination.

3.2

Volatile Hydrocarbons

■ Introduction

Objectives. The volatile organic compounds ( VOCs) in soils primarily orig-
inate from petroleum products and solvents. The spectra of the VOCs de-
pend on their source. The analysis of benzene, toluene, and ethylbenzene
and xylenes (BTEX) is widely used as an indicator of contamination with
light petroleum products, e.g., petrol and kerosene. Furthermore, the gaso-
line additives MTBE and TAME as well as halogenated volatile compounds
can be analyzed with this method.

Principle. A soil sample is extracted with methanol. A defined volume of the
methanol extract is transferred into water and the water sample is heated
to 80 °C in a headspace vial. When equilibrium is established between the
gaseous and liquid phases, an aliquot of the gaseous phase is injected on

100 K.S. j0rgensen et al.

a column of a gas chromatograph and the VOCs are determined with a mass
selective detector.

Theory. VOCs are a group of compounds that have a boiling point from 20
to 220 °C and usually they have two to ten C atoms. They are mainly un-
substituted or substituted monoaromatics and short-chain aliphatic com-
pounds that differ in solubility and in toxicity. The individual compounds
are quantitatively determined using this method, as can also be the diaro-
matic compound naphthalene. We do not recommend measuring the sum
of VOCs because such a sum is unspecific and depends on the compounds
included.

The sampling (ISO 10381-1 1994; ISO 10381-2 1994; Owen and Whittle
2003) is a crucial step in the analysis of VOCs. In order to prevent their loss
during preparative steps, field-moist samples are used (ISO 14507 2003).
The sample is added into a pre-weighed glass container containing a known
amount of methanol. To control the quality of the determination, field du-
plicates, a procedural blank, and a control sample are analyzed. The two
main methods of analysis of VOCs are static headspace/gas chromatogra-
phy (e.g., ISO/PRF 22155 in prep.) and purge and trap/gas chromatography
(e.g., ISO 15009 2002). In the analysis of volatile aliphatic and aromatic
hydrocarbons a mass selective detector (MSD) is used. VOCs can also be
detected with a photo ionization detector (PID), a flame ionization detec-
tor (FID), and an electron capture detector (ECD; Owen and Whittle 2003).
The identification of target compounds (ISO/DIS 22892 in prep.) is easy
with a MSD, and a possible matrix effect can be eliminated. The method
described here is that using static headspace/gas chromatography (MSD)
and is based on the proof of a new international standard ISO/PRF 22155
and has earlier been described by Salminen et al. (2004).

■ Equipment

• Usual laboratory glassware, free of interfering compounds

• Shaking machine

• Headspace analyzer and gas chromatograph with a mass selective detec-
tor (MSD)

- Oven temperature program: maintain 35 °C for 2min, then steadily
raise by 14°C/min up to 90 °C. Maintain 90 °C for 5min, then raise
by 12°C/min up to 190 °C. Maintain 190 °C for 1 min, then raise by
40 °C/min up to 225 °C, and maintain at 225 °C for 1 min.

- Carrier gas: helium.

- Gas flow: lOmL/min.

- Split ratio (gas flow rate through split exit: column flow rate): 5.7:1.

3 Quantification of Soil Contamination 101

• Column: stationary phase non-polar or low polar fused silica capillary
column; film thickness 1.4 p.m; column length 30 m; internal diameter
0.25 mm

■ Reagents

• Methanol

• Internal standards, e.g., toluene-d 8 , a,a,a-trifluorotoluene

• Helium

• Synthetic air

• Volatile aromatic and halogenated hydrocarbons for standard solutions:
MTBE, TAME, benzene, ethylbenzene, toluene, m-xylene, p-xylene,
o-xylene, styrene, naphthalene, dichloromethane, chloroform, carbon
tetrachloride, 1 ,2-dichloroethane, 1,1,1 -trichloroethane, cis- 1 ,2-di-
chloroethene, trichloroethene, tetrachloroethene, chlorobenzene, 1,2-
dichlorobenzene, 1 ,4-dichlorobenzene, 1 ,2,3 -trichlorobenzene, 1 ,2,4-
trichlorobenzene, 1 ,3,5-trichlorobenzene

• Standard stock solutions

- Standard solutions: for each analyte, lOmg/mL of methanol

- Internal standard (see above) solution, lOmg/mL of methanol

• Working standard solutions

- Standard solutions: 1 mg mixed analyte solution/mL of methanol

- Internal standard (see above) solution, 10p.g/mL of methanol

• Calibration solutions: at least five different concentrations by suitable

dilutions of the working standard solutions within the range of 0.05-
lOpg/L

■ Sample Preparation

In the field, approximately 20 g of field-moist soil sample is taken directly
into a pre-weighed headspace vial containing 20 mL of methanol. No sieving
of the samples is recommended. A separate sample is taken for dry mass
determination in a glass jar leaving no headspace.

■ Procedure

1. Weigh the vial containing the soil sample and methanol.

2. Shake the vial containing sample and methanol for 30 min with the
shaking machine.

3. Allow the vial to stand for 10-15 min to settle the solid material.

102 K.S. j0rgensen et al.

4. Pipette 10 mL of water, 100 pi of methanol extract, and 5pL of the
working internal standard solution into a headspace vial.

5. Place the vial in the headspace system and heat the sample at 80 °C for
lh.

6. Use headspace injection for gas chromatographic analysis.

7. Detect the compounds with the mass selective detector (MSD).

8. Identify the peaks of the internal standards by using the absolute re-
tention times.

9. Determine the relative retention times for all the other relevant peaks
in the gas chromatogram. These retention times should be determined
in relation to those of the internal standards.

10. Determine the dry mass content, e.g., by using the method described
in ISO 11465(Chapt.2)

11. Calculate the concentrations of the analytes.

To prepare a calibration curve, treat the calibration standards as the soil
samples:

1. Add 100 pL of calibration solution to a headspace vial containing 10 mL
of water.

2. Add a known amount of working internal standard solution into the vial.

3. Close the vial and treat it according to the procedure.

■ Quality Control

1. Procedural blank determination: add 100 pL of methanol and 5 pi of the
working internal standard solution to 10 mL of water. Treat this mixture
as the soil sample.

2. Control sample determination: add a known amount of working stan-
dard solution to a pristine soil sample that contains neither VOCs nor
methanol. Treat the control sample as the soil sample and calculate the
recovery (%) of the analytes. Mark the recovery on the quality- control
chart.

■ Calculation

Concentration of analytes is quantified with respect to the internal standard
using the following formula:

_ C iw X Vte X V,
Cm,i — TA

m dm x V,

(3.1)

3 Quantification of Soil Contamination 103

c m> i content of the analyte cc i" in the sample (mg/kg soil dry mass)

C[ w mass concentration of the analyte "i" in the spiked water sample
obtained from the calibration curve (pg/L)

Vte total volume of the extract (methanol added to the soil sample + water
in the sample obtained from the determination of dry mass content;

mL)

V w volume of the spiked water sample for headspace measurement (mL)

^dm dry mass of the test sample used for extraction (g)

V a volume of the aliquot of methanol extract used for the spiking of
water sample for headspace measurement (pi)

■ Notes and Points to Watch

• Assure that compounds do not evaporate during sample handling.

• Exposure of samples to air, even during sampling, shall be avoided as far
as possible.

• The use of plastics, other than PTFE, shall be avoided.

• Samples shall be analyzed as soon as possible.

• Store the samples in the dark at 4 ± 2 °C no longer than 4 days.

• The standard and calibration solutions can be stored for 1 year at -18 °C.

• The internal standard solutions can be stored for several years at -18 °C.

• Avoid direct skin contact and inhalation of vapors from standards and
samples.

3.3

Hydrocarbons in the Range Ci to C 40

■ Introduction

Objectives. Petroleum derivatives such as diesel fuel, heating oil, and lu-
brication oil are widely used in human activities and thus are common
pollutants in the soil environment. These petroleum products are complex
mixtures of hundreds of various hydrocarbons. The analytical method
described here (modified ISO 16703 2004 Salminen et al. 2004) allows
a quantitative and a composition pattern determination of all hydrocar-

104 K.S. j0rgensen et al.

bons (that is, n-alkanes from CnH 2 2 to C 39 H 80 , isoalkanes, cycloalkanes,
alkyl benzenes, and alkyl naphthalenes) with a boiling range of 196 to
518 °C. Gasolines cannot be quantified using this method. Furthermore,
high concentrations of polyaromatic hydrocarbons (PAHs) may interfere
with the analysis.

Principle. A soil sample is extracted by sonication with n-heptane-acetone
including the internal standards (n-decane and n-tetracontane). To sepa-
rate the organic phase, water is subsequently added. The extract is washed
with water and the polar constituents and water are removed from the ex-
tract with Florisil (U.S. Silica Co., Berkeley Springs WV, USA) and sodium
sulfate, respectively. Hydrocarbons in the range from C 10 to C 40 are deter-
mined from an aliquot of the purified extract with a gas chromatograph
equipped with a flame ionization detector (FID). For the quantification of
all the hydrocarbons in this range, the total peak area between the internal
standards n-decane and n-tetracontane is measured.

Theory. Petroleum derivatives are complex mixtures of various hydrocar-
bons with different characteristics (e.g., volatility, water solubility, biode-
gradability). In the assessment of petroleum hydrocarbon contamination
and the effects of microbial activity (past, present, or future) on the fate
of these contaminants in soil, it is essential to know the quantity and the
composition of the contaminating agents. This information is of high value
when, for instance, a bioremediation process is followed over a span of
time. Moreover, as hydrocarbons differ in their amenability to microbial
degradation, this information is of a remarkable value.

In the past, gravimetric or infrared spectrometric methods have been
extensively used for the determination of hydrocarbons in soil. While these
methods can be used for quantification of a range of hydrocarbons, they
do not provide any information of the their quality, that is, of their com-
pound composition pattern. To obtain this information, more sophisticated
methods such as gas chromatographic analyses, are employed.

The extraction of hydrocarbons shall be performed in such a manner
that the broad spectrum of the compounds of interest is included in the
analysis. Moreover, it is essential that the extraction procedure is suitable
for field-moist soil samples in which hydrocarbons may be attached to
soil particles, and in which soil water present in the samples may impede
the extraction of the non-polar hydrocarbons. Thus, a mixture of polar
(acetone) and non-polar (n-heptane) solvents is used. On the other hand,
polar compounds have to be removed from the extract as they interfere
with the gas chromatographic analysis, and to avoid the inclusion of po-
lar compounds other than petroleum hydrocarbons in the analysis. It is
to be noted that PAHs and volatile compounds have to be analyzed sepa-
rately.

3 Quantification of Soil Contamination 105

■ Equipment

• Usual laboratory glassware free of interfering compounds

• Sonicator

• Laboratory centrifuge

• Gas chromatograph (GC) with a non-discriminating injection system
and a flame ionization detector (FID), helium as a carrier gas

• Pre-column (in case on-column injection is used)

• Capillary column specifications: 5% phenyl polysilphenylene-siloxane
stationary phase, e.g., SGE BPX5 capillary column, 5 m length and 1.4-
l^m film thickness

■ Reagents

• n-Heptane

• Acetone

• Ion- exchanged water

• n-Decane (Ci H 22 ), n-eicosane (C 2 oH 42 ), n-triacontane (C 30 H 62 ), n-pen-
tatriacontane (C 35 H 72 ), and n-tetracontane (C 40 H 82 ) - n-decane and n-
tetracontane being used as the integration window and the latter also as
an internal standard

• Florisil (150-250 p.m, 60-100 mesh) (Activated Florisil is stored in a des-
iccator and is usable for a week after the activation. Note: the activity of
Florisil will gradually decrease after the activation.)

• Anhydrous sodium sulfate (Na 2 S0 4 ) must be kept at 550 °C for at least
2 h prior to its use

• Diesel fuel and lubrication oil standards free of additives

• Helium

• Hydrogen

• Synthetic air

• Control soil sample

• Standard stock solutions

- Standard extraction solution (0.15 mg/mL of C 10 H 22 and 0.20 mg/mL
of C 40 H 82 ): weigh 20 p.L of n-decane and 20 mg of n-tetracontane and
dissolve in 100 mL of n-heptane. Prepare the solution in a volumetric

106 K.S. j0rgensen et al.

flask by weighing and calculate the accurate concentrations of the
internal standards n-decane and n-tetracontane in the solution. Store
the solution at 4 °C in the dark. The solution is usable for at least 6
months if stored in a tightly closed (Teflon-capped) glass vial.

- Working standard extraction solution: dilute the standard extraction
solution 1:9 (v/v) in n-heptane. Prepare the solution in a volumetric
flask by weighing and calculate the accurate concentrations of the
internal standards n-decane and n-tetracontane in the solution. The
solution is usable for 1 week if stored in a tightly closed (Teflon-
capped) glass vial.

- Calibration stock solution (20 mg hydrocarbons/mL): Weigh 100 mg
of diesel fuel and 100 mg of lubrication oil and dissolve in lOmL of
n-heptane. Prepare the solution in a volumetric flask by weighing and
store the solution at 4 °C in the dark. The solution is usable for at least
6 months if stored in a tightly closed (Teflon-capped) glass vial.

- Working calibration solutions: prepare at least five solutions with
final hydrocarbon concentration ranging from 0.1 to 2-3 mg/mL.
Prepare the solution by diluting the calibration stock solution with
n-heptane to obtain a final volume of 10 mL. Weigh the amounts of
solutions used to calculate the exact hydrocarbon concentrations in
the working calibration solutions. The solution is usable for at least
6 months if stored in a tightly closed (Teflon-capped) glass vial.

- Stock solution for testing the performance of the gas chromatograph:
weigh 5.0 mg each of n-decane (C10H22), n-eicosane (C 2 oH 42 ), n-tria-
contane (C 30 H 62 ), n-pentatriacontane (C35H72), and n-tetracontane
(C 40 H 82 ) and dissolve them in 10 mL of heptane. Prepare the solution
in a volumetric flask by weighing the mass of the added heptane to
calculate the exact concentration of the individual n-alkanes in the
solution. Store the solution at 4 °C in the dark. The solution is usable
for at least 6 months if stored in a tightly closed (Teflon capped) glass
vial.

- Working solution for testing the performance of the gas chromato-
graph: dilute the test stock solution in n-heptane in a ratio of 1:9 (v/v).
Prepare the solution in a volumetric flask by weighing to calculate the
exact concentration of the individual n-alkanes in the solution.

■ Sample Preparation

Sampling should be performed according to good practices (ISO 10381-1
1994; ISO 10381-2 1994). For the analysis, a homogenized field-moist soil

3 Quantification of Soil Contamination 107

sample is used (ISO 14507 2003). However, if the water content of the sample
is extraordinarily high, separation of the organic phase may occur prior to
the extraction (that is, at the time of the introduction of the sample into the
extraction solution). In such case, the sample has to be pre-dried overnight
at room temperature prior to the extraction.

■ Procedure

Prior to Analysis

1. Calibrate the gas chromatograph by running aliquots of the working
standard solutions.

2. An aliquot of the working test solution should be run on the GC and the
yields of the individual n-alkanes calculated. The ratio between C 2 oH 42
and C 40 H 82 should not exceed 1.2.

Analytical Procedure

1. Weigh 10 g of a sample into an extraction vial.

2. Weigh 5-10 g of a control sample with a known concentration into
a separate vial.

3. Add 10 mL of working standard solution and 20 mL of acetone into each
of these vials.

4. Prepare a blank determination: add 1 mL of working standard solution
and 20 mL of acetone but omit the sample. The blank and the control
sample are treated in a similar manner to the (unknown) samples.

5. Mix the samples gently and sonicate for 30 min. Add ice into the soni-
cator to keep the samples cool.

6. Add 30 mL of water and shake for 1 min.

7. Centrifuge the samples (2,500 rpm, 5 min).

8. Transfer the organic phase into a 25-mL test tube with a Teflon-lined
screw cap, add 10 mL of water, and shake for 1 min.

9. Transfer the organic phase into another test tube with a Teflon-lined
screw cap and add approx. 0.5 g of Na 2 S0 4 and shake.

10. Add approx. 1.5 g of Florisil into the tube and shake for 10 min in
a mechanical shaker.

11. Centrifuge the tubes (2,000 rpm, 1 min).

12. Transfer an aliquot of the purified extract into a GC vial. Avoid the
introduction of Florisil into the GC vial.

108 K.S. j0rgensen et al.

13. Run all the samples by GC.

14. Solvent blank should be subtracted from the sample chromatogram.
Integrate the total area between the peaks of Ci H 2 2 and C 40 H 82 to obtain
the hydrocarbon concentration of the extracts from the calibration
extracts.

15. Integrate the total area of the C 40 H 82 peak to obtain the recovery of
C 40 H 82 in the analysis.

■ Quality Assurance

1. The hydrocarbon concentration in the blank extract should be below
0.025 mg/mL.

2. The recovery of the internal standard n-tetracontane should be calculated
in each extract. The yield should be 100 ± 20% of the theoretical value
of C 40 H 82 in the extraction solution.

3. The hydrocarbon content of the control soil sample should be monitored
over time and the results ought to be analyzed according to general good
quality procedures.

■ Calculation

The concentration of hydrocarbons in the range from C 10 H 2 2 to C 40 H 82
(c H c) i n the sample is calculated as follows:

c 2C x 10 x 1000 x /

Chc = -£ - A (3.2)

m x d s

chc concentration of hydrocarbons in the range from Ci H 22 to C 40 H 82
in the sample (mg/kg dry mass)

c gc hydrocarbon concentration of the extract calculated from the cali-
bration equation (mg/mL)

10 volume of the organic solvent used in the extraction (10 mL of hep-
tane)

1,000 conversion factor of the soil mass (1 kg = 1,000 g)

/ dilution factor (if applicable)

m wet mass of the sample (g)

d s content of dry substance in the field-moist sample (g/g), determined
according to ISO 11465 (1993)

3 Quantification of Soil Contamination 109

■ Notes and Points to Watch

• The samples should be analyzed as soon as possible. If this is not feasible,
the samples should be stored at -20 °C.

• Hydrocarbons are subjected to biodegradation both under aerobic and
anaerobic conditions. Therefore, storage of the samples at temperatures
above °C should be avoided (Salminen et al. 2004).

• The efficacy of each Florisil stock has to be tested prior to its use in the
analysis.

• Weighing of the liquid, viscous standard compounds gives very precise
solutions.

• Avoid skin contact and inhalation of vapors from standards and samples.

3.4

Polyaromatic Hydrocarbons (PAHs)

■ Introduction

Objectives. Polycyclic aromatic hydrocarbons (PAHs) are often found at
contaminated sites, particularly in connection with tar contamination at
former gasworks. They also exist as diffuse contamination in urban areas
and alongside roads. Furthermore, wood impregnation with creosote and
incomplete combustion of hydrocarbons are major sources of PAHs in soil.
Polyaromatic hydrocarbons are a group of more than 100 different com-
pounds. This method describes the determination of a small selection of
the many PAHs found in the environment. The US Environmental Protec-
tion Agency (EPA) has chosen 16 of these PAHs to be the most important
ones to be analyzed (EPA priority list 1982).

Principle. A field-moist sample is extracted twice with acetone, and then
hexane is added to the acetone extract. The extract is washed twice with
water and the organic layer is dried with anhydrous sodium sulfate. When
necessary, the extract is cleaned up by adsorption chromatography on
a silica gel. The (purified) extract is analyzed by capillary gas chromatog-
raphy with mass selective detection, using appropriate deuterated PAHs as
internal standards.

Theory. Polycyclic aromatic hydrocarbons occur ubiquitously in the en-
vironment. Sixteen PAHs (Table 3.1) were chosen by the US EPA to be
analyzed in environmental samples because they are the most abundant at
hazardous waste sites and more information is available on these than on
other PAHs. Moreover, the chosen compounds exhibit harmful effects that

110

K.S. j0rgensen et al.

Table 3.1. Native and deuterated PAHs with their specific ions (target ion with qualifier ion
in parentheses)

Native PAH

Mass number

Deuterated PAH

Mass number

(amu)

Naphthalene

128(129)

D 8 -Naphthalene

136

Acenaphthene

Acenaphthylene

Fluorene

154(153)
152(151)
166(165)

Dio -Acenaphthene

164

Anthracene
Phenanthrene
Fluoranthene
Pyrene

178 (89)
178(179)
202(101)
202(101)

D io -phenanthrene

188

Benz(a)anthracene
Chrysene

228(114)
228(114)

Di2-chrysene

240

Benzo(b)fluoranthene
Benzo(k)fluoranthene
Benzo(a)pyrene
Indeno(l,2,3-cd)

252 (253)
252 (253)
252 (253)
276(138)

D 12 -perylene

264

pyrene

Dibenzo(ah)anthracene

Benzo(ghi)perylene

278(139)
276(138)

are representative of PAHs and exposure to these is more frequent than that
to other PAHs.

The most common analytical methods are based on liquid chromatogra-
phy with fluorescence detection or UV detection (HPLC/FL or UV), or on
gas chromatography with mass selective detection (GC/MSD). Both tech-
niques have their benefits. Generally, HPLC has less resolution, which is
problematic when the studied samples contain complex PAH mixtures. On
the other hand, the UV and the fluorescence detection are highly sensitive
and specific. Mass spectrometry is a powerful tool for identifying individ-
ual compounds. The sensitivity of the GC/MSD can be increased if the mass
spectrometer is operated in selected ion monitoring (SIM) mode. In the lit-
erature, there are numerous applications available for analyzing PAHs that
differ as to, e.g., extraction techniques, solvents used, and clean-up meth-
ods. The method presented here is based on GC/MSD technology and on
the draft international standard method (ISO/DIS 18287 in prep.), but uses
hexane instead of petroleum ether as the solvent. The method is relatively
fast and is applicable to all types of soils, covering a wide range of PAH
contamination. With this method, other PAHs than those in the Table 3.1

3 Quantification of Soil Contamination 111

can be determined as well. A detection limit of 0.01 mg/kg dry mass can be
ensured for each PAH.

■ Equipment

• Usual laboratory glassware free of interfering compounds

• Shaking machine

• Laboratory centrifuge

• Gas chromatograph (GC) with a mass selective detector (MSD)

- Oven temperature program: maintain 60 °C for 2min, then steadily
raise by 20 °C/min up to 180 °C, then raise by 8 °C/min up to 280 °C
and keep at that temperature for 10 min.

- Splitless injection (split closed for 2 min) of 1 p.L.

- Carrier gas: helium 1 mL/min.

• Capillary column specifications: medium polar stationary phase, e.g.,
HP-5MS, film thickness 0.25 p.m, length 30 m, internal diameter 0.25 mm.

■ Reagents

• Acetone

• n-Hexane

• Ion- exchanged water

• Anhydrous sodium sulfate (Na 2 S0 4 ), must be kept at 550 °C for at least
2 h prior to its use

• Silica gel 60 (particle size 60-200 pm), deactivated (Heat silica gel 60 for
5h at 130 °C in a drying oven. Allow to cool down in a desiccator and
add 10% water (w/w) in a flask. Shake for 5 min by hand until all lumps
have disappeared, and then shake for 2 h in a shaking machine. Store
deactivated silica gel in absence of air. It can be used for 1 week.)

• Helium

• Nitrogen

• Quality control soil sample (e.g., certified reference material or in-house
reference material)

• Calibration stock solutions

- Native PAHs (PAHs to be determined): commercially available cer-
tified standard stock solution can be used with a concentration of
approx. 100p.g/mL for each native PAH (e.g., Dr. Ehrenstorfer PAH
Mix 9, X20950900CY).

112 K.S. j0rgensen et al.

- Deuterated PAHs (internal standards): commercially available cer-
tified standard stock solution can be used with a concentration of
approx. 1,000 pg/mL for each deuterated PAH (e.g., Dr. Ehrenstorfer
PAH Mix 31, YA20953100TO). It is recommended that at least five
deuterated PAHs be used as internal standards. The internal stan-
dards are chosen to resemble the physical and chemical properties of
the compounds to be analyzed (see Table 3.1).

• Calibration working solution

- Native PAHs: transfer 5 mL of the calibration stock solution contain-
ing the native PAHs stock solution into a 25-mL volumetric flask and
fill up to the mark with hexane (20 pg/mL)

- Deuterated PAHs: transfer 1 mL of the calibration stock solution con-
taining the deuterated PAHs stock solution to a 25-mL volumetric
flask and fill up to the mark with hexane (40 pg/mL)

• Calibration standard solutions: prepare a series of calibration standards
over a suitable range (e.g., 0.2-10 pg/mL) by transferring 0.1-5 mL of
the native PAH calibration working solution into a 10-mL volumetric
flask and fill up to the mark with hexane. Transfer 1 mL of the standard
solution into a GC vial and add 100 pL of the deuterated PAH calibration
working solution. Each of the calibration standards nominally contains
4 pg/mL of each of the deuterated PAHs. However, laboratories should
determine their own concentration range depending on the samples to
be analyzed.

■ Sample Preparation

Sampling should be performed according to good practices (ISO 10381-1
1994, ISO 10381-2 1994). For the analysis, a homogenized field-moist soil
sample is used (ISO 14507 2003). Stones and other bigger materials ob-
viously not contaminated should not be analyzed. Large particles with
expected contamination should be reduced in size and analyzed with the
finer sample material.

■ Procedure

Extraction Procedure

1. Weigh 10 g of a field-moist (or air-dried-overnight) sample into an ex-
traction flask equipped with a Teflon inlay (a conical flask or a centrifuge
tube with a capacity of 100 mL).

2. Add 25 mL of acetone.

3 Quantification of Soil Contamination 113

3. Close the flask with a screw cap and extract by shaking for 15 min in
a shaking machine.

4. After settling, separate the organic phase into a shaking funnel of
500 mL either by decanting or by using a centrifuge (2,500 rpm, 5 min).

5. Repeat the extraction with 25 mL of acetone.

6. Add 50 mL of hexane to the combined acetone extracts, and remove the
acetone and other polar compounds by shaking with 100 mL of water.
Discard the water and perform another wash in the same manner.

7. If necessary, concentrate the extract on a water bath at 40 °C to about
10 mL using a gentle stream of nitrogen at room temperature. Record
the final volume of the extract and dry the concentrated extract over
anhydrous sodium sulfate.

8. Transfer 1 mL of the dried extract into a GC vial and add 100 p.L of
the deuterated PAH calibration working solution. The sample then
nominally contains 4p.g/mL of each of the deuterated PAHs.

9. Prepare a blank determination in a similar manner but without any soil
sample.

10. Perform an extraction of the quality control soil sample in the same
manner as of the test sample.

Clean-Up Procedure

1. If necessary, the extract can be cleaned with a silica gel adsorption
column. Prepare the column by placing a small plug of glass wool on the
bottom of the column, add 4 g of deactivated silica gel and then about
1 cm of anhydrous sodium sulfate to the top.

2. Condition the column by eluting 10 mL of hexane. When the eluant
reaches the top of the column packing, transfer an aliquot (lmL) of
the concentrated extract containing the internal standards to the top
of the column. Elute with 50 mL of hexane and collect the extract in
a point-shaped test tube.

3. Concentrate the purified extract in a water bath at 40 °C to about 1 mL
using a gentle stream of nitrogen at room temperature.

4. Transfer the purified extract into a GC vial.

Gas Chromatographic Analysis

1. Set the gas chromatograph in such a manner that optimum separation of
the PAHs is achieved. Special attention should be paid to benzo(b)fluor-
anthene and benzo(k)fluoranthene separation.

1 14 K.S. j0rgensen et al.

2. Run the working standard solutions and all the samples by a GC with
mass selective detection in the scan mode (mass range from 50 to
300 amu).

■ Quality Assurance

1. The blank measurement of the total method should be carried out with
each series of soil samples. The PAH concentration in the blank should
be carefully studied, and if traces of contamination are found, the source
of contamination should be investigated.

2. The quality control sample should also be analyzed with each series of
soil samples. The results should be monitored over time and the results
treated statistically.

■ Calculation

For the quantitative analysis, a calibration curve of the ratio of the PAH
determined to the internal standard peak area against the mass of PAH in
the sample injected is constructed using the data handling system. Prepare
these calibration curves for each native PAH using the specific ions (target
ion as the quantitation ion and another ion as the qualifier ion), and the
appropriate deuterated PAH as an internal standard (see Table 3.1).

The amount of PAH in the GC vial (A PA h in p.g/mL) can be obtained from
the calibration curve. Hence, the concentration of the native PAH in the
soil sample can be calculated by the following equation:

c n = x V x / (3.3)

m x d s

c n content of an individual PAH in the sample (mg/kg soil dry mass)

Apah amount of PAH in the GC vial, obtained from the calibration curve
(pig/mL)

V volume of the concentrated extract (mL)

/ dilution factor

m mass of the sample (g wet mass)

d s content of dry mass in the field-moist sample, determined according
to ISO 1 1465 (g dry mass/g wet mass)

■ Notes and Points to Watch

• The samples should be analyzed as soon as possible. If not feasible, the
samples should be stored at -20 °C.

3 Quantification of Soil Contamination 115

• Certain PAHs are carcinogenic and all the samples and standard solu-
tions should be handled with extreme care.

• The efficacy of each silica gel batch has to be tested prior to its use in the
analysis.

• For highly polluted soil samples, clean-up and concentration steps may
not be necessary.

3.5

Heavy Metals

■ Introduction

Objectives. Most heavy metals are of geological origin, but contamination
with them may be due to industrial, mining, agricultural, waste handling
or other activity. Often a mixture of such metals occurs. The most common
contaminants are arsenic, cadmium, chromium, copper, cobalt, lead, mer-
cury, nickel, uranium, and zinc. In contrast to organic contaminants, heavy
metals cannot be degraded by microbes or plants. Thus the bioremediation
strategy is based on the movement of metals, e.g., from soil to plants as
in phytoremediation, or on bioloeaching (see Chapt. 6). Some metals can
undergo microbial oxidation-reduction or become methylated. Different
ionic species of a heavy metal may have different toxicity, e.g., As 3+ is much
more toxic than As 5+ . The method described here gives total concentra-
tion of each metal, but does not give any information on the speciation.
For that purpose separation of the ionic species may be achieved, e.g.,
by ion chromatography, followed by induced plasma mass spectrometry
(ICP-MS).

Principle. Soil samples are freeze dried, homogenized, sieved, digested in
cone. HN0 3 in a microwave oven, and analyzed using ICP-MS.

Theory. Traditional methods for heavy metals' extraction have been based
on digestion in aqua regia (ISO 1 1466 1995) before determination by atomic
absorption spectrometry (AAS), or more recently by ICP-MS. Destruction
with hydrofluoric acid (ISO 14869-1 2001) is being used for some metal
samples, e.g., in geological research. These extraction procedures give the
highest yield of the metal content in a soil sample. However, these agents
pose occupational health risks and alternative digestion using HNO3 has
become common. The yield obtained using this method has been consid-
ered sufficient in many countries for the determination of contamination
with heavy metals (Karstensen et al. 1998). The method described here
employs digestion with HNO3 and analysis by ICP-MS and has earlier been
described by Salminen et al. (2004).

116 K.S. j0rgensen et al.

ICP-MS is a mult i- element analytical technique that can be used to
measure the concentration of several elements simultaneously. The sample
solution is nebulized into the plasma. A large percentage of atoms are
ionized and a fraction of these ions are captured in the interface region of the
system and channeled into the mass spectrometer. The mass spectrometer
serves as a mass filter, and selectively transmits ions according their mass-
to -charge ratio.

The common elements to be analyzed by ICP-MS in soils are Al, As, Cd,
Cr, Cu, Mn, Ni, Pb, Zn, B, Ba, Cs, Fe, Se, Sr, Ti, U, and V. Mercury is best
determined by using the technique of direct combustion, which decom-
poses the sample in an oxygen-rich environment and removes interfering
elements. A dual-path-length cuvette/spectrophotometer specifically de-
termines mercury over a wide dynamic range. The method for mercury
requires no pretreatment other than freeze-drying, but a special piece of
equipment is needed (e.g., an AMA254 Advanced Mercury analyzer); it is
not described here in further detail.

■ Equipment

• Freeze drier

• Microwave oven with Teflon tubes and a cooling system

• Inductively coupled plasma mass spectrometer (ICP-MS)

• Centrifuge and centrifuge tubes

• Polystyrene tubes

■ Reagents

• Water: grade 1 as specified in ISO 3696

• Digestion solution: cone. HN0 3 density 1.42 kg/L (69%)

• Calibration standards: Single or multi-element (SPEX CertiPrep, Metu-
chen, NJ, USA)

- Multi-element solution 2(10 mg/L; Ag, Al, As, Ba, Be, Bi, Ca, Cd, Co,
Cr, Cs, Cu, Fe, Ga, In, K, Li, Mg, Mn, Na, Ni, Pb, Rb, Se, Sr, Tl, U, V,
Zn). Standards are commercially available in 5% HN0 3 .

- Multi-element solution 4(10 mg/L; B, Ge, Mo, Nb, P, Re, S, Si, Ta, Ti,
W,Zr)

• Optimization solution: Mg, Ba, Rh, Pb, and Ce in 1% HN0 3 (10 }ig/L)

• Internal standard: rhodium (1 mg/L)

3 Quantification of Soil Contamination 117

• Control material (NIST, Gaithersburg, MD, USA; SRM NIST 2709 San
Joaquin Soil: Al, Ca, Fe, Mg, P, K, Si, Na, S, Ti, Sb, As, Ba, Cd, Cr, Co, Cu,
Pb, Mn, Hg, Ni, Se, Ag, Sr, Th, V, Zn)

■ Sample Preparation

Use freeze-dried, homogenized, and sieved (< 2 mm) soil samples.

■ Procedure

1. Dry a frozen sample in a freeze-drier.

2. Homogenize the dried sample manually.

3. Sieve the sample (< 2 mm).

4. Weigh accurately 0.25-0.5 g of the dried sample into a digestion tube.

5. Add 5 mL of cone, nitric acid.

6. Set one blank, one reference sample, and one duplicate sample to each
batch.

7. Digestion program: step 1: 250 W, 5min; step 2: 400 W, 5min; step 3:
500 W, lOmin.

8. Cool the digestion tubes to room temperature in a water bath.

9. Open the tubes and transfer each digested solution quantitatively to
a 30 mL plastic tube and dilute with water to a volume of 25 mL.

10. If the samples are not clear, centrifuge at 3,000 rpm for 3 min.

11. Dilute each sample (1:10 or 1:100) to a volume of 10 mL with water and
add the internal standard (100 p.L of rhodium solution).

12. The sample is ready for analysis.

To perform a calibration, proceed as follows:

1. Calibrate the instrument using two calibration solutions, namely, blank
and 50p.g/L of standard solution. Normally multi-element standard so-
lutions are used.

2. Prepare 10 mL of the calibration solution and add the internal standard
as described for soil samples.

3. Perform the calibration and analyze the samples.

118 K.S. j0rgensen et al.

■ Calculation

The mass concentration for each element is determined with the aid of the
instrument's software. Enter the value of the dry mass of each sample into
this software, and it calculates results directly in mg/kg soil dry mass.

■ Notes and Points to Watch

• Pay attention to the interference between/among different metals.

• See that the acid concentration is same in the calibration and the sample
solutions.

• The instrument must be located in a laboratory free of contaminants.

References

Douglas DJ, Houk RS (1985) Inductively Coupled Plasma Mass Spectrometry. Prog Anal.
Atom Spectrosc 8:1

EPA Method 3015 (1994) Microwave assisted acid digestion of aqueous samples and ex-
tracts for total metals analysis by FLAA, Furnace AA, ICP Spectrometry and ICP Mass
Spectrometry

ISO 10381-1 (1994) Soil quality - Sampling - Part 1: Guidance on the design of sampling
programmes

ISO 10381-2 (1994) Soil quality - Sampling - Part 2: Guidance on the design of sampling
techniques

ISO 11465 (1993) Soil quality - Determination of dry matter and water content on a mass
basis - Gravimetric method

ISO 11466 (1995) Soil quality - Extraction of trace elements soluble in aqua regia

ISO 13877 (1998) Soil quality - Determination of polynuclear aromatic hydrocarbons
(PAH) - Method using high performance liquid chromatography

ISO 14507 (2003) Soil quality - Pretreatment of samples for determination of organic
contaminants

ISO 14869-1 (200 1 ) Soil quality - Dissolution for the determination of total element content -
Part 1: Dissolution with hydrofluoric and perchloric acids

ISO 15009 (2002) Soil quality - Gas chromatographic determination of the content of
volatile aromatic hydrocarbons, naphthalene and volatile halogenated hydrocarbons -
Purge-and-trap method with thermal desorption

ISO 15587-2 (2002) Water quality - Digestion for the determination of selected elements in
water - Part 2: Nitric acid digestion

ISO 16703 (2004) Soil quality - Determination of content of hydrocarbon in the range Cio
to C40 by gas chromatography

ISO 17294-1 (2004) Water quality - Application of inductively coupled plasma mass spec-
trometry (ICP-MS) - Part 1: General guidelines

ISO 17294-2 (2003) Water quality - Application of inductively coupled plasma mass spec-
trometry (ICP-MS) - Part 2: Determination of 62 elements

ISO 3696 (1992) Water Quality - Water for analytical laboratory use - Specification and test
methods

3 Quantification of Soil Contamination 119

ISO/DIS 18287 (in preparation) Soil quality - Determination of polycyclic aromatic hy-
drocarbons (PAH) - Gas chromatographic method with mass spectrometric detection
(GC-MS)

ISO/PRF 22155 (in preparation) Soil quality - Gas chromatographic determination of the
content of volatile aromatic and halogenated hydrocarbons and selected ethers - static
headspace method

ISO/DIS 22892 (in preparation) Soil quality - Guideline for GC/MS identification of target
compounds

ISO/TR 11046 (1994) Soil quality - Determination of mineral oil content - method by
infrared spectrometry and gas chromatographic method

j0rgensen KS, Puustinen J, Suortti A-M (2000) Bioremediation of petroleum hydrocarbon-
contaminated soil by composting in biopiles. Environ Pollut 107:245-254

Karstensen KH, Ringstad O, Rustad I, Kalevi K, j0rgensen K, Nylund K, Alsberg T, Olafs-
dottir K, Heidenstam O, Solberg H (1998) Methods for chemical analysis of contaminated
soil samples - tests of their reproducibility between Nordic laboratories. Talanta 46:423-
437

Laine MM, j0rgensen KS (1997) Effective and safe composting of chlorophenol-
contaminated soil in pilot scale. Environ Sci Technol 31:371-378

MLS- 1200 MEGA Microwave digestion system with MDR Technology; Operator Manual
(1992) Milestone, Sorisole, Italy

Owen S, Whittle P (2003) Volatile organic compounds. In: Thompson KC, Nathanail CP
(eds) Chemical analysis of contaminated land. Blackwell Publ, CRC Press, pp 177-188

Reference Manual Elan 5000 (1992) Perkin Elmer, Norwalk, Connecticut, USA

Salminen JM, Tuomi PM, Suortti A-M, j0rgensen KS (2004) Potential for aerobic and anaer-
obic biodegradation of petroleum hydrocarbons in boreal subsurface. Biodegradation
15:29-39

User's Manual Elan 5000 (1992) Perkin Elmer, Norwalk, Connecticut, USA

Immunotechniques as a Tool
for Detection of Hydrocarbons

Grazyna A. Plaza, Krzysztof Ulfig, Albert J. Tien

4.1

RaPID Assay Test System

■ Introduction

Objectives. Immunoassays (IMAs) are now being seen as useful analytical
tools, and supplement to conventional analytical methods such as gas chro-
matography and high performance liquid chromatography. The main IMA
principle can be illustrated by the following reaction: Ab + Ag + Ag* ^>
AbAg + AbAg* (Ab = antibody, Ag = antigen, Ag* = labeled antigen).

Immunochemical methods provide rapid, sensitive, and cost-effective
analyses for a variety of environmental contaminants (van Emon and
Mumma 1990; van Emon and Lopez- Avila 1992; Marco et al. 1995). The
driving force in the development of immunochemical methods is the need
for rapid, simple, sensitive, and cheap tests that can be performed on-
site without requiring sample transfer to an analytical laboratory. The
increasing popularity of field IMA analyses can, in large part, be ascribed
to portable equipment and minimal set-up requirements (van Emon and
Gerlach 1995). Table 4.1 shows advantages and disadvantages of IMAs for
environmental analyses.

The following IMA techniques can be used in environmental studies:
D TECH (Strategic Diagnostics, Newark, DE, USA), PETRORISC (EnSys,
Research Triangle Park, NC, USA), EnviroGard (Millipore, Billerica, MA,
USA), and RaPID (Ohmicron, Newtown, PA, USA) assays. Table 4.2 presents
the properties of these systems and their application matrices and detection
limits.

Principle. The RaPID assay uses magnetic particles as the solid-support
component of the ELISA (enzyme-linked immunosorbent assay). Attach-
ing antibodies to microscopically small magnetic particles facilitates the
chemical reaction between antibody and contaminant. The concentration
of the compound to be detected is quantified after a color reaction.

Grazyna A. Plaza, Krzysztof Ulfig: Institute for Ecology of Industrial Areas, 40-844 Katowice,
6 Kossutha, Poland, E-mail: pla@ietu.katowice.pl

Albert J. Tien: Holcim Group Support Ltd Corporate Social Responsibility Occupational
Health and Safety, Im Schachen, 5113 Holderbank, Switzerland

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

122

G.A. Plaza et al.

Table 4.1. Some advantages and disadvantages of IMAs for environmental analysis (accord-
ing to Sherry 1992; Sherry 1997)

Advantages

Disadvantages

Wide applicability

Sensitive, specific, and highly selective

Rapid and easy to use

Reduced preparation

Rapid with high sample throughput

Ideal for large sample loads;
easily automated

Suited to laboratory and field use

Cost-effective analysis
of small-volume samples

Development costs

Hapten synthesis can be difficult

Can be vulnerable to cross reacting
compounds and non-specific
interferences

Requires independent confirmation

Not suited to small sample loads
or multi-residue determinations

Lack of acceptance

Theory. One of the most common enzyme immunoassay (EIA) modifica-
tions, sometimes termed "double antibody sandwich techniques," is ELISA
(van Emon and Mumma 1990). ELISA is based on combining selective an-
tibodies with sensitive enzyme reactions to produce analytical systems
capable of detecting very low levels of chemicals. The RaPID system uti-
lizes covalent binding of antibodies to magnetic particles that are made
of silanized iron oxide (Fig. 4.1). The first stage is the immunochemical
reaction between antibodies/magnetic particles and a chemical compound
as antigen. The second stage is separation of magnetic particles from the
antigen by applying a magnetic field. After washing, the color reagent is
added and the concentration of the colored product is measured (RaPID
assay environmental user's guide 1996; Plaza et al. 1999). The assay steps
are presented in Fig. 4.2.

Equipment

RPA-I RaPID analyzer (spectrophotometer): laboratory bench-top-
based, single wavelength, microprocessor-controlled analyzer

Magnetic rack composed of two parts: the top rack holds the test tubes
in place and the bottom base contains the magnets

Portable balance

Test tubes

Vortex mixer

Timer

4 Immunotechniques as a Tool for Detection of Hydrocarbons

123

Table 4.2. A comparison of immunological test systems (EnviroGard Protocol 2004b,
www.sdix.com)

DTECH

EnSys

EnviroGard

RaPID Assay

Technology

Latex

Coated

Magnetic

particle

tube

particle

Result type

Qualitative

and semi-

quantitative

Sample throughput

1-4

1-10

1-17

1-50

samples/h

Analy

sis time

20 min/run

30 min/run

60 min/run

EPA SW-846 method

4030, 4035

Storage

' shelf life

Ambient

Refrigerated

1 year

Training level

Low;

Medium;

no training

training

required

recommended

Instrument

DTECHTOR

Photometer

RPA-1 Analyzer

Analyzer or

color card

Application matrix and detection limits

Analyte

Application

BTEX

Soil

2.5-35 ppm

—

2 ppm

0.9 ppm

Water

0.6-10 ppm

—

0.1 ppm

0.09 ppm

TPH

Soil

—

10 ppm

5 ppm

10 ppm

Water

—

0.1 ppm

1 ppm

PAH

Soil

—

1 ppm

0.2 ppm

Water

—

15 ppb

2 ppb

0.9 ppb

Carcino-

Soil

—

10 ppb

genic PAH

Water

—

0.2 ppb

Reagents

All the reagents (Extraction Solution, Enzyme Conjugate, Antibody-
Coupled Magnetic Particles, Color Reagent, Washing Solution, and Stop-
ping Solution) are supplied by Ohmicron, Newtown, PA, USA, and their
composition is under protection.

Sampling and Sample Preparation

Collect water and soil samples from the contaminated area in 500 mL
wide-mouth bottles (Nalgene; Nalge Nunc, Naperville, IL, USA).

124

G.A. Plaza et al.

TRADITIONAL
IMMUNOASSAYS

MAGNETIC PARTICLE-BASED
TECHNOLOGY

Antibody

Antibodies are typically
coated on plastic test tubes
or plates

Magnetic particle
with antibody attached

Antibodies are linking to
microscopically small
magnetic particles which
speed up the chemical
reaction between the antibody
and the contaminant

Fig. 4.1. Comparison of traditional IMAs and magnetic-particle-based technology (RaPID
assay environmental user's guide 1996)

Test directly samples or store them at 4 °C; water content in soil samples
should not be more than 20-25%.

Extract soil samples before testing:

- Weigh 10 g of soil into the soil collection tube (Fig. 4.3) and add 20 mL
of the extraction solution; screw the cap on tightly.

- Shake vigorously and continuously for at least 60 s.

- Remove the screw cap and attach the filter cap, then attach the plunger
rod to the plunger of the soil collector, and filter the extract into the
Extract Collection Vial.

- Fill with 0.5-1 mL of the filtrate and cap the vial.

■ Procedure

1. Mix 200 pL of soil extract or water sample with 250 pL of the Enzyme
Conjugate and 500 pL of antibody-coupled magnetic particles; incubate
the mixture for 15 min at room temperature.

2. Put all tubes into the magnetic rack and wait 2 min for the particles to
separate.

4 Immunotechniques as a Tool for Detection of Hydrocarbons

125

Immunological reaction

o — «] — *

Separation

Stepl

Color Development

Step 3

Legend

o — < Magnetic particle with antibody

<\ — * Antigen conjugate with

^ Antigen

□ Chromogen substrate

■ Colored Product

Fig. 4.2. Principle of RaPID assay (RaPID assay environmental user's guide 1996; Plaza et al.
1999)

Soil collector

TOP _ Screw cap

BOTTOM

Luer cap

Plunger rod Plunger

Luer cone

Fig. 4.3. Soil collection tube (RaPID assay environmental user's guide 1996)

126 G.A. Plaza et al.

3. Add 1 mL of washing solution, vortex each tube and wait 2 min. Repeat
this step.

4. Add 500 pL of color reagent. S^'^^-tetramethylbenzidine is used as
the chromogen. Incubate for 20 min at room temperature.

5. Add 500 pL of stopping solution (0.5% sulfuric acid).

6. Within 15 min after adding the stopping solution, transfer 1 mL of solu-
tion to cuvettes and read absorbance of standard solutions and samples
at 450 nm using the RPA-I analyzer.

■ Calculation

1. The concentration of the colored product is directly proportional to the
concentration of the labeled compounds.

2. The RPA-I analyzer can perform mathematical computations, and sam-
ple concentrations with statistics are obtained. Results are directly re-
ported in ppb (ng/g) or ppm (pg/g).

3. The hydrocarbon concentration in soil is calculated according to the
following formula, taking into account the concentration calculated by
the analyzer (ppb or ppm), the volume of extraction solution (20 mL),
and the mass of soil used for extraction (g dry mass):

concentration x volume

Hydrocarbon concentration (ppb or ppm) =

soil mass

■ Notes and Points to Watch

• Temperature control is required for reagent storage (4-8 °C) and during
the performance of the assay (room temperature: 15-30 °C).

• Use specific test kits for specific hydrocarbons; do not mix the reagents.

• All the reaction steps should be done exactly according to the assay
protocol.

• Do not use test kit components after the expiration date.

4.2

EnviroGard Test System

■ Introduction

Objectives. Environmental IMAs have been developed and evaluated for
analyses including major classes of pesticides, organic compounds such

4 Immunotechniques as a Tool for Detection of Hydrocarbons 127

as polychlorinated biphenyls (PCB), polyaromatic hydrocarbons (PAH),
pentachlorophenols (PCP), benzene/toluene/ethylbenzene/xylene (BTEX),
total petroleum hydrocarbons (TPH), dioxins and furans, microbial tox-
ins, as well as inorganic compounds such as cadmium, lead, and mer-
cury (Vanderlaan et al. 1988; van Emon and Mumma 1990; Sherry 1992;
van Emon and Lopez- Avila 1992; Knopp 1995; van Emon and Gerlach
1995; Gerlach et al. 1997). The EnviroGard test systems are quick and
reliable with semi-quantitative results allowing screening at various lev-
els of contamination (Table 4.2). They can be used during site remedia-
tion to detect contaminants and monitor the cleanup and industrial pro-
cesses.

Principle. EnviroGard uses coated polystyrene test tubes as the solid sup-
port component of the ELISA. The system is based on the use of polyclonal
antibodies, immobilized on the test tube walls, that can bind specific con-
taminants.

Theory. The EnviroGard IMA kit uses 12 x 75 mm polystyrene test tubes
coated with an antibody against the target contaminant (analyte). Coated
polystyrene test tubes allow screening for various contaminations like PCB,
PAH, TPH, BTEX, and PCP. When analytes are present in the sample, they
compete with the specific Enzyme Conjugate (labeled analyte) for a limited
number of binding sites on the antibodies (EnviroGard Protocol 2004a,
2004b). According to the principles of competitive IMAs, the absorbance
signal (or optical density) of the final reaction mixture is inversely pro-
portional to the concentration of the contaminant (analyte) present in the
test sample. After the immunological reaction, the unbound molecules are
washed away and a chromogenic substance is added to the test tube. In
the presence of bound specific enzyme conjugate, the clear substance is
converted to a blue color.

Equipment

SDI Sample Extraction Kit contains devices to process 12 samples, i.e.,
12 each: extraction jars with screw caps, filter modules, ampule crack-
ers, wooden spatulas, weigh canoes, disposable transfer pipettes, and
ampules each containing 10 mL of 100% methanol

20 antibody-coated test tubes (12 x 75 mm), 1 vial of negative control
(methanol), calibrator vials, 1 vial of Hydrocarbons-Enzyme Conjugate,
1 vial of Substrate Solution, 1 vial of Stop Solution, pipettes and tips

Portable balance

Spectrophotometer

128 G.A. Plaza et al.

• Test tube rack

• Timer

■ Reagents

• Methanol (100%)

• Stop solution: 1 N HC1

■ Sampling and Sample Preparation

• Collect soil without excess twigs, rocks, or pebbles in labeled 500 mL
plastic containers.

• Dry soil samples with a water content of 30% or more (by mass) before
testing.

• Store soil samples at 4 ° C in tightly sealed containers to avoid evaporative
losses.

• Use 10 mL of methanol to extract hydrocarbons from 10 g soil. When
extracting clay samples add an additional 10 mL of methanol to the
sample and shake vigorously for 1-2 min.

• Filter the extract using Whatman #1 filter paper. A clay sample may soak
up all the methanol, leaving little or no excess liquid to filter.

■ Procedure

1. Incubate all reagents at room temperature (at least 1 h) before use.

2. Remove the antibody-coated test tubes (20 tubes/assay) from the foil
pouch and label as negative control (NC), calibrators (CI, C2, etc.), and
samples (SI, S2, etc.).

3. Place the test tubes in the test tube rack, and add 100 pi of sample extract.

4. Add 200 pi of specific enzyme conjugate to all test tubes.

5. Gently shake the rack to mix for 10-15 s, and leave the tubes undisturbed
for 10 min.

6. Shake out the test tube contents vigorously into a sink, and wash the test
tubes with double distilled water. Repeat this wash step three times.

7. Add 500 pi of substrate solution to all test tubes, briefly shake, and then
incubate for 5 min at room temperature.

8. Add 500 pi of stop solution to all test tubes.

4 Immunotechniques as a Tool for Detection of Hydrocarbons 129

9. The results can be interpreted visually within 5 min after adding the
substrate solution, or can be obtained more precisely with a spectropho-
tometer.

■ Calculation

1. One enzyme molecule can convert many substrate molecules. Every
test tube has the same number of antibody-binding sites and receives
the same number of the specific enzyme conjugate molecules. Color
development is inversely proportional to the hydrocarbon concentration,
i.e., dark color indicates low concentration and light color indicates high
concentration.

2. Visual interpretation: compare the sample test tube to the calibrator test
tubes against a white background. If a sample test tube contains a darker
color than the calibrator test tube, the sample contains hydrocarbons at
a concentration lower than the calibrator. If the sample test tube is less
colored than the highest calibrator test tube, the soil extract should be
diluted in methanol and the assay performed again.

3. Photometric interpretation: place the negative control test tube into the
spectrophotometer. Record the optical density (OD) at 450 nm for the
negative control, and then measure the OD of calibrator #1. Repeat this
step to determine the OD for each of the remaining calibrators and for
samples.

An example of data is given here:
Tube OD 450 nm Interpretation

NC (negative control)

1.00

CI (2ppm)

0.87

C2(10ppm)

0.52

C3 (50 ppm)

0.25

SI (sample 1)

0.45

> 10 ppm, < 50 ppm

S2 (sample 2)

0.72

> 2 ppm, < 10 ppm

Notes and Points to Watch

The EnviroGard test is a screening test only.

The test system can be used in the laboratory and in the field.

Store all test kit components at 4-8 °C when not in use.

Storage all reagents at ambient temperature (18-27 °C) on the day before
using.

130 G.A. Plaza et al.

• Use only reagents or test tubes from one kit; do not mix the components
from different test kits.

• Do not expose substrate to direct sunlight.

• Do not freeze test kit components or expose them to temperature greater
than 37 °C.

• Use gloves and protective clothing during the experiment.

References

EnviroGard™ Protocol (2004a) Petroleum fuels in soil. Test kit 70040000. Strategic Diag-
nostics Inc, USA

EnviroGard™ Protocol (2004b) Remediation assessment and industrial testing. Strategic
Diagnostics Inc, USA (www.sdix.com)

Gerlach RW, White RJ, O'Leary WFD, van Emon JM (1997) Field evaluation of an immunoas-
say for benzene, toluene and xylene (BTX). Wat Res 31:941-945

Knopp D (1995) Application of immunological methods for the determination of environ-
mental pollutants in human biomonitoring. A review. Anal Chem Acta 311:383-392

Marco MP, Gee S, Hammock BD (1995) Immunochemical techniques for environmental
analysis. II Antibodies production and immunoassay development. Trends Anal Chem
14:415-428

Plaza G, Ulfig K, Bevolo AJ (1999) Application of the immunoassay techniques for the
determination of PAHs and BTEX in soil. Int Environ Tech 9: 9-10

RaPID assay environmental user's guide (1996), OHMICRON, USA

Sherry J (1992) Environmental chemistry. The immunoassay option. Crit Rev Anal Chem
23:217-300

Sherry J (1997) Environmental immunoassays and other bioanalytical methods. Overview
and update. Chemosphere 34:1011-1025

Van Emon JM, Gerlach CL (1995) A status report on field-portable immunoassay. Environ
SciTechnol 29:312-317

Van Emon JM, Lopez- Avila V (1992) Immunochemical methods for environmental analysis.
Anal Chem 64:79-88

Van Emon JM, Mumma RQ (1990) Immunochemical methods for environmental analysis.
ACS Symp Series 442, Am Chem Soc, Washington, DC

Vanderlaan M, Watkins BE, Stanker L (1988) Environmental monitoring by immunoassay.
Environ Sci Technol 11:247-254

Feasibility Studies for Microbial Remediation
Hydrocarbon-Contaminated Soil

Ajay Singh, Owen P. Ward, Ramesh C. Kuhad

5.1
Introduction

While bioremediation processes are considered to be advantageous in terms
of their relatively low cost, process flexibility, benign nature environmen-
tally, and on-site utility, there have also been many instances where the
processes have failed to achieve the required low contaminant concentra-
tion criteria (Mandelbaum et al. 1995; Iwamoto and Nasu 2001; Grommen
and Verstraete 2002). These failures have reduced consumer confidence
in bioremediation and consequently the technology only garners a small
portion of the US$ 7-8 billion US annual remediation market (Srinivasan
2003). Bioremediation processes must comply with generally accepted good
operating principles and have predictable end-points. The processes must
be validated in advance such that they do not fail (Ward 2004). Feasibil-
ity studies are therefore critical for the implementation of a successful
bioremediation technology.

Contaminated sites never exhibit identical characteristics and the expe-
rience from one site can only be exploited at another to a limited extent.
The biodegradation process in soil is complex, involving diffusion of con-
taminants in the soil matrix, adsorption to the surface of soil particles, and
biodegradation in the biofilms existing on the soil particles, in pores, and
in the bound and free water after desorption from the soil surfaces. A va-
riety of complex biodegradation patterns result from physical interactions
between pollutants and soil matrix and from biological interactions among
different organisms. Numerous factors such as soil moisture, pH, temper-
ature, aeration, nutrient sources, type of soil, type(s) of contaminant(s),
and interplay between these factors, affect the ecology of the microbial
population and degradation of hydrocarbons in contaminated soil.

For full-scale bioremediation applications, several important points need
to be considered. In particular, how low can the concentration of the con-

Ajay Singh, Owen P. Ward: Department of Biology, University of Waterloo, Waterloo, On-
tario, Canada N2L 3G1, E-mail: ajasingh@sciborg.uwaterloo.ca

Ramesh C. Kuhad: Department of Biotechnology, Kurukshetra University, Kurukshetra -
136 119, Haryana, India

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

132 A. Singh et al.

taminant be obtained during treatment considering: bioavailability and
microbial activity, the fate of the contaminant in terms of mineralization,
biotransformation, evaporation, build-up of microbial biomass, sorption
to soil - and also considering indicators of time needed to obtain the set
goal such as degradation rate for achieving the target level of contaminant,
and finally the capital and operating costs?

Both slurry bioreactors and land treatment technologies have success-
fully been used to remove hydrocarbons from petroleum-contaminated
soils. Naturally occurring or introduced microbial populations convert
hydrocarbons to carbon dioxide, water, biomass, and humic material. For
a successful bioremedial treatment of soil, it is important to consider several
factors, including type and extent of contamination, bacterial population
present, duration since contamination, optimal microbiological conditions,
soil characteristics, proper bioremediation technique, and appropriate an-
alytical method. Proper planning for the execution of a bioremediation
technology based on the above criteria is of utmost importance to minimize
risk of failure in terms of effort, time, and money. Protocols for conducting
feasibility studies are, therefore, required to evaluate the effectiveness of
a planned treatment method.

In this Chapter, methods commonly used for determination of biodegra-
dation potential in feasibility or biotreatability studies on bioremediation
of contaminated soils and sludges are discussed. During feasibility stud-
ies certain environmental and nutritional parameters are optimized for
achieving accelerated and complete biodegradation of hydrocarbons.

5.2

Determination of Biodegradation Potential

The data from the feasibility studies of hydrocarbon-contaminated soil is
used to design a suitable full-scale bioremediation technology. Hence, it is
important to carefully plan the biotreatability studies. Successive stages of
planning and execution of feasibility studies are shown in Fig. 5.1.

5.2.1

Sampling and Soil Preparation

Preparation of soil samples for site characterization or biodegradation
experiments can alter the physico-chemical and biological properties of the
original soil present in the field. In order to assess the extent of the overall
soil contamination, proper field sampling procedures are required to obtain
representative samples. The spatial variability of soil characteristics maybe

5 Feasibility Studies for Microbial Remediation

133

Hydrocarbon-contaminated site/soil/sludge

Assessment of contaminated waste or site and sampling

Analysis of samples and characterization of waste

Feasibility or biotreatability studies planning

Microcosms or Mesocosms or Composting technique

Determination of biodegradation potential

Optimization of environmental and nutritional conditions

Analysis and process monitoring

Evaluation and interpretation of results

Measurements of success

Selection of the appropriate bioremediation technology

Cost-benefit analysis

Design and implementation of the technology

Fig. 5.1. Successive stages for planning and implementation of feasibility studies

considered for determination of the location and number of representative
samples in heterogeneous and very large sites. The number of samples to be
collected and analyzed usually depends on the overall sampling objective,
contaminant distribution, size of the contaminated site and sampling and
analytical costs (Huesemann 1994a).

1. Samples may be collected from a uniform depth, using proper soil sam-
pling equipment such as a core or a split-spoon sampler at 20-30 lo-
cations and pooled. Top 10-15 cm of soil is often sampled. There are
many procedures for choosing a sampling location. The most popular
ones are either randomly selecting field locations for periodic sam-
pling or placing a hypothetical square or rectangular grid over the site
and taking samples at the center of each grid. Samples should be ho-
mogenized prior to subsampling and submission for analysis (see also
Chapt. 1).

2. Soil samples are stored in either plastic bags or in glass or other non-
reactive containers in a cooler on ice for immediate analysis. For long
term storage, the samples should be at the field moisture levels and

134 A. Singh etal.

stored in glass or other non-reactive container around 4°C. Air drying
of samples should be avoided.

3. For biodegradation experiments, a relatively homogeneous sample
should be prepared by sieving soil through a 2-4 mm sieve and soil
moisture level determined so that all the experiments are carried out
using the same conditions. Depending on the aim of the experiment,
contaminant level, water content, pH, organic matter, nutrient levels,
and type of soil are determined by standard methods (see Chapt. 2).

5.2.2

Selective Microbial Enrichment

Successful application of a bioremediation technique requires the iden-
tification of favorable conditions for the useful hydrocarbon-degrading
microorganisms to actively grow and metabolize contaminants. Classical
isolation methods, selective enrichment of specific microbes, and genetic
approaches can be used to obtain a single microbial species or a group
of different microorganisms (a consortium; see Chapt. 13). Selective en-
richment is the most practical approach for large scale applications where
the enrichment process is designed to increase the population of specific
microorganisms. Suitable conditions and selective pressures are applied to
encourage growth of microbes capable of degrading a particular substrate
or a mixture of potential contaminants, in the growth medium, that are the
targets for biodegradation or bioremediation as the sole sources of carbon
(Vecchili etal. 1990).

The initial inoculum can be obtained from the contaminated soil, sludge,
or wastewater with known degradative activity. A consortium of hydrocar-
bon-degrading microorganisms can be obtained by adding about 1 or 2 g
of hydrocarbon-contaminated soil or sludge to 100 mL mineral medium in
Erlenmeyer flasks. Culture can be further maintained in a flask by routinely
transferring a 2% inoculum into a fresh medium at weekly intervals.

Cyclone fermenters (Liu 1989) with a 1-L working volume can also be
used to develop and maintain hydrocarbon-degrading cultures (Fig. 5.2).
The cultures obtained by selective enrichment in the flasks can be inoc-
ulated into a liter of mineral medium in a cyclone fermenter. The culture
is maintained once weekly by removing about 50% of the volume of the
culture and replacing with fresh medium and a known amount of the
hydrocarbon as a sole carbon source.

Soil columns may be used to enrich for hydrocarbon-degrading cultures.
Glass soil columns with an inner diameter of 40 mm and length of 350 mm
packed with 50 g air-dried soil and slightly moist pre-washed quartz sand
between two layers of glass wool can be used to enrich desired strains

5 Feasibility Studies for Microbial Remediation

135

Fig. 5.2. Cyclone fermenters used for main-
tenance of hydrocarbon-degrading mi-
croorganisms

(Pfarl et al. 1990). The columns are rinsed with mineral medium several
times prior to enrichment process and air is provided by using compressed
air bubbled through distilled water at 1-2 L/h.

5.2.3
Controls

In bioremediation experiments, sterile controls should frequently be used
to demonstrate the biological activity and biodegradation process, since
abiotic loss mechanisms such as adsorption and volatilization can occur
simultaneously. Soil sterilization can be achieved by various physical and
chemical methods. Methods include using chemicals such as mercuric
chloride and sodium azide, as well by as autoclaving and providing gamma
radiation. For inoculation of sterile soil, autoclave and gamma radiation
are the most suitable because no residual chemical is left after sterilization.
However, any sterilization method will alter the soil properties. Wolf et al.
(1989) found that mercuric chloride had the least effect on soil properties
such as pH, surface area, and release of Mn among various sterilization
methods/agents tested and compared such as gamma radiation, microwave,
dry heat, propylene oxide, sodium azide, mercuric chloride, chloroform,
and antibiotics.

In bioaugmentation studies, where the experiments are conducted with
added inoculum to the soil, a killed-culture inoculum treatment should
be included as control for possible nutrient effects of dead microbial cells.
Mass balance of the existing or spiked contaminant should be determined
at the conclusion of the experiment to evaluate contaminant disappearance
due to biodegradation or abiotic mechanism.

136 A. Singh et al.

5.2.4

Soil Microcosms

One of the simplest methods requiring minimal equipment for soil biode-
gradation studies is with use of a biometer flask (Bellco, Vineland, NJ, USA).
The United Sates Environmental Protection Agency (US EPA) and Organi-
zation of Economic Cooperation and Development (OCED) have also rec-
ommended this method (OECD 1981; McFarland et al. 1991; Skladany and
Baker 1994). Biodegradation activity can be evaluated by directly monitor-
ing the loss of the target compounds or indirectly by measuring by-products
of biodegradation or electron acceptor consumption.

A biometer flask, a 250-mL Erlenmeyer flask with a side arm contain-
ing potassium hydroxide to trap C0 2 evolved during biodegradation, is
used in batch experiments to monitor degradation of the target compound
present in or added to the contaminated soil. For biodegradation feasibil-
ities studies, around 20% (w/v) aqueous soil suspension is recommended.
Flasks are incubated with or without C0 2 -free air and periodically KOH
solution is withdrawn and titrated with a standard acid solution to de-
termine the amount of C0 2 produced. The matrix can be analyzed at the
end of the test for organic and inorganic compounds. The biometer flasks
can be modified to investigate specific problems related to specific types
of contaminants and challenges in studying a given biodegradation. This
flask system can be used to study biodegradation of both semi- volatile and
volatile compounds, and to screen commercial inoculates as well.

An electrolytic respirometer, designed to measure the oxygen uptake or
rate of respiration by microbes in soil and sludge has been used by the
US EPA for evaluation of commercial products for use in Prince William
Sound, Alaska (Venosa et al. 1992). The respirometer consists of a reactor
module connected to an electrolytic oxygen generator. The depletion of
oxygen by microbes creates a vacuum that triggers the oxygen generator.
The electricity used to generate the oxygen is proportional to the amount
of oxygen (mg/L), while the C0 2 produced by microbial activity is trapped
in KOH solution. The decision to choose a better amendment is based on
high oxygen uptake rate, growth of degraders, and significant degradation
of aliphatic and aromatic hydrocarbons.

Another method to quickly determine biotreatability of hydrocarbon-
contaminated soils and sludges is to simply use 250 mL Erlenmeyer flasks
with working volumes of 50 mL containing 20% (w/v) soil or sludge slurry.
For petroleum-contaminated soil or sludge samples, total petroleum hy-
drocarbon (TPH) content is determined as hexane-extractable material.

1. Set up at least 6 flasks for each test.

2. Add a known mass of sludge or contaminated soil to the flask in order

5 Feasibility Studies for Microbial Remediation 137

to obtain less than 20% solids and 10% TPH concentration in a total
working volume of 50 mL.

3. Add 45 mL of the nutrient medium and 1.25 mL (0.25% final concentra-
tion) of a non-ionic surfactant (10% w/v stock solution).

4. Adjust pH of the contents to 6.8-7.2 using 5 N NaOH or 5 N HC1.

5. Inoculate the flask with 2.5 mL (5%, v/v) of a microbial inoculum.

6. Incubate the flasks at 30 °C for 14 days on a shaker (200 rpm).

7. Extract the whole contents of 2 flasks with equal volume of n-hexane at
the starting time of the test to determine initial TPH content. Extract
contents of 2 flasks each with hexane after 7 and 14 days to determine
residual TPH contents.

8. After determination of TPH (Chapt. 3), dissolve the residue in a known
volume of hexane for gas chromatographic analysis of hydrocarbons.

In the above method, at least duplicate flasks should be set up for each
sampling point and the contents of whole flasks should be extracted to
determine residual hydrocarbons. Appropriate, controls for abiotic losses
should be also be set up as described above.

5.2.5

Slurry Bioreactors

The slurry bioreactor approach is to suspend and mechanically mix soil
in aqueous solutions in a contained vessel or tank. Land-based systems
usually require very long treatment times due to lack of control of envi-
ronmental factors such as seasonal variation in temperature, pH, moisture,
as well as of natural microbial activity, and mixing and circulation limita-
tions. These problems can be eliminated in bioreactor systems, which are
characterized by much higher rates and extents of degradation due to the
minimization of mass-transfer, increased desorption of contaminants by
continuous mixing, and control of environmental and nutritional factors
such as pH, temperature, and moisture, bioavailability of nutrients and oxy-
gen in order to promote rapid microbial growth and activity (Singh et al.
2001; Van Hamme et al. 2003).

Process conditions in bioreactors can be optimized for biodegradation
depending on the nature of contaminant. Desired temperature and pH can
be consistently maintained throughout the process and suitable amend-
ments such as nutrients, surfactants, and microbial cultures can be sup-
plied. Several examples of slurry reactors can be found in the literature.

138 A. Singh etal.

A method developed in the authors' laboratory and successfully scaled up
for field applications is described here.

1. Depending on the availability, use a 1-5 L or even larger volume biore-
actor fitted with pH, temperature, and dissolved oxygen control for
biotreatability studies. Alternatively, construct an inexpensive bioreac-
tors by putting an air sparger in a glass or metal beaker or container.

2. For biotreatability studies in the bioreactor, and depending on the soil
and sludge composition, mix a sample of about 20% solids by mass
with aqueous nutrient medium.

3. Depending on the critical micellar concentration, add a non-ionic sur-
factant with a hydrophilic-lipophilic balance (HLB) value 12-13 to ob-
tain final concentration of 0.05-0.25%.

4. Adjust the pH of the medium to around 7.0 using NaOH or HC1 solu-
tions.

5. Add the inoculum, prepared and maintained in a cyclone fermenter as
described in Sect. 5.2.2, at the level of 10% (v/v) to the bioreactor.

6. Maintain an aeration level of 0.1-0.2 wm (volume per volume per
minute) during the process to avoid oxygen limitation in the system.
Dissolved oxygen concentration should be maintained above 2 mg/L.

7. A small mixer can also be used at about 200-300 rpm to achieve better
mixing of the reactor contents.

8. Keep the temperature at between 28 and 32 °C using a water bath or
heater.

9. Monitor the pH regularly and maintain it between 6.5 and 7.5 through-
out the process.

10. Compensate for any losses due to evaporation of water by adding water
to the working volume level.

11. Total microbial count and hydrocarbon-degrading bacteria can be de-
termined at regular intervals to monitor the progress of biodegradation.

12. Monitor biodegradation of hydrocarbons at periodic intervals for 2-
4 weeks.

The experimental design and data analysis during a biotreatability study
will depend on the specific aim of the study. The slurry reactor experiments
should be repeated to ensure consistent results. While sub-sampling over
an extended period in a bioreactor experiment, care should be taken to
ensure that the volume in the reactor is not drastically reduced.

5 Feasibility Studies for Microbial Remediation 139

5.2.6

Land Treatment

A set of laboratory experiments using contaminated soil can be carried
out in order to investigate the feasibility of land treatment of such soil.
Biodegradation potential of a particular hydrocarbon waste can be de-
termined by the extensive chemical characterization of the petroleum-
contaminated soil. Huesemann (1994b) has provided useful guidelines on
carrying out laboratory feasibility studies on potential of land treatment of
petroleum-contaminated soil.

Laboratory mesocosms to study biodegradation of petroleum hydrocar-
bons in contaminated soil can be prepared in open glass or metal trays as
follows:

1. Trays containing 5-10 kg of contaminated or spiked soil are prepared.

2. Oil and grease or TPH content is determined and adjusted in the range
of 5-7% by diluting with clean soil.

3. To obtain optimal soil moisture content for the microbial activity, soil
moisture is adjusted to between 50 and 80% of the field capacity (water-
holding capacity), usually between 10 and 16 g of water per 100 g of dry
soil.

4. Adjust the pH to around 7.0 using lime, caustic soda, elemental sulfur
or ammonium sulfate.

5. The trays should be incubated at the optimum temperature range for
microbial degradation of 25-35 °C.

6. For each 100 kg of oil to be degraded, 1 kg of nitrogen and 0.2 kg of
phosphorus should be added as nutrient fertilizer to obtain an oil:N:P
ratio of 100:1:0.2.

7. The duration of the biotreatability study depends on the overall ob-
jective of the project. In general, it is recommended to run for 3-6
months.

8. Oil and grease or TPH content, moisture and pH should be periodically
monitored.

9. The soil should be lightly raked or mixed at 1-2-week intervals to
provide proper aeration, mixing, and moisture control.

10. The moisture content should be monitored at 1- or 2-week intervals and
the soil sprayed with water to adjust to the optimum moisture content.

Monitoring the disappearance of oil and grease or TPH, as well as mois-
ture, pH, and nitrogen is important during the treatability studies. Total

140 A. Singh et al.

heterotrophic or hydrocarbon-degrading microbial counts may also be
monitored to evaluate the biodegradation process. It is important to use
the same sampling strategy and methods throughout the treatment period.

5.2.7
Composting

While composting of yard and municipal wastes has been performed
for decades, composting of hydrocarbon-contaminated soils represents
an emerging ex-situ biological technology. Composting has been demon-
strated to be effective in biodegrading explosives and polycyclic aromatic
hydrocarbons (PAHs) in soils (USEPA 1996, 1998). In the composting of
contaminated soil, organic amendments including manure, sewage sludge,
compost, yard wastes, and food processing wastes are often added to supple-
ment the amount of nutrients and readily degradable organic matter in soil.
Sewage sludge and compost containing abundant nitrogen, organic mat-
ter, and high microbial diversity, with total microbial populations higher
than fertile soils, have great potential in bioremediation. A small-scale
biotreatability method (Van Gestel et al. 2003) for composting technology
is described here:

1. Two insulated composting bins can be used, one filled with biowaste
(vegetable, fruit, garden, and paper waste) only, and the other filled
with a mixture of biowaste and petroleum-oil-contaminated soil at
a 10:1 ratio (fresh mass).

2. Dewatered sewage sludge or matured compost can be used instead of
biowaste.

3. Spruce bark can be used as a bulking agent at the ratio of soil to bulking
agent, 1:3 on a volume basis.

4. The soil should be collected from the top 15 cm of the soil surface and
air dried and sieved to pass a 2-4 mm sieve.

5. The soil can be spiked with commercial crude oil or diesel oil at a con-
centration to obtain a concentration of 5-10 g/kg after mixing with the
biowaste.

6. The initial pH is adjusted about 7.0-7.4.

7. The composting process is controlled using airflow and moisture con-
tent.

8. Aerobic composting can be performed for 12 weeks.

5 Feasibility Studies for Microbial Remediation 141

9. At regular time intervals, the content should be turned to avoid prefer-
ential aeration pores.

10. Compost samples for chemical and microbiological analyses should be
taken every time the compost is mixed.

11. Microbial counts, dry matter content, pH, temperature, electrical con-
ductivity, and exhaust gas composition should be regularly monitored.

12. Microbial composition of the biowaste-only composting bin serves as
a reference for the composting process of contaminated soil.

13. To investigate the degradation rate of oil in soil alone, a soil-only exper-
iment (without organic amendments) should also be run as a control.

Composting technologies can be applied to cleanse contaminated soil ex
situ. By adding an organic matrix to contaminated soil the general microbial
activity is enhanced and also the activity of specific degraders, which may
be found in the contaminated soil or introduced along with the organic
material. Biodegradation rates in composting systems have been found to
be slightly higher than in land treatment of hydrocarbons and lower than
in slurry reactors.

5.2.8
Scale-Up

The data obtained from the small-scale biodegradation experiments can
be used to design full-scale biotreatment systems. In most cases slurry
bioreactors can be directly scaled up. The US EPA has suggested a three-
tier approach before a full-scale application of the technology in the field
(US EPA, 1991; McFarland et al. 1991):

1. Laboratory screening to establish the occurrence and rate of biodegra-
dation and establishing optimum process parameters

2. Bench-scale testing to establish performance of the process parameters
and cost estimate for the scale-up of appropriate technologies

3. Pilot testing on the most promising technology to establish system design
and detailed cost structure

Land- or reactor-based full-scale bioremediation systems have been suc-
cessfully used to clean up hydrocarbon-contaminated soils and sludges.
More information on the scale-up of bioremediation technologies can be
obtained in the literature (Huesemann 1994; Cutright 1995; Crawford and
Crawford 1996; Loehr and Webster 1996; Von Fahnestock et al. 1998; Alle-
man and Leeson 1999; Stegmann et al. 2001; Singh and Ward 2004).

142 A. Singh et al.

5.3

Process Monitoring and Evaluation

It is important to make sure that system operation and monitoring plans
have been developed for the land treatment operation. Regular monitor-
ing is necessary to ensure optimization of biodegradation rates, to track
constituent concentration reductions, and to monitor vapor emissions, mi-
gration of constituents into soils beneath the landfarm (if unlined), and
groundwater quality. If appropriate, ensure that monitoring to determine
compliance with storm water discharge or air quality permits is also pro-
posed.

1. Molecular composition of a petroleum contaminant can be useful in
estimating the biodegradation potential of the contaminated soil. Gas
chromatography (GC) analysis (Chapt. 3) may identify easily biodegrad-
able compounds such as straight chain alkanes. GC analysis of various
volatile (benzene, toluene, ethyl benzene, and xylenes) and semi- volatile
(polynuclear aromatic hydrocarbons, PAHs) compounds are required
by the regulatory agencies. However, gravimetric determination of oil
and grease or TPH content following Soxhlet extraction can be used to
design and optimize a reactor or land-based treatment process.

2. Since abiotic processes such as dilution, adsorption, and volatilization
can be responsible for hydrocarbon disappearance, criteria other than
simple hydrocarbon disappearance should be used to assess biodegra-
dation by microorganisms. Increase in the number of hydrocarbon-
degrading bacteria as the bioremediation progresses provides evidence
of biodegradation. Formation of colonies on the surface of a solidified
mineral salts medium with silica gel, incubated in vapors of volatile
hydrocarbons (Walker and Coleman 1976), can be used to enumerate
hydrocarbon-degrading bacteria. Bacteria capable of degrading semi-
volatile hydrocarbons (e.g., PAHs) can be enumerated by examining
colonies on agar plates for their ability to visibly alter a layer of pre-
cipitated insoluble hydrocarbon (Bogardt and Hemmingsen 1992). The
modified most probable number (MPN) technique can be used for non-
volatile hydrocarbons either by applying a floating sheen of oil to the
surface of mineral medium or by placing hydrocarbons dissolved in
a solvent in 24- or 96-well microtiter plates (Brown and Braddock 1990;
Steiber et al. 1994; Haines et al. 1996). The presence of hydrocarbon-
utilizing bacteria is detected by the emulsification or dispersion of sheen,
by reduction of added iodonitrotetrazolium violet, or by the appearance
of colored metabolites in the medium (see also Chapt. 13).

5 Feasibility Studies for Microbial Remediation 143

3. Since microbial communities play a significant role in biogeochemi-
cal cycles, it is important to analyze the community structure and its
changes during bioremediation processes (Chaps. 10 and 12). The tem-
poral and spatial changes in bacterial populations and the diversity of
the microbial community during bioremediation can be determined us-
ing sophisticated molecular methods (van Elsas et al. 1998; Widada et al.
2002).

4. Biodegradation potential of a hydrocarbon-contaminated soil can be
estimated by its chemical characterization and the relative biodegrad-
ability of the contaminants. Mono aromatic compounds such as ben-
zene and alkyl benzene and low molecular weight n-alkanes are eas-
ily biodegradable as compared to high molecular weight and highly
branched molecules. While PAHs with four or more rings are consid-
ered recalcitrant, two or three ring PAHs can be degraded by different
microbial species.

5. The volatile constituents present in petroleum-contaminated soils tend
to evaporate during biotreatment, particularly during tilling or plowing
operations in land treatment and aeration of the bioreactors, rather
than being biodegraded by bacteria. For compliance with air quality
regulations, the volatile organic emissions should be estimated based on
initial concentrations of the petroleum constituents present. Depending
upon specific regulations for air emissions, control of VOC emissions
may be required. Control involves capturing vapors and then passing
them through an appropriate treatment process before being vented to
the atmosphere. Control devices range from an erected structure such
as a greenhouse or plastic tunnel to a simple cover such as a plastic sheet
for land treatment and a carbon filter or biofQter for a slurry reactor.

6. Solid-phase microextraction (SPME) has been used to monitor biodegra-
dation of semivolatile hydrocarbons in diesel-fuel-contaminated water
and soil (Eriksson et al. 1998) and of volatile hydrocarbons during bac-
terial growth on crude oil (Van Hamme and Ward 2000). Although the
method requires external calibration with several standard calibration
curves, SPME was proven to be a rapid and accurate method for monitor-
ing volatile and semivolatile hydrocarbons in petroleum biodegradation
systems.

5.4
Bioaugmentation

Bioaugmentation can be denned as the introduction of a large number
of exogenous microorganisms into the environment of a biotreatment sys-

144 A. Singh et al.

tern. Diverse microorganisms, including many species of bacteria and fungi
are known to degrade hydrocarbons. The most prevalent bacterial hy-
drocarbon degraders belong to the genera Pseudomonas, Achromobacter,
Flavobacterium, Rhodococcus, and Acinetobacter. Penicillium, Aspergillus,
Fusarium, and Cladosporium are most frequently isolated hydrocarbon
degrading filamentous fungi. Among the yeasts Candida, Rhodotorula,
Aureobasidium, and Sporobolomyces are the hydrocarbon degraders most
often reported (Van Hamme et al. 2003). Environmental and nutritional
factors influence the presence, survival, or activity of microorganisms in
contaminated soils.

There are at least four different routes that result in the development of
microbes capable of degradation of hydrocarbons at a certain site:

1. The indigenous microflora are exposed to the contaminant long enough
for genetic evolution to create a capacity to degrade the compound(s).

2. The indigenous microflora, adapted to the local conditions, are exposed
to one or more contaminating xenobiotic compounds. The bacteria ac-
quire genes and degradation pathways from bacterial cells immigrating
from elsewhere.

3. The indigenous, well-adapted microflora are maintained ex-situ and
then artificially supplied with the required degradative capacity.

4. A bacterium that is thought to be competitive at the contaminated site is
chosen. This may be a strain that is known to degrade the contaminant
or one that is specifically constructed for this purpose.

Bioaugmentation-related experiments can be conducted in slurry biore-
actors described above. Bio augmentation studies can be carried out either
using mixed cultures or individual pure strains. The effect of initial popula-
tion size on biodegradation of contaminants can be determined by varying
inoculum densities. The inoculum size can be varied from 10 5 to 10 9 CFU/g
of soil in the bioaugmentation studies. The effect of a commercial or selec-
tively developed inoculum on the rate of biodegradation, C0 2 evolution,
time of lag phase after inoculation, and microbial population dynamics
during biodegradation process can be monitored.

5.5

Effect of Surfactants

The biodegradation rate of a contaminant depends on the rate of contam-
inant bioavailability, uptake, and mass transfer. Bioavailability of a con-
taminant in soil is influenced by a number of factors such as desorption,

5 Feasibility Studies for Microbial Remediation 145

diffusion, and dissolution. Use of chemical- or bio-surfactants in contam-
inated soil can help overcome bioavailability problems and accelerate the
biodegradation process.

Biosurfactants, surface-active substances synthesized by living cells,
have the properties of reducing surface tension, enhancing the emulsifi-
cation of hydrocarbons, stabilizing emulsions, and solubilizing hydrocar-
bon contaminants to increase their availability for microbial degradation.
Biosurfactant-producing microbes play an important role in the acceler-
ated bioremediation of hydrocarbon-contaminated sites (Rahman et al.
2003; Shin et al. 2004). The low-molecular-weight biosurfactants (glycol-
ipids, lipopeptides) are more effective than those of high molecular weight
(amphipathic polysaccharides, proteins, lipopolysaccharides, lipoproteins)
in lowering the interfacial and surface tensions (Mulligan 2005).

Some simple laboratory experiments to study biosurfactant production
and application in bioremediation are described here.

5.5.1

Screening of Microbial Cultures for Biosurfactant Production

Different microbial cultures can be screened for biosurfactant production
using the following method:

1 . Prepare a series of 250-mL flasks containing 50 mL of sterile YPG medium
(composition per L: 5 g peptone, 5 g yeast extract, 10 g glucose, pH 7.0)
and incubate on a shaker (200 rpm) at 30 °C after inoculation with indi-
vidual cultures.

2. Add 1% glycerol after 24 h.

3. Measure biomass content, biosurfactant production, surface tension, and
emulsification activity at 12-24 h intervals.

4. For biomass determination, filter the culture broth using GF/C filters,
place the filters at 110 °C for 24 h, and weigh to calculate biomass (dry
mass).

5. Surface-active compounds can be extracted by liquid-liquid extraction
using 10 mL of chloroform:methanol (2:1 ) mixture from 10 mL of the cell-
free culture broth acidified with 1 N HC1 to pH 2. Concentrate the organic
extracts by drying them overnight in a drying chamber at a temperature
around 44 °C, and measure the mass of the biosurfactant.

For purification of the biosurfactant to determine its properties and
application, the culture broth is filtered through a centrifuge filter with
lOkDa molecular weight cut-off at 6,000 g until the minimal amount of

146 A. Singh et al.

retentate is achieved. The retentate is diluted in 50% methanol in order
to dissociate the micelles and filtered at 6000 g again. After collection of
filtrate, methanol is evaporated under vacuum in a rotary evaporator at
65 °C and the aqueous solution of the purified biosurfactant is lyophilized.

Surface tension (mN/m) can be measured using a standard commercial
tensiometer. The emulsification activity can be determined by adding a hy-
drocarbon (xylene, benzene, n-hexane, kerosene, gasoline, diesel fuel, or
crude oil) to the same volume of cell-free culture broth, vortexing for 2 min
and letting stand for 24 h. The emulsification activity is determined as the
percentage of height of the emulsified layer divided by the total height of
the liquid column (Rahman et al. 2003).

A blood agar lysis method can also be used for screening cultures for
their biosurfactant-producing capabilities (Youssef et al. 2004). Culture is
streaked onto blood agar plates and incubated for 48 h at 37 °C. The zones of
clearing around the colonies indicate biosurfactant production. The diam-
eter of the clear zones depends on the concentration of the biosurfactant.

5.5.2

Effect of Biosurfactants

Biosurfactant preparations can be purchased from a commercial chemical
supplier or purified from the culture broth as described above. For different
hydrocarbons, a biosurfactant is added to the cultures to obtain concen-
trations above and below the critical micelle concentration (CMC). The
CMC value is determined by measuring surface tension in different dilu-
tions of a 4 g/L solution of the biosurfactant. The value of CMC, expressed
in mg/L, is obtained from the plot of the surface tension versus the loga-
rithm of the concentration. A rhamnolipid biosurfactant concentration of
50-2,000 mg/L is generally useful in biodegradation studies.

The biodegradation experiment to study effect of biosurfactants can be
conducted in 250 mL Erlenmeyer flasks containing 50 mL of the culture
medium described before. Appropriate controls, such as no-biosurfactant
and abiotic controls, are run along with the flasks containing different
concentrations of the biosurfactant. Cultures are incubated on a shaker for
7-14 days at 30 °C.

5.5.3

Effect of Chemical Surfactants

Properties of chemical surfactants that influence their efficacy include
charge (nonionic, anionic, or cationic), hydrophilic-lipophilic balance
(HLB, a measure of surfactant lipophilicity), and CMC (the concentration

5 Feasibility Studies for Microbial Remediation 147

at which surface tension reaches a minimum and surfactant monomers ag-
gregate into micelles). However, there is always a concern that the surfactant
may get used preferentially as a carbon source instead of the contaminant.
Hence, there is a need to provide a perspective as to when or how surfac-
tants may be exploited in petroleum hydrocarbon degradation processes
to improve rates and extents of degradation.

Typical surfactant concentrations for washing of contaminant soil are
1-2%, whereas the same contaminants may be solubilized in an aqueous
solution at a surfactant concentration of 0.1-0.2%. Non-ionic surfactants
within the HLB range of 11 to 15 can optimally support microbial degra-
dation of hydrophobic contaminants. Nonylphenol ethoxylated surfactants
with HLB 12 and 13 can substantially enhance biodegradation of hydro-
carbons at surfactant concentrations greater than CMC value. Different
groups of nonionic surfactants should be tested at different concentrations
greater than their CMC during feasibility studies in soil microcosms or
slurry reactors.

5.6

Optimization of Environmental Conditions

The procedures described in the previous sections on different technologies
can be used in studies of the factors affecting biodegradation rates and
determining appropriate biotreatment strategy for contaminated soil.

1. The optimum soil pH for hydrocarbon bioremediation in soil ranges
from 6 to 8. Methods for adjusting pH usually include periodic appli-
cation of lime and/or sulfur. The requirement of acid or alkaline solu-
tions/solids for pH control is developed in biotreatability studies and
the frequency of their application is modified during land treatment or
slurry reactor operation as needed. In case of acidic soil (pH < 6), lime
or calcium carbonate may be added to increase the pH to the required
optimum range. For alkaline soil (pH > 8), elemental sulfur, ammonium
sulfate, or aluminum sulfate may be added to lower the pH.

2. Optimum temperature range for microbial degradation is 25 to 35 °C.
Biodegradation rates are expected to slow considerably below 15 °C or
above 40 °C. However, temperature cannot be maintained for land appli-
cation. Land treatment of hydrocarbon-contaminated soils is difficult to
operate in temperate and arid zones. Slurry bioreactors are always more
useful in such places because environmental conditions can be more
precisely maintained and with relative ease.

3. During land treatment, soil microorganisms can only biodegrade petro-
leum hydrocarbons within a limited range of favorable soil moisture

148 A. Singh et al.

conditions. If the soil is too dry, bacterial growth and metabolisms will
be greatly reduced or even inhibited. Alternatively, if the soil is too wet or
flooded, soil aeration will be greatly impaired which, in turn, will result
in anaerobic conditions that are not conducive to hydrocarbon biodegra-
dation. Since the moisture content at field capacity is strongly dependent
on the soil type (clay and high organic matter soils retain comparatively
higher moisture content), it is important to determine the moisture re-
tention profile for each soil to be studied. The optimum moisture content
for stimulating petroleum hydrocarbon biodegradation ranges from 50
to 80% of the moisture content at field capacity. For example, if the soil
moisture at field capacity was determined to be 20 g of water per 100 g of
dry soil, the soil moisture content should be maintained between 10 and
16 g of water per 100 g of dry soil.

4. In order to limit the demand of oxygen by soil bacteria, it is important
not to overload the soil with too high levels of oil contamination during
land application. As outlined below, the optimum contaminant load-
ing level for land treatment is about 5% (by weight) of oil. Maximum
degradation rates are typically observed in the 10-15 cm upper plow
layer if hydrocarbon concentrations are maintained around 5%. Addi-
tion of peroxygen compounds may also help slowly release oxygen into
the soil and thereby enhance the aerobic biodegradation of petroleum
hydrocarbons.

5. There are other processes such as volatilization, leaching, sorption and
photo-oxidation that may cause the removal of certain hydrocarbon
compounds or classes during biotreatment. It has been estimated that
between 15 and 60% of fuel hydrocarbons (diesel, jet fuel, and heat-
ing oil) can be lost during soil bioremediation by land treatment solely
due to evaporation (Salanitro 2001). At room temperature (20 °C), most
hydrocarbons with carbon numbers up to Q 5 or Ci6 readily evaporate
from soil if in free contact with air. Even heavier hydrocarbons (> Ci 6 )
including three- and four-ring PAHs are likely to volatilize in intense
sunshine. These competing loss mechanisms during field or laboratory
bioremediation studies should be measured or estimated either by cal-
culating a complete mass balance or by carrying out proper microbial
control experiments.

5.7

Optimization of Nutritional Factors

For biotreatment of petroleum hydrocarbons, bacteria that are both aerobic
and heterotrophic are the most important in the biodegradation process.

5 Feasibility Studies for Microbial Remediation 149

Since microorganisms require organic and inorganic nutrients such as ni-
trogen, phosphorus, magnesium, calcium, iron, and trace metals to support
cell growth and sustain biodegradation processes, nutrients need to be sup-
plemented during biotreatment in bioreactors or land in order to maintain
active bacterial populations. However, excessive amounts of certain nutri-
ents such as phosphate and sulfate can repress microbial metabolism. Im-
portant nutrient sources for biotreatability or feasibility studies are shown
Table 5.1.

1. Nutrients are added for the growth and maintenance of microorgan-
isms. By providing an appropriate balance of nutrients it is possible to
achieve high level of growth of hydrocarbon-degrading bacteria and thus
accelerated rates of hydrocarbon degradation. The typical non-carbon
elemental composition of major bacterial components is nitrogen 12.5%;
phosphorus 2.5%; potassium 2.5%; sodium 0.8%; sulphur 0.6%; calcium
0.6%; magnesium 0.3%; copper 0.02%; manganese 0.01%, and iron 0.01%
(Rehm 1993). Use of appropriate concentrations and ratios of nutrients
can avoid a situation where growth is limited by depletion of one essential
nutrient while all other nutrients may be present in excess.

2. An oil carbon content of 80% can be assumed for the purpose of cal-
culating C:N or C:P ratio. Although a wide range of C:N and C:P ratios
has been recommended in the literature, an oil:N:P ratio of 100:1:0.2
can be used for the feasibility studies. Thus, for each 100 kg of oil to be
degraded, 1 kg of nitrogen and 0.2 kg of phosphorus can be added as
nutrient fertilizer in the preliminary studies. Optimum C:N ratio can

Table 5.1. Important nutrient sources for biotreatability or feasibility studies

Nutrient source Examples

Defined medium

Nitrogen KN0 3 , NH4NO3, NH 4 C1

Phosphorus KH2PO4, sodium tripolyphosphate (NasPaOio)

Potassium KNO3

Calcium CaCl 2 • 2H2O

Magnesium MgS0 4 • 7H 2

Iron FeCl 3 • 6H 2

Trace metals MnS0 4 • H 2 0, CuS0 4 • 5H 2 0, ZnCl 3 • 4H 2 0, H3BO3, CoCl 2 • 6H 2 0,

Na 2 Mo0 4 • 2H 2

Complex medium

Nitrogen Yeast extract, Peptone, Urea, NPK fertilizer

Phosphorus NPK fertilizer

Magnesium NPK fertilizer

Trace metals Yeast extract, NPK fertilizer

150 A. Singh etal.

determined in microcosms or slurry bioreactors by varying C:N ratio
from 10 to 100.

3. For land treatment, nutrient supply methods usually include periodic
application of solid fertilizers, while tilling to blend soils with the solid
amendments, or applying liquid nutrients using a sprayer. For bioslurry
reactors, a blend of solid nutrients can be added, which is quickly dis-
solved in the medium due to continuous mixing. The composition of
nutrients is developed in lab treatability studies.

4. The inability of microbes to completely mineralize a contaminant and
transform it to other organic compounds means that these organisms
require other substrates to support their growth. The contaminants
are transformed by "co-metabolic" processes, where a second substrate
serves as primary energy or carbon source.

5. Using a naturally selected and acclimated indigenous bacterial culture
originating from the sludge is supplemented with a carefully designed
blend of nutrients containing sources of nitrogen, phosphate, a complex
protein, essential minerals, and a surfactant. The bioslurry reactor sys-
tem can promote growth of a highly active microbial population and
rapid conversion of the petroleum hydrocarbons at the rate of about 1%
petroleum hydrocarbons degraded per day (Ward et al. 2003).

6. Generally high molecular weight PAHs (five-ring) are only biodegraded
in the presence of other hydrocarbons such as lower molecular weight
PAHs or complex hydrocarbon mixtures such as crude oil. If these neces-
sary co-substrates are absent, the co-metabolic biodegradation of higher
molecular weight PAHs cannot proceed.

5.8
Conclusions

Bioremediation is a cost effective and environmentally friendly hydrocar-
bon-contaminated soil remediation technology. The successful bioreme-
diation of contaminated soils depends on numerous environmental pa-
rameters and operational factors, which need to be optimized in order to
achieve maximum treatment benefits. Even under optimal conditions it is
unlikely that all contaminants will be removed from the soil. This incom-
plete biodegradation may be acceptable if the residual hydrocarbons can
be shown to have no significant impact on ecological receptors and do not
pose a risk to groundwater resources.

The effectiveness of bioremediation depends on the success in identi-
fying the rate-limiting factors and optimizing them in the feasibility and

5 Feasibility Studies for Microbial Remediation 151

biotreatability studies. Feasibility studies are essential and may have enor-
mous impact on the cost of the full-scale operation. Depending on the
site, nature of contamination, and type of soil, various methods for fea-
sibility studies are currently available. These methods can be modified to
accommodate the lab facilities and equipment availability. Sometimes it
is difficult to extrapolate the results directly from the laboratory to the
field. Nevertheless, successful bench- or pilot-scale test results are mostly
useful in designing the full-scale bioprocessing system for bioremediation
of hydrocarbon-contaminated soil.

One of the main barriers to greater effective adoption of bioremediation
technologies is the perception that the processes are very project-specific,
requiring much customization. There is a need to develop more robust and
technologically versatile processes that do not require significant research
and development for each project (Ward 2004). Government funding initia-
tives and the market favor use of more controlled and accelerated processes
and that are typically more predictable (Srinivasan 2003). Hughes et al.
(2000) have provided guidance with regard to selection of bioremediation
configuration for treatment of different classes of chemicals.

The choice of technology configuration based on application of such
principles should precede the design of a feasibility study, and the latter
then used to confirm and validate the effectiveness of the technology.

References

Alleman BC, Leeson A (eds) (1999) Bioreactor and ex situ biological treatment technologies.
Battelle Press, Columbus, pp 31-36

Bogardt AH, Hemmingsen BB (1992) Enumeration of phenanthrene-degrading bacteria by
an overlayer technique and its use in evaluation of petroleum contaminated sites. Appl
Environ Microbiol 58:2579-2582

Brown EJ, Braddock JF (1990) Sheen screen, a miniaturized most-probable number method
for enumeration of oil-degrading microorganisms. Appl Environ Microbiol 56:3895-
3896

Crawford RL, Crawford DL (eds) (1996) Bioremediation: Principles and Applications. Cam-
bridge Univ Press, Cambridge, UK

Cutright (1995) A feasible approach to the bioremediation of contaminated soil: from lab
scale to field test. Fresenius Environ Bull 4:67-73

DECHEMA (1992) Laboratory methods for the evaluation of biological soil clean-up pro-
cesses. Interdisciplinary Working Group, Environmental Biotechnology- Soil. Deutsche
Gesellschaft fuer Chemisches Apparatewesen, Chemische Technik und Biotechnologie,
Germany

Eriksson M, Swartling A, Dalhammer G (1998) Biological degradation of diesel fuel in water
and soil monitored with solid-phase micro-extraction and GC-MS. Appl Microbiol
Biotechnol 50:129-134

Grommen R, Verstraete W (2002) Environmental biotechnology: the ongoing quest.
J Biotechnol 98:113-123

152 A. Singh et al.

Haines JR, Wrenn BA, Holder EL, Strohmeier KL, Herrington RT, Venosa AD ( 1 996) Measure-
ment of hydrocarbon-degrading microbial populations by a 96-well plate most-probable
number procedure. J Ind Microbiol 16:36-41

Huesemann MH (1994a) Guidelines for the development of effective statistical soil sam-
pling strategies for environmental applications. In: Calabrese EJ, Kostecki PT (eds)
Hydrocarbon-contaminated soils and groundwater. Lewis, Boca Raton, pp 47-96

Huesemann MH (1994b) Guidelines for land-treating petroleum hydrocarbon contaminated
soils. J Soil Contamin 3:299-318

Hughes JB, Neale CN, Ward CH (2002) Bioremediation. Encyclopedia of Microbiology, 2 nd
edn, Academic Press, New York, pp 587-610

Iwamoto T, Nasu M (2001) Current bioremediation practice and perspective. J Biosci Bioeng
92:1-8

Jurgensen KS, Puustinenl J, Suortti A-M (2000) Bioremediation of petroleum hydrocarbon-
contaminated soil by composting in biopiles. Environ Pollut 107:245-254

Lang S, Wullbrandt D (1999) Rhamnose lipids - biosynthesis, microbial production and
application potential. Appl Microbiol Biotechnol 51:22-32

Liu D (1989) Biodegradation of pentachlorophenol and its commercial formulation. Bull
Environ Contam Toxicol 4: 115-127

Loehr RC, Webster MT (1996) Performance of long-term, field-scale bioremediation pro-
cesses. J Haz Mat 50:105-128

Mandelbaum RT, Allan DL, Wackett LP (1995) Isolation and characterization of a Pseu-
domonas sp that mineralizes the s-triazine herbicide atrazine. Appl Environ Microbiol
61: 1451-1457

McFarland MJ, Sims RC, Blackburn JW (1991) Use of treatability studies in developing
remediation strategies for contaminated soils. In: Sayler GS, Fox R, Blackburn JW (eds)
Environmental biotechnology for waste treatment. Plenum, New York, pp 163-174

Mulligan CN (2005) Environmental applications for biosurfactants. Environ Pollut 133:
183-198

Noha H, Youssef NH, Duncana KE, Naglea DP, Savagea KN, Knappb RM, Mclnerneya MJ
(2004) Comparison of methods to detect biosurfactant production by diverse microor-
ganisms. J Microbiol Meth 56:339- 347

OECD (1981) Inherent biodegradability in soil. In: OECD Guidelines for testing of chemicals,
Part 304A, Organization of Economic Co-operation and Development, Paris, France,

pp i-ii

Pfarl C, Ditzelmuller M, Loidle M, Streichsbier F (1990) Microbial degradation of xenobiotic

compounds. FEMS Microbiol Ecol 73: 255-262
Rahman KSM, Rahman TJ, Lakshamanaperumalsamy P, Marchant R, Banat IM (2003) The

potential of bacterial isolates for emulsification with a range of hydrocarbons. Acta

Biotechnol 23: 335-345
Rehm HJ (1993) Biotechnology Vol 3, VCH, Weinheim, pp 131

Salanitro JP (2001) Bioremediation of petroleum hydrocarbons in soil. Adv Agron 72:53-105
Shin K-H, Kim K-W, Seagren EA (2004) Combined effects of pH and biosurfactant addition

on solubilization and biodegradation of phenanthrene. Appl Microbiol Biotechnol 65:

336-343
Shulga A, Karpenko E, Vildanova-Martsishin R, Turovsky A, Soltys M (1999) Biosurfactant

enhanced remediation of oil-contaminated environments. Adsorpt Sci Technol 18: 171-

176.
Singh A, Mullin B, Ward OP (2001) Reactor-based process for the biological treatment

of petroleum wastes, PN # 200. Proceedings, Middle East Petrotech 2001 Conference,

Bahrain, pp 1-13

5 Feasibility Studies for Microbial Remediation 153

Singh A, Ward OP (eds) (2004) Applied bioremediation and phytoremediation. Soil Biology

Series, Volume 1, Springer- Verlag, Germany
Skladany GJ, Baker KH (1994) Laboratory biotreatability studies. In: Baker KH and Herson

DS (eds) Bioremediation, McGraw-Hill, New York, pp 97-172
Srinivasan U (2003) US Bioremediation Markets. Frost and Sullivan Research Report No

7857. Palo Alto California, pp 1-300
Stegmann R, Brunner G, Calmano W, Matz G (2001) Treatment of contaminated soil: fun-
damentals, analysis, applications. Springer, Berlin
Steiber M, Haesler F, Werner P, Frimmel FH (1994) A rapid screening method for microor-
ganisms degrading polycyclic aromatic hydrocarbons in microplates. Appl Microbiol

Biotechnol 40: 753-755
USEPA (1991) Guide for conducting treatability studies under CERCLA: aerobic biodegrada-

tion remedy screening - interim guidance, USEPA publication no. EPA/540/2-91 /013A
USEPA (1996) Engineering bulletin: Composting, EPA/540/S-96/502
USEPA (1998) An analysis of composting as an environmental remediation technology,

EPA530-R-98-008
van Elsas JD, Duarte GF, Rosado AS, Smalla K (1998) Microbiological and molecular bi-
ological methods for monitoring microbial inoculants and their effects in the soil

environment. J Microbiol Meth 32:133-154
Van Gestela K, Mergaertb J, Swings J, Coosemansa J, Ryckeboer J (2003) Bioremediation of

diesel oil- contaminated soil by composting with biowaste. Environ Poll 125: 361-368
Van Hamme J, Singh A, Ward OP (2003) Recent advances in petroleum microbiology.

Microbiol Mol Biol Rev 67:503-549
Van Hamme JD, Ward OP (2000) Development of a method for the application of solid-phase

microextraction to monitor biodegradation of volatile hydrocarbons during bacterial

growth on crude oil. J Ind Microbiol Biotechnol 25:155-162
Vecchili GI, Delpanno MT, Painceira MT (1990) Use of selected autochthonus soil bacteria

to enhance degradation of hydrocarbons in soils. Environ Pollut 67:249-258
Venosa AD, Haines JR, Nisasamaneepong W, Gouind R, Pradhan S, Siddique (1992) Efficacy

of commercial products in enhancing oil biodegradation in closed laboratory reactor.

J Ind Microbiol 10:13-23
Von Fahnestock FM, Wickramanayake GB, Kratzke RJ, Major WR (1998) Biopile design,

operation, and maintenance handbook for treating hydrocarbon-contaminated soils,

Battelle Press, Columbus, Ohio
Walker JD, Colwell RR (1976) Enumeration of petroleum-degrading microorganisms. Appl

Environ Microbiol 31:198-207
Ward OP (2004) The industrial sustainability of bioremediation processes. J Ind Microbiol

Biotechnol 31:1-4
Ward OP, Singh A, Van Hamme JD (2003) Accelerated biodegradation of petroleum waste.

J Ind Microbiol Biotechnol 30: 260-270
Widada J, Nojiri H, Omori T (2002) Recent developments in molecular techniques for

identification and monitoring of xenobiotic-degrading bacteria and their catabolic genes

in bioremediation. Appl Microbiol Biotechnol 60:45-59
Wolf DC, Dao TH, Scott HD, Lavy TL (1989) Influence of sterilization method on selected

soil microbiological, physical and chemical properties. J Environ Qual 18:39-44
Youssef NH, Duncan KE, Nagle DP, Savage KN, Roy M, Knapp RM, Mclnerney MJ (2004)

Comparison of methods to detect biosurfactant production by diverse microorganisms.

J Microbiol Meth 56:339- 347

Feasibility Studies forMicrobial Remediation
of Metal-Contaminated Soil

Franz Schinner, Thomas Klauser

■ Introduction

Objectives. Heavy metal contamination of soil is widespread due to metal
processing industries, tannery, combustion of wood, coal and mineral oil,
traffic, and plant protection. The toxic effects of heavy metals result mainly
from the interaction of metals with proteins (enzymes) and inhibition of
metabolic processes. In contrast to organic pollutants, metals are not min-
eralized by microorganisms but can be oxidized or reduced, transformed
to different redox stages, or complexed by organic metabolites.

Besides excavation and deposition, a conventional treatment for decon-
tamination of metal-polluted soil is extraction using mineral acids. The
disadvantages of such a treatment are the destruction of soil, high costs
of acids, and low acceptance. Alternative remediation strategies to reduce
bioavailability of metals are: (1) immobilization with repeated addition of
substances such as carbonate, phosphate, apatite, zeolite, clay minerals,
peat, or humus: and (2) bioleaching with heterotrophic microorganisms,
preferably fungi. The latter method represents a sustainable remediation
treatment of metal-polluted soils (Wasay et al. 1998, Schinner et al. 2000).
Autotrophic bacteria as known from ore leaching cannot be recommended
for this treatment due to the high buffer capacity of soil compared to ore.
The most limiting factor of heterotrophic leaching is the availability of
inexpensive carbohydrates, such as molasses or the refuse from process-
ing sugar, fruits, or white wine. Heterotrophic leaching can be done off
site, on site, and in situ, and is an alternative treatment for the decon-
tamination of metal- containing filter dusts (Schinner and Burgstaller 1989;
Burgstaller et al. 1992) and industrial sludges.

Principle. Metal-contaminated soil is supplemented with carbohydrates to
increase the excretion of organic acids by autochthonous (procedure A)
or inoculated (procedure B) fungi. Organic acids mobilize metals that are
eluted from soil by percolation with water.

Theory. In natural ecosystems fungi play an important role in the mobiliza-
tion of nutrients and trace elements from soils. Autochthonous soil fungi

Franz Schinner, Thomas Klauser: Institute of Microbiology, Leopold Franzens University,
Technikerstrafte 25, 6020 Innsbruck, Austria, E-mail: franz.schinner@uibk.ac.at

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

156 R Schinner, T. Klauser

but also inoculated single strains or mixed populations increase organic
acid production after the addition of carbohydrates. Fungi mobilize met-
als from mineral soils by excretion of acidifying protons (acidolysis), by
excretion of organic acids forming cyclic organometal complexes (complex-
olysis), or by redoxolysis with organic acids. Some of the Deuteromycetes,
especially members of the genera Aspergillus and Penicillium, produce
various organic acids, such as citric, oxalic, tartaric, gluconic, succinic,
formic, and amino acids (Burgstaller and Schinner 1993; Gadd 1999). The
metal- containing eluate obtained after percolation of soil with water can
be regenerated by conventional metal recovery, such as precipitation, ion
exchange, or biosorption.

■ Equipment

• Percolator: a filtration unit (funnel and suction flask; e.g., Nalgene,
500 mL; Nalge Nunc, Naperville, IL, USA), with gauze mat (e.g., Schlei-
cher & Schull TG 100; Schleicher & Schull, Dassel, Germany) instead of
filter

• Vacuum pump

• Multichannel peristaltic pump

• Timer (for circuit switching peristaltic pump and vacuum pump)

• Atomic absorption spectrometry (AAS) or inductively- coupled plasma-
atomic emission spectrometry (ICP-AES) for metal analyses

• High performance liquid chromatography (HPLC) for analyses of or-
ganic acids (optional): e.g., column AMINEX-HPX 87H (Bio-Rad, Her-
cules, CA, USA); flow rate 0.6 mL/min; column temperature 41 °C, wave-
length 210 nm, eluant 4 mM H 2 S0 4 (Womersley et al. 1985)

■ Materials and Reagents

• Quartz sand

• Sawdust

• Grain (e.g., barley, wheat, rye)

• Organic-acid-producing fungi, e.g., Aspergillus sp. or Penicillium sp.

• Complex substrate: e.g., dried refuse of sugar production, fruit, or white
wine processing

• Molasses solution: 150 g (< 75% dry mass)/L of water

6 Feasibility Studies for Microbial Remediation of Metal- Contaminated Soil 157

• Sterile KC1 solution: 10 g/L

• Deionized water

■ Sample Preparation

Use air-dried, sieved (< 5 mm) soil.

■ Procedure

Procedure A: Metal Leaching with Autochthonous Microorganisms

1. Assemble percolation units, pour about 10 mm quartz sand onto the
gauze mat. Connect tubes to flasks containing molasses and water, and
install pumps and timer. Prepare 3-4 replicates.

2. Weigh 150 g of sieved soil, 15 g of sawdust, and 15 g of complex substrate
into a beaker, mix, and pour it into the percolator.

3. Add 60 mL of molasses solution, 50 mL of KC1 solution and 70 mL of
water to rewet substrate.

4. Start percolation with molasses solution, use the peristaltic pump at
a flow rate of 20mL/h for 15 h every day. Repeat this percolation on
days 2, 3, and 6.

5. On days 3, 5, and 7, use water instead of molasses solution as percolation
fluid.

6. For the suction of the fluid substrate and water and for additional aera-
tion, start the vacuum pump every 90 min for 25 min without interrupt-
ing the percolation (from the beginning to the end of experiment).

7. To measure the leaching efficiency, take daily samples for metal detection.
After 15 h of percolation with molasses or water add 120 mL of water onto
the soil, suck off each percolator for 5 min, centrifuge 5 mL of eluate for
15 min at 10,000 g. The supernatant is used for quantification of metals
and organic acids. Repeat this procedure every day in the same way.

8. After 3 days add 15 g of complex substrate and mix it into the topsoil of
each percolator.

Procedure B: Metal Leaching with Bioaugmentation

1. Prepare the inoculum for bioaugmentation as follows:

1.1. Sterilize 30 g of grain together with 30 mL of water in a 500-mL
Erlenmeyer flask for 35 min at 121 °C.

158 R Schinner, T. Klauser

1 .2. Inoculate the substrate with organic-acid-producing fungi. Use 5 mL
of spore suspension or collect spores from a stock culture to which
sterile ringer solution is added to wash away spores.

1.3. Incubate the inoculated grain 7-10 days at 25 °C to produce spores.

1.4. Add 200-300 mL of sterile KC1 solution and shake thoroughly.
Transfer the spore-containing suspension to a sterile flask. About
10 8 spores/mL are required.

1.5. Store spore suspensions for the preparation of further inoculates at
-20 °C.

2. Perform the procedure for metal leaching as described for procedure A,
except for step 3: To rewet and inoculate soil and substrate, add 70 mL
of spore suspension instead of 70 mL of water to 60 mL of molasses
solution and 50 mL of KC1 solution. After 3 days (step 8) add 20 mL of
spore suspension together with 15 g complex substrate and mix it into
the topsoil of each percolator.

Monitoring Metal Remediation

Determine the pH value (Chapt. 2), the heavy metal content (using AAS
or ICP; Chapt. 3), and eventually organic acids in the eluate using HPLC
(Womersley et al. 1985). Additionally, soil enzyme activity (Chapt. 17),
microbial biomass (Chapt. 14), or fungal biomass of soil (Rossner 1996)
can be analyzed.

■ Calculation

Calculate the amount of leached metals from the sum of metal contents
measured each day, considering soil dry mass and extraction volume.

■ Notes and Points to Watch

• Metal-leaching experiments with soil percolation attain decontamina-
tion rates of 40-80%.

• Inoculation with organic-acid-producing strains can result in a higher
leaching efficiency.

• Mixed cultures of fungi are more efficient than single strains.

• The clay fraction of soils contains more metals than the silt and sand
fraction.

• The leaching efficiency of soil microorganisms depends strongly on the
soil buffer capacity. Acidification of soil (pH < 6) may be necessary.

6 Feasibility Studies for Microbial Remediation of Metal- Contaminated Soil 159

• The leaching efficiency of soil microorganisms depends on the quality
and quantity of soil organic matter, and on the aeration of soil.

• The duration of percolation and the addition of molasses and water must
be optimized for each soil.

• In situ bioleaching of metal-contaminated soils needs effective drainage
systems.

• Depending on the soil material needed for monitoring analyses, a larger
volume of soil (for example, 300 g soil in a 1 L percolator) can be used.

References

Burgstaller W, Schinner F (1993) Leaching of metals with fungi. J Biotechnol 27:91-116
Burgstaller W, Strafter H, Wobking H, Schinner F (1992) Solubilization of zinc from filter

dust with Penicillium simplicissimum: Bioreactor leaching and stoichiometry. Environ

Sci Technol 26:340-346
Gadd G.M (1999) Fungal Production of citric and oxalic acid: Importance in metal speciation,

physiology and biogeochemical processes. Adv Microb Phys 41:47-92
Rossner H (1996) Fungal biomass by ergosterol content. In: Schinner F, Kandeler E, Ohlin-

ger R, Margesin R (eds) Methods in soil biology. Springer, Berlin Heidelberg New York,

pp 49-51
Schinner F, Burgstaller W (1989) Extraction of zinc from industrial waste by a Penicillium

sp. Appl Environ Microbiol 55:1 153-1 156
Schinner F, Huber W, Atzwanger M, Klauser T (2000) Metodo per il trattamento di suoli

inquinati da composti da metalli pesanti (Method for the treatment of soils polluted

by heavy metal compounds and related plant of treatment). Patents: IT/01. 12.00/IT

RM000631, IT/01. 12.00/IT RM000632
Wasay S.A, Barrington S.F, Tokunaga S (1998) Using Aspergillus niger to bioremediate soil

contaminated by heavy metals. Bioremediation J 2:183-190
Womersley C, Drinkwater L, Crowe JH (1985) Separation of tricarboxylic acid cycle acids

and other related organic acids in insect haemolymph by high-performance liquid

chromatography. J Chromatogr 318:112-116

7 Feasibility Studies for Phytoremediation
' of Metal-Contaminated Soil

Aleksandra Sas-Nowosielska, Rafal Kucharski,
Eugeniusz Malkowski

7.1
Introduction

Phytoremediation, which is the use of herbaceous plants and trees to stabi-
lize, recover, or volatilize pollutants in contaminated soil, is considered an
emerging new technology. The application of phytoremediation is said to
be environmentally friendly, relatively low in cost, and high in public accep-
tance. However, there are still a number of limitations that affect its imple-
mentation on a large scale. The most considerable limitations are: narrow
range of contaminant concentrations within which the method can be ap-
plied (potential of plant toxicity), dependence on weather, time-dependent
growing season, and requirement for management of by-products. Until
recently, the most commonly applied phytoremediation methods have been
phytoextraction and phytostabilization, particularly for soils polluted with
heavy metals.

Phytoremediation is more a biological than a technical approach, and
it is difficult to create a definitive protocol that could be applied to any
polluted site. The limiting factors differ from site to site, and therefore each
project protocol, must be customized to site-specific conditions.

7.2
Phytoextraction

Phytoextraction is a biological method that utilizes properties of specific
species of plants to take up and accumulate pollutants from soil. Certain
species, called hyperaccumulators, may accumulate metals up to several
percent of their dry mass (Brooks 1998; McGrath et al. 2000). Unfortu-
nately, the practical use of these plants for phytoextraction is limited due

Aleksandra Sas-Nowosielska, Rafal Kucharski: Land Management Department, Insti-
tute for Ecology of Industrial Areas, Kossutha 6 St, 40-833 Katowice, Poland, E-mail:
sas@ietu.katowice.pl

Eugeniusz Malkowski: Department of Plant Physiology, Faculty of Biology and Environ-
mental Protection, University of Silesia, Jagiellonska 28 St, 40-032 Katowice, Poland

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

162 A. Sas-Nowosielska et al.

to sparse production of biomass and problems with mechanical harvest-
ing. Nevertheless, genetic research is being conducted to increase biomass
production.

A compromise to the problem of low accumulation properties is to use
plant species with extensive biomass production to compensate for the
lower metal accumulation rates. Such plants remove certain amounts of
metals, but the process is very slow. The addition of chelators to con-
taminated soil enhances metal uptake, an approach known as "induced
phy to extraction" (Salt et al. 1998). Depending on local climate and chem-
istry of pollutants being removed, the most commonly used species for
heavy-metal extraction are Brassica and Helianthus.

A schematic diagram of the processes in an induced phytoextraction
project aimed at cleaning up soils moderately contaminated with lead,
cadmium, and/or zinc is presented in Fig. 7.1. The description contains
step-by-step procedures necessary to perform a phytoextraction project
including theory, legal considerations, technical aspects, as well as logistic
issues and equipment.

7.2.1
Treatability Study

Site Characterization

Site characterization includes the following information:

• Site contaminants (targets: Pb, Zn, and/or Cd)

• Existing vegetation (indicating potential for plant growth)

• Proximity to water (for irrigation)

• Proximity to electrical supply

• Site accessibility for vehicles and farm equipment

• Field observations

• Historical site activities

• Summary of regional hydrology/geology

The purpose of a treatability study is to identify optimal conditions for
metal uptake into the aboveground portion of the plants and to determine
if the soil to be treated will support plant growth. The further objectives
are to evaluate and select the appropriate plant species and soil amend-
ments, and to optimize plant growth for maximal removal of metals from
soils. Treatability studies include short-term investigations for evaluating
the growth and metal uptake potential of selected plant species under con-
trolled conditions in a growth chamber or greenhouse.

7 Feasibility Studies for Phytoremediation of Metal-Contaminated Soil

163

CONTAMINATED SITE
CHARACTERIZATION

TREATABILITY STUDY

STREAMLINE TEST

SITE
PREPARATION

PLANTING

BIOMASS
PRODUCTION

HARVESTING OF
METAL-CONTAINING CROP

SAFE CROP
DISPOSAL

ECONOMICAL
CONSIDERATIONS

RISK ASSESSMENT

LEGAL CONSIDERATIONS

PLANT CARE
FERTILIZING

METAL - MOBILIZING
SOIL AMENDMENTS

CROP
PRE-TREATMENT

Fig. 7.1. Induced phytoextraction process

164 A. Sas-Nowosielska et al.

A bridge between routine treatability studies and full-scale field appli-
cations is the Streamline Test (Sas-Nowosielska et al. 2001), which in com-
bination with a routine laboratory study allows for a rapid and inexpensive
assessment of soil features across the entire site to be treated.

Performance of Treatability Study

1. Soil from the site being investigated should be collected from the top
0-25 cm depth horizon and be well homogenized and sieved to pass
a 4 mm sieve. The soil is placed inside 400-cm 3 plastic pots filled previ-
ously with drainage of approx. 100 g of clean pea/river gravel (2-8 mm
diameter) placed in the bottom. Each pot should be filled with 300 g of
the sieved soil and watered with 100 mL of distilled water prior to plant-
ing the seeds. The seeds should be placed on the surface of the soil in
a circular pattern, covered with a thin layer of soil and moistened with
an additional 40 mL of water.

2. Pots are then placed inside the growth chamber or greenhouse on sep-
arate plastic saucers in order to prevent leaching of metals and to avoid
cross contamination. The seeds should be kept wet during germination
to avoid additional compaction of the soil surface and exposure of seeds.

3. Plants should be adequately fertilized 10 days after germination using
commercial mixtures, and 14-16 days after germination the seedlings
should be thinned as needed.

4. Application of soil amendments should be completed approx. 1 week
prior to harvest. Amendments should be administered in a single dose,
or in three doses if necessary.

Five replicates of each treatment are recommended.

Sampling and Analytical Procedures

An accurate chemical analysis of soil and plants by an accredited laboratory
is a key requirement for conducting a successful treatability study. The
results should be reliable, as they will indicate the effectiveness of the
process and what changes maybe needed to enhance the phytoremediation
process. For most of the sampling and analytical activities described in this
chapter, ISO Standards are recommended (see References and Chapts. 1-3).
Samples should be collected to determine the concentration (spatial
variation) of the target metals at the site, initially and during the phytore-
mediation activities. The actual number and location of samples should be
based on the final layout of the field. Samples collected should be extracted
and analyzed following the ISO Standards (see References).

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 165

Plant material should be washed with tap water in an ultrasonic washer
to remove soil particles and then dried at 70 °C. Approximately 1 g of dried
ground material should be wet-ashed using concentrated nitric acid in
a microwave system. Concentrations of metals should be analyzed by flame
atomic absorption spectrophotometer (FAAS) or by inductively coupled
plasma spectroscopy (ICP-AES).

A chain of custody should be maintained during sampling activities and
Quality Assurance/Quality Control procedures are to be followed. Analyt-
ical work in phytoremediation projects focuses on soil and plant analyses,
preceded by the sampling process. The related regulations are listed below.
In the case of soil, the following information on the investigated material
is required:

• Soil texture (hydrometric method)

• Soil pH in 1 N KC1, H 2 0, and 0.01 M CaCl 2 ; soil-to-solution ratio of 1:2.5
(Chapt. 2)

• Soil electroconductivity; soil to solution ratio of 1:2.5

• Soil organic matter content by loss on ignition (Chapt. 2; Houba et al.
1995)

• Cation exchange capacity (CEC), according to ISO 13536 (1995)

• Content of NO" and NH+ (Chapt. 2; Houba et al. 1995)

• P content in water extract and in an ammonium lactate-acetic acid extract
(Houba etal. 1995)

• K content soluble in 8 M KC1 (Houba et al. 1995)

• Amorphous Al and Fe content, using an ammonium oxalate -oxalic acid
extract method (Houba et al. 1995), and obtaining the concentration of
Al and Fe with ICP analysis (Houba et al. 1995)

• Metal extraction and determination (optional):

- Total heavy-metal and other major cation concentrations are deter-
mined after extraction with aqua regia. Soil should be ground to pass
a 0.25 mm sieve; concentrations of metals is analyzed by FAAS or
ICP-AES (Chapt. 3).

- Bioavailable fraction: 5 g of air-dried soil ground to < 0.25 mm is
extracted with 50 mL of 0.01 M CaCl 2 for 5 h and the concentration of
metals is analyzed in the extract using FAAS or ICP-AES.

- Potentially available metal fraction: 4 g of air-dried soil ground to
pass a < 2-mm sieve are extracted with 40 mL of 0.43 N HNO3 for 4 h.

166 A. Sas-Nowosielska et al.

The concentration of metals is analyzed in the extract using FA AS or
ICP-AES.

- Exchangeable cations: extraction is according to ISO 13536 (1995) as
for CEC; concentration is measured by AAS.

Streamline Test

Site characterization and treatability studies currently are conducted se-
quentially, prior to the initiation of full-scale planting. The purpose of
these activities is to describe the nature and extent of contamination at
the target site, and to determine if, and under what conditions, proposed
plant species will extract the target contaminants. The present approach is
time consuming, expensive, and may not lead to a successful scale-up for
field scale application of phytoextraction. The traditional treatability study
is conducted in greenhouse conditions with controlled air temperature,
light, water regime, and homogenized soil. These carefully controlled con-
ditions often do not mimic real world conditions. A streamline test (ST)
is an attempt to combine the treatability study and site characterization
into an integrated single effort (Sas-Nowosielska et al. 2001). The concept
of the ST was based on a geostatistical assumption that an adequately dis-
tributed number of soil samples may describe the distribution of metals
across an investigated site. The variability of lead and cadmium contents
in soil was estimated in previous field scale phytoextraction experiments
(Kucharski et al. 1998). Based on these findings, it was assumed that two
crossed strips covering approx. 20% of the total site surface would be suffi-
cient to represent the entire area for site characterization purposes. Topsoil
samples were taken outside and inside the strips, and analyzed for con-
tents of metal. Comparison of average concentrations of lead, cadmium,
and zinc in the soil inside and outside showed no significant differences.
It was concluded that the ST better reflects the "real world" conditions as
compared to the usual treatability study. The ST provides an early indica-
tion/screening of the suitability of the site for the application of phytore-
mediation.

7.2.2

Full-Scale Application

Seedbed Preparation and Plant Protection

Plant protection consists of applying herbicides for weed control and in-
secticides to combat herbivore insects. The principle of application should
follow the rules of good agriculture practice. Once the fertilizer and insec-
ticides have been applied, the seedbed will be prepared for planting.

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 167

The site preparation activities in general require the site to be cleared,
cleaned and the soil developed into a condition that will allow planting.
The following are examples of the kinds of obstacles that should be taken
into consideration:

- Quantity and extent of surface debris

- Depth to water table

- Potential for flooding from off the site

- Location of depressions in the soil that will collect water and drown
plants

- Location of a reliable water source for irrigation

Fertilization

Most plant species have varying nutritional needs. Fertilization protocols
should be prepared individually after a soil analysis indicating the required
nutrients for the species used. An example of this was demonstrated when
Brassica was used to clean lead-contaminated soil. Brassica sp. have lower
phosphorus and potassium requirements than other species, but require
abundant nitrogen and supplemental sulfur to promote rapid vegetative
growth. Thus, in accord with the findings of the soil fertility analysis, the
site had to be fertilized with nitrogen, phosphorus, potassium, and sulfur.
Generally, other nutrients are not found to be deficient. Fertilizer type,
placement, and quality play an important role in the success of the crop
development. For example, Brassica sp. are very sensitive to salt damage
from fertilizer placed too close to the seed or with the seed.

Irrigation

Irrigation is used to achieve maximum plant growth pertaining to soil
moisture. The objective is to maintain the recommended soil moisture
levels for each individual crop (depending on plant species and/or cultivar)
during the project's first and second crops. The soil moisture has to be
kept at the optimum level. According to local conditions, various irrigation
systems can be used (dripping, overhead sprinkling, wand-style spraying).
The initial irrigation after planting should wet the soil profile to a depth
of 15 cm. Care should be taken to not apply too much water. Brassica sp.
do not respond well to standing water. The soil should be kept damp but
not saturated until the seedlings emerge. This may require irrigation every
day for sandy soils and every 5-7 days for heavy soil types. The site should
be checked daily to determine if the plants need irrigation. Timing of
irrigations will depend on many variables, such as the size of the plants,

168 A. Sas-Nowosielska et al.

rainfall, temperature, soil type, and the rate of evapotranspiration, to name
a few. A simple tool that can be used to aid a project's irrigation needs is
a tensiometer.

Soil Amendments

• For Brassica sp.: K 3 EDTA (2.5 mmol/kg of soil; stock solution concentra-
tion 50%) and acetic acid (5 mmol/kg of soil; stock solution concentration
80%).

• For Helianthus sp.: K 3 EDTA (5 mmol/kg of soil; stock solution concentra-
tion 50%) and acetic acid (5 mmol/kg of soil; stock solution concentration
80%).

Doses of amendments should be calculated individually for each kind
of soil and applied as a diluted solution through the irrigation system or
tractor driven sprayer.

Species Used for Phytoextraction

• Brassica juncea, the species commonly used for lead phytoextraction,
grows well on fertile, well-drained soils. Successful B. juncea establish-
ment requires a fine, firm seedbed that is free of weeds and rubble. All
vegetation that will compete with the phytoremediation crop should be
removed. The objective is to produce a seedbed that will give the newly
seeded crop maximum opportunity to germinate and grow. The addi-
tion of high quality clean organic matter should be made only when it
is essential. B. juncea has a low tolerance for high salinity and poorly
aerated soils. A well-drained soil provides optimum conditions for rapid
germination and uniform emergence. The seeding rate for B. juncea is
15 kg/ha, and the ideal plant population is 1 10-160 plants/m 2 , which pro-
duces 1.5-3.0 g seeds/m 2 . Planting depth will depend on soil moisture.
The seeds must have good contact with moist soil to achieve maximum
germination and emergence. Ideal planting depth is 1-2 cm. Under dry
sandy conditions the depth may have to be adjusted to 2.5 cm, but plant-
ing deeper than 2.5 cm can result in poor emergence and reduced plant
population.

• Other plant species, such as sunflower, can be used for a phytoextraction
process. This plant can produce a high amount of biomass (60 t/ha), but
heavy-metal accumulation is rather low (about 100 mg Pb/kg dry soil).

• Some plants, termed "hyperaccumulators," take up toxic elements in sub-
stantial amounts, resulting in concentrations in aboveground biomass
over 100 times of those observed in conventional plants. It is econom-
ically hard to grow plants hyperaccumulating toxic metals because of

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 169

their very low biomass production and difficulty of harvesting (Blaylock
etal. 1997).

Harvesting

All aboveground biomass should be removed at harvest at the right time
of plant maturity, after adding the chelating amendment. Application of
the soil amendment should be made in a manner so that as little of the
amendment as possible contacts the plant. This can be done with a hand-
held application wand that directs the liquid amendment at the soil below
the plants leaves, or by using an automatic dispenser (Kucharski et al. 2000).

The physical removal of the plants can be started 7-10 days after the
application of the amendment. The plants should be cut as close to the
ground as possible. This can be accomplished with a fodder harvester,
which cuts plants into 3-5 cm pieces loaded onto the adjacent trailer.

After harvest, the soil must be retilled and prepared for the next crop's
planting. In general, the techniques for the first crop are used for the subse-
quent crops, but less fertilizer should be used for them. The recommended
rates must be determined.

Crop Disposal

Important steps after harvest are reduction of the crop volume and re-
moval of excess water. These will improve technical parameters of harvested
biomass in terms of further processing and reduce transport costs to the
treatment or disposal site. Volume reduction of contaminated plant mate-
rial can be achieved by composting, compaction, or pyrolysis processes.

Composting and compaction should be considered as pre-treatment
steps, since a large volume of contaminated biomass will still exist after
both processes. Total dry-mass loss of contaminated plant biomass is an
advantage of composting as a pretreatment step. It will reduce costs of
transportation to a hazardous waste disposal facility and of deposition,
or of transportation to other facilities where final crop disposal is to take
place. Compaction does not result in total dry mass loss of plant biomass
but works faster than composting. Pyrolysis is also considered a pretreat-
ment step, since metal-contaminated material (coke breeze) is one of the
end-products. Significant more volume/mass reduction of contaminated
plant biomass is observed than from composting or compaction. More-
over, pyrolytic gas is recovered in the process (Sas-Nowosielska et al. 2004).

For final disposal, incineration of contaminated plant material in non-
ferrous or cement rotary kilns or in a municipal waste incineration plant
is considered the most promising method because it significantly reduces
the biomass of harvested plant material. Deposition in hazardous waste
disposal facilities seems to be the simplest way to dispose of contaminated

170 A. Sas-Nowosielska et al.

crops. However, it is not completely adequate since significant amount of
heavy-metal contaminated material will remain in the environment and
costs of its disposal are high (Sas-Nowosielska et al. 2004).

Monitoring

Routine monitoring should include collection and analysis of the following:
soil, airborne deposition, plants, vadose zone moisture, ground water, and
irrigation water. In addition, routine soil chemistry and weather monitoring
should be conducted. Weekly visits to the site are recommended to examine
the growth of plants, soil moisture, appearance of pests, etc.

7.2.3
Conclusions

The following remarks attempt to summarize the up-to-date observations
concerning phytoextraction:

• The method is applicable to cleansing sites contaminated to a moderate
or medium degree.

• Although described in literature as a lead-extraction method, significant
amounts of zinc and cadmium can thus be extracted. High concentra-
tions of zinc in soil can impair plant growth.

• Results of laboratory and bench studies should be cautiously transferred
to the field; conclusions carelessly drawn from experiments on pollutant-
spiked soil can be a source of very serious errors.

• Each new clean-up project has to be custom-tailored due to significantly
varying soil conditions and pollutant distribution.

• Induced phytoextraction, where costs of reagents contribute to the major
portion of all project expenditures, should for economic reasons only be
used on particularly valuable areas.

• Continuous phytoextraction, which may be considered a type of natural
contaminant attenuation by selected species of plants, is applicable to
the sites where time is not a driving factor.

• It is highly recommended to use indigenous species for phytoextraction,
as they are considerable cheaper than exotic species, and do not create
adaptation problems.

• Although EDTA, a commonly used metal- chelating agent, was found not
to exert any adverse effects on soil bacterial and fungal life, it should be
very carefully applied considering its potential for mobilization of metal
into other compartments of the environment.

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 171

7.3

Phytostabilization Potential for Soils

Highly Contaminated with Lead, Cadmium and Zinc

The use of certain plant species to immobilize contaminants in the soil
and ground water through accumulation and absorption by roots, adsorp-
tion onto roots' epidermis, or precipitation within the root zone is called
phytostabilization. This term further infers a physical stabilization of soil.
Phytostabilization does not remove contaminants from the soil, but reduces
the hazards to human health and environment.

Plants are used to cover the soil surface to prevent erosion, reduce
water percolation, serve as a barrier to prevent direct contact with the
soil-immobilized contaminants, and to control soil pH, gases, and redox
conditions (Vangronsveld et al. 1995). Plant roots may change soil pH by
release of exudates or through the production of C0 2 during root respira-
tion.

Phytostabilization is a site stabilization technique that reduces the risk
of soil contaminants through the use of soil amendments that induce the
formation of insoluble contaminant species. The method essentially con-
sists of a combination of the use of immobilizing soil additives to reduce
the bioavailability of heavy metals in contaminated soil and the creation
of a dense vegetation cover. Bioavailability as a function of remediation
treatment can be quantified based on the contaminant enrichment factor
(EF), which is the ratio of contaminant metal concentration in plant tissue
to the total concentration in the soil.

7.3.1

Evaluation of Site Contaminants

The investigator must establish a procedure, using less aggressive extrac-
tion, for evaluating the level of contaminant initially associated with the
solid phase. The relative extractability of the contaminant of interest is
then evaluated before and after a given stabilization treatment. In essence,
such techniques are an abbreviated sequential extraction and subjected to
the same empirical limitations with respect to interpretation as is the full
procedure.

Sampling Setup

The first sampling takes place before the technology is started. All contam-
inated areas should be sampled by using the procedures described in ISO
Standards. The following general soil parameters are quantified:

172 A. Sas-Nowosielska et al.

• Soil texture (hydrometric method)

• Soil pH in 1 N KC1, H 2 and 0.01 M CaCl 2 ; soil to solution ratio of 1:2.5
(Chapt. 2)

• Soil electroconductivity; soil to solution ratio of 1:2.5

• Soil organic matter content by loss on ignition (Chapt. 2; Houba et al.
1995);

• Cation exchange capacity (CEC), according to ISO 13536 (1995);

• Content of NO" and NH+ (Chapt. 2; Houba et al. 1995);

• P content in water extract and in an ammonium lactate-acetic acid extract
(Houba etal. 1995);

• K content soluble in 8 M KC1 (Houba et al. 1995);

• Amorphous Al and Fe content, by an ammonium oxalate-oxalic acid
extract method (Houba et al. 1995), t concentrations of Al and Fe being
analyzed with ICP (Houba et al. 1995)

• Metal extraction and determination (optional; see Sect. 7.2.1)

7.3.2

Logistic Considerations

As in case of phytoextraction, the following is to be examined before the
technical activity in the project is started:

• Existing vegetation (indicating potential for plant growth)

• Proximity to water (for installation of irrigation system)

• Site accessibility for vehicles and farm equipment

• Field observations

• Historical site activities

• Summary of regional hydrology/geology

7.3.3
Additives

Soil amendments for phytostabilization should inactivate metal contam-
inants, reducing their bioavailability, and preventing leaching and plant
uptake. Some amendments, e.g., phosphate fertilizer, have secondary ben-
efits such as supplying plant nutrients.

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 173

Limestone and organic materials (Li and Chaney 1998), natural and
synthetic zeolites, phosphate minerals (apatite, calcium phosphate, am-
monium polyphosphate, etc.), iron and manganese oxides, are suggested
for metal stabilization (Knox et al. 2000, 2001)

The application rates of the additives generally range from 0.5-5% by
soil mass. The largest amounts of additives are generally needed for highly
contaminated areas with high a percentage of bioavailable forms of con-
taminant.

7.3.4
Plants

Desirable features of species used for land phytostabilization are as follows
(Berti et al. 1998; Vangronsveld and Cunningham 1998):

- Tolerance to high concentrations of pollutant (Li and Chaney 1998)

- Ability to create a dense root mat

- Ability to accumulate pollutants in a non-edible underground part

- Low maintenance requirements (watering, pest and weed control)

- Resistance to the local climatic extremes

In the course of investigation of very highly metal-polluted areas it has
been found that Deschampsia caespitosa ecotype Warynski, a weed growing
spontaneously on the soil close to zinc smelter dumps, creates very dense
and durable plant cover upon being supplied with necessary nutrients. This
ecotype has appeared to be relatively strong and healthy, in spite of poor soil
conditions and inadequate watering. Species such as Brassica juncea, some
cultivars of grasses (Agrostis tenuis, Festuca rubra) and some cultivars of
hybrid poplars are also suggested for this purpose.

7.3.5

Full-Scale Application

The methodology presented has focused on highly metal- contaminated
areas with poor plant cover. Chemostabilization combined with phytosta-
bilization is meant to prevent pollutant migration via wind, water erosion,
and leaching.

Agronomic input includes the nutrients necessary for vigorous growth
of vegetation and rhizosphere microbes. It should be done based on a local
Good Agriculture Practice. Before the soil contaminants are to be stabi-

174 A. Sas-Nowosielska et al.

lized, inorganic (nitrogen, phosphorus, potassium), and organic fertilizers
(manure, compost, etc.) have to be applied.

For using 5% superphosphate, lime at the rate of about 12 t/ha should be
applied and mixed to a depth of 20 cm. After liming, the amendment should
be mixed with the upper 10-cm layer of soil. Two weeks after amendment
application, about 60 kg/ha of D. caespitosa seeds should be planted. This
concentration eliminates creation of tufts and increases density of plant
cover.

7.3.6

Effectiveness of Technology

Phytochemostabilization can be achieved by creation of appropriate plant
cover in combination with soil amendments. The approach we suggest
is suitable for areas heavily polluted with bivalent heavy metals, where
commercially available species for revegetation cannot survive. The overall
goal is complete coverage of the contaminated surface with a plant canopy,
whose growth is enhanced by chemicals, and which will simultaneously
immobilize pollutants and continue to support plant growth.

In particularly complex deterioration of soil, where chemical soil dam-
ages are followed by its mechanical destruction, the concerted action of
phyto- and chemostabilization may yield positive results. Evaluation of
hazard reduction must be made to validate the effectiveness of phytostabi-
lization.

7.3.7
Monitoring

Phytostabilization does not remove contaminants from the soil. There are
two objectives of monitoring during a phytostabilization process:

• To evaluate the long term effectiveness of immobilization in revegetated
areas, which leads to an estimation of the reduction of the long term
leaching potential and the influence of vegetation on leaching risks

• To evaluate plant uptake potential in relation to potential transfer of
heavy metals into the food chain

Monitoring the fate of contaminants, as well as the presence of additives
over a long period of time is required, particularly in areas with strong
impact from acid rain and possible changes in soil redox potential. Immo-
bilization effectiveness and plant uptake determinations should be carried
out during the growing seasons of several years.

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 175

7.3.8
Conclusions

The combined chemo-phytostabilization method has the following advan-
tages:

• Phosphate as used in the method decreases the concentration of bivalent
heavy metals in roots and shoots, and their bioavailable fraction in
leachates, and also improves plant cover density.

• Further, phosphate thus introduced in soil may facilitate the propagation
of Deschampsia in the third year of growth by enhancing production of
seeds, which germinate on bare soil between the tufts.

• The procedure supports the growth of the root system and makes it
stronger, resulting in increases of up to 70% water retention and reduced
metal migration.

• The growth of D. caespitosa is improved in the process at the expense of
the growth rate of Cardaminopsis sp. This is a positive phenomenon, be-
cause high heavy-metal accumulation rates in Cardaminopsis sp. shoots
results in a potential introduction of heavy metals into the food chain.

• Metal migration to lower soil levels is decreased by the procedure as
a result of metal-chemical binding and the development of a strong plant
cover.

• An optimization study to evaluate phosphorus addition to the soil and
satisfactory plant growth remains to be done, and the price of the additive
is also a matter of concern.

• Phosphate used as a fertilizer for metal contaminated soils in very high
concentration is considered disadvantageous as it causes saturation with
phosphate in the upper soil layers. This can lead to phosphate leaching.
Phosphate use is therefore limited to areas with a deep water table where
groundwater pollution by phosphate is unlikely, and where the greater
benefit of obtaining healthy plant cover is unlikely to be achieved.

• Phosphate is not recommended for arsenic-polluted soils, as competition
between arsenate and phosphate can provoke increased arsenic levels in
plants, causing risks of food-chain propagation and accumulation.

Acknowledgements. The authors wish to express their thanks to Mr. Laymon
Gray of Florida State University for his editorial contribution to this paper.

176 A. Sas-Nowosielska et al.

References

Berti WR, Cunningham SD, Cooper EM (1998) Case studies in the field - in-place inac-
tivation and phytorestoration of Pb-contaminated sites. In: Vangronsveld J and Cun-
ningham SD (eds) Metal-contaminated soils: in situ inactivation and phytorestoration.
Springer- Verlag, Berlin Heidelberg and RG Landes Co, Georgetown, TX, USA, pp 235-
248

Blaylock MJ, Salt DE, Dushenkov S, Zakharova O, Gussman C, Kapulnik Y, Ensley BD,
Raskin I (1997) Enhanced accumulation of Pb in Indian mustard by soil-applied chelat-
ing agents. Environ Sci Technol 31:860-865

Brooks RR (1998) Phytochemistry of hyperaccumulators. In: Brooks RR (ed) Plants that
hyperaccumulate heavy metals. Cab International, Wallingford, Oxon, UK, pp 15-53

Houba VJG, Van der Lee J J, Novozamsky I (1995) Soil analysis procedures, other procedures
(Soil and plant analysis, Part 5b). Dept Soil Sci Plant Nutr, Wageningen Agricultural
University, pp 217

ISO 11265 (1994) Soil quality - Determination of the specific electric conductivity

ISO 11464 (1993) Soil quality - Pretreatment of samples for physico-chemical analyses

ISO 13536 (1995) Soil quality - Determination of the potential cation exchange capacity and
exchangeable cations using barium chloride solution buffered at pH = 8.1

ISO 7888 (1985) Water quality - Determination of electrical conductivity

ISO/CD/10381-5 (1995) Soil quality - Sampling

ISO/DIS 10390 (1993) Soil quality - Determination of pH

ISO/DIS 11047 (1994) Soil quality - Determination of cadmium, chromium, cobalt, copper,
lead, manganese, nickel and zinc. Flame and electromatic thermal atomic absorption
spectrometric methods

ISO/DIS 11466 (1995) Soil quality - Extraction of trace metals and heavy metals soluble in
aqua regia

Knox AS, Seaman J, Adriano DC, Pierzynski G (2000) Chemophytostabilization of metals in
contaminated soils. In: Wise DL, Trantolo DJ, Cichon EJ, Inyang HI, Stottmeister U (eds)
Bioremediation of contaminated soils. Marcel Dekker, Inc, New York, Basel, pp 811-
836

Knox AS, Seaman JC, Mench MJ, Vangronsveld J (2001) Remediation of metal- and
radionuclides-contaminated soils by in situ stabilization techniques. In: Iskandar IK
(ed) Environmental restoration of metal-contaminated soils. Lewis Publ, Boca Raton,
London, New York, Washington, DC, pp 21-60

Kucharski R, Sas-Nowosielska A, Dushenkov S, Kuperberg JM, Pogrzeba M, Malkowski E
(1998) Technology of phytoextraction of lead and cadmium in Poland. Problems and
achievements. In: Symposium Proceedings, Warsaw'98, Fourth Int Symposium and
Exhibition on Environmental Contamination in Central and Eastern Europe, pp 55

Kucharski R, Sas-Nowosielska A, Krynski K (2000) Amendment application technology
for phytoextraction. In: Symposium Program, Prague 2000, Fifth Int Symposium and
Exhibition on Environmental Contamination in Central and Eastern Europe, Abstract,
p376

Kucharski R, Sas-Nowosielska A, Kuperberg M, Bocian A (2004) Survey and assessment.
How urbanization and industries influence water quality. In: Integrated watershed man-
agement - ecohydrology & phytotechnology, manual. UN Educational, Scientific and
Cultural Organization, Venice, Italy, pp 45-60

Li YM, Chaney L (1998) Case studies in the field - industrial sites: phytostabilization of zinc
smelter- contaminated sites: the Palmerton case. In: Vangronsveld J, Cunningham SD
(eds) Metal-contaminated soils: in situ inactivation and phytorestoration. Springer-
Verlag Berlin Heidelberg, and RG Landes Co, Georgetown, TX, USA, pp 197-210

7 Feasibility Studies for Phytoremediation of Metal- Contaminated Soil 177

McGrath SP, Dunham SJ, Correl RL (2000) Potential for phytoextraction of zinc and cadmium
from soils using hyperaccumulator plants. In: Terry N, Banuelos G (eds) Phytoremedi-
ation of contaminated soil and water. Lewis Publ, Florida, pp 1-13

Salt DE, Smith RD, Raskin I (1998) Phytoremediation. Annu Rev Plant Physiol Plant Mol
Biol 49:643-668

Sas-Nowosielska A, Kucharski R, Korcz M, Kuperberg M, Malkowski E (2001) Optimizing
of land characterization for phytoextraction of heavy metals. In: Gworek B, Mocek A
(eds) Element cycling in the environment, bioaccumulation - toxicity - prevention,
Monograph, vol 1. Instytut Ochrony (Erodowiska, Warsaw, Poland, pp 345-348

Sas-Nowosielska A, Kucharski R, Malkowski E, Pogrzeba M, Kuperberg M, Krynski K (2004)
Phytoextraction crop disposal - an unsolved problem. Environ Pollution 128:373-379

Vangronsveld J, Cunningham SD (1998) Introduction to the concept. In: Vangronsveld J,
Cunningham SD (eds) Metal- contaminated soils: in situ inactivation and phytorestora-
tion. Springer- Verlag, Berlin Heidelberg, and RG Landes Co, Georgetown, TX, USA,
pp 1-15

Vangronsveld J, Van Assche F, Clijsters H (1995) Reclamation of a bare industrial area
contaminated by non-ferrous metals: in situ metal immobilization and revegetation.
Environ Pollution 87:51-59

Quantification of Hydrocarbon
Biodegradation Using Internal Markers

Roger C. Prince, Gregory S. Douglas

■ Introduction

Objectives. Soil contamination is invariably heterogeneous, and monitor-
ing the loss of contaminant during bioremediation is often frustrated by
this heterogeneity. But if the initial source of contamination was relatively
homogeneous, it is possible to identify biodegradation as the selective loss
of the most biodegradable components, while more recalcitrant molecules
are conserved. Measuring the concentrations of a series of compounds us-
ing gas chromatography (GC) coupled with mass spectrometry (MS), often
in the selected ion monitoring (SIM) mode, allows this to be achieved with
high precision.

Hopanes have proven to be useful conserved internal markers for fol-
lowing the biodegradation of crude oil contamination (Prince at al. 1994),
trimethylphenanthrenes for following the biodegradation of diesel fuel
(Douglas et al. 1992), and 2,2,3,3-tetramethylbutane and 1,1,3-trimethyl-
cyclopentane for following the anaerobic biodegradation of gasoline and
condensate (Townsend et al. 2004). Undoubtedly, there are many other
compounds that could be used. Even if the "conserved" internal marker
is itself eventually degraded, this will have the effect of underestimating
the extent of biodegradation of compounds referred to it, making the ap-
proach a conservative one. The principal requirements are that the samples
under consideration initially had the same contaminant, and that the com-
pound chosen as the "conserved" internal standard be amongst the least
degradable in the mixture under study, and be present at a high enough
concentration to be measured with good precision.

Principle. Depending on the type of contamination, which can be deter-
mined from the hydrocarbons present (Stout et al. 2002), the least biode-
graded sample is identified, and candidate conserved species are identified.
The ratios of various analytes to these species are then followed over time,
and biodegradation is identified from their coherent loss. The concentra-
tion of the conserved species (e.g., hopane) on an oil-weight basis may

Roger C. Prince: ExxonMobil Research and Engineering Co., Annandale, New Jersey 08801,
USA, E-mail: Roger.C.Prince@ExxonMobil.com

Gregory S. Douglas: NewFields Environmental Forensic Practice LLC, Rockland, Mas-
sachusetts 02370, USA

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

180 R.C. Prince, G.S. Douglas

also be used to estimate the total quantity of oil that has been degraded
(Douglas et al. 1994) within a sample.

Theory. The biodegradation of hydrocarbons has been studied for al-
most a century, and the overall process is quite well understood (Prince
2002). Under aerobic conditions, n-alkanes and simply substituted mono-
aromatic species are amongst the most readily biodegraded hydrocarbons,
followed by the iso- and monocyclic alkanes, benzene and the simply alky-
lated two and three-ring aromatics (Solano-Serena et al. 1999). More highly
alkylated species, four- ring and larger aromatics (Douglas et al. 1994), and
compounds containing tertiary carbons are more resistant to biodegrada-
tion (Prince et al. 1994). Similar patterns are seen under methanogenic and
sulfate-reducing conditions, with the apparent distinction that some cyclic
alkanes are very readily degraded under these conditions (Townsend et al.
2004). The biodegradation of at least some hydrocarbons, e.g., toluene,
occurs under other anaerobic conditions as well (Chakraborty and Coates
2004).

Inevitably some analyte in any complex mixture is its least biodegrad-
able compound. Referring the concentrations of other analytes to this com-
pound provides a ready index of the extent of biodegradation of that analyte,
and removes much of the variability in the absolute concentration of the an-
alyte in soil and sediment samples. This is shown graphically in the figures.
Figure 8.1 shows the biodegradation of 2-methylhexane over 100 days in
samples from a condensate-contaminated anaerobic aquifer amended with
a small amount of gasoline and incubated under sulfate-reducing condi-
tions (Townsend et al. 2004). The raw data are seen in Fig. 8.1 A, the data
referred to 1,1,3-trimethylcyclohexane as a conserved internal marker in
Fig. 8. IB. Similarly, Fig. 8.2 shows the biodegradation of the sum of the
USEPA priority pollutant polycyclic aromatic hydrocarbons (PAHs; Keith
and Telliard 1979) in a historically contaminated refinery soil over a time
span of 1.5 years (Prince et al. 1997). The raw data are seen in Fig. 8.2 A, the
data referred to 17a(H),21/?(H)-hopane as a conserved internal marker in
Fig. 8.2B. In both cases, the biodegradation of the target compound(s) is
much more apparent in the B panels.

■ Procedure

The precise recipes for extracting and analyzing samples will depend on
many site-specific variables, and we give only a broad description of the pro-
tocols involved. Measurements made for regulatory compliance are usually
specifically mandated by the regulators involved, and we do not discuss
them here. Rather we focus on measurements made to assess whether
biodegradation is proceeding, and whether bioremediation protocols are

8 Quantification of Hydrocarbon Biodegradation Using Internal Markers

181

100 ■

arb.
units

20 ■

100

°/c

■

40 80

Days

120

20 ■

Days

120

Fig. 8.1. A The biodegradation of 2-methylhexane under sulfate-reducing conditions in sam-
ples collected from a condensate-contaminated aquifer, amended with 1 uL of gasoline (per
50 g sediment, 75 mL groundwater) and incubated in the laboratory under sulfate-reducing
conditions (Townsend et al. 2004). The individual incubations were carefully assembled
with equal weights of sieved sediments in each bottle, yet the raw data are still very hetero-
geneous. B The data and referenced to the concentration of 1,1,3-trimethylcyclohexane in
each sample

140

arb.
units

200 400

Days

200 400

Days

600

Fig. 8.2. Biodegradation of the 16 USEPA Priority Pollutant PAHs in a refinery soil. The data
(the sum of the concentrations) were collected after a bioremediation protocol of adding
slow release nutrients was initiated (Prince et al. 1997). A Although the soil was tilled during
the treatment, and individual samples were sieved prior to analysis, the raw data are still
very heterogeneous. B The data referenced to the concentration of 17a(H),21/?(H)-hopane
in each sample

indeed stimulating the process. This is best done by comparing samples
from a site undergoing active bioremediation with samples from a similarly
contaminated site with no intervention. Unfortunately, this is often impos-
sible, and samples collected during active bioremediation protocols have
to be compared with samples taken at the beginning of the remediation. In
either case, absolute amounts of contaminants in "replicate" samples are
likely to be log-normally distributed (Limpert et al. 2001 ), and changes due

182 R.C. Prince, G.S. Douglas

to biodegradation will be difficult to detect unless the conserved-marker
approach is used.

Sample Preparation

Sample preparation is fundamentally different if the compounds of concern
are in the gasoline or diesel and higher range. For soils, sediments, and water
samples contaminated with gasoline, the appropriate extraction procedure
is "purge-and-trap" analysis (Uhler et al. 2003). For soils contaminated with
kerosene, diesel, heating, or crude oil it is more appropriate to extract the
hydrocarbons into a solvent and inject the solvent-hydrocarbon mixture
directly into the GC (Douglas et al. 1992, 2004).

Internal Standards

Often it is appropriate to add surrogate internal standards prior to extrac-
tion. These may be added for two fundamentally distinct reasons. One is
to assess the efficiency of the extraction protocol: fluorobenzene is often
used for "purge-and-trap" analyses, while o-terphenyl is often used in sol-
vent extractions. The second is to add compounds to check that the mass
spectrometer is working correctly: deuterated compounds are often used
(Uhler et al. 2003; Douglas et al. 1992, 1994, 2004).

"Purge-and-Trap"

"Purge-and-trap" protocols for the extraction of volatile hydrocarbons
are described in USEPA methods 5030B: "Purge-and-Trap for Aqueous
Samples," and 5035: "Closed-System Purge-and-Trap and Extraction for
Volatile Organics in Soil and Waste" (USEPA 2003). Although the technical
aspects are discussed in the EPA Method, the target analytes to which this
method is applied includes only eight hydrocarbons present in gasoline
(benzene, toluene, ethylbenzene, m-, p-, and o-xylene, styrene, and naph-
thalene). This is inadequate for detailed characterization of gasoline and
other light hydrocarbon products and for measuring conserved species. Uh-
ler et al. (2003) have modified Method 8260 to quantitatively measure more
than 100 diagnostic gasoline-related compounds ranging from isopentane
to dodecane in nonaqueous phase liquid products, water, and soil. Due
to the wide range of solubilities and volatilities of these compounds (e.g.,
benzene versus dodecane), caution must be exercised when analyzing these
additional compounds by the purge-and-trap methods and careful calibra-
tion and monitoring of analyte-recovery efficiencies should be performed
(Uhler et al. 2003).

In essence, an appropriate amount of sample to give a response within
the calibrated range of the GC system is flushed (purged) with an inert gas
to transfer the analytes of interest to a trap. When the purging is complete,

8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 183

which usually takes several minutes, the trap is rapidly heated to transfer
the sample into the GC column. If the sample is a soil sample, sufficient
clean water is added prior to the purging to make a fluid slurry. The initial
sampling must be done rapidly and into tightly sealed vessels to prevent
any loss of volatile components during sample collection and storage. In
our hands, samples containing about 1 pi of gasoline are appropriate for
analysis (Townsend et al. 2004).

Solvent Extraction

Solvent extraction protocols are described in USEPA method 3500B: "Or-
ganic extraction and sample preparation" (USEPA 2003). Soil or sediment
samples are dried by mixing them with enough anhydrous sodium sul-
fate to make a freely flowing dry mixture. Typical samples may require an
equal weight of sodium sulfate, and it is important to mix thoroughly and
for some time (perhaps 20 min) to allow the drying agent to hydrate and
dry the sample. Samples are then serially extracted, at least three times,
with an appropriate solvent (e.g., methylene chloride or methylene chlo-
ride/acetone 1 + 1), perhaps in a Soxhlet extraction device, by accelerated
solvent extraction (ASE), or by supercritical fluids.

The extracts are dried with sodium sulfate, filtered, and then concen-
trated as appropriate. It is important that this solvent-evaporation be done
carefully to minimize the loss of lighter volatile components, such as the
two-ring aromatics. Only in rare cases where it is known that there are no
volatile compounds should it be allowed to proceed to dryness. Automated
devices are available, but solvent-evaporation can be done manually under
a gentle stream of dry nitrogen gas at ambient temperature.

Depending on the minimum detection limits required (Douglas et al.
2004), and the presence of interfering compounds, it maybe appropriate to
process the solvent extract on an alumina or silica column to isolate "clean"
fractions of saturate, aromatic, and polar compounds. This is described in
detail in USEPA method 361 1: "Alumina column cleanup and separation of
petroleum wastes" and USEPA method 3630 "Silica Gel Cleanup" (USEPA
2003). Often the two hydrocarbon fractions (saturate and aromatic hy-
drocarbons) are combined, concentrated to an appropriate volume, and
amended with additional internal standards to allow quantitation; again
deuterated compounds are often used. In our hands, 1 p.L injections of
samples containing about 5 mg of crude oil/mL solvent are appropriate for
analysis (Douglas et al. 1992, 2004).

Gas Chromatography and Mass Spectrometry (GC/MS)

This requires an appropriate high-resolution capillary column equipped
with a mass spectrometer (McMaster and McMaster 1998; Hubschmann

184 R.C. Prince, G.S. Douglas

2000). USEPA methods 8260 and 8270D (USEPA 2003) provide GC/MS
protocols for the measurement of volatile and semi-volatile hydrocar-
bons, respectively. As noted above, the EPA protocols are not designed
for petroleum product analysis and have been modified by various inves-
tigators to increase the number of petroleum-specific target compounds
(Douglas and Uhler 1993; Uhler et al. 2003) and improve the sensitivity of
the methods (Douglas et al. 1994, 2004).

For the modified EPA Method 8260 (Uhler et al. 2003) compounds are
identified and quantified using full-scan mass spectrometry (typically from
m/z = 35-300) for the extended volatile hydrocarbon target analyte list (109
gasoline-specific compounds). The advantage of full-scan analysis is that
additional compounds can always be evaluated, and extracted ion plots of
compound classes (e.g., alkylcyclohexanes, Townsend et al. 2004) can be
obtained to determine that the products are derived from the same source.
Although the full-scan GC/MS approach is not as sensitive as selected ion
monitoring (SIM), it is generally adequate for volatile hydrocarbon analysis.

In contrast, it is essential to use selected ion monitoring (SIM) in the
modified EPA Method 8270 (Douglas et al. 1992, 2004). This protocol al-
lows the measurement of the major paraffins and isoparaffins, the aro-
matics on the USEPA list of priority pollutants (Keith and Telliard, 1979)
and their alkylated forms, and the steranes and hopanes that are so valu-
able in discriminating different crude oils (Peters et al. 2004). The most
significant modifications of the USEPA Method are the inclusions of the
dibenzothiophenes, alkylated PAHs, steranes and hopanes that provide
petroleum source identification and bioremediation efficacy information
(Douglas et al. 2002).

Analytes are identified by the retention times of authentic standard com-
pounds, and by reference to mass spectral libraries such as those distributed
by NIST/EPA/NIH (NIST 2004). It is always appropriate to use more than
one ion to identify analytes in the initial samples to assess whether there
are any interfering species present, and if so, how to account for them.

For research purposes it is usually possible to arrange the concentra-
tions of analytes to fall into the linear range of detectability, which should
be determined with a range of calibration standards. A lot of work has gone
into optimizing detection limits for the analysis of complex environmental
samples for forensic applications (Douglas et al. 2004), but only the simplest
precautions are needed for most studies quantifying biodegradation. Cer-
tainly the mass spectrometer should be tuned with an appropriate standard,
such as decafluorotriphenylphosphine, before every batch of samples, and
standard samples and blanks should be included in every group of samples.
Of course, if the analytical variability is large then the ability to detect an
impact of a bioremediation protocol is reduced. Therefore, it is preferable
to measure all the samples for a particular study at one time, or at least to

8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 185

include control and reference samples with every batch. This may require
that early samples be preserved until analysis; careful freezing or acidifica-
tion to pH 2 with HC1 both work well. Furthermore, it is appropriate to set
some "quality control" values that the standard samples must satisfy before
the data are considered suitable for analysis. Guidelines for suitable control
values are given in USEPA method 8270D (USEPA 2003) and in Page et al.
(1995).

■ Calculation

We can calculate the percent of an analyte remaining (Figs. 8.1 and 8.2)
from the equation:

(Aq/Cs)
% Remaining = — — - x 100 (8.1)

8 (Ao/Q)

A s concentration of the target analyte in the sample

Cs concentration of the conserved compound in the sample

A concentration of the target analyte in the initial sample

C concentration of the conserved compound in the sample

Alternatively the percent depletion of biodegradable analytes within the
oil (Fig. 8.3) can be calculated using the equation:

(Aq/Cq) - (As/Cs) ....

%Loss = x 100 (8.2)

(Ao/Co)

Note that these equations work equally well in absolute concentration
terms, or in arbitrary units, as long as the latter are obtained under identical
conditions for all samples.

Notes and Points to Watch

The approach outlined here relies on the initial source of contamination
being reasonably homogeneous. This is readily achieved in laboratory
studies, and often pertains to acute contamination accidents such as oil
spills. But chronic contamination may prove too heterogeneous for this
approach to work without subdividing areas under consideration (e.g.,
Prince et al. 1997). For example, the composition of gasoline has changed
over the years as more effective refinery processes have been introduced,
and as the molecular composition has come under regulatory oversight.

186

R.C. Prince, G.S. Douglas

Similarly, contamination at town gas sites and refineries may be from
a mixture of sources. It is thus essential to take enough samples of the
contamination prior to any remediation activities to delineate areas of
similar and distinctly different contamination.

It is important to minimize evaporative losses prior to analysis. This
means carefully sealed sample vials for "purge-and-trap" analyses, and
care during evaporative solvent removal from extracts. Including appro-
priate surrogate compounds in the analysis can assess such losses.

Biochemical intuition and published work will help identify potential
analytes to be used as conserved internal compounds. Consistently neg-
ative values for the % depletion of other analytes with respect to the
"conserved" one will indicate that the "conserved" compound is in fact
more degradable than the other analytes, and allow selection of a better
standard compound (e.g., see Fig. 8.3)

The simple analysis of Figs. 8.1 and 8.2 may be all that is needed to
demonstrate that biodegradation is occurring, but more complicated
models for biodegradation, taking into account the amount of oil, its

100

depletion

-100

Referred to C 3 -phenanthrenes
Referred to hopane

/7C 1(

Co-phenanthrenes

hopane

pristane

phytane

naphthalene

C-fluorenes

chrysene

C 3 -chrysenes

phenanthrene

benzo[b]fluoranthene

Fig. 8.3. Percent depletion plot for some alkanes, PAHs, and hopane in a degraded
Alaskan North Slope crude oil (Douglas et al. 1994). The hatched series repre-
sents the percent depletion of each analyte based on the C3-phenanthrenes (the
trimethyl, methyl- ethyl, propyl and isopropylphenanthrenes) as the conserved inter-
nal marker. Note that some compounds have a negative apparent depletion, indi-
cating that the C3-phenanthrenes are less conserved than those analytes. The solid
series represents the percent depletion based on the more biodegradation resistant
17a(H),21^(H)-hopane. (Prince et al. 1994)

8 Quantification of Hydrocarbon Biodegradation Using Internal Markers 187

prior weathering, and the amount of available fertilizer, have been used to
demonstrate the effectiveness of bioremediation in the field (Bragg et al.
1994).

• Biodegradation can be identified by the loss of biodegradable com-
pounds, as discussed above. The loss of photochemically labile species
can also be followed (Garrett et al. 1998; Douglas et al. 2002), as can the
loss following extensive washing and evaporation (Douglas et al. 2002;
Prince et al. 2002) and the increase of pyrogenic compounds following
partial oil combustion (Garrett et al. 2000). Providing a sample of the
initially spilled oil is available, these environmental processes can then
be identified in samples collected from historical spills (Prince et al.
2003).

• The general approach can also be used to follow the biodegradation of
any complex mixture of contaminants, such as polychlorinated biphenyls
(Abramowicz 1995).

References

Abramowicz DA (1995) Aerobic and anaerobic PCB biodegradation in the environment.

Environ Health Perspect 103 Suppl 5:97-99
Bragg JR, Prince RC, Harner EJ, Atlas RM (1994) Effectiveness of bioremediation for the

Exxon Valdez oil spill. Nature 368:413-418
Chakraborty R, Coates JD (2004) Anaerobic degradation of monoaromatic hydrocarbons.

Appl. Microbiol. Biotechnol. 64:437-446
Douglas GS, Burns WA, Bence AE, Page DS, Boehm P (2004) Optimizing detection limits for

the analysis of petroleum hydrocarbons in complex environmental samples. Environ

Sci Technol 38:3958-3964
Douglas GS, McCarthy KJ, Dahlen DT, Seavey JA, Steinhauer WG, Prince RC, Elmendorf DL

(1992) The use of hydrocarbon analyses for environmental assessment and remediation.

J Soil Contam 1:197-216
Douglas GS, Owens EH, Hardenstine J, Prince RC (2002) The OSSAII pipeline spill: the

character and weathering of the spilled oil. Spill Sci Technol Bull 7:135-148
Douglas GS, Prince RC, Butler EL, Steinhauer WG (1994) The use of internal chemical

indicators in petroleum and refined products to evaluate the extent of biodegradation.

In: Hinchee RE, Alleman BC, Hoeppel RE, Miller RN (eds) Hydrocarbon remediation.

Lewis Publ, Boca Raton, FL, pp 219-236
Douglas GS, Uhler AD (1993) Optimizing EPA methods for petroleum contaminated site

assessments. Environ Test Anal 2:46-53
Garrett RM, Guenette CC, Haith CE, Prince RC (2000) Pyrogenic polycyclic aromatic hy-
drocarbons in oil burn residues. Environ Sci Technol 34:1934-1937
Garrett RM, Pickering IJ, Haith CE, Prince RC (1998) Photooxidation of crude oils. Environ.

Sci. Technol. 32:3719-3723
Hubschmann, H.-J. (2000) Handbook of GC/MS: fundamentals and applications. Wiley-

VCH, Weinheim, Germany
Keith LH, Telliard WA, (1979) Priority pollutants I. - a perspective view. Environ Sci Technol

13:416-423

188 R.C. Prince, G.S. Douglas

Limpert E, Stahel WA, Abbt M (2001) Log- normal distributions across the sciences: Keys

and clues. Bioscience 51:341-352
McMaster M, McMaster C (1998) GC/MS: A practical user's guide. Wiley- VCH, New York
NIST (2004) NIST/EPA/NIH mass spectral library www.nist.gov/srd/mslist.htm
Page DS, Boehm PD, Douglas GS, Bence AE (1995) Identification of hydrocarbon sources in

benthic sediments of Prince William Sound and the Gulf of Alaska following the Exxon

Valdez oil spill. In: Wells PG, Butler JN, Hughes JS (eds) Exxon oil spill: Fate and effects

in Alaskan waters, ASTM Special Technical Publication #1219, American Society for

Testing and Materials, Philadelphia, pp 41-83
Peters KE, Walters CC, Moldowan JM (2004) The Biomarker guide, biomarkers and isotopes

in petroleum exploration and earth history, vol 1-2, 2nd edn. Cambridge Univ Press,

New York
Prince RC (2002) Biodegradation of petroleum and other hydrocarbons. In: Bitton G (ed)

Encyclopedia of environmental microbiology. Wiley, New York, pp 2402-2416
Prince RC, Drake EN, Madden PC, Douglas GS (1997) Biodegradation of polycyclic aromatic

hydrocarbons in a historically contaminated site, in: Alleman BC, Leeson A (eds) In situ

and on-site bioremediation 2. Battelle Press, Columbus, OH, pp 205-210
Prince RC, Elmendorf DL, Lute JR, Hsu CS, Haith CE, Senius JD, Dechert GJ, Douglas GS,

Butler EL (1994) 17a(H),21/?(H)-hopane as a conserved internal marker for estimating

the biodegradation of crude oil. Environ Sci Technol 28:142-145
Prince RC, Garrett RM, Bare RE, Grossman MJ, Townsend GT, Suflita JM, Lee K, Owens EH,

Sergy GA, Braddock JF, Lindstrom JE, Lessard RR (2003) The roles of photooxidation

and biodegradation in long-term weathering of crude and heavy fuel oils. Spill Sci

Technol Bull 8:145-156
Prince RC, Stibrany RT, Hardenstine J, Douglas GS, Owens EH (2002) Aqueous vapor ex-
traction: a previously unrecognized weathering process affecting oil spills in vigorously

aerated water. Environ Sci Technol 36:2822-2825
Solano-Serena F, Marchal R, Ropars M, Lebeault JM, Vandecasteele JP ( 1 999) Biodegradation

of gasoline: kinetics, mass balance, and fate of individual hydrocarbons. J Appl Microbiol

86:1008-1016
Stout SA, Uhler AD, McCarthy KJ, Emsbo-Mattingly S (2002) Chemical fingerprinting of

hydrocarbon. In: Murphy B, Morrison R (eds) Introduction to environmental forensics.

Academic Press, New York, pp 135-260
Townsend GT, Prince RC, Suflita JM (2004) Anaerobic biodegradation of alicyclic con-
stituents of gasoline and natural gas condensate by bacteria from an anoxic aquifer.

FEMS Microbiol Ecol 49:129-135
Uhler RM, Healey EM, McCarthy KJ, Uhler AD, Stout, SA (2003) Molecular fingerprinting

of gasoline by a modified EPA 8260 gas chromatography-mass spectrometry method.

Int J Environ Anal Chem 83:1-20
USEPA (2003) Index to EPA test methods, http://www.epa.gov/epahome/index/

Assessment of Hydrocarbon Biodegradation
Potential Using Radiorespirometry

Jon E. Lindstrom, Joan E Braddock

■ Introduction

Objectives. Following environmental exposure to petroleum, acclimation
of microbial communities to hydrocarbon metabolism may occur through
selective enrichment of member populations possessing hydrocarbon cata-
bolic pathways, induction or repression of enzymes, or genetic mutations
resulting in new metabolic capabilities (Leahy and Colwell 1990). Measure-
ments of carbon substrate mineralization in vitro can be used to assess the
hydrocarbon biodegradative potential of microbial communities in envi-
ronmental samples previously exposed to oil contamination in situ (Walker
and Colwell 1976; Lindstrom et al. 1991; B0rresen et al. 2003).

Using 14 C-labeled hydrocarbon substrates, mineralization of specific hy-
drocarbon compounds can be tracked, and low levels of mineralization
activity are detectable if sufficiently high specific activity substrates are
employed. Model compounds can indicate the degree of a community's
acclimation to various hydrocarbon classes (e.g., hexadecane for linear
alkanes, toluene for monoaromatic hydrocarbons, or phenanthrene for
polycyclic aromatic hydrocarbons (PAHs; Bauer and Capone 1988). By
appropriately manipulating experimental conditions, this method may be
used to assess the prior exposure of environmental samples to hydrocarbon
contamination (Braddock et al. 1996; Braddock et al. 2003), or the effects of
fertilization or other field treatments used to enhance in situ hydrocarbon
degradation (Lindstrom et al. 1991). In addition, manipulation of nutri-
ent levels or other amendments in the assay may be used in bench-scale
treatability studies prior to initiating field-scale bioremediation efforts.

Principle. A 14 C-labeled hydrocarbon substrate is added to a soil sample
suspended in sterile diluent contained in a sealed volatile organic anal-
ysis (VOA) vial. The sample is incubated under appropriate conditions
(dictated by the experimental question), and microbial metabolism of the
added substrate is measured by recovery of 14 C-labeled C0 2 evolved during

Jon E. Lindstrom: Shannon & Wilson, Inc., 2355 Hill Road, Fairbanks, Alaska 99709, USA,
E-mail: JEL@shanwil.com

Joan F. Braddock: College of Natural Science and Mathematics, University of Alaska Fair-
banks, Fairbanks, Alaska 99775, USA

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

190 J.E. Lindstrom, J.R Braddock

incubation. Microbial activity is halted by adding a strong base at the end
of the incubation period, which sequesters the C0 2 generated by microbial
substrate mineralization as carbonates in solution. The 14 C-labeled C0 2
is subsequently recovered by acidifying the suspension, then stripping the
C0 2 from solution with nitrogen gas, and capturing it in a basic scintilla-
tion cocktail. The 14 C0 2 derived from mineralization of the added labeled
substrate is counted by liquid scintillation, and its radioactivity compared
to that added with the labeled substrate.

Theory. Petroleum is a complex mixture of hydrocarbons, and nitrogen-,
sulfur- and oxygen-containing organic compounds; and the hydrocarbon
fraction itself may be composed of hundreds of aliphatic, alicyclic, and
aromatic compounds (National Research Council 1985). Heterotrophic
biodegradation of the organic substrates in petroleum therefore occurs
via a diversity of pathways, with metabolic intermediates funneled to cen-
tral metabolic pathways leading to the production of microbial biomass
and carbon dioxide (Wackett and Hershberger 2001). The fate of carbon in
the substrate metabolized varies depending on the organism, the pathways
used, and other factors. For example, biomass incorporation of glucose
was approximately twice that of phenolic compounds in taiga forest floor
samples, while respiration of C0 2 in these samples was significantly higher
for phenolic compounds (Sugai and Schimel 1993). Despite the variation in
carbon allocation among substrates and microbial communities, respira-
tion of carbon dioxide is useful for monitoring biodegradation of organic
substrates, particularly when the source of the carbon may be tracked by
radioactive labeling.

The protocol described here assesses the respiration activity of organ-
isms in environmental samples. The procedure is designed to minimize the
many factors affecting the actual mineralization activity in situ, except for
the in situ microbial biomass and its potential to biodegrade the hydrocar-
bons tested. The rate of 14 C0 2 production (r*, Bq/day) from a radiolabeled
substrate is a function of the overall rate of C0 2 production (R) and the
specific activity of the added label (Brown et al. 1991):

r* = x R (9.1)

(Sn+A)

A* radioactivity of the labeled substrate added to the sample (Bq/g soil)

S n in situ substrate concentration (|^g/g soil)

A concentration of substrate added with the radiolabeled substrate (|ig/g
soil)

R rate of C0 2 production (|^g/day) from carbon sources in the sample

9 Assessment of Hydrocarbon Biodegradation 191

By adding to the sample an amount of the tested substrate (A) that is
large compared to S n> the value of r* will mainly depend on A, rather than S n
(Brown et al. 1991). As the amount of substrate added to the sample must be
greater than the in situ concentration, and conditions in vitro are designed
to minimize the various other factors affecting in situ mineralization rates,
the value of r* reflects the microbial community's biodegradation potential
only and is not a measure of in situ mineralization rates.

The choice of incubation conditions may be used to assess the degree of
a microbial community's acclimation to a given hydrocarbon substrate in
the environmental sample, evaluate the effectiveness of field treatments, or
establish optimum growth conditions for the community being studied. As
the in situ mineralization rate may be attenuated due to nutrient deficien-
cies or other environmental factors, radiorespirometric assays conducted
with added nutrients or other amendments are useful for assessing the
degree of community acclimation (suggesting prior exposure; Braddock
et al. 1996; Braddock et al. 2003) to the hydrocarbon substrate or class of
substrates (e.g., alkanes, monoaromatics, PAHs) being tested, since such
environmental limitations are removed.

A lag period following substrate addition is observed in the assay, with its
duration commonly varying as a function of the solubility and molecular
structure of the substrate (Brown et al. 1991). To measure the activity of
the extant biomass present in the sample on collection, an appropriate
incubation period must be chosen that is short enough to avoid in vitro
acclimation of the native biomass to the added substrate, but long enough
to detect its mineralization (see below).

Equipment

Incubators equilibrated to temperatures dictated by experimental re-
quirements

Apparatus for collecting C0 2 evolved from the soil suspension follow-
ing incubation and liquid scintillation counter to detect the radioac-
tivity associated with mineralization of the added labeled substrate.
[A schematic of an apparatus suitable for stripping and capturing C0 2
evolved from the soil suspension is shown in Fig. 9.1: Nitrogen gas is
bubbled through the acidified soil suspension via a spinal needle (10-
cm, 18-gauge deflected-point, non-coring, septum-penetrating needle
with standard hub and stainless steel cannula; Popper and Sons, New
Hyde, NY, USA) that pierces the silicone septum of the VOA vial. The gas
stream strips the C0 2 from the suspension, and is conveyed to a Harvey
trap (R.J. Harvey Instruments, Hillsdale, NJ, USA) containing acidified
toluene via Tygon tubing attached to a 1-mL syringe sleeve cut to fit in

192

J.E. Lindstrom, J.R Braddock

Gas flow direction

Source of N

40-mL VOA vial
containing acidified
soil suspension

Harvey trap containing toluene
acidified with 12 N HCI

20-mL scintillation vial
containing C0 2 -sorbing
scintillation cocktail

Fig. 9.1. Schematic diagram of stripping apparatus used to collect 14 C02 from samples
following incubation. Nitrogen gas is bubbled through the sample, and the gas stream flows
through a Harvey trap containing acidified toluene to trap any volatile hydrocarbons in
the gas stream. Finally, 14 C02 is collected in a vial containing a C02-sorbing scintillation
cocktail

the tubing and equipped with a 16-gauge needle that pierces the VOA
vial septum. The gas stream is bubbled through the acidified toluene in
the Harvey trap to capture any labeled organic substrate that may have
been stripped from the soil suspension. The gas stream containing the
labeled C0 2 is then conveyed to a 20-mL scintillation vial fitted with
a two-hole rubber stopper and glass tubing (a 1-mL glass pipette cut to
a 5 cm length works well here for the glass tubing, as it provides a tapered
and polished tip). The influent gas stream is bubbled through a 10 mL
scintillation cocktail containing /?-phenylethylamine (PEA) to capture
the C0 2 . Following a 15-min stripping period, the gas flow is stopped,
the rubber stopper removed, and the scintillation vial capped and placed
in a scintillation counter to determine the amount of recovered radioac-
tivity. The stripping apparatus may be modified so that a number of
samples may be run simultaneously. This requires a manifold equipped
with valves and multiple sets of the apparatus described above. A single
nitrogen tank can be connected to the manifold and used to strip 14 C0 2
evolved from several soil suspensions in parallel.]

9 Assessment of Hydrocarbon Biodegradation 193

• Sterile and pre-cleaned or combusted 40 mL borosilicate VOA vials
equipped with Teflon-lined, 0.125-mm-thick, silicone septa (e.g., I-Chem
Brand; Nalge Nunc, Rochester, NY, USA)

• Sterile 10-mL pipettes

• 100-|iL syringe (Hamilton, Reno, NV, USA)

• Syringes fitted with an 18-gauge needle

■ Reagents

• Sterile diluent: modified Bushnell-Haas broth (mineral nutrient; from
Atlas 1993, but modified to contain l/10th strength FeCl 3 ) or Ringer's
solution (Collins et al. 1989)

• Hydrocarbon test substrate: Prepare a solution of non-labeled hydro-
carbon substrate (hexadecane, benzene, phenanthrene, etc.) in acetone
(2 g/L). Then add 14 C-labeled hydrocarbon substrate with sufficient spe-
cific activity to obtain a final radioactivity of about 20 Bq/pL.

• Toluene, acidified by adding HC1: Approximately 5-mL aliquots of toluene
are used in the Harvey trap of the stripping apparatus (Fig. 9.1); add
0.1 mL of 12 N HC1 to 5 mL of toluene placed in the trap.

• Scintillation cocktail (Cytoscint ES; MP Biomedicals, Irvine, CA, USA)
containing PEA to sorb C0 2 . Add 2.5 mL PEA to 7.5 mL Cytoscint and
shake to mix; the PEA cocktail needs to be mixed within about 1 h of use.

• ION NaOH to terminate incubation, and sequester evolved 14 C0 2 in
solution

• 12 N HC1 to release 14 C0 2 for recovery and counting

■ Sample Preparation

Use fresh soil samples. If samples must be stored, refrigerate them following
collection. Sieve soil samples (2-mm mesh) to homogenize.

■ Procedure

Assay Preparation

Soil samples are prepared as a suspension in a sterile aqueous diluent,
determined by the experimental question. Modified Bushnell-Haas broth
is used as diluent if assaying nutrient- optimized mineralization potential
(to assess acclimation of the microbial population to the target substrate).
Ringer's solution is used as diluent if assaying the mineralization potentials

194 J.E. Lindstrom, J.R Braddock

of field-treated soils (e.g., fertilized versus unfertilized). Ringer's solution
may also be amended with macronutrients (N, P), vitamins, or anaerobic
terminal electron acceptors for bench-scale treatability studies.

1. Prepare a nominal 1:10 dilution (w/v) of soil in diluent based on soil wet
mass, preparing a volume sufficient for distribution into several assay
vials. For example, if three or four replicates are desired per sample, add
5 g soil to 45 mL diluent. Collect a portion of the soil sample for a dry
mass determination. The final measured potential will be adjusted per
gram dry mass accordingly.

2. Distribute 10 mL of the soil suspension into VOA vials. Prepare a min-
imum of three replicates for each substrate/treatment combination to
obtain a mean value for the sample's mineralization potential. Securely
replace the caps on the vials to avoid gas leakage during incubation and
C0 2 recovery.

3. Prepare killed controls ("time zero" samples) to be used for subtracting
background radioactivity counts from assay samples. Inject 1 mL ION
NaOH solution through the septum of each control vial. This should
result in a solution pH above 12 in the vial, halting microbial activity.
At least three controls should be prepared for each substrate/treatment
combination, and the mean value is used to "correct" the final result, as
described below.

4. Accurately inject 50p.L radiolabeled substrate solution though the sep-
tum of each vial. Careful measurement is required at this step to assure
reproducibility of the assay. The injection results in addition of 100 pg
substrate to the soil suspension. Briefly swirl or shake the vial to mix,
and incubate under conditions dictated by the experimental design.

5. Sample microbial activity is terminated at the end of the incubation pe-
riod (determined from time-course experiments, described below) by
injecting 1 mL ION NaOH through the septum of each sample vial, as
described for the time zero controls. Swirl the sample vial to distribute
the NaOH. Samples may be stored after treatment with NaOH; the high
pH conditions in the vial sequester the carbon dioxide generated by mi-
crobial mineralization as carbonates in aqueous solution, preventing loss
of C0 2 from the vial pending processing to recover the 14 C-labeled C0 2 .
The samples can be stored for at least a month after this step if necessary.

Recovery of Evolved 14 C0 2

Radiolabeled C0 2 evolved from the soil suspension during the incubation
period is captured by stripping it from solution and capturing it in a basic
medium. PEA is used to trap the C0 2 .

9 Assessment of Hydrocarbon Biodegradation 195

1. Following the incubation period, the soil suspension is acidified by
adding HC1 to release the C0 2 previously sequestered in solution by
addition of NaOH. Inject 1.5 mL 12 N HC1 through the septum into the
VOA vial, and swirl briefly to distribute into solution.

2. Using the apparatus shown in Fig. 9.1, place the two-hole rubber stopper
with glass tubing and Tygon on a scintillation vial containing 1 mL PEA
scintillation cocktail.

3. Place the influent tubing attached to the scintillation vial on the effluent
side of the Harvey trap containing acidified toluene.

4. Attach Tygon tubing to the influent side of the Harvey trap, and attach
a needle to the other end of the Tygon.

5. Pierce the septum of the VOA vial with the needle, making certain the
tip of the needle is above the liquid level in the VOA vial.

6. Pierce the VOA vial septum with the spinal needle attached to a source
of N 2 gas. There should be no gas flow until all connections have been
checked for tightness.

7. Turn on the N 2 gas source and adjust the gas flow rate to approx.
lOmL/min.

8. Strip the C0 2 from the soil suspension for 15min, then stop the gas
flow through the apparatus.

9. Remove the stopper from the scintillation vial, place a cap on the vial,
and determine the amount of radioactivity using a liquid scintillation
counter.

1 0. Rinse all glass tubing tips that contacted scintillation cocktail by dipping
in distilled water several times and wiping clean with a lab wipe; follow
with an acetone rinse.

1 1 . Periodically check for radioactivity carryover between samples by run-
ning method blanks (distilled water in VOA vials) treated as though
they were samples, except without addition of NaOH or HC1.

12. After running each sample, check for clogged needles. Spinal needles
can be cleared with a fine-gauge wire.

Volatile Versus Nonvolatile Substrates

If assaying the mineralization potential of a volatile substrate (e.g., benzene,
toluene, etc.), it is necessary to remove unmetabolized substrate from the
suspension prior to recovering the C0 2 . This is accomplished by bubbling
N 2 gas through the suspension after adding the NaOH, but before adding

196 J.E. Lindstrom, J.R Braddock

the HC1. The C0 2 will still be sequestered in solution in carbonate form, and
will not be lost while volatilizing the substrate from the suspension. After
removing the volatile substrate from the suspension, the C0 2 -stripping
apparatus is assembled, the sample is acidified, and C0 2 recovery proceeds
as described.

Determining Incubation Period

Relatively high concentrations of both labeled and non-labeled substrate
are added to the soil suspensions in this assay to avoid interferences from
field-derived hydrocarbon substrates (Brown et al. 1991). It is therefore
necessary to detect significant substrate mineralization in a reasonable
time frame, while avoiding artifacts associated with in vitro acclimation
of the microbial community assayed. This is accomplished by conducting
time-course assays with samples prepared as described above. A minimum
of three replicate assays should be conducted for each incubation period.

Depending on the substrate chosen, sample incubation times should be
distributed evenly from time zero to the longest reasonable incubation time.
Relatively labile substrates (e.g., linear alkanes up to Ci6, low molecular
weight aromatics up to naphthalenes) may be incubated up to 2 weeks,
with incubations of, e.g., 0, 3, 7, 10, and 14 days. Anaerobic incubations,
more recalcitrant substrates, colder temperatures, etc., may dictate time
courses of longer duration.

Following completion of the time series, plot and inspect the data. Choose
an incubation time longer than the observed lag period, but the shortest
possible time that yields 14 C0 2 recoveries significantly above background
(time zero data).

■ Calculation

The radioactivity recovered from each vial is normalized to a dry soil
basis, using the data from the portion of soil sample collected for dry mass
determination. The mean value of the radioactivity recovered from the time
zero control samples prepared at the beginning of the incubation period
is then determined, and subtracted from the associated treatment samples
to obtain a "corrected" radioactivity value for each vial. Note that 1 g wet
mass of soil is added per vial; thus, each vial represents 1 g wet mass of soil.

v ^(sample) — ^-(time zero controls) , v

^(corrected) — TT - ; v^»W

soil dry mass

X( corrected) sample radioactivity corrected (Bq/g soil dry mass)

^(sample) sample radioactivity recovered as C0 2 (Bq)

9 Assessment of Hydrocarbon Biodegradation 197

■X(time zero controls) mean radioactivity of controls recovered as C0 2 (Bq)
Soil dry mass (g dry soil/g wet soil)

A mean value of radioactivity recovered as 14 C0 2 for each sample can
be calculated from the corrected Bq/g dry soil data. The radioactivity
recovered as 14 C0 2 is then compared to that supplied with the added
labeled substrate. The results maybe expressed as a percentage of substrate
added that was mineralized in the assay by the formula:

^ = ^(corrected) ^ m (93)

^(substrate)

Si substrate mineralized (%/g soil dry mass)

X( corrected) radioactivity corrected (Bq/g soil dry mass)
X( substrate) total radioactivity added to microcosms (Bq)

Alternatively, the data maybe converted to |ig substrate mineralized. The
addition of 50 p.L of the substrate solution (2 g/L) results in 100 pg substrate
being added to the microcosms. The mass of substrate mineralized per
gram dry soil may be calculated by the formula:

So x Si ,

S 2 = 9.4

100

S 2 substrate mineralized (|ig/g soil dry mass)
S initial substrate concentration (100 p.g)

Si substrate mineralized (%/g soil dry mass)

Depending on the experimental question, results among field treatments
maybe assessed for significant treatment effects (using unamended diluent
in the assay). Nutrient-amended assays can be used to demonstrate the
prior acclimation of microbial communities to hydrocarbon degradation,
as nutrient limitations potentially present in the field are removed in the
laboratory assay. Alternatively, a comparison between nutrient-amended
assays and unamended assays, or among various amendments, may be
conducted as a treatability study prior to implementing field treatment.

■ Notes and Points to Watch

• To assure no gas leakage occurs from the stripping apparatus, all Tygon
tubing connections (i.e., to glass tubing, Harvey trap, 16-gauge and spinal
needles) should be secured with several wraps of wire. Tygon tubing can
be protected from being cut by the wire by wrapping the tubing with
a piece of laboratory tape before securing with wire.

198 J.E. Lindstrom, J.R Braddock

• As noted above, it is important to periodically check for obstructions in
the various needles and glass tubing used in the stripping apparatus, and
to clean the glass tubing that comes in contact with scintillation cocktail
to prevent carryover of radioactivity from previously stripped samples.

• Carryover of radioactivity from previous samples run on the stripping
apparatus should be checked periodically by running distilled water
method blanks. If excessive radioactivity (i.e., significantly above back-
ground) is recovered from the method blank, change the toluene in
the Harvey trap, and rerun a method blank. If excessive radioactivity
persists, it may be necessary to change the Tygon tubing.

• When using volatile substrates in the assay, the volatile compounds
require removal prior to recovering the 14 C0 2 , as described. As the
volatile substrate is radioactive, the exhaust gas from this process must
be properly captured (e.g., activated carbon filter) and disposed.

• It is not uncommon to observe substantial variance among samples using
this assay; careful adherence to the protocol will reduce the variance sub-
stantially. We recommend preparing as many replicate assays as possible
in order to obtain a lower standard error for the mean mineralization
potentials determined.

References

Atlas RM (1993) Handbook of microbiological media. CRC Press, Boca Raton, FL
Bauer JE, Capone DG (1988) Effects of co-occurring aromatic hydrocarbons on degrada-
tion of individual polycyclic aromatic hydrocarbons in marine sediment slurries. Appl

Environ Microbiol 54:1649-1655
B0rresen M, Breedveld GB, Rike AG (2003) Assessment of the biodegradation potential of

hydrocarbons in contaminated soil from a permafrost site. Cold Regions Sci Technol

37:137-149
Braddock JF, Lindstrom JE, Prince RC (2003) Weathering of a subarctic oil spill over 25 years:

the Caribou-Poker Creeks Research Watershed experiment. Cold Regions Sci Technol

36:11-23
Braddock JF, Lindstrom JE, Yeager TR, Rasley BT, Brown EJ (1996) Patterns of microbial

activity in oiled and unoiled sediments in Prince William Sound. Proceedings of the

Exxon Valdez Oil Spill Symposium, Feb. 1993. Am Fish Soc Symp 18:94-108
Brown EJ, Resnick SM, Rebstock C, Luong HV, Lindstrom J (1991) UAF radiorespirometric

protocol for assessing hydrocarbon mineralization potential in environmental samples.

Biodegradation 2:121-127
Collins CH, Lyne PM, Grange JM (1989) Collins and Lyne's microbiological methods, 6 th edn,

Butterworths, London
Leahy JG, Colwell RR (1990) Microbial degradation of hydrocarbons in the environment.

Microbiol Rev 54:305-315
Lindstrom JE, Prince RC, Clark JC, Grossman MJ, Yeager TR, Braddock JF, Brown EJ (1991)

Microbial populations and hydrocarbon biodegradation potentials in fertilized shore-

9 Assessment of Hydrocarbon Biodegradation 199

line sediments affected by the T/V Exxon Valdez oil spill. Appl Environ Microbiol
57:2514-2522

National Research Council (1985) Oil in the sea: inputs, fates, and effects. National Academy
Press, Washington, DC

Sugai SF, Schimel JP (1993) Decomposition and biomass incorporation of 14 C-labeled glu-
cose and phenolics in taiga forest floor: effect of substrate quality, successional state,
and season. Soil Biol Biochem 25:1379-1389

Wackett LP, Hershberger CD (2001) Biocatalysis and biodegradation: microbial transfor-
mation of organic compounds. ASM Press, Washington, DC

Walker JD, Colwell RR (1976) Measuring the potential activity of hydrocarbon-degrading
bacteria. Appl Environ Microbiol 31:189-197

Molecular Techniques for Monitoring
and Assessing Soil Bioremediation

Lyle G. Whyte, Charles W. Greer

10.1

General Introduction

Classical culture-dependent microbiological methods have succeeded in
culturing ~1% of the microbial species in a given environmental sample.
In reality, this is due to the fact that most isolation procedures are too gen-
eral, and a wider variety of methods must be developed to recover a larger
representation of microorganisms from most natural environments. Never-
theless, our knowledge of microorganisms is largely based on the represen-
tatives that have been cultured in the laboratory and studied in vitro. Since
approx. 1990, significant advances in molecular biology techniques have
transformed environmental microbiology and microbial ecology. These
techniques bypass the major limitations of culture-dependent microbio-
logical methods by extracting nucleic acids directly (DNA and RNA) from
terrestrial or aquatic samples (soils, waters, wastewaters, etc.) and which
theoretically represent 100% of the microbial species in a given sample.
A variety of techniques are then used to manipulate and subsequently
characterize individual DNA and RNA molecules from complex microbial
communities with a relatively high degree of sensitivity and specificity.
These techniques have been applied to contaminated soil and aquatic sys-
tems and have greatly aided in characterizing and monitoring pollutant
biodegrading microbial populations within these systems. In addition, the
knowledge gained from using these molecular techniques has helped iden-
tify novel biodegradation pathways and opened up new perspectives in
bioremediation processes and pollution abatement. The following survey
presents an overview of the prominent molecular techniques that are cur-
rently being utilized for environmental microbiology with a specific focus
on soil microbiology. The overview is summarized in Fig. 10.1. Several
specific techniques that include total DNA extraction, polymerase chain

Lyle G. Whyte: Dept. of Natural Resource Sciences, McGill University, Macdonald Cam-
pus 21, 111 Lakeshore Road, St. Anne de Bellevue, Quebec, Canada H9X 3V9, E-mail:
whyte@nrs.mcgill.ca

Charles W. Greer: Biotechnology Research Institute, National Research Council of Canada,
6100 Royalmount Ave., Montreal, Quebec, Canada H4P 2R2

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

202 L.G. Whyte, C.W. Greer

reaction (PCR) analyses, and community characterization using denatur-
ing gradient gel electrophoresis, are covered in detail in this chapter.

10.2

Extraction and Purification of Nucleic Acids (DNA) from Soil

■ Introduction

Objectives. All of the current molecular methods crucially rely on the suc-
cessful extraction and purification of sufficient amounts of nucleic acids
from environmental samples. Consequently, many methodologies have
been and continue to be developed for extracting nucleic acids from soils
and sediments, and improvements are constantly being reported in the
literature. The soil-DNA-isolation methodologies vary considerably with
respect to reliability, yield, purity, and degree of shearing, and only recently
have some of these methods been compared (Yeates et al. 1998; Martin-
Laurent et al. 2001; Schneegurt et al. 2003; Kauffmann et al. 2004; Mumy and
Findlay 2004). Several commercial extraction kits are also available such as
MoBio Laboratories Ultraclean Soil DNA Kit (Mobio Laboratories, Solona
Beach, CA, USA) and the BiolOl FastDNA soil kit (La Jolla, CA, USA); they
feature short extraction times and the potential for reduced variability
(Mumy and Findlay 2004) and have become quite popular despite their rel-
atively high cost and limitation to the extraction of nucleic acids from 1 g or
less of sample. In addition, extraction efficiency and resulting nucleic acid
quality are strongly influenced by the source of the sample, and there are
numerous co-extracted interfering substances (humics, pollutants, heavy
metals, etc.). The method described below is routinely used in our lab-
oratories for isolating nucleic acids from soil and sediments from both
contaminated and pristine environments, and has yielded fairly uniform
quantities and qualities of nucleic acids.

Principle. An initial soil washing step prior to DNA extraction helps solu-
bilize and reduce contaminants when high quality DNA is required (Fortin
et al. 2004). Total community DNA is isolated from soil and sediments us-
ing a direct DNA extraction procedure based on chemical/enzymatic lysis
(lysozyme and proteinase K in combination with SDS). Released micro-
bial DNA is ethanol precipitated and the total community DNA is purified
by a polyvinylpolypyrrolidone (PVPP) spin column filtration step which
removes contaminating soil organic compounds such as fulvic and humic
acids (Fortin et al. 2004).

Theory. The starting point in the analysis of total community DNA from
environmental samples is the efficient extraction of nucleic acids of suf-

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation

203

Soil Sample

Extract Nucleic Acids
(total community DNA)

PCR Dependent Analyses

PCR Amplification

16S rRNA PCR Fragments

- Identify, characterize,
monitor complex
microbial populations

Clone Library

Sequence

Identification
of phylotypes

Catabolic Genotypes

- Detect, monitor,
quantitate degradative
genotypes in
contaminated soils

ARDA

DGGE

TGGE

RISA

RAPD

- Sequence
major bands

PCR Independent Analyses

Metagenomics

Metagenomic
Libraries

Clone and sequence
large fragments (BAC
Fosmids, cosmids)

Environmental
Genome Shotgun
Cloning

Clone and sequence
small fragments

Environmental Microarrays

Phylogenetic, functional gene
microarrays to identify,
characterize, monitor complex
microbial populations

Fig. 10.1. Molecular techniques used in environmental microbiology

ficient quality for subsequent molecular analyses. Two general method-
ologies are commonly employed: (1) direct lysis of microbial cells within
the environmental sample by chemical treatments, sonication, freeze-thaw,
or bead beating protocols; or (2) extraction of the bacterial cells from
the environmental sample followed by cell lysis. Humic and fulvic acids
are often co-extracted from soil with the nucleic acids; they must be re-
moved as they can seriously interfere with subsequent molecular reactions
(DNA polymerase amplification, DNA-DNA hybridizations, DNA labeling,
restriction nuclease digestion). The released soil DNA usually can be puri-

204 L.G. Whyte, C.W. Greer

fied by a variety of methods/techniques such as chromatography and silica
gel or PVPP spin-filter columns. The quality and quantity of the purified
soil DNA extract is generally verified by agarose gel electrophoresis and/or
spectrophotometry (Abs. 260/280 nm).

■ Equipment

Water bath incubators (30 °C, 37 °C, 85 °C)

Microcentrifuge (13,600-15,800 g) at4°C

Platform shaker

Speedvacuum (optional)

Gel electrophoresis apparatus, ultraviolet light source, and camera or gel
documentation system

Spectrophotometer, quartz cuvettes

Vortex

Pipettors (10-20, 100, 1,000 pL) and tips

Microcentrifuge tubes (0.5, 1.5, 2.0 mL)

MicroSpin columns (Amersham Biosciences, Baie d'Urfe, Que., Canada)

Reagents

Buffer 1: 50 mM Tris-HCl, pH 8.3, containing 200 mM NaCl, 5 mM EDTA,
and 0.05% Triton X-100 (Fisher Scientific, Nepean, Ont. Canada)

Buffer 2: 50 mM Tris-HCl, pH 8.3, containing 200 mM NaCl and 5mM
EDTA

Buffer 3: 10 mM Tris-HCl, pH 8.3, containing 0.1 mM EDTA

Distilled water

250 mM Tris-HCl, pH 8.0, with 5 mg/mL lysozyme (freshly prepared)

Proteinase K (20 mg/mL)

20% SDS

7.5 M ammonium acetate

2-propanol

70% ethanol

TE buffer: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 205

• TAE buffer: 40 mM Tris-acetate, 1 mM EDTA, pH 8.0

• Acid-washed PVPP spin columns (preparation below)

• 200 mM potassium phosphate buffer, pH 7.0

• Cone. HC1

Note: All solutions should be sterilized by filtration or autoclave. The recipes
for many of the solutions described throughout this chapter can be found
in Sambrook and Russell (2001).

■ Sample Preparation

Soil samples to be extracted should be frozen (-20 or -80 °C) as soon as
possible upon collection and stored frozen to minimize changes in micro-
bial communities because of the sampling process and/or degradation of
nucleic acids.

■ Procedure

Soil-Washing Step

1. Add 1 mL of buffer 1 to 0.5 g of soil or sediment in a microcentrifuge
tube. Mix by vortexing (speed 4) for 30 s, then by inverting for 1.5 min.

2. Centrifuge 5 min at 4 °C at 3,000 g. Remove the supernatant with a pipette.

3. Add 1 mL of buffer 2 to sediment. Mix by vortexing (speed 4) for 30 s,
then by inverting for 1.5 min.

4. Centrifuge 5 min at 4 °C at 3,000 g. Remove the supernatant with a pipette.

5. Add 1 mL of buffer 3 to sediment. Mix by vortexing (speed 4) for 30 s,
then by inverting for 1.5 min.

6. Centrifuge 5 min at 4 ° C at 3,000 g. Remove the supernatant with a pipette.

Nucleic Acid Extraction Step

1. Add 450 p.L of sterilized distilled water to the 0.5 g of sample in a 1.5 mL
microcentrifuge tube. Vortex (moderate speed) for approx. 2 s to dis-
lodge the pellet.

2. Add 50 pL of 250 mM Tris-HCl, pH 8.0, containing lysozyme (5 mg/mL).

3. Incubate at 30 °C, mixing by inversion, for 30 min.

4. Transfer to a 37 °C water bath and incubate for another 30 min, mixing
by inversion every 10 min.

206 L.G. Whyte, C.W. Greer

5. Add 5 pL of proteinase K (20 mg/mL). Incubate for 1 h at 37 °C, mixing
by inversion every 10 min.

6. Add 50 pi of 20% SDS. Incubate at 85 °C for 30 min, mixing gently by
inversion every 10 min.

7. Centrifuge 10 min at room temperature at 13,600 g. Transfer the super-
natant to a fresh 1.5 mL microcentrifuge tube.

8. Add 1/2 volume of 7.5 M ammonium acetate. Mix gently by inversion,
and incubate on ice for 15 min.

9. Centrifuge 5 min at 4 °C, at 13,600 g. Transfer and split the supernatant
in two fresh 1.5 mL microcentrifuge tubes and treat each tube sepa-
rately.

10. Add 1 volume of cold 2-propanol, and precipitate DNA overnight at
-20 °C.

11. Centrifuge 15 min at 4 °C at 15,800 g. Discard the supernatant.

12. Wash the pellet with 500 pL of cold 70% ethanol. Gently tap the tube or
mix by inversion.

13. Centrifuge 5 min at 4 °C at 15,800 g. Discard the supernatant.

14. Dry the pellet in speedvacuum for 5 min or air dry the pellet (approx.
30 min).

15. Add 100 pi of TE buffer to the pellet in each tube, place the sample
on ice on a shaking platform and let the pellet slowly dissolve (approx.
1 h). Combine the DNA extracts from the two tubes.

16. Warm up the DNA combined extract for 10 min at 37 °C.

17. Purify 50 pL of total community DNA on a PVPP column.

DNA Purification with PVPP Columns (Modified from Berthelet et al. 1996)

1. Prepare acid washed PVPP. Pour 1,034 mL of cone. HC1 (11.6M) slowly
with stirring into 2,966 mL of MilliQ (Qiagen Inc., Hilden, Germany)
water; this will result in ca. 4 L of 3 M HC1. Add 150 g PVPP and suspend
with stirring at room temperature for 12-16 h. Leave the suspension
to settle for 30-60 min, then aspirate or decant the supernatant. Again
suspend the PVPP, now in approx. 3.5 L of 200 mM potassium phosphate
buffer (pH 7.0) and stir 1-2 h. Repeat the aspiration/decant and suspen-
sion twice more until the supernatant pH is close to 7.0 (check aliquot
with pH meter). Then repeat the aspiration/decant and suspension two
more times with approx. 3.5 L of 20 mM potassium phosphate buffer

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 207

(pH 7.0). Aliquot the final suspension into small bottles and autoclave
(15-20 min, 121 °C). Store at 4°C.

2. Acid washed PVPP (0.9 mL) slurried in 20 mM potassium phosphate
(pH 7.0) is added to empty sterile MicroSpin columns placed inside 2.0-
mL tubes and centrifuged for 3 min at 735 g at room temperature. If the
top of the column is still immersed following centrifugation, remove the
liquid from the collection tube, and re-spin the column.

3. Load the "crude" DNA extract (50-100 p.L) onto the center of the column
being careful not to touch the side of the column. This ensures that all
of the sample will pass through the column and be cleaned, and not run
down the side of the column.

4. Place the loaded columns in the microcentrifuge, ensuring that the slop-
ing face of the packed column is facing the middle of the centrifuge.
Centrifuge the columns for 3 min at 735 g at room temperature and
collect the filtrate.

5. The "clean" DNA extract is then stored at -20 °C and is ready for PCR.
The used PVPP is discarded and the MicroSpin columns washed for
reuse.

Agarose Gel Electrophoresis

We check the quality and quantity of ca. 5 pL of purified soil DNA extract
by both agarose gel electrophoresis in 0.7% gels in TAE buffer (stained with
ethidium bromide and visualized by ultraviolet light) and spectrophotom-
etry (Abs. 260/280 nm) using standard methods as described by Sambrook
and Russell (2001).

Notes and Points to Watch

This methodology can be readily scaled up to 10 g soil samples as de-
scribed in Fortin et al. (2004).

All molecular biology methodologies are notoriously variable. Incuba-
tion temperatures and durations, volumes, centrifugation parameters,
etc. should be vigorously adhered to.

Lysis of the microbial cells during DNA extraction represents a critical
step in PCR-mediated approaches (von Wintzingerode et al. 1997). Each
physical, chemical, and biological step involved in the preparation and
analysis of an environmental sample is a source of bias which might give
a distorted view of a given ecological niche (von Wintzingerode et al.
1997). It is often a question of whether there was sufficient or preferential

208 L.G. Whyte, C.W. Greer

disruption of microbial cells. Rigorous conditions maybe required to lyse
Gram-positive cells but also may cause excessive shearing of nucleic acids
of the Gram-negative cells, potentially biasing the reported diversity
of the sample as well as possibly creating artifacts and chimeric PCR
products (Liesack et al. 1991). Therefore, checking the soil DNA extract
by agarose gel electrophoresis will indicate the quality of the extracted
DNA and the extent of shearing.

• Quantification of the soil DNA extract by spectrophotometry is often
inaccurate and does not appear to correlate with other methods such as
agarose gel electrophoresis using known concentrations of DNA stan-
dards or PicoGreen (Molecular Probes, Leiden, Netherlands).

10.3

Amplification of Catabolic Genotypes

and 16S rDNA Genotypes by PCR

■ Introduction

Objectives. Many of the molecular methodologies used in environmental
microbiology rely on a PCR amplification step and are therefore considered
PCR dependent. The objective of PCR is to amplify target gene sequences
from total community DNA extracted from an environmental sample. The
amplified sequences can then be characterized by a variety of molecular
methodologies as shown in Fig. 10.1. In soil biodegradation studies, the
target gene sequences can be either catabolic (biodegradative) genes of in-
terest or a phylogenetic gene (almost always the 16S rDNA gene). In the case
of catabolic genes, the PCR amplification step is generally used to detect
the presence or absence of various catabolic genotypes in contaminated
soils. Determining the prevalence and composition of specific biodegrada-
tive genotypes, and hence microbial populations, in contaminated soils
significantly aids in assessing the feasibility of using biotreatment and in
developing appropriate bioremediation strategies for a particular contam-
inated site, as well as in monitoring the effects on specific populations
during bioremediation operations. For example, we routinely use PCR
screening for hydrocarbon-degradative genotypes to perform biotreatabil-
ity assessments of contaminated soils (Whyte et al. 1999; Soloway et al. 200 1 ;
Whyte et al. 2001) and to monitor bioremediation treatments (Whyte et al.
2003). The prevalence of various alkane monooxygenase genotypes and
other degradative genotypes in hydrocarbon-contaminated and pristine
soils from a variety of Arctic, Antarctic, and alpine contaminated soils was
also determined by PCR screening (Whyte et al. 2002; Margesin et al. 2003;
Luz et al. 2004).

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 209

In comparison, PCR amplification is currently the most widely used
method to obtain 16S rDNA genotypes for detailed characterization of
microbial communities. Unlike PCR amplification of degradative geno-
types, however, PCR amplified 16S rDNA genes from a total commu-
nity DNA extract must be further characterized by one or more of the
molecular methodologies [clone and sequencing, denaturing gradient gel
electrophoresis (DGGE; see Sect. 10.4, below), temperature gradient gel
electrophoresis (TGGE), etc.], shown in Fig. 10.1, to obtain meaningful
information on the amplified 16S rDNA PCR product. These molecular
methods have been used to characterize cold-adapted microbial popula-
tions in hydrocarbon-contaminated soils originating in northern Canada
( Juck et al. 2000), to determine the effect of oil contamination and a biostim-
ulation treatment on Pseudomonas diversity in soil microcosms (Evans et
al. 2004), to monitor microbial population changes in beach sediments dur-
ing an experimental oil spill (Macnaughton et al. 1999), and to monitor the
impact on microbial community composition during the bioremediation
of hydrocarbon-contaminated soils (Mills et al. 2003).

Principle. During PCR, double-stranded DNA (from total community DNA
soil extracts) is separated into single strands at high temperature (denat-
uration). Two oligonucleotide primers then anneal (at a lower annealing
temperature) to complementary regions (which flank the target sequence)
of the single-stranded DNA. A heat-stable DNA polymerase synthesizes
a new strand of DNA by extending the primer using the complementary
strand as a template, thus creating a duplicate copy of the target sequence.
This cycle is repeated 20-30 times resulting in an exponential amplification
(2 20 -2 30 fold) of the target sequence.

Theory. PCR is the simplest and currently the most widely used method
to detect/obtain catabolic genotypes or 16S rDNA genotypes for detailed
downstream characterization of soil microbial communities. These proce-
dures are increasingly being utilized to perform biotreatability assessments
of contaminated soils, to monitor the effects of soil bioremediation treat-
ments on microbial populations, and to identify and characterize important
and/or novel biodegradative microbial strains or groups of microorganisms
in contaminated soils. This Section describes PCR procedures for the am-
plification of catabolic genotypes and 16S rDNA genes for cloning and
sequencing.

Equipment

PCR work station chamber (UV hood; optional but recommended for
16S rDNA PCR)

210 L.G. Whyte, C.W. Greer

• Microcentrifuge

• Thin walled 0.2-mL PCR tubes

• Pipettes (10, 100 pL) and tips (filtered tips are recommended for 16S
rDNA PCR)

• PCR thermocycler

• Gel electrophoresis apparatus, ultraviolet light source, camera or gel
documentation system

• PCR cleanup kit (QIAquick PCR Purification Kit; Qiagen Inc.)

• PCR cloning kit (Promega pGem-T Easy Cloning Kit; Promega Corpo-
ration, Madison, WI, USA)

• Petri dishes

• Incubator (37 °C)

• Water bath incubators (37, 42 °C)

■ Reagents

Catabolic Genotype PCR Amplification

• Catabolic target gene forward and reverse oligonucleotide primers (for-
ward and reverse; 0.4-0.8 mM stock solutions)

• Soil DNA extract

• DNA from reference organisms

• 100 bp DNA ladder (Fermentas, SM0241, Invitrogen, Carlsbad, CA, USA)

• DNA polymerase (Taq DNA polymerase is often used)

• DNA polymerase buffer: 10 mM Tris-HCl, pH 9.0, containing 50 mM KC1
and 15mMMgCl 2

• 25 mM MgCl 2

• 1.25 mM stock solution of each deoxynucleoside triphosphate (dNTP),
namely dATP, dCTP, dGTP, dTTP

16S rDNA PCR Amplification

• General ("universal") Bacteria primers:

- 27F(10pM)5 / -GGTTACCTTGTTACGACTT

- 758R (10 pM) y-CTACCAGGGTATCTAATCC

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 211

Soil DNA extract

DNA from reference organisms (control DNA)

100 bp DNA ladder (Fermentas, SM0241)

Taq DNA polymerase (5 U/pL; Invitrogen)

10 x PCR buffer: 200 mM Tris-HCl/500 mM KC1

50 mM MgCl 2

10 mM stock solution of each dNTP (dATP, dCTP, dGTP, dTTP)

Bovine serum albumin (BSA; 10 mg/mL in sterile UV irradiated ddH 2 0)

Sterile UV irradiated H 2

Store all PCR reagents at -20 °C.

Cloning and Sequencing of 16S rDNA PCR Amplicons: Transformation

• Lac~ competent E. coli cells (DH5a)

• Isopropyl-B-D-thiogalactopyranoside (IPTG) solution: 0.1 M, filter ster-
ilized (store at 4 °C)

• Ampicillin (Amp) solution: 10 mg/mL, filter sterilized (store at 4°C)

• 5-bromo-4-chloro-3-indoyl-/J-d-galactoside (X-Gal) solution: 50 mg/mL
in N, N'-dimethyl-formamide, make fresh for each transformation (store
at-20°C)

• Luria broth (LB) medium (per L): 10 g tryptone, 5g yeast extract, 5g
NaCl, adjust to pH 7.0 with NaOH

• LB plates with Amp/IPTG/X-Gal: To 1 L of autoclaved LB medium, add
5 ml of IPTG solution, 10 ml of ampicillin solution, and 1.6 mL of X-Gal
solution.

Note: All solutions should be sterilized by filtration or autoclave. The recipes
for many of the solutions described throughout this chapter can be found
in Sambrook and Russell (2001).

■ Sample Preparation

Total community DNA is extracted and purified as described in Sect. 10.2.
The soil DNA extracts should always be stored frozen at -20 °C or -80 °C
and kept on ice during the procedure to minimize nucleic acid degradation,
which will occur at greater rates at higher temperatures.

212 L.G. Whyte, C.W. Greer

■ Procedure

Primer Design

An important key to PCR is optimal design of oligonucleotide primers
specific to the desired gene target of interest. It is the specificity of the
primers that allows PCR to amplify catabolic or 16S rDNA genes that can
be in low abundance in complex environmental samples.

Catabolic Gene PCR Primers

We have designed and utilized a variety of primers for PCR amplification
of catabolic genes; the specific oligonucleotide sequences of these primers
are available for a variety of hydrocarbon degradative genes (Whyte et al.
2002; Margesin et al. 2003; Luz et al. 2004) and dehalogenation of chlori-
nated organics (Fortin et al. 1998). In general, we design PCR primers as
follows. Gene sequences for key enzymes from known bacterial biodegrada-
tive pathways are identified and searched for in databases such as the nu-
cleotide database (GenBank) at the NCBI web site (www.ncbi.nlm.nih.gov).
The DNA sequences of all corresponding genes encoding the key enzyme
are retrieved for comparative DNA and protein alignments using appropri-
ate molecular biology sequence software. PCR forward and reverse primers
for each catabolic gene sequence are then selected from the alignments by
PCR primer software and/or manually by identifying homologous regions
shared by the selected DNA sequences. For construction of the oligonu-
cleotide primers, long sequences originating from within the coding region
of the catabolic genes and having a high G+C content are preferred to ensure
specificity. In addition, we use the following general criteria for designing
catabolic gene primers:

- Generally 20-30 nt in length.

- At least 5 nt at both ends of the primer exhibiting exact match pairing
with the target DNA sequence,

- Ideally, if there are mismatches, they should be in the middle of the
primer sequence.

- Ideally, the ends of the primer should terminate with 2-3 G or C.

- Both forward and reverse primers should possess similar G+C content,
with an average G+C content of 40-60%, with limited stretches of poly-
purines or polypyrimidines to ensure specificity.

- The PCR products generated should be ca. 200- 1,000 nt in length and
thus produce an easily detectable band by agarose gel electrophoresis.

Specificity of the selected primer sequences for the gene of interest is
then verified by Fasta and blastn search programs available at the NCBI
website.

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 213

16S rDNA PCR Primers

Given the varying degrees of conservation of thel6S rDNA gene for Bacteria
and Archaea, 16S rDNA primers or probes can be designed with any degree
of specificity for groups, ranging from both domains (Ward et al. 1992),
to a single domain (Battin et al. 2001), or to various subgroups all the way
down to the species and sub-species level (Ahn et al. 2002). Generally, most
16S rDNA PCR-based studies rely on using a set of general or "universal"
primers specific for the Bacterial domain and/or less often, the Archaeal
domain. These general primer sets are readily available in the literature but
should be updated periodically as the 16S rDNA database grows daily. One
can also design new conserved 16S rDNA primer sets by accessing the Ribo-
somal Database Project (RDP) website (http://rdp.cme.msu.edu/index.jsp)
algorithm that can be used in choosing the proper primer set for the PCR
amplification in question.

PCR Amplification, Cloning, and Sequencing
PCR Amplification of Catabolic Genotypes

1. For each PCR reaction, set up the following reaction in a 0.2-mL micro-
centrifuge tube:

- 1-5 pi of total community DNA soil extract (ca. lOOngofDNA)

- 2 pL of each oligonucleotide primer (0.4-0.8 mM stock; final concen-
tration 0.2 pM)

- 5 pL of DNA polymerase buffer

- 2pLof25mMMgCl 2

- 8 pL of 1.25 mM dNTP (200 pM of each dATP, dCTP, dGTP, and dTTP
solution)

- 2.0-5.0 U of DNA polymerase

2. The final volume in the tube is brought to 50 pL with sterile distilled
water. Prior to the addition of DNA polymerase, the samples are boiled
for 2 min and then transferred to ice.

3. Negative control: The same mixture is used for the negative control
except that the total community DNA soil extract is replaced with sterile
distilled water.

Positive Control: The same mixture is used for the positive control except
that the total community DNA soil extract is replaced with a genomic
DNA extract from the appropriate control reference organism. Template
DNA for PCR from the reference organism can be obtained by resus-
pending 2-3 colonies in 500 pL of sterile distilled water and boiling for

214 L.G. Whyte, C.W. Greer

10 min. The sample is cooled on ice, centrifuged in a microcentrifuge for
2 min at 12,000 g, the supernatant collected, and stored at -20 °C.

4. PCR is conducted using an appropriate PCR thermal cycler. We generally
are successful using the following PCR parameters:

- 30 cycles of 1 min at 94 °C (denature)

- 1 min at 60 °C (anneal)

- 1 min at 72 °C (extend)

- A final extension of 3 min at 72 °C.

5. To determine the presence or absence of the appropriately sized PCR
fragment, ca. 5-10 pL of the PCR reaction mixture of soil DNA extracts
and the corresponding positive and negative controls, and a 100 bp DNA
ladder are analysed by agarose gel electrophoresis (1-1.4% agarose gels
using TAE buffer) and visualized by ethidium bromide staining essen-
tially as described by Sambrook and Russell (2001). There should be
a single band of the same size in both the positive and sample lanes, with
no band in the negative control lane.

6. To confirm that DNA had been successfully extracted from the soils and
could be amplified by PCR, general ("universal") 16S rDNA bacterial
primers are used as a positive PCR amplification control for all soil DNA
extracts (Whyte et al. 2002).

PCR Amplification of 16S rRNA for Downstream Cloning and Sequencing

The protocol given is to PCR amplify 16S rDNA from an environmental
sample.

1. All subsequent work should be conducted in an enclosed PCR work
station chamber. This will help eliminate contamination of plastic ware
with extraneous 16S rDNA that is present ubiquitously. Latex gloves
should be worn throughout the procedure.

2. A PCR master mix (enough for three PCR reactions) is prepared in
a 1.5 mL microcentrifuge tube by combining the following reagents:

- 7.5 pL of each oligonucleotide primer (10 pM stock; final concentra-
tion 0.5 pM)

- lpLoflOx PCR buffer

- 4.5 pL of 50 mM MgCl 2 (final concentration 1.5 mM)

- 3pL of 10 mM dNTP stock solution (final concentration 200 pM of
each dATP, dCTP, dGTP, and dTTP)

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 215

- 1.9pLofBSA(10mg/mL)

- 92.6 pL of sterile, irradiated water

3. Briefly microcentrifuge the PCR master mix tube at 15,000 g for ca. 10 s.

4. Add 3 pL of Taq polymerase to the PCR master mix tube, briefly vortex
(2-4 s) to mix and microcentrifuge at 15,000 g for ca. 10 s.

5. Label the following 0.2 mL PCR reaction tubes: positive control, negative
control, sample. To each reaction tube add 45 pL of master mix.

6. Add 5 pL of E. coli (or other positive control) genomic DNA (ca. 50-
100 ng) to the positive control. To the negative control, add 5 pi of water.
To the sample tube, add 1-5 pL of total community DNA soil extract (ca.
100 ng of DNA).

7. Briefly vortex (2-4 s) the PCR reaction tubes to mix. Microcentrifuge the
tubes at 15,000 g for ca. 10 s.

8. PCR is conducted in an appropriate thermal cycler. We are generally
successful using the following PCR parameters:

The first 10 cycles are conducted using a "touchdown protocol" from
65-55 °C, with the annealing temperature decreasing by 1 °C at each
cycle.

- lminat94°C

- lmin at 65-55 °C

- 3minat72°C

The subsequent 20 cycles are performed with an annealing tempera-
ture of 55° C.

9. The presence or absence of the appropriately sized PCR fragment (73 1 bp
for Bacteria 16S rDNA) is determined by agarose gel electrophoresis
(0.8%) of 5 pL of the PCR reaction mixture of soil DNA extracts, posi-
tive and negative controls, and a 100 bp DNA ladder, and visualized by
ethidium bromide staining essentially as described by Sambrook and
Russell (2001). There should be a single band of the same size in both
the positive and sample lanes, with no band in the negative control lane.

Cloning and Sequencing of 16S rDNA PCR Products

1 . We generally use a spin column purification system such as the Qiaquick
PCR purification kit to clean the PCR reaction prior to cloning. It is
important to purify PCR reaction products as unbound primers and
unincorporated nucleotides can be inhibitory to the ligation reaction.

2. The purified PCR product is quantified by spectrophotometry (Abs.
260/280 nm; Sambrook and Russell 2001).

216 L.G. Whyte, C.W. Greer

3. Because of their ease of use and reliability, commercial PCR cloning
kits, such as the pGEM-T Easy Vector (Promega Corp.), are commonly
used for ligating 1 6S rDNA PCR amplicon libraries into a cloning vector.
A ligation reaction is set up as described in the pGEM-T Easy Vector
technical manual. If this is a first-time attempt at cloning with this
PCR product, it may be necessary to optimize the vectoninsert molar
ratio. We generally optimize the reaction with 1:3, 1:1, and 3:1 ratios.
To calculate the amount of PCR product to include in the reaction use
the following formula:

ng insert

(50 ng of vector) x (size of PCR product) x (insert:vector ratio)
kb size of vector (3.0 kb for pGEM-T Easy)

4. Incubate the ligation reaction at 4 °C overnight.

5. Transformation protocols of E. coli competent cells with recombinant
vector (16S rDNA inserted into the pGEM-T Easy Vector) can be found
in the pGEM-T Easy technical manual or Sambrook and Russell (2001).
Competent cells can be provided with the pGEM-T Easy Vector system;
we have had success using E. coli DH5a (made chemically competent
as described by Sambrook and Russell 2001). We generally follow the
protocol provided in the manual. Always ensure that positive and neg-
ative controls are included in the analysis. The positive control consists
of cells transformed with the vector DNA alone; the negative control
cells are treated the same as cells being transformed, but with no added
DNA.

6. Spread plate 100 p.L of each transformation (in duplicate) and controls
onto appropriately labeled LB/Amp/X-Gal/IPTG plates.

7. Incubate plates overnight at 37 °C.

8. Score blue and white colonies. White colonies arise from insertion of
a cloned product into the pGEM-T Easy Vector. More than 60% white
colonies should be observed.

9. White colonies are either directly sequenced or screened for unique
clones prior to sequencing by amplified ribosomal DNA restriction
analysis (ARDRA; sometimes called restriction fragment length poly-
morphism, RFLP; see Massol-Deya et al. 1997 for a typical ARDRA
protocol for 16S rDNA amplicons). Sequencing of the 16S rDNA in-
serts in the pGEM-T Easy Vector system is conducted with primers as
described by that system's technical manual; ensure that when ream-
plifying the cloned inserts to use the pGemT-Easy primers T7 and Sp6

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 217

to avoid amplifying E. coli 16S rDNA genes. Sequencing is usually per-
formed by commercial laboratories or by in-house sequencing facilities
commonly found in most large research institutions.

10. Sequences are submitted for comparison and identification to the Gen-
Bank databases using the NCBI Blastn algorithm, the EMBL databases
using the Fasta algorithm (http://www.ebi.ac.uk/fasta33/nucleotide.
htmL) and/or the Ribosomal Database Project (RDP) using its Sequence
Match. Sequences that demonstrate strong homology are then aligned
to reference sequences and phylogenetic trees commonly constructed
( Juck et al. 2000). Sequences that demonstrate uncertain alignments are
checked for chimeras using the CHECK_CHIMERA software program
function at the RDP site.

Notes and Points to Watch

A key limitation to 16S rDNA PCR amplifications is contamination of
DNA introduced by unintentional tube-to-tube contamination or con-
taminated reagents. For this reason, false-positive signals and false-
negative amplifications are not uncommon due to the extreme sensitivity
of the 16S rDNA PCR reaction, and the ubiquity of 16S rDNA genes in
almost all biological materials. Fortunately, this problem can be avoided
simply by using good laboratory techniques as indicated above.

We often perform PCR amplification on both undiluted soil DNA extracts
(as described here) and diluted extracts (1/10, 1/100). Diluting the DNA
extract can result in the parallel dilution of undesired contaminants that
inhibit the PCR reaction; it is not uncommon to observe successful PCR
amplification from the diluted samples but not the undiluted sample.

To minimize the loss of nucleic acids from small sample volumes, ad-
ditives such as BSA and T4 gene 32 (gp32) can be used to reduce the
inhibitory effect of contaminants (Kreader 1996).

The PCR procedures described here should be considered qualitative
rather than quantitative. Differences in band intensity do suggest dif-
ferences in the relative amounts of the genotypes in the original sam-
ples, but keep in mind that PCR reactions are very sensitive to reaction
conditions. Quantitative PCR protocols (real-time quantitative PCR or
RT-qPCR) have been recently developed and are being applied to con-
taminated soils.

Very similar nucleic acid sequences can also affect amplification of to-
tal community DNA, especially during 16S rDNA PCR amplifications.
Chimeric sequences result from the heterologous combination of two

218 L.G. Whyte, C.W. Greer

non-identical but similar strands of DNA, but do not generally exist
in the sample being investigated. However, chimeric sequences can be
formed at frequencies of several percent during PCR (Liesack et al. 1991).
The resultant PCR artifacts can affect subsequent analyses by erroneously
suggesting the existence of novel taxa from these hybrid sequences. The
binding of heterologous DNA into chimeric structures has also been
shown to compete with the binding of specific primers during the an-
nealing step (Meyerhans et al. 1990; Ford et al. 1994; Wang and Wang
1996). As well, DNA damage such as that caused by mechanical and
chemical shearing has been suggested to contribute to the formation of
chimeric DNA during PCR (Paabo et al. 1990).

• Another pitfall of PCR is the production of minute errors by Taq DNA
polymerase, which lacks the ability to proofread. (Ford et al. 1994).
However, this is only a potential problem when sequencing the resulting
PCR products.

• For cloning into the pGEM-T Easy Vector system, it is essential to use
a thermostable polymerase that lacks 3 / -5 / exonuclease activity in the
initial 16S rDNA amplification step of soil DNA extracts. This will in-
sure that a 3'A overhang is present on the PCR product and will greatly
improve the efficiency of the ligation process, as well as avoiding cir-
cularization of PCR products. Common polymerases that lack the 3 f -5 f
exonuclease activity are Taq, Tfl, and Tth.

10.4

DGGE Analysis Soil Microbial Communities

■ Introduction

Objectives. Denaturing gradient gel electrophoresis (DGGE) is a very ver-
satile method for screening the total microbial community DNA from
a complex sample. Our limited knowledge of the total microbial com-
munity composition and function in complex environmental samples has
necessitated the development of techniques like DGGE to enable us to look
more directly at the representative microorganisms, independent of the bi-
ases introduced by culturing. In the last 10 years, more than 1,000 articles
have been published using DGGE for the analysis of various environmental
samples.

DGGE analysis of microbial communities produces a complex profile
or banding pattern, which can be quite sensitive to spatial and temporal
sampling variations (Murray et al. 1998). The classic means of analyzing this
variability has been visual, reporting differences between samples in band

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 219

intensity, or the presence or absence of specific bands. However, a recent
study suggests that the results of denaturing gradient methods are readily
amenable to statistical analysis, provided there is sufficient standardization
of analytical procedures (Fromin et al. 2002). This would provide the rigor
of statistical validation of observations and permit a broader range of
comparisons to be made between different samples and between different
experimental or environmental parameters.

DGGE is a useful method for visualizing the major members of a micro-
bial community, but several factors must be considered when interpreting
the data. The limit of resolution of this method is about 1% of the total
community population (Muyzer et al. 1993; Murray et al. 1996), and in very
complex samples, more bands may be produced than can be resolved. Ini-
tial calibration to ensure optimal gradient and electrophoretic conditions
is also important (Muyzer et al. 1993; Muyzer and Smalla 1998). DGGE
requires rather large quantities of DNA for reliable visualization, possibly
as much as 500 ng for environmental samples (Nakagawa and Fukui 2002).
Also, DGGE is typically limited to fragments of no more than 500 bp (My-
ers et al. 1985), which limits the amount of sequence information that can
ultimately be retrieved. Some ambiguity can exist in associating a single
band in a DGGE profile with a single microbial species, since it is possible
that multiple amplicons co-migrate to the same location in the gel, and
similarly, multiple bands may be produced by a single species since multi-
ple copies of 16S rDNA do exist in the same microorganism (Nubel et al.
1997).

Principle. DGGE separates a mixture of PCR-amplified DNA fragments
according to differences in sequence G-C content, based on their differential
mobility through a DNA-denaturing gel. Once separated, the individual
fragments can be recovered from the gel and the nucleotide sequences
determined and compared against existing databases (GenBank, Ribosome
Database Project) to identity microorganisms in the sample.

Theory. DGGE, which is based on the early work of Fischer and Lerman
(1979), is one of the most commonly used methods for the characterization
of complex microbial communities, and was pioneered by Muyzer et al.
(1993) for environmental samples. In a manner similar to the other PCR-
based characterization techniques, samples for DGGE analysis are prepared
either directly from PCR-amplified environmental DNA (Ahn et al. 2002;
Ibekwe et al. 2002), from clone libraries constructed from PCR-amplified
environmental samples (Liu et al. 2002), or in some cases from colonies
obtained from enrichment cultures (Bonin et al. 2002). Total community
DNA is extracted, purified and used as a PCR template for the amplification
of specific target molecules. The most common target molecule is the 16S
rDNA gene which is used as a phylogenetic marker to assess biodiversity

220 L.G. Whyte, C.W. Greer

and eventually to identify individual members within the community. Gen-
eral 16S rDNA primers, often referred to as universal primers, are used to
amplify the total community DNA. This produces a mixture of fragments
derived from the individual microorganisms in the sample. Because each
fragment has a different internal sequence, the fragments can subsequently
be separated based on their melting behavior in a denaturing gradient, usu-
ally composed of urea and formamide. As the double-stranded PCR frag-
ments move through the gel from low to high denaturant concentration,
they begin to separate into single strands, which reduces their mobility.
Complete strand separation is prevented by incorporating a GC rich region
(ca. 40 bases), referred to as a GC clamp, at the 5'-end of one of the PCR
primers. The DNA comes to rest when it is almost fully denatured. The
position along the gradient at which the DNA stops is determined primar-
ily by the relative proportions of G+C and A+T in a given amplicon, since
G-C bonds are more difficult to denature than A-T bonds. Properly cali-
brated, DGGE is sensitive enough to detect even single base-pair differences
between amplicons (Miller et al. 1999). The result in complex samples is
typically a banding pattern that is representative of the molecular diversity
in the sample. The individual bands can subsequently be extracted from
the DGGE gel and sequenced to potentially identify individual microor-
ganisms.

■ Equipment

• See Sect. 10.3 for equipment for PCR amplification

• Gradient mixer (BioRad Model 385 Gradient Former, BioRad Laborato-
ries Inc., Mississauga, Ont. Canada)

• BioRad Dcode Universal Mutation Detection System (BioRad Labs.; or
equivalent)

• Fluorlmager system, model 595 (Molecular Dynamics Inc., Sunnyvale,
CA, USA; or equivalent)

• PCR clean up kit (QIAquick PCR Purification Kit, Qiagen Inc.; containing
PB, EB, PE buffer, and column- collection tubes)

Reagents

50 x TAE (per L): 242 g Tris base, 57.1 mL glacial acetic acid, 100 mL 0.5 M
EDTA, pH 8.0. To prepare 1 x TAE, dilute 1:50 with distilled water.

Acrylamide-denaturant solutions: the acrylamide solutions are only sta-
ble for 1 month. All glassware should be rinsed with ultrapure water.

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 221

- 8% acrylamide/0% denaturant: To make 100 mL of solution, mix
20 mL of 40% Acrylamide/Bisacrylamide (37.5:1; BioRad); 2mL of
50 x TAE buffer prepared with ultrapure water and ultrapure reagents,
and 78 mL ultrapure water. Filter through a 0.22-|im filter. Mix and
degas for 10-15 min. Store at 4°C in a brown bottle for approx. 1
month.

- 8% acrylamide/80% denaturant: To make 100 mL of solution mix
20 mL of 40% acrylamide/bisacrylamide (37.5:1); 2mL of 50 x TAE
buffer prepared with ultrapure water and ultrapure reagents; 32 mL
deionized formamide; 33.6 g ultrapure urea, and adjust volume to
100 mL. Filter through a 0.22- |im filter. Mix and degas for 10-15 min.
Store at 4 °C in a brown bottle for approx. 1 month.

• Ammonium persulfate (APS) 10% (w/v) solution: Add 100 mg of dry APS
to 1 mL of distilled water, vortex to dissolve. This is used immediately
and then discarded.

• TEMED

• Gel Loading Dye 2X (BioRad's recipe, final concentration): 0.05% bro-
mophenol blue/0.05% xylene cyanol/70% glycerol. Prepare a 2% bro-
mophenol blue and a 2% xylene cyanol solution. Mix 0.25 mL of each
solution with 7.0 mL of 100% glycerol, add 2.5 mL of distilled water to
make volume up to 10.0 mL. Store at room temperature.

• Glycogen solution (20mg/mL; Roche Diagnostics 901393, Laval, Que.,
Canada)

• 3 M sodium acetate (pH 5.2)

• 100%ethanol

• Vistra Green (Amersham Biosciences) solution: Dissolve 25 p.L of Vistra
Green in 250 mL of lx TAE buffer (1:10,000 dilution). Store solution at
4 °C for 3-4 days.

• 100 bp molecular weight ladder (Fermentas SM0241)

■ Sample Preparation

1. PCR amplification of extracted and purified total community DNA:
It may be necessary to dilute the soil DNA extract (preparation see
Sect. 10.2) 1:10 or 1:100 to optimize PCR yield.

2. A typical PCR reaction (total volume 50 pL) is composed of the following:

- 1.0 pL of template DNA (or dilution)

222 L.G. Whyte, C.W. Greer

- 1.0 pL U341GC#2 primer (25 pmol); sequence:
5^^- GCGGGCGGGGCGGGGGCACGGGGGGCGCGGCGGGC
GGGGCGGGGG CCTACGGGAGGCAGCAG-3' (GC clamp underlined)

- 1 .0 pL of U758 primer (25 pmol); sequence:
5 / 758 _ 740 -CTACCAGGGTATCTAATCC-3 /

- 0.5pLofl00mMMgCl 2

- 8.0pLofl.25mMdNTPs

- 32.4 pL of sterile deionized water

- 0.625 pL of BSA (lOmg/mL; optional, but often improves the PCR
when using DNA recovered from soils with high organic content)

3. In a separate tube add lOx DNA polymerase buffer (5 pL per reaction) and
DNA polymerase (0.5 pL per reaction). We typically use rTaq polymerase
for this work. It is easier to prepare this mixture to accommodate all
planned reaction tubes, and add 5.5 pL of the mixture to each reaction.

4. For a "hot start" the tubes are put in the thermal cycler and heated to
96 °C for 5 min. The temperature is then reduced to 80 °C and the DNA
polymerase buffer/DNA polymerase mix is added to each tube.

5. PCR is conducted using the following conditions:

The first ten cycles use a "Touchdown protocol" from 65-55 °C, with the
annealing temperature decreased by 1 °C at each cycle.

- lminat94°C

- 1 min at 65-55 °C

- 3 min at 72 °C

The subsequent 20 cycles are performed with an annealing temperature
of55°C.

7. The PCR reactions are analyzed by agarose gel electrophoresis using
5-10 pL of reaction in a 1.4% agarose gel using TAE buffer (Sambrook
and Russell 2001). Several dilutions (i.e., 1, 2, and 4pL of a 1:10 di-
lution) of a 100 bp molecular weight ladder (Fermentas SM0241) are
electrophoresed in the gel as well to quantify the amount of PCR prod-
uct. For complex environmental samples it is advisable to prepare up to
500 ng of PCR product to apply to each lane of the DGGE.

■ Procedure

Denaturant Gradient Gel (after Fortin et al. 2004)

1. Assemble the glass plates with spacers and clamps and secure to the
casting stand.

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 223

2. Clamp (or tape) needle outlet from the gradient mixer between the
glass plates (middle/top) so that it will inject the gel solution between
the plates.

3. To prepare a 30-70% gradient, add 7.2 mL of 8% acrylamide/0% denat-
urant solution and 4.3 mL of 8% acrylamide/80% denaturant solution
to one 50-mL Falcon tubes (Fisher Scientific; label "Low") and add
1.4 mL of 8% acrylamide/0% denaturant solution and 10.1 mL of 8%
acrylamide/80% denaturant solution to another 50 mL Falcon tubes
(label "High").

4. Add 115yiL of 10% fresh APS solution to each tube. Mix gently by
inversion. Be careful not to introduce air into the solution.

5. Add 1 1 .5 p.L of TEMED to each tube. Mix gently by inversion. Be careful
not to introduce air.

6. Add the low denaturant solution gently to the left chamber (Low) of the
gradient mixer. Remove air bubble from transfer tube by opening the
valve stem quickly until the transfer tube between the two chambers is
just full of low denaturant solution.

7. Add the high denaturant solution gently to the right chamber (High)
of the gradient mixer, turn on the mixer and the pump, open the out
valve on the right side, and transfer the entire solution to the plates.

8. Gently layer 1 mL of water on top of the gel to stop it from drying out.

9. Let the gel polymerize for 1.5 h at room temperature.

Buffer

10. Add 6 L of 1 x TAE to gel tank (i.e., fill to the FILL line).

11. Insert the lid and turn on. Let the buffer warm up until the temperature
reaches 60 °C. This takes more than 1 h, so you should do this 30min
after pouring the gel.

Spacer Gel

12. Using filter paper, remove the water on top of the polymerized gel.

13. Insert gel comb fully.

14. Mix 3.75 mL of 8% acrylamide/0% denaturant with 1.25 mL of 1 x TAE
and with 45 iiL of 10% (w/v) APS and 4.5 \iL TEMED.

15. Add this to the top of the denaturant gradient with a pipette.

16. Let polymerize for 0.5 h.

224 L.G. Whyte, C.W. Greer

Loading and Running Gel

17. Remove the comb and any excess polyacrylamide from the gel.

18. Assemble the plates on the core. Pour approx. 350 mL of 1 x TAE in the
upper chamber to check the integrity of the seal. If buffer is leaking,
discard the buffer, disassemble, lubricate the gasket, reassemble the
plates onto the core and test again. Insert into tank containing 1 x TAE
buffer prewarmed to 60 °C.

19. Let equilibrate for 15 min.

20. Wash wells with syringe using the 1 x TAE buffer from the tank.

21. Load wells with samples diluted in 2 x gel-loading buffer.

22. Run gel at 80 V for 16 h at 60 °C.

Staining Gel

23. Stain gel for 0.5 h in 1:10,000 dilution of Vistra Green solution with
gentle shaking.

23. De-stain for 0.5 h in 250 mL of 1 x TAE with gentle shaking.

24. Scan the gel on a glass plate using a Fluorlmager, save an image of the
gel for printing (image the same size as the gel) to use as a template for
selecting bands for excision and sequencing.

Excising DGGE Bands and Purification for Nucleotide Sequencing

1. Transfer the gel onto a sheet of Plexiglas under which has been placed
the printed image of the stained gel.

2. Cut bands of interest from gel using a scalpel or razor blade, and transfer
into microcentrifuge tubes.

3. Add 60 pi of sterile deionized water to each DGGE fragment and elute
overnight in a 37 °C incubator.

4. Centrifuge at 1 6,000 g at room temperature for 1 min and transfer super-
natant to a fresh tube. Purify 50 pi with the QIAquick PCR purification
kit.

5. Add 5 volumes of Qiagen PB buffer to the 50 pi of supernatant. Vortex,
and apply the sample to the QIAquick column-collection-tube assem-
bly.

6. Centrifuge at 16,000 g for 1 min (binding step).

7. Discard flow-through. Place the column back in the collection tube.

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 225

8. To wash, add 750 pi of Qiagen PE buffer to the column, and centrifuge
at 16,000 g for 1 min.

9. Discard flow- through. Place the column back in the collection tube.

10. Centrifuge again at 16,000 g for lmin, and transfer the column to
a 1.5 mL microcentrifuge tube.

11. Add 50p.L of prewarmed (5 min at 50 °C) Qiagen elution buffer (EB;
10 mM Tris-HCl, pH 8.5) to the center of the membrane.

12. Incubate for 5 min at room temperature.

13. Centrifuge at 16,000 g for 1 min (elution step).

14. Store the purified DNA at -20 °C.

15. Re-amplify the purified DNA: It is advisable to prepare 200 ng for se-
quencing both strands. (To obtain good sequence results it is important
to optimize PCR conditions to obtain a single band, in an agarose gel,
of re-amplified product.) Several PCR reactions can be pooled and pre-
cipitated: Add 1 p.1 of glycogen solution, 1/10 volume of 3M sodium
acetate (pH 5.2) and 2.5 volumes of 100% ethanol. Precipitate at -20 °C
for 1.5 h or overnight to accumulate sufficient product.

16. Purify the re-amplified product using a QIAquick PCR purification kit
or GENE CLEAN II (Qbiogene) or a GFX punification kit (Amersham
Biosciences).

17. Quantify the product on an agarose gel with a dilution series of a molec-
ular standard, as described above, prior to submitting for sequencing.

■ Notes and Points to Watch

• The DGGE gel plates should be carefully assembled and checked for
leaks.

• The minimum and maximum denaturant solutions can be varied in
concentration to change the resolution of the gel. We have found that
a gradient from 35 to 65% denaturant gives the best resolution for many
environmental samples.

• When preparing denaturant solutions and adding the APS and TEMED,
care should be taken not to introduce air. This can be accomplished by
adding all the ingredients to separate 50-mL Falcon tubes, and gently
mixing by inversion before adding to the gradient mixer.

• It is important to ensure that the printed image of the scanned gel is
identical in dimensions to the gel itself, since bands are being excised "in

226 L.G. Whyte, C.W. Greer

the blind" and the image is the template for removing the bands. When
scanning the gel ensure that all four corners of the gel are scanned to
facilitate subsequent alignment with the printed image. After removing
the bands, the gel is re-scanned to ensure that the correct bands have
been successfully recovered.

10.5

Genomics in Environmental Microbiology

Environmental microbiology has only recently entered the genomics era.
Genomics in its broadest sense entails the complete sequencing of an or-
ganism's entire complement of DNA (made up of the 4 bases, A, T, C,
and G). The sequence of DNA for a particular gene is the genetic code,
or blueprint, that is translated into specific proteins, the key components
in assembling all the organism's structures, regulating its functions, and
consequently its behavior and physiology. Since the cost of genome se-
quencing has decreased substantially, an ever increasing number of mi-
crobial genomes are being sequenced, including important microorgan-
isms from industrial or environmental perspectives. Many of the publicly
available genome sequencing projects directed towards these latter or-
ganisms are sponsored by the US Department of Energy's (DOE) Microbial
Genome Project (www.sc.doe.gov/production/ober/microbial.html) in col-
laboration with other partners. The DOE projects are targeting microor-
ganisms involved in, for example, bioremediation, carbon sequestering,
energy production, cellulose degradation biotechnology, and technology
development. Presently 43 microorganisms with biodegradation capabili-
ties have been/are being fully sequenced to hopefully identify new micro-
bial processes involved in bioremediation and lead to the development of
novel technologies and methodologies (i.e., genomic approaches, molecu-
lar monitoring tools) for studying the structure and function of complex
microbial communities associated with contaminated environments.

As sequencing costs have diminished, PCR- independent methodologies
(Fig. 10.1), including metagenomic libraries and environmental genome
shotgun cloning approaches, have also emerged as novel ultra-high through-
put methods to characterize complex environmental microbial communi-
ties. Although still quite expensive, the PCR-independent methodologies
overcome some of the limitations of the PCR-dependent methodologies,
including the inherent bias of primer specificity in PCR amplification and
the relatively limited amount of sequence information obtained from the
small PCR gene targets amplified and sequenced (ca. 300- 1 ,000 nt). Metage-
nomic libraries are created by extracting total genomic DNA from an en-
vironment and cloning relatively large fragments (5,000-300,000 nt) into

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 227

lambda, cosmid, fosmid, or bacterial artificial chromosome (BAC) vec-
tors. The metagenomic libraries created are then screened for functional
and/or genetic diversity that allows for clones of interest to be singled out
and sequenced (Eyers et al. 2004). For example, Rondon et al. (2000) con-
structed metagenomic libraries from total DNA extracted from two soils
that contained more than 1 Gbp of DNA. Shotgun cloning is the process
by which total community DNA is extracted from a sample, broken up to
reduce the size, and the fragments (ca. 2,000-6,000 nt) then ligated into
cloning vectors. Each cloning vector and the fragment of community DNA
it carries is then amplified separately by growth in a bacterial host. The
entire assemblage of the clone library is randomly sequenced and then
reassembled in a procedure termed direct shotgun sequencing. Venter et
al. (2004) performed direct shotgun sequencing (1.045 billion base pairs)
of the microorganisms in the Sargasso Sea and identified hundreds of new
bacterial species and 1.2 million new genes! These studies have clearly
demonstrated the enormous biodiversity present in the environment, and
that we have only begun to identify the vast majority of microorganisms
out there.

Environmental microarrays are considered an emerging technology with
tremendous potential in the field of environmental genomics (Greer et al.
2001; Rhee et al. 2004; Stahl 2004; Zhou and Thomson 2002). Success-
ful application of microarray technology, which uses high-density, high-
throughput techniques, promises to revolutionize our understanding of
microbial diversity and microbial ecology, as thousands of potential gene
probes can be printed on an array and hybridized to labeled total nucleic
acids extracted from environmental samples. Environmental microarray
technology is at a developmental stage where significant problems regard-
ing specificity, sensitivity, and quantitation remain to be resolved (Eyers et
al. 2004; Rhee et al. 2004; Stahl 2004). Nevertheless, application-specific
environmental microarrays were recently used to detect sulfate-reducing
bacteria (Loy et al. 2002), methanotrophs (Bodrossy et al. 2003), and
biodegradative populations (Rhee et al. 2004) in environmental samples.
Two types of environmental microarrays are presently being developed.
Functional gene microarrays (FGMA) contain a variety of catabolic, biogeo-
chemical cycling, heavy metal transformation genes, etc., as gene targets.
Phylogenetic gene microarrays (PGMA) contain taxonomic gene targets,
usually the 16S rDNA genes representing most genera of Bacteria and Ar-
chaea. Environmental microarrays will be increasingly used to detect and
characterize complex microbial communities in contaminated soils as well
as to monitor degradative populations during bioremediation treatments.
This will lead to a better understanding of important processes such as
biogeochemical cycles and bioremediation in soils that are associated with
mixed microbial populations in natural environments.

228 L.G. Whyte, C.W. Greer

Acknowledgements. The authors gratefully acknowledge the technical ex-
pertise and contributions of Nathalie Fortin, David Juck, Diane Labbe,
Danielle Ouellette, Sylvie Sanschagrin, Blaire Steven, Dan Speigelman, and
Gavin Whissell.

References

Ahn J, Daidou T, Tsuneda S, Hirata A (2002) Characterization of denitrifying phosphate-
accumulating organisms cultivated under different electron acceptor conditions using
polymerase chain reaction-denaturing gradient gel electrophoresis ASSA Water Res
36:403-412

Battin TJ, Wille A, Sattler B, Psenner R (2001) Phylogenetic and functional heterogeneity
of sediment biofilms along environmental gradients in a glacial stream. Appl Environ
Microbiol 67:799-807

Berthelet M, Whyte LG, Greer CW (1996) Rapid, direct extraction of DNA from soils for PCR
analysis using polyvinylpolypyrrolidone spin columns. FEMS Microbiol Lett 138:17-22

Bodrossy L, Stralis-Pavese N, Murrell JC, Radajewski S, Weilharter A, Sessitsch A (2003)
Development and validation of a diagnostic microbial microarray for methanotrophs.
Environ Microbiol 5:566-82

Bonin PC, Michotey VD, Mouzdahir A, Rontani J-F (2002) Anaerobic biodegradation of
squalene: Using DGGE to monitor the isolation of denitrifying bacteria taken from
enrichment cultures. FEMS Microbiol Ecol 42:37-49

Evans FF, Seldin L, Sebastian GV, Kjelleberg S, Holmstrom C, Rosado AS (2004) Influence of
petroleum contamination and biostimulation treatment on the diversity of Pseudomonas
spp. in soil microcosms as evaluated by 16S rRNA based-PCR and DGGE. Lett Appl
Microbiol 38:93-98

Eyers L, George I, Schuler L, Stenuit B, Agathos SN, El Fantroussi S (2004) Environmental
genomics: exploring the unmined richness of microbes to degrade xenobiotics. Appl
Microbiol Biotechnol 66:123-130

Fisher SG, Lerman LS (1979) Length-independent separation of DNA restriction fragments
in two dimensional gel electrophoresis. Cell 16:191-200

Ford JE, McHeyzer- Williams MG, Lieber, MR ( 1 994) Chimeric molecules created by gene am-
plification interfere with the analysis of somatic hypermutation of murine immunoglob-
ulin genes. Gene 142:279-283

Fortin N, Beaumier D, Lee K, Greer CW (2004) Soil washing improves the recovery of total
community DNA from polluted and high organic content sediments. J Microbiol Meth
56:181-191

Fortin N, Fulthorpe RR, Allen DG, Greer CW (1998) Molecular analysis of bacterial iso-
lates and total community DNA from Kraft pulp mill effluent treatment systems. Can
J Microbiol 44:537-546

Fromin N, Hamelin J, Tarnawski S, Roesti D, Jourdain-Miserez K, Forestier N, Teyssier-
Cuvelle S, Gillet F, Aragno M (2002) Statistical analysis of denaturing gel electrophoresis
(DGE) fingerprinting patterns. Environ Microbiol 4:634-643

Greer CW, Whyte LG, Lawrence JR, Masson L, Brousseau R (2001) Current and future
impact of genomics-based technologies on environmental science. Environ Sci Technol
35:360A-366A

Ibekwe AM, Kennedy AC, Frohne PS, Papiernik SK, Yang C-H, Crowley DE (2002) Microbial
diversity along a transect of agronomic zones. FEMS Microbiol Ecol 39:183-191

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 229

Juck D, Charles T, Whyte LG, Greer CW (2000) Polyphasic microbial community analysis

of petroleum hydrocarbon contaminated soils originating from 2 northern Canadian

communities. FEMS Microbiol Ecol 33:241-249
Kauffmann IM, Schmitt J, Schmid RD. (2004) DNA isolation from soil samples for cloning

in different hosts. Appl Microbiol Biotechnol 64:665-70
Kreader CA (1996) Relief of amplification in PCR with bovine serum albumin or T4 gene 32

protein. Appl Environ Microbiol 62:1102-1106
Liesack W, Stackebrandt E (1992) Occurrence of novel groups of the domain Bacteria as

revealed by analysis of genetic material from an Australian terrestrial environment.

J Bacteriol 174:5072-5078
Liesack W, Weyland H, Stackebrandt E (1991) Potential risks of gene amplification by PCR

as determined by 16S rDNA analysis of a mixed culture of strict barophilic bacteria.

Microb Ecol 21:191-198
Liu W-T, Chanc O-C, Fang HP (2002) Characterization of microbial community in granular

sludge treating brewery wastewater. Water Res 36:1767-1775
Loy A, Lehner A, Lee N, Adamczyk J, Meier H, Ernst J, Schleifer KH, Wagner M (2002)

Oligonucleotide microarray for 16S rRNA gene-based detection of all recognized lin-
eages of sulfate-reducing prokaryotes in the environment. Appl Environ Microbiol

68:5064-5081
Luz AP, Pellizari VH, Whyte L.G, Greer CW (2004) A survey of indigenous microbial

hydrocarbon degradation genes in soils from Antarctica and Brazil. Can J Microbiol

50:323-333
Macnaughton S, Stephen JR, Chang YG, Peacock A, Flemming AM, Leung KT, White DC

(1999) Characterization of metal-resistant soil eubacteria by polymerase chain reaction

in denaturing gradient gel electrophoresis with isolation of resistant strains. Can J

Microbiol 45:116-124
Margesin R, Labbe D, Schinner F, Greer CW, Whyte LG (2003) Characterization of

hydrocarbon-degradative microbial populations in contaminated and pristine alpine

soils. Appl Environ Microbiol 69:3985-3092
Martin-Laurent F, Philippot L, Hallet S, Chaussod R, Germon JC, Soulas G, Catroux G (200 1 )

DNA extraction from soils: old bias for new microbial diversity analysis methods. Appl

Environ Microbiol 67:2354-2359
Massol-Deya A, Weller R, Rios-Hernandez L, Zhou JZ, Hickey RF, Tiedje JM (1997) Succes-
sion and convergence of biofilm communities in fixed-film reactors treating aromatic

hydrocarbons in groundwater. Appl Environ Microbiol 63:270-276
Meyerhans A, Vartanian JP, Wain-Hobson SW (1990) DNA recombination during PCR

Nucleic Acids Res 18:1697-1691
Miller KM, Ming TJ, Schulze AD, Withler RE (1999) Denaturing gradient gel electrophoresis

(DGGE): a rapid and sensitive technique to screen nucleotide sequence variation in

populations. BioTechniques 27:1016-1024
Mills DK, Fitzgerald K, Litchfield CD, Gillevet PM (2003) A comparison of DNA profiling

techniques for monitoring nutrient impact on microbial community composition during

bioremediation of petroleum-contaminated soils. J Microbiol Methods 54:57-74
Mumy KL, Findlay RH (2004) Convenient determination of DNA extraction efficiency using

an external DNA recovery standard and quantitative-competitive PCR. J Microbiol

Methods 57:259-68
Murray AE, Preston CM, Massana R, Taylor LT, Blakis A, Wu K, DeLong EF (1998) Seasonal

and spatial variability of bacterial and archaeal assemblages in the coastal waters near

Anvers Island, Antarctica. Appl Environ Microbiol 64:2585-95

230 L.G. Whyte, C.W. Greer

Muyzer G, Smalla K (1998) Application of denaturing gradient gel electrophoresis (DGGE)
and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie van
Leeuwenhoek 73:127-141

Muyzer G, Waal EC, Uitterlinden AG (1993) Profiling of complex microbial populations by
denaturing gradient gel analysis of polymerase chain reaction-amplified genes coding
for 16S rRNA. Appl Environ Microbiol 59:695-700

Myers RM, Fischer SG, Lerman LS, Maniatis T (1985) Nearly all single base substitutions
in DNA fragments joined to a GC-clamp can be detected by denaturing gradient gel
electrophoresis. Nucleic Acids Res 13:3131-3145

Nakagawa T, Fukui M (2002) Phylogenic characterization of microbial mats and streamers
from a Japanese alkaline hot spring with a thermal gradient. J Gen Appl Microbiol
48:211-222

Niibel U, Garcia- Pichel F, Muyzer G (1997) PCR primers to amplify 16S rRNA genes from
cyanobacteria. Appl Environ Microbiol 63:3327-3332

Paabo S, Irwin DM, Wilson AC (1990) DNA damage promotes jumping between templates
during enzymatic amplification. J Biol Chem 265:4718-4721

Rondon MR, August PR, Bettermann AD, Brady SF, Grossman TH, Liles MR, Loiacono
KA, Lynch BA, MacNeil IA, Minor C, Tiong CL, Gilman M, Osburne MS, Clardy J,
Handelsman J, Goodman RM. (2000) Cloning the soil metagenome: a strategy for
accessing the genetic and functional diversity of uncultured microorganisms. Appl
Environ Microbiol 66:2541-2547

Sambrook J, Russell DW (2001) Molecular Cloning: A laboratory manual. Cold Spring
Harbor Laboratory Press, New York

Schneegurt MA, Dore SY, Kulpa CF Jr. (2003) Direct extraction of DNA from soils for studies
in microbial ecology. Curr Issues Mol Biol 5:1-8

Soloway DA, Nahir M, Billowits ME, Whyte LG (2001) In-situ bioremediation of diesel-
contaminated soil in Canada's Arctic territory: a case study at the Whitehorse Interna-
tional Airport, Yukon Territory. Polar Record 202:267-272

Stahl DA (2004) High-throughput techniques for analyzing complex bacterial communities.
Adv Exp Med Biol 547:5-17

Venter JC, Remington K, Heidelberg JF, Halpern AL, Rusch D, Eisen JA, Wu D, Paulsen I,
Nelson KE, Nelson W, Fouts DE, Levy S, Knap AH, Lomas MW, Nealson K, White O,
Peterson J, Hoffman J, Parsons R, Baden-Tillson H, Pfannkoch C, Rogers YH, Smith HO
(2004) Environmental genome shotgun sequencing of the Sargasso Sea. Science 304:66-
74

von Wintzingerode F, Gobel UB, Stackebrandt E (1997) Determination of microbial diversity
in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microbiol Rev
21:213-229

Wang GC, Wang Y (1996) The frequency of chimeric molecules as a consequence of PCR
co-amplification of 16S rRNA genes from different bacterial species. Microbiology
142:1107-1114

Ward DM, Bateson MM, Weller R, Ruff- Roberts AL (1992) Ribosomal RNA analysis for
microorganisms as they occur in nature. Adv Microbiol Ecol 12:219-286

Whyte LG, Bourbonniere L, Bellerose C, Greer CW (1999) Bioremediation assessment of
hydrocarbon-contaminated soils from the high Arctic. Bioremediation J 3:69-79

Whyte LG, Goalen B, Hawari J, Labbe D, Greer CW, Nahir M (200 1 ) Bioremediation treatabil-
ity assessment of hydrocarbon contaminated soils from Eureka, Nunavut. Cold Regions
Sci Technol 32:121-132

Whyte LG, Labbe D, Goalen B, Buchko J, Nahir M, Billowits M, Greer CW (2003) In-
situ bioremediation of hydrocarbon contaminated soils in the high Arctic. ARCSACC
Conference. Edmonton, May 4-6, pp 245-256

10 Molecular Techniques for Monitoring and Assessing Soil Bioremediation 231

Whyte LG, Schultz A, van Beilen JB, Luz AP, Pellizari D, Labbe D, Greer CW (2002) Prevalence
of alkane monooxygenase genes in Arctic and Antarctic hydro carbon- contaminated and
pristine Soils. FEMS Microbiol Ecol 41:141-150

Zhou J, Thompson, DK (2002) Challenges in applying microarrays to environmental studies.
Curr Opin Biotechnol 13:204-207

Bioreporter Technology

for Monitoring Soil Bioremediation

Steven Ripp

11.1

General Introduction

Bioreporters refer to intact, living cells that have been genetically engi-
neered to produce a measurable signal transcriptionally induced in re-
sponse to a specific chemical or physical agent in their environment. Biore-
porters contain three essential genetic elements, a promoter sequence,
a regulatory gene, and a reporter gene. In the wild-type cell, the promoter
gene is transcribed upon exposure to an inducing agent, leading to subse-
quent transcription of downstream genes that encode for proteins that aid
the cell in either adapting to or combating the agent to which it has been ex-
posed. In the bioreporter, the downstream genes, or portions thereof, have
been removed and replaced with a reporter gene. Consequently, transcrip-
tion of the promoter gene activates the reporter gene, reporter proteins are
produced, and some type of measurable signal is generated. These signals
can be categorized as either colorimetric, fluorescent, luminescent, chemi-
luminescent, electrochemical, or amperometric. Although each bioreporter
functions differently, the end product is always the same - a measurable
signal that is, ideally, proportional to the concentration of the specific
chemical or physical agent to which they have been exposed (Fig. 11.1).

Bioreporters can also be constructed without such inherent specificity.
These bioreporters rely on reporter genes that are induced by a group of
substances rather than just one or a few. Their primary use is for the detec-
tion of toxic substances, which, upon exposure to the bioreporter, induce
a stress-response gene that is fused to a reporter gene. Thus, an increase in
signal intensity indicates toxicity, but the substance that initiated the signal
cannot be uniquely identified. Reporter systems can also be designed to
operate in the reverse, where a decrease in signal intensity indicates tox-
icity. These bioreporters contain a constitutively expressed reporter gene
that always remains on. Upon toxin exposure, the bioreporters either die or
their metabolic activities are severely reduced, thereby causing a reduction
in signal strength.

Steven Ripp: The University of Tennessee, Knoxville, Tennessee, 37996, USA, E-mail:
saripp@utk.edu

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

234 S. Ripp

Promoter Transcription Translation
Element q

-^^zzza- +zz ►oj^

Reporter mRNA

Gene Keporter

Protein

o ^

q O Inducer

Fig. 11.1. Anatomy of a bioreporter organism. Upon exposure to a specific inducer, the
promoter/reporter gene complex is transcribed into messenger RNA (mRNA) and then
translated into a reporter protein that is ultimately responsible for signal generation

Although all data generated by a bioreporter can be obtained much
more accurately using conventional analytical techniques such as gas chro-
matography and mass spectrometry (GC/MS), bioreporters offer a distinct
advantage in that they report not only on a chemical's presence but on its
bioavailability and overall effect on a living system. Bioreporters are also
significantly cheaper, faster, and easier to use than typical analytical meth-
ods. Additionally, for some select bioreporter systems, the bioassay can
be performed in situ, continuously, on-line, and in real time. Such traits
make bioreporters particularly well suited for bioremediation application.
Typically, in any bioremediative design, the first step is to identify and
quantify the contaminants present, which is best achieved using analytical
techniques such as GC/MS. But after site characterization, bioreporters can
play useful roles in the frequent monitoring that ensues, for example, when
mapping the site to assess contaminant distribution prior to and during the
remediation process, as well as in post-closure monitoring. Bioreporters
can also be used to report on the status of environmental parameters impor-
tant to successful bioremediation, such as nutrient levels, pH, and dissolved
oxygen; or they can provide general biomass measurements (via ATP quan-
tification) as an indication of overall microbial activity. As well, where the
bioremediation strategy entails using an enhanced, bioengineered microor-
ganism, the bioremediation practitioner must provide a means of tracking
the microbe to ensure containment and monitor potential recombinant
gene transfer events in the indigenous microbial population. This visual
tagging can often be provided through bioreporter technology. A series of
excellent reviews on bioreporter systems are available (Daunert et al. 2000;
Keane et al. 2002; Belkin 2003).

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 235

11.2

An Overview of Reporter Systems

for Soil Bioremediation Application

/^-Galactosidase (lacZ)

The lacZ gene derived from Escherichia coli encodes a /J-galactosidase
(/J-gal) that catalyzes the hydrolysis of /J-galactosides. Traditional lacZ
bioreporters are assayed colorimetrically. The substrate o-nitrophenyl-
j8-D-galactoside (ONPG) is added to permeabilized bioreporter cells, af-
ter inducer exposure, to generate a yellow by-product whose intensity
correlates with /J-gal activity to provide an estimate of target chemi-
cal concentration. The assay is simple and highly reliable, and has be-
come integral to commercially available genotoxicity test kits such as
the SOS Chromotest (Institute Pasteur, Paris). Due to low sensitivities
and narrow dynamic ranges, however, the colorimetric test is largely
being replaced by other detection methods. By simply using different
/J-galactoside substrates, fluorescent, luminescent, or chemiluminescent
assays are possible. A major disadvantage remains, however, in that the
reporter cells must be lysed or undergo a membrane disruption step in
order to quantify /J-gal activity. Thus, data is obtained only incremen-
tally and results are delayed, sometimes by several hours, in relation
to the time required to complete the /J-gal assay. Newer electrochem-
ical and amperometric assays are beginning to solve this problem by
measuring /J-gal activity either directly or indirectly in an on-line, near
real-time format. However, the endogenous presence of /J-gal in natural
environments and its potentially high background activity must always
be taken into account when performing any of these assays. Table 11.1
provides examples of lacZ-based bioreporters for environmental monitor-
ing.

Catechol 2,3-Dioxygenase (xylE)

The xylE encoded catechol 2,3-dioxygenase is part of the pWWO plasmid
of Pseudomonas putida and is involved in the degradation of aromatic
compounds. Catechol 2,3-dioxygenase catalyzes the cleavage of colorless
catechol to produce the yellow compound 2-hydroxymuconic semialde-
hyde, forming the basis of this reporter assay. Reporter systems for xylE
have been developed primarily for studying gene regulation, and their
utility as environmental reporters is rather limited to the tagging of mi-
croorganisms destined for environmental release. In this task xylE serves
well since its endogenous activity in environmental systems is extremely
low, as compared to lacZ.

236

S. Ripp

Table 11.1. The lacZ-based bioreporters

Analyte

Reporter gene

Time for induction

Concentration

Antimonite, arsenite

arsR

30min

10 -15 M

Biphenyls

bphk

ImM

Cadmium

zntk

<lh

25 nM

Chlorocatechol

clcR

5 min

10 _8 M

Chromate

chr

luM

Copper

pcoE

0.01 mM

CUPl

25 min

0.5-2 mM

Salmonella sulk

0.025 pg/mL

dink, B y D

< 30 min

1 pg/mL

umuC

0.05 pg/mL

reck

< 10 min

1 pg/mL

sfik

< 1 ng/mL

Mercury

mer

0.2 ng/mL

Nickel

cnr

128 pM

Pesticide toxicity

HSP104

1.5 h

0.1 mg/L

Phenols

dmpR

0.5 mM

Zinc

smtk

12 pM

^-Lactamase (bid)

/J-Lactamase cleaves /J-lactam rings in certain antibiotics. Synthetic sub-
strates have been developed that can also be cleaved by ^-lactamase to form
colorimetric or fluorescent products. As with catechol 2,3-dioxygenase,
/J-lactamase is routinely used for gene regulation studies but rarely as
a reporter for environmental assessment. However, /Mactamase-derived
reporters for mercury, arsenic, and cadmium are available, although their
operational capacity under environmental conditions is unknown.

Green Fluorescent Protein (GFP)

Green fluorescent protein (GFP) is a photoprotein isolated and cloned
from the jellyfish Aequorea victoria (Misteli and Spector 1997). Variants
have also been isolated from the sea pansy Renilla reniformis. GFP pro-
duces a blue fluorescent signal without the addition of an exogenous sub-
strate. All that is required is an ultraviolet light source to activate the
fluorescent properties of the photoprotein. This ability to autofluoresce
makes GFP highly desirable in biosensing assays since it can be used
on-line and in real time to monitor intact, living cells. Additionally, the
ability to alter GFP to produce light emissions besides blue (i.e., cyan,
red, and yellow) allows it to be used as a multianalyte detector. Conse-
quently, GFP has been incorporated into bioreporters for the detection

1 1 Bioreporter Technology for Monitoring Soil Bioremediation

237

Table 11.2. GFP-based bioreporters

Analyte

Reporter gene

Time for induction

Concentration

Arsenic

arsR

lppb

Benzene derivatives,

tbu

3.3 uM

branched alkenes

Biocides

TEF

25 min

100 ug/mL

Cadmium

Cd-binding peptide

0.5 uM

Iron

pvd

Unknown

10" 4 M

Mercury

mer

16 h

< 50 ng/mL

Nitrate

nar

0.05 mM

Octane

alkB

1-2.5 h

0.01-0.1 uM

Tetracyclines

tetR

50 min

< 10 ng/mL

Toluene

tbuAl

0.2 uM

Table 11.3. Representative examples of GFP used

as a visual tag

Application

Matrix

Monitoring Arthrobacter
Monitoring

Pseudomonas pseudoalcaligenes
Monitoring Alcaligenes faecalis
Monitoring Pseudomonas fluorescens
Monitoring Pseudomonas putida
Monitoring Pseudomonas sp.
Survival of Pseudomonas sp.
Survival of Pseudomonas resinovorans
Survival of Moraxella sp.
Temperature effects

on Arthrobacter chlorophenolicus
TOL plasmid expression
Transport of Pseudomonas putida

4-Chlorophenol-contaminated soil

PCB-contaminated soil

Phenol-contaminated soil

3-Chlorobiphenyl-contaminated root rhizosphere

Activated sludge

PAH -contaminated soil

2,3-Dichlorobiphenyl-contaminatedsoil

2,3-Dichlorodobenzo-p-dioxin-contaminatedsoil

p-Nitrophenol-contaminated soil

Agricultural soil

Biofilm

Groundwater

of various heavy metals (Table 11.2) and as a visual tag within bacterial,
yeast, nematode, plant, and mammalian hosts for monitoring purposes
(Table 11.3).

Uroporphyrinogen (Urogen) III Methyltransferase (UMT)

UMT catalyzes a reaction that yields two fluorescent products that produce
a red-orange fluorescence in the 590-770 nm range when illuminated with
ultraviolet light (Sattler et al. 1995). So as with GFP, no addition of exoge-
nous substrates is required. UMT has been used for whole-cell sensing of
antimonite, arsenite, and arsenate (Feliciano et al. 2000).

238

S. Ripp

Luciferases

Insect Luciferase (luc). Firefly luciferase catalyzes a reaction that produces
visible light in the 550-575 nm range. A click-beetle luciferase is also avail-
able that produces light at a peak closer to 595 nm. Both luciferases require
the addition of an exogenous substrate (luciferin) for the light reaction
to occur. Examples of luc-based bioreporters constructed for the detec-
tion of inorganic and organic compounds of environmental concern are
presented in Table 11.4. Visual tagging of microorganisms with luc has
also been performed in, for example, 4-chlorophenol-contaminated soils
to track bioremediation progress.

Bacterial Luciferase (lux). Luciferase is a generic name for an enzyme that
catalyzes a light- emitting reaction. Luciferases can be found in bacteria,
algae, fungi, jellyfish, insects, shrimp, and squid, and the resulting light
that these organisms produce is termed bioluminescence. In bacteria, the
genes responsible for the light-emitting reaction (the lux genes) have been
isolated and used extensively in the construction of bioreporters that emit
a blue-green light with a maximum intensity at 490 nm (Meighen 1994).
Three variants of lux are available, one that functions at < 30 °C, another
at < 37 °C, and a third at < 45 °C. The lux genetic system consists of five
genes, luxA, luxB, luxC, luxD, and luxE. Depending on the combination
of these genes used, several different types of bioluminescent bioreporters
can be constructed.

Table 11.4. The /wc-based bioreporters

Analyte

Reporter Gene

Time for Induction

Concentration

Arsenite

ars

10 nM

Arsenite, antimonite,

ars

33 nM (antimonite)

cadmium

Benzene, toluene,

xylR

30min

3 uM (xylene)

xylene

Cadmium, lead,

cadA

2-3 h

1 nM (antimony)

antimony

Chromate

chr

50 nM

Copper, lead, mercury

Drosophila Mtn
promoter

48 h

3-19 ppm

Environmental

ERE

10-12 h

10 -7 M (DDT)

estrogens

Herbicides

tac-luc-luxAB-aphll

> 30 min

ppm levels

Mercury

mer

100 nM

Organomercurials

mer

0.2 nM (methyl-
mercury chloride)

Zinc

znt

40 uM

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 239

Luciferase AB (luxAB). The luxAB bioreporters contain only the luxA and
luxB genes, which together are responsible for generating the light signal.
However, to fully complete the light- emitting reaction, a substrate must be
supplied to the cell. Typically, this occurs through the addition of the chem-
ical decanal at some point during the bioassay procedure. Numerous luxAB
bioreporters have been constructed within bacterial, yeast, insect, nema-
tode, plant, and mammalian cell systems and have been applied toward
detection of various environmental contaminants, monitor and control of
bioremediation process, assays of toxicity, application of visual tags, and
estimation of microbial biomass.

Luciferase CDABE (luxCDABE). Instead of containing only the luxA and luxB
genes, bioreporters can contain all five genes of the lux cassette, thereby
allowing for a completely independent light generating system that requires
no extraneous additions of substrate nor any excitation by an external light
source. In these bioassays, the bioreporter is simply exposed to a target an-
alyte and a quantitative increase in bioluminescence results, often within
less than 1 h. Due to their rapidity and ease of use, along with the ability to
perform the bioassay repetitively in real time and on-line, luxCDABE biore-
porters have become extremely attractive for environmental monitoring.
Additionally, the recent development of microluminometers for detecting
the bioluminescent signal reduces this assay down to a miniaturized for-
mat (Nivens et al. 2004). Table 11.5 illustrates the widespread application
of luxCDABE-based bioreporters.

Non-specific luxCDABE. Nonspecific lux bioreporters are typically used for
the detection of chemical toxins. They are usually designed to continuously
bioluminesce. Upon exposure to a chemical toxin, either the cell dies or
its metabolic activity is retarded, leading to a decrease in bioluminescent
light levels. Their most familiar application is in the Microtox assay (Azur
Environmental, Newark, DE, USA) where, following a short exposure to
several concentrations of the sample, decreased bioluminescence can be
correlated to relative levels of toxicity (Hermens et al. 1985). The Vitotox
test (Flemish Institute for Technological Research, Mol, Belgium) operates
similarly (Verschaeve et al. 1999).

Mini-Transposons as Genetic Tools in Bioreporter Constructions

A transposon is a discrete genetic element capable of translocating from
a donor site within the DNA molecule into one of many non-homologous
target sites under the assistance of a transposase enzyme. Their use as re-
porter elements was first applied in gene regulation studies using a phage
Mu transposable element containing a promoterless lac gene. This was
followed by similar constructs using primarily the Tn5 and TnlO family

240

S. Ripp

Table 11.5. The luxCDABE-based bioreporters

Analyte

Reporter

Time for induction

Concentration

2,3-Dichlorophenol

reck

(stress promoter)

50 mg/L

2,4,6-Trichlorophenol

reck

(stress promoter)

lOmg/L

2,4-D

tfdRP

20-60 min

2 uM-5 mM

3 -Xylene

xyl

hours

3uM

4-Chlorobenzoate

fcbk

380 uM-6.5 mM

4-Nitrophenol

reck

(stress promoter)

0.25 mg/L

Anatoxin Bl

Various stress
promoters

45 min

1.2 ppm

Ammonia

hao

30 min

20 uM

BTEX

tod

1-4 h

0.03-50 mg/L

(benzene, toluene,

ethylbenzene,

xylene)

Cadmium

cupS

19 mg/kg

Chlorodibromo-

reck

(stress promoter)

20 mg/L

methane

Chloroform

reck

(stress promoter)

300 mg/L

Chromate

chrk

10 uM

Cobalt

cnr

4-6 h

9uM

Copper

Not specified

1 uM-1 mM

Hydrogen peroxide

katG

20 min

0.1 mg/L

Iron

pupk

hours

lOnM-luM

Isopropyl benzene

ipb

1-4 h

1-100 uM

Lead

pbr

4,036 mg/kg

Mercury

mer

70 min

0.025 nM

Naphthalene

nahG

8-24 min

12-120 uM

Nickel

cnr

4-6 h

0.1 uM

Nitrate

narG

0.05-50 uM

Organic peroxides

katG

20 min

Not specified

PCBs

bph

1-3 h

0.8 uM

p-Chlorobenzoic acid

fcbk

40 min

0.06 g/L

Pentachlorophenol

reck

(stress promoter)

0.008 mg/L

Phenol

reck

(stress promoter)

16 mg/L

Salicylate

nahG

15 min

36 uM

Silver

zntkp

0.1 uM

Tetracycline

tet

50 min

< lOng/mL

Trichloroethylene

tod

1-1.5 h

5-80 uM

Zinc

smtk

0.5-4 uM

of transposons as well as a variety of others such as Tn3/Tnl, Tn916/917,
and TnlOOO. Although powerful mutagenic tools, natural transposons had
several disadvantages, especially in environmental applications; they re-
quired an antibiotic resistance marker for selection and were composed of

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 241

inverted repeat elements that promoted unwanted genetic rearrangements
and inherent instability (secondary transposition). They were also large
and difficult to work with genetically and were subject to transposition im-
munity, which prevented multiple transposon insertions within the same
bacterial strain, severely limiting their cloning value. The development of
mini-transposons solved many of these problems. Mini-transposons are
shortened hybrids of natural transposons, usually Tn5 and TnlO, in which
the transposase gene is placed outside the boundaries of the inverted re-
peats. In this formation, the mobile element undergoes insertion into the
target site but the transposase does not, thus preventing any further re-
arrangements. Mini-transposons are also not affected by transposition
immunity, thereby allowing for multiple insertions of foreign inserts in
the same strain, provided that each insert has its own unique selectable
marker. Additionally, mini-transposons typically maintain an origin of
replication that allows for delivery into a broad range of hosts. Various
mini-transposons customized with reporter genes have been developed for
simplified construction of bioreporter organisms. By inserting a genetic
promoter element into a unique cloning site within the mini-transposon
vector, one can theoretically engineer any of the bioreporter classes dis-
cussed above. Furthermore, the ability to stably insert the mini-transposon
into the host chromosome makes these systems ideal for environmental
applications, since the necessity for antibiotic selection can be reduced.
Newer mini-transposons based on heavy-metal-resistance determinants
make antibiotic selection obsolete. Methods for constructing and using
mini-transposons are expertly described by de Lorenzo andTimmis (1994).

11.3

Single Point Measurements of Soil Contaminants

■ Introduction

Objectives. The application of bioreporters to soil bioremediation mon-
itoring can be applied in several different formats. This can range from
simply adding bioreporters to soil extracts to detect chemical presence to
being so multifaceted as to use in flow-cell formats for on-line, continuous
monitoring. With the numerous types of bioreporter systems available, the
bioremediation practitioner has a wide range of options to choose from for
the particular monitoring needs being addressed. For the sake of simplicity,
the protocols described below will relate to a luxCDABE-based bioreporter
system, but other systems can be substituted with corresponding substi-
tutions in growth conditions, types of substrate added (if required), and
monitoring instrumentation. The reviews by Belkin (2003), Daunert et al.
(2000), and Keane et al. (2002) should be addressed for further direction.

242 S. Ripp

Principle. Bioreporter cells containing all five genes of the lux cassette are
exposed to a soil suspension containing the target analyte, and a quantita-
tive increase in bioluminescence is measured.

■ Equipment

• Centrifuge and Corex glass centrifuge tubes (Corning Inc., Corning, NY,
USA) with Teflon screw cap lids

• 25 mL mineralization vials with Teflon screw cap lids

• Rotating shaker

• Instrument capable of monitoring bioluminescence [Perkin-Elmer Vic-
tor Multilabel reader (Wellesley, MA, USA), Azur Environmental Delta-
tox, Zylux Femtomaster (Oak Ridge, TN, USA), Wallac Microbeta (Welles-
ley, MA, USA), etc.]

■ Reagents

• YEPG medium (per L): 0.2 g yeast extract, 2 g polypeptone, 1 g glucose,
0.2gNH 4 NO 3 ,pH7.0

• Mineral salts medium (MSM; per L):0.1gMgSO 4 x 7H 2 O,0.2gNH 4 NO 3 ,
100 mL phosphate buffer, 0.1 mL trace elements solution

- Phosphate buffer: 0.5 M K 2 HP04/NaH 2 P04 mixture, pH 7.0; added
to MSM after autoclaving separately

- Trace elements (per L distilled water): 10.0 g MgO, 2.94 g CaCl 2 , 5.4 g
FeCl 3 x 6H 2 0, 1.44 g ZnS0 4 x 7H 2 0, 0.25 g CuS0 4 , 0.062 g H 3 B0 4 ,
0.49 g Na 2 Mo0 4 x H 2 0; added to MSM after filter sterilizing

■ Sample Preparation

Obtain field-moist soil samples or soil cores from test site.

■ Procedure

1. Preparation of bioreporter (description is for the luxCDABE bioreporter
Pseudomonas fluorescens HK44 (Ripp et al. 2000); other bioreporter
growth conditions will differ):

1.1. Inoculate 100 mL of YEPG medium from a frozen stock of biore-
porter cells. Grow overnight at 30 °C, with shaking at 200 rpm.

1.2. The next day, inoculate 1:10 into 100 mL fresh YEPG medium. Grow
at 30 °C with shaking to an optical density at 546 nm of 0.35.

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 243

2. Soil preparation

2.1. Divide soil into 10 g portions into clean 25 mLCorex glass centrifuge
tubes. Perform in triplicate.

2.2. Add 7 mL MSM and shake at room temperature and 200 rpm for
lh.

2.3. Centrifuge at 7,500 g and 25 °C for 10 min to remove large particu-
lates.

2.4. Remove 2 mL of supernatant into a 25 mL mineralization vial.

3. Bioluminescent assay

3.1. Add 2 mL of bioreporter culture (OD 546 = 0.35) to 2 mL soil extract
in a mineralization vial.

3.2. Transfer bioreporter/soil mixture to an appropriate holding device
based on type of light reader being used (microtiter plate, glass vial,
cuvette, etc.). Monitor light output for approximately 1 h.

4. Preparation of standard curve

4.1. Perform assay as described above with sterilized, uncontaminated
soil to which known concentrations of contaminant have been
added.

5. Controls

5.1. Perform assay as described above with sterilized, uncontaminated
soil. Bioreporters will generate a background level of signal that
must be subtracted from signals obtained in test samples.

■ Calculation

Subtract background light levels from test sample light levels and plot
results on the standard curve to determine contaminant concentrations.
Light levels are expressed using the arbitrary unit of relative light unit
(RLU).

■ Notes and Points to Watch

• The optimal temperature for lux bioluminescent activity can be modified
by using different lux cassettes. Vibrio fischeri lux functions at 30 °C,
V^ harveyi at 37 °C, and Photorhabdus luminescens at 42 °C.

• The lux reaction requires oxygen and will not operate efficiently under
anaerobic or low oxygen conditions.

244 S. Ripp

• Any of the bioreporter systems described above could generate false
positive signals due to non-specific induction by non-target inducers.
Appropriate controls must be incorporated into experimental protocols.

• For statistical purposes, assays should be performed in triplicate.

11.4

Continuous On-Line Vapor Phase Sensing
of Soil Contaminants

■ Introduction

Objectives. Bioremediation processes often require a quick "snapshot" of
soil contaminant concentrations to verify that environmental conditions are
conducive for optimal bioremediation and to provide an overall assessment
of where the contaminants are located. Bioreporters offer a very rapid
assessment technology that can be economically applied to contaminated
sites of interest.

Principle. A flow- through chamber containing alginate-encapsulated biore-
porter cells is inserted into a borehole and the presence of contaminants in
the vapor phase is continuously monitored via bioluminescent signals.

■ Equipment

• Flow-through bioreporter chamber (Fig. 11.2): The chamber consists of
a porous stainless steel tube (10 cm long x 2.3 cm diameter). Into the
bottom of the tube are packed bioreporter cells encapsulated in alginate.
Into the top of the tube is inserted a 1-mm diameter fiber-optic cable.
The other end of the cable truncates into a photomultiplier tube [PMT;
Hamamatsu model R-4632 Hamamatsu, Hamamatsu City, Japan), for
example] that measures the bioluminescent signals. For complete details,
see Ripp et al. (2000).

• 26-gauge needle

• Disposable 10-mL syringes

■ Reagents

• YEPG medium (Sect. 11.3)

• Sterile saline solution: 0.85% NaCl

1 1 Bioreporter Technology for Monitoring Soil Bioremediation

245

HK44 deriverd
bioluminescence

Porous
housing

Fiber optic
cable

Alginate

encapsulated

HK44

bioreporter

cells

Fig. 1 1.2. Device for monitoring volatile PAHs using immobilized P. fluoresce ns HK44 biore-
porters. The HK44 bioreporters bioluminesce in response to PAH exposure and the biolu-
minescent signal is transduced through a fiber-optic cable to an external light detector

• Sterile 3.5% (w/v) low viscosity alginic acid (Sigma- Aldrich, St. Louis)
solution in distilled water (this must be stirred overnight, then auto-
claved the next day)

• Sterile 0.1 M SrCl 2 solution

■ Sample Preparation

Holes (approx. 4-cm diameter) need to be bored on-site, to extend into the
zone of suspected contamination, for insertion of flow- through bioreporter
chambers. A control hole should also be bored in a known uncontaminated
area.

■ Procedure

1. Preparation of alginate-encapsulated bioreporter cells

1.1. Grow bioreporter culture in YEPG to OD 546 = 0. 35, as described in
Sect. 11.3.

1.2. Centrifuge (ca. lOmin at 3,000 g) cells and wash with an equal
volume of saline solution. Recentrifuge and again suspend in an
equal volume of saline solution.

246 S. Ripp

1.3. Gently mix one part of cell suspension with two parts of alginic acid
solution. Hold on ice.

1.4. Gently stir 1 L of refrigerated SrCl 2 solution in a sterile 1.5-L beaker
on a stirplate.

1.5. Transfer the cell/ alginic acid mixture to a 10-mL disposable syringe.
Attach a 26-gauge needle and gently push mixture through needle
in dropwise fashion, allowing drops to fall into the beaker with the
SrCl 2 solution. Drops will solidify into beads upon contact with
SrCl 2 . To ensure complete solidification, allow beads to gently stir
in SrCl 2 solution for approx. 45 min.

1.6. Remove the beads by decanting through sterile cheesecloth.

1.7. Store in a closed container at 4.0 °C for up to 3 months.

2. Preparation of flow- through bioreporter chamber

2.1. Pack 5 g of alginate-encapsulated cells into the bottom of the cham-
ber.

2.2. Insert fiber-optic cable directly above the encapsulated cells.

2.3. The chamber, suspended by the fiber-optic cable, can nowbe dropped
into the boreholes. Vapor phase detection of contaminant presence
is then monitored continuously by the PMT and data downloaded
to a laptop computer. Encapsulated cells must be replenished on
a weekly basis. Newer generation sensors have reduced the bulky
PMT modules down to an integrated circuit format, and it is now
possible to construct wireless chip-based devices, to wit, biolumi-
nescent bioreporter integrated circuits (BBICs), for remote sentinel
detection of target analytes (Nivens et al. 2004).

2.4. Background light levels should be obtained from a bioreporter
chamber installed in a control borehole.

■ Calculation

Subtract light levels obtained from the control bore hole to correct for back-
ground base levels of bioluminescence emanating from the bioreporter.
Plot bioluminescence (RLU) versus time to illustrate trends in biolumi-
nescence output. Based on these trends, the overall bioremediation process
can be evaluated and monitored to determine the effectiveness of the biore-
mediation program (i.e., low light levels will indicate unfavorable growth
conditions for the bioremediative microbes, which should result in the
implementation of analytical and microbiological assays and/or localized
treatments to diagnose and correct the existing problem).

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 247

■ Notes and Points to Watch

• See Sect. 11.3

• Avoid using phosphate buffers since they will degrade the alginate ma-
trix.

• The target contaminant(s) must produce an adequate vapor phase to be
detected.

11.5

Quantification of Soil-Borne lux-lagged
Microbial Populations Using
Most-Probable-Number (MPN) Analysis

■ Introduction

Objectives. Microorganisms genetically engineered for optimal biodegra-
dation of target contaminants can be introduced at a contaminated site to
enhance the bioremediation process. After release, these microbes must be
monitored to ensure that they remain within site boundaries. As well, their
population numbers must be monitored to verify that they remain viable
and metabolically active. The lux genes serve as excellent markers here
because their endogenous presence in soil ecosystems is negligible, thus
negating background interferences. The lux bioluminescent signal is also
easily measured. The Zwx-MPN assay described below requires insertion of
the luxCDABE cassette within the bioremediative microbe. For examples
on how this is accomplished, see King et al. (1990). Other bioreporter sys-
tems are applicable as well, but heightened background interferences will
occur. For information on the introduction and release of engineered mi-
crobes into soil and other environmental ecosystems, see Sayler and Ripp
(2000).

Principle. Soil inoculated with a lux-tagged bacterium is serially diluted.
The addition of sodium salicylate results in the induction of the lux operon,
and the intensity of bioluminescence can be correlated to cell numbers.

Equipment

PMT-based light reader capable of monitoring in 96-well microtiter plate
formats [Perkin-Elmer Victor Multilabel reader, Wallac Microbeta, BMG
Labtech Lumistar (Offenburg, Germany), etc.]

248 S. Ripp

■ Reagents

• Sterile sodium pyrophosphate solution: 0.1% (w/v)

• Sterile saline solution: 0.85% NaCl

• Sterile sodium salicylate solution: 6 mg/mL

■ Sample Preparation

The contaminated soil is first inoculated with the bioremediation-enhanced,
lux-tagged microbe. There are several methods for accomplishing this, and
one can refer to Sayler and Ripp (2000) for general guidelines. After inocu-
lation, soil samples (> 1 g) are removed from within areas and at depths that
received inoculant and transported to the lab on ice. Again, P. fluorescens
HK44 (Ripp et al. 2000) is used as an example.

■ Procedure

1. In a sterile test tube, add 1 g of soil to 9mL sodium pyrophosphate
solution and vortex 1 min at top speed to remove microbes from soil
particles.

2. Add 100 pi of soil suspension to the first column of a 96-well black, solid
bottom microtiter plate (Dynex Technologies, Chantilly, VA, USA).

3. Dilute 1:2 in 100 pL saline solution throughout columns 2 through 12.

4. Add 20 pL of sodium salicylate solution to all wells. In this example,
sodium salicylate serves as the inducer of the lux operon in P. fluorescens
HK44. Other bioreporters will use different inducers, but all act similarly
to turn on the bioreporter signal.

5. Prepare a duplicate control plate containing saline dilutions from 1 g of
sterile soil mixed with sodium pyrophosphate solution. Add 20 pL of
sodium salicylate solution to all wells. This plate will provide a measure
of background bioluminescence, if any, that needs to be subtracted from
bioluminescence counts in the sample plate.

6. Seal plates with transparent plate sealer (Perkin-Elmer Topseal) and
incubate at room temperature (23-28 °C) with gentle shaking for 16 h.
This permits maximum induction of bioluminescence from HK44 cells.
Use of other bioreporters will require optimization of incubation times.

7. Measure photon emission from wells with a microtiter-based light reader,
such as the Perkin-Elmer Victor instrument. Read each plate in triplicate
for statistical verification.

1 1 Bioreporter Technology for Monitoring Soil Bioremediation 249

■ Calculation

Input data into any variety of MPN software programs (see, for example,
Klee 1993). These programs will use Poisson statistics to estimate cell
numbers based on where bioluminescence is first observed within the
dilution series.

■ Notes and Points to Watch

• See Sect. 11.3.

• Accurate MPN population estimates require accurate dilutions. Ensure
that all dilution series are performed as carefully as possible.

References

Belkin S (2003) Microbial whole-cell sensing systems of environmental pollutants. Curr
Opin Microbiol 6:206-212

Daunert S, Barrett G, Feliciano JS, Shetty RS, Shrestha S, Smith-Spencer W (2000) Genetically
engineered whole-cell sensing systems: coupling biological recognition with reporter
genes. Chem Rev 100:2705-2738

de Lorenzo V, Timmis KN (1994) Analysis and construction of stable phenotypes in gram-
negative bacteria with Tn5- and TnlO-derived minitransposons. Methods Enzymol
235:386-405

Feliciano J, Liu Y, Ramanathan S, Daunert S (2000) Fluorescence-based sensing system for
antimonite and arsenite using cob A as the reporter gene. 219th ACS National Meeting

Hermens J, Busser F, Leeuwangh P, Musch A (1985) Quantitative structure-activity relation-
ships and mixture toxicity of organic chemicals in Photobacterium phosphoreum: the
Microtox test. Ecotoxicol Environ Saf 9:17-25

Keane A, Phoenix P, Ghoshal S, Lau PCK (2002) Exposing culprit organic pollutants: a review.
J Microbiol Meth 49:103-119

King JMH, DiGrazia PM, Applegate B, Burlage R, Sanseverino J, Dunbar P, Larimer F,
Sayler GS (1990) Rapid, sensitive bioluminescence reporter technology for naphthalene
exposure and biodegradation. Science 249:778-781

Klee AJ (1993) A computer program for the determination of most probable number and
its confidence limits. J Microbiol Meth 18:91-98

Meighen EA (1994) Genetics of bacterial bioluminescence. Annu Rev Genet 28:117-139

Misteli T, Spector DL (1997) Application of the green fluorescent protein in cell biology and
biotechnology. Nat Biotechnol 15:961-964

Nivens DE, McKnight TE, Moser SA, Osbourn SJ, Simpson ML, Sayler GS (2004) Biolu-
minescent bioreporter integrated circuits: potentially small, rugged and inexpensive
whole-cell biosensors for remote environmental monitoring. J Appl Microbiol 96:33-46

Ripp S, Nivens DE, Ahn Y, Werner C, Jarrell J, Easter JP, Cox CD, Burlage RS, Sayler GS (2000)
Controlled field release of a bioluminescent genetically engineered microorganism for
bioremediation process monitoring and control. Environ Sci Technol 34:846-853

Sattler I, Roessner CA, Stolowich NJ, Hardin SH, Harris-Haller LW, Yokubaitis NT,
Murooka Y, Hashimoto Y, Scott AI (1995) Cloning, sequencing, and expression of the
uroporphyrinogen-III methyltransferase cob A gene ofPropionibacteriumfreudenreichii
(shermanii). J Bacterid 177:1564-1569

250 S. Ripp

Sayler GS, Ripp S (2000) Field applications of genetically engineered microorganisms for

bioremediation processes. Curr Opin Biotechnol 11:286-289
Verschaeve L, Van Gompel J, Thilemans L, Regniers L, Vanparys P, van der Lelie D (1999)

VITOTOX bacterial genotoxicity and toxicity test for the rapid screening of chemicals.

Environ Mol Mutagen 33:240-248

^ ^ Interpretation of Fatty Acid Profiles
' ^ of Soil Microorganisms

David B. Hedrick, Aaron Peacock, David C. White

12.1

Obtaining Fatty Acid Profiles from Soil Samples

This work focuses on the calculations performed on the peak areas obtained
by gas chromatography (GC). All the steps of soil sampling, lipid extraction
and fractionation, derivatization, and capillary GC have been repeatedly re-
viewed, and will only be briefly mentioned (a bibliography of work done in
this laboratory is available at http://cba.bio.utk.edu/director_peerfull.html,
and an extensive bibliography of methods is provided by Dr. William
Christie's group, Mylnefield Research Services Ltd. at http://www.
lipidlibrary.co.uk/lit_surv.html).

Sampling is the most important step in sample analysis, and is often
delegated to the most junior member of the lab or to site specialists not
associated with the lipid laboratory, such as a subsurface sediment drilling
crew. Besides sampling location, the sample's consistency, integrity, and
appearance should be recorded. In order to obtain deep subsurface sam-
ples, the use of drilling equipment and drilling mud is usually required,
and methods have been developed to prevent and detect drilling mud con-
tamination of samples (Griffin et al. 1997; Phelps et al. 1989).

Capillary GC with flame ionization detection (FID) is a powerful ana-
lytical method - simpler in operation, of greater linear range, and more
sensitive, reliable, and reproducible than most analytical instrumentation
available. The users' manuals for the chromatograph and data system are
the primary references for their operation. If you won't read the manual,
you shouldn't touch the equipment. There are also many excellent reviews
of capillary chromatography of polar lipid fatty acids (PLFA) available (for
example, Grob and Barry 1995).

Capillary GC-MS is a necessary adjunct to GC-FID for the identification
of fatty acid peaks (Christie 2003). Various chemical methods are also avail-
able to help with specific identification problems such as silver ion chro-
matography to separate saturates, monounsaturates, and polyunsaturates

David B. Hedrick, Aaron Peacock, David C. White: Center for Biomarker Analysis, University
of Tennessee, 10515 Research Drive, Suite 300, Knoxville, Tennessee 37932-2575, USA,
E-mail: dwhitel@utk.edu

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

252 D.B. Hedrick et al.

(Momchilova and Nikolova-Damyanova 2000), and special derivatization
methods to determine the position and geometry of monounsaturation,
such as MS of dimethyldisulfide adducts (Nichols et al. 1986). MS of picol-
inyl esters provides more informative fragmentations than GC-MS of the
methyl ester (Christie et al. 1991; Harvey 1992).

This work presupposes some knowledge of Microsoft Excel (Microsoft
Corp., Redmond, WA), which is used to manipulate chromatographic re-
sults in many laboratories. The on-line help system is the basic reference
for Excel, such as it is. A novice user will benefit from one of the many intro-
ductory books available at a bookstore. Also assumed is some background
in the statistical procedures commonly applied to PLFA data, including
analysis of variance (ANOVA) and factor analysis.

12.2

Transforming Fatty Acid Peak Areas

to Total Microbial Biomass

Gas chromatography provides a peak area proportional to the amount of the
compound in the sample responsible for the peak. A known concentration
of an internal standard, usually 19:0 or 21:0, is added to the sample before
analysis to allow calculation of absolute amounts (see Sect. 12.5 for the
naming of fatty acids). The equation used to calculate the total amount of
fatty acids in a sample is,

(sum A F a/Ats) x IS x X ,

FA=- ^— — 12.1

FA total picomoles of fatty acids per gram dry mass of sample (pmol/g

dry mass)

sum A F a sum of the areas of all identified fatty acid peaks excluding the
internal standard

A IS area of the internal standard peak

IS concentration of internal standard used (50 pmole/p.L)

X volume of internal standard used to dilute the fatty acid methyl

esters (p.L)

Y mass of sample extracted (g soil dry mass). In some instances,

rather than grams dry mass as the divisor, it will be volume of
water (L), surface area in meters squared, or some other extensive
variable.

12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 253

Many analysts calculate the pmol/g dry mass for each fatty acid, then
add them together to get the total pmole/g dry mass. This is not good
practice, since the pmol/g dry mass for each fatty acid is not then of use in
further analysis, and the more complicated calculation makes more work
and opportunities for error.

The total moles of membrane fatty acids is proportional to the total
microbial biomass. The constant of proportionality used in our laboratory
is 2.5 x 10 4 cells/pmol PLFA (Balkwill et al. 1988; White et al. 1996 and
references therein). This conversion factor was derived from measurements
on laboratory cultures, so the number of cells will be underestimated for
environments populated by smaller bacterial cells, such as oligotrophic
environments.

Researchers who count cells, with automated cell counting instruments
or by microscopy, are often uncomfortable with measurements of viable
biomass expressed as moles of PLFA or grams dry mass of cells. In order
to estimate cell counts from moles of PLFA requires knowledge of the
distribution of cell sizes in the sample and the amount of PLFA per cell for
different sizes, information which is not usually available. It makes more
sense to transform cell counts to moles PLFA or from the latter to grams
dry weight of cells, since the cell counting can provide the data on cell size
distribution.

For most sample sets, the biomass will not be normally distributed, that
is, a histogram of the biomass data will be skewed with a long tail toward
the higher biomasses. This can be tested for by using the standard f-test
for normality. Also, in most biomass data sets, the variance of biomass
increases with the absolute value of the biomass. This violates the assump-
tions of parametric statistics, including ANOVA and factor analysis, and
lowers the power of any statistical test employed. These problems can be
solved by a log(X + A) transformation, where X is the mole percent of the
fatty acid, and A is a small constant. The small constant is added so that
zero values give a real solution when the log transform is applied. The
most common value used for A is one, which gives a value of zero for the
transform when X is zero, since log(0 + 1) = 0.

There are two approaches to proving the value of applying a log trans-
form to biomass data, the theoretical and the practical. The theoretical
explanation involves the scaling of the forces affecting microbial biomass
(Magurran 1988) and the fractal structure of microbial environments (Man-
delbrot 1982), and is beyond the scope of this work. The practical reason
for the log transform is that it works; applying a log transformation to the
data is perfectly legitimate, and results in more significant differences on
statistical tests.

254

D.B. Hedrick et al.

12.3

Calculation and Interpretation of Community Structure

After the biomass, the next most important information to extract from
a PLFA profile is the community structure. But where the biomass is a single
value for each sample with a straightforward interpretation, the commu-
nity structure data is multivariate with many options in its interpretation.
A "standard" method for presenting community structure data, how to
create a custom method for community structure, and factor analysis will
be presented.

12.3.1

Standard Community Structure Method

In the standard method for community structure analysis of PLFA pro-
files, chemically related fatty acids are grouped as in Table 12.1. A PLFA
profile may contain, for example, from 18 to 92 fatty acids. The standard
community structure approach summarizes that in six variables, which are
just the sum of the mole percents of each of the fatty acid groups. The use
of a standard community structure analysis method allows comparison
between/among experiments.

Table 1 2. 1 . Groups of chemically related fatty acids used in the standard community structure
analysis

Group name

Rule

Examples

Microbiota represented

Saturates

Saturated straight-

12:0, 13:0, 14:0,

All organisms

chain fatty acids

15:0, 16:0, 17:0,
18:0

Monounsaturates

Fatty acids with

14:lcc>5c,

Proteobacteria

a single unsaturation

16:1g)7c,

plus cyclopropyls

I6:loo7t,
\S:lco7c

Mid-chain branched

Any mid- chain

10Mel6:0,

Actinomycetes,

branched fatty acid

10Mel8:0

sulfate-reducers

Terminally branched

Iso- and anti-iso-

il4:0,il5:0,

Gram positive bacteria

branched saturated

al5:0,il6:0,

fatty acids

il7:0,al7:0

Polyunsaturates

Any fatty acid with

18:2a)6c,

Eukaryotes

more than one

1S:3co3c

unsaturation

Branched unsaturates

Any branched
monounsaturate

H7:lco7c

Anaerobes

12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 255

The standard community structure breakdown was originally devel-
oped on marine sediments, and has been successfully applied to microbial
communities from many environments, including, for example, marine
macrofaunal burrows (Marinelli et al. 2002), a subsurface zero-valent iron
reactive barrier for bioremediation (Gu et al. 2002), marine gas hydrates
(Zhang et al. 2002), soils contaminated with jet fuel (Stephen et al. 1999),
and to a comparison of subsurface environments (Kieft et al. 1997).

12.3.2

Custom Community Structure Methods

When examination of the chromatograms or the mole percent table shows
differences with treatment, but no significant differences are found in the
standard community structure groups, some other way of grouping the fatty
acids maybe more useful. For example, if samples differ in the proportions
of Cyanobacteria and Eukaryotic algae, it may be useful to separate the
polyunsaturates with 18 or fewer carbons characteristic of Cyanobacteria
(0ezanka et al. 2003) from those typical of Eukaryotic algae with 20 or
more carbons (Erwin 1973).

There are several methods for developing alternative community struc-
ture groups. The manual method uses the pattern recognition power of the
human eye. The PLFA chromatograms are printed on the same scale and
spread out on a large table. Similar-looking chromatograms are grouped
together and different-looking ones are placed in separate groups. While
very low- tech, this works remarkably well. This same approach can be ap-
plied to a mole percent table by printing it out, cutting out a strip for each
sample, and sorting the samples by similarity. Once the samples have been
sorted into similar groups, the fatty acids responsible are summed to form
new community structure groups.

Given access to statistical software, a triangular table of Pearson's r
correlation coefficients is usually available as an output option. Visual
examination of this table will locate fatty acids with high correlations, which
are then grouped together to form new community structure groups.

12.3.3

Factor Analysis

Factor analysis includes several related methods, including principal-com-
ponents analysis. The virtue of this method is that it automatically con-
structs fatty acid groups reflecting the differences in community structure,
rather than applying a preconception of fatty acid groups. The data deter-
mines the fatty acid groups, rather than the analyst. Factor loadings greater

256 D.B. Hedrick et al.

than 0.7 indicate fatty acids with "significant" effects on the results. The
factor scores are new variables that are linear combinations of the origi-
nal values. These new variables can be submitted to statistical tests such
as ANOVA like any other variable. Examples of the application of factor
analysis to PLFA profiles include storage perturbation of soil microbial
communities (Haldeman et al. 1995; Brockman et al. 1997), soils at differ-
ent temperatures (Zogg et al. 1997), and soils from different ecosystems
(Myers etal. 2001).

The results of factor analysis are usually improved by applying the log(X+
1) transformation to the mole percent data before factor analysis. A rough
method to determine whether the mole percent data is normally distributed
is to calculate the maximum, average, and the minimum not equal to zero
for each fatty acid. The formulas for these in Excel are "= max(b2.b45)", "=
average(b2.b45)", and "= min(if(b2.b45 = 0, 100,b2.b45))", where b2.b45
is the range containing the data. The formula for min is what Excel
terms an array formula; you have to hold down the Shift and Control keys
while you press Enter to enter the formula. If the difference between the
maximum and average is greater than the difference between the average
and the minimum for most of the fatty acids, then the data is not normally
distributed and the log(X + 1 ) transformation will probably improve results.

There are theoretical reasons to advocate the arcsin[square root(X)]
transformation over the log(X + 1 ) transformation, but very little difference
is found in practice, and the log(X + 1) is simpler to apply and explain.
Similarly, there are theoretical reasons to prefer factor analysis sensu stricto
over principal components analysis, and vice versa, which can, and have
been, argued for days to no conclusion. In practice, the two methods give
very similar results.

12.4

Calculation and Interpretation

of Metabolic Stress Biomarkers

The membrane of the bacterial cell handles all of its interactions with
its environment, and bacteria have many strategies to deal with stressful
environmental conditions, including modifying the fatty acids used in the
membrane. This is illustrated in Eq. (12.2), where S stands for the substrate
fatty acid and P for the product fatty acid induced by metabolic stress,
namely, a trans monounsaturate or cyclopropyl fatty acid.

S^ P

cis monounsaturate — >► trans monounsaturate (12.2)

cis monounsaturate —> cyclopropyl

12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 257

The stress biomarkers are then calculated as the ratio of the mole percents
of the product to the substrate fatty acids, as in Eq. (12.3):

BMs tr ess = P/S (12.3)

where BM Str ess is the value of the stress biomarker. The most common trans-
formations are !6:lco7c^ \6:lco7 t, 16:l(x>7c^Cyl7:0, lS:lco7c^ 18:loo7 t,
andl8:lco7c^Cyl9:0.

There are problems with the application of the stress biomarkers. The
first type of problem is when the stress-induced product fatty acid is only
detected in a minority of the samples. This will most likely prevent detection
of statistically significant differences. The second problem is when the
substrate fatty acid is not detected, but the stress-induced fatty acid is; this
has been seen in hot acid environments such as hydrothermal systems.
Since division by zero is undefined in standard algebra, undefined results
appear that standard statistical programs are unable to use. This problem
can be solved by a modification of Eq. (12.3),

BMs tr ess = P/(S + 1) (12.4)

The metabolic stress biomarkers have been applied to, for example, tap
water biofilms (White et al. 1999), and soils contaminated with jet fuel
(Stephen et al. 1999).

12.5

Naming of Fatty Acids

Creating clear, consistent, and unambiguous names for microbial fatty acids
is challenging due to the wide variety of possible structures. At the same
time, it is essential for understanding the data and communicating results.
The IUPAC rules for naming chemical compounds are supposed to provide
unambiguous names, but there are problems with this approach. The most
important is that IUPAC counts carbons from the opposite end of the fatty
acid molecule from most of the enzymes that modify the fatty acid.

The need for a compact notation has led to the development of the
omega system for naming fatty acids. Fatty acids are named according to
the pattern of AiBcoC. The A stands for the number of carbon atoms in the
fatty acid backbone, B is the number of double bonds, and C is distance
of the nearest unsaturation from the aliphatic (co) end of the molecule.
This can be followed by a "c" for cis or a "t" for trans configuration of
the unsaturation. The prefixes "i," "a," and "br" stand for iso, anti-iso,
and unknown branching position of the carbon chain, respectively. Mid-
chain branching is noted by a prefix "lOMe" for a 10-methyl fatty acid, and

258 D.B. Hedrick et al.

cyclopropyl fatty acids by prefix "Cy." For example: \S:lco7c is 18 carbons
long with one double bond occurring at the 7th carbon atom from the co
end, and the unsaturation is in the cis conformation. Also, 16:0, il6:0, al6:0,
and brl6:0 are all 16-carbon fatty acids, while 10Mel6:0 and Cyl7:0 both
contain a total of 17 carbons, not counting the carbon of the methyl ester
moiety.

References

Balkwill DL, Leach FR, Wilson JT, McNabb JF, White DC (1988) Equivalence of micro-
bial biomass measures based on membrane lipid and cell wall components, adenosine
triphosphate, and direct counts in subsurface sediments. Microbial Ecol 16:73-84

Brockman FJ, Li SW, Fredrickson JK, Ringelberg DB, Kieft TL, Spadoni CS, White DC,
McKinley JP (1997) Post-sampling changes in microbial community composition and
activity in a subsurface paleosol. Microbial Ecol 36:152-164

Christie WW (2003) Lipid analysis; isolation, separation, identification and structural anal-
ysis of lipids, 3rd edn. Oily Press, Bridgwater, UK

Christie WW, Brechany EY, Lie Ken Jie MSF, Bakare O (1991) MS characterization of picolinyl
and methyl ester derivatives of isomeric thia fatty acids. Biol Mass Spectrom 20:629-635

Erwin JA (1973) Fatty acids in eukaryotic microorganisms. In: Erwin JA (ed) Lipids and
biomembranes of eukaryotic microorganisms. New York, Academic Press, pp 41-143

Griffin WT, Phelps TJ, Colwell FS, Fredrickson JK (1997) Methods for obtaining deep
subsurface microbiological samples by drilling. In: Amy PS and Haldeman DL (eds)
The microbiology of the terrestrial and deep subsurface. CRC Press, Boca Raton, pp 23-
43

Grob RL Barry EF (1995) Modern practice of gas chromatography. Wiley, New York

Gu B, Zhou J-Z, Watson DB, Philips DH, Wu L, White DC (2001) Microbiological character-
ization in a zero-valent iron reactive barrier. Appl Environ Microbiol 77:293-309

Haldeman DL, Amy PS, Ringelberg DB, White DC (1994) Changes in bacteria recoverable
from subsurface volcanic rock samples during storage at 4 °C. Appl Environ Microbiol
60:2679-2703

Harvey DJ (1992) Mass spectrometry of picolinyl and other nitrogen-containing derivatives
of fatty acids. In: Christie WW (ed) Advances in lipid methodology, vol 1. Oily Press,
Ayr, UK, pp 19-80

Kieft TL, Murphy EM, Amy PS, Haldeman DL, Ringelberg DB, White DC (1997) Laboratory
and field evidence for long-term starvation survival of microorganisms in subsurface
terrestrial environments. In: Proceed instruments, methods, and missions for the in-
vestigation of extraterrestrial organisms, 27 July to 1 August. Int Soc Optical Engin, San
Diego, CA

Magurran AE (1988) Chapt. 2. Diversity indices and species abundance models. In: Magur-
ran AE (ed) Ecological diversity and its measurement. Princeton Univ Press, Princeton,
NJ

Mandelbrot B (1982) The fractal geometry of nature. Freeman, San Francisco, CA

Marinelli RL, Lovell CR, Wakeham SG, Ringelberg D, White DC (2000) An experimental in-
vestigation of the control of bacterial community composition in macrofaunal burrows.
Marine Ecol Prog Series 235:1-13

Momchilova S, Nikolova-Damyanova B (2003) Stationary phases for silver ion chromatog-
raphy of lipids: Preparation and properties. J Sep Sci 26:261-270

12 Interpretation of Fatty Acid Profiles of Soil Microorganisms 259

Myers RT, Zak DR, Peacock A, White DC (2001) Landscape-level patterns of microbial
community composition and substrate, use in forest ecosystems. Soil Sci Soc Am J
65:359-367

Nichols PD, Guckert JB, White DC (1986) Determination of monounsaturated double bond
position and geometry for microbial monocultures and complex consortia by capillary
GC-MS of their dimethyl disulphide adducts. J Microbiol Meth 5:49-55

Phelps TJ, Fliermans CB, Garland TR, Pfiffner SM, White DC (1989) Recovery of deep
subsurface sediments for microbiological studies. J Microbiol Meth 9:267-280

0ezanka T, Dor I, Prell A, Dembitsky VM (2003) Fatty acid composition of six freshwater
wild cyanobacterial species. Folia Microbiol 48:71-75

Stephen JR, Chang Y-J, Gan YD, Peacock A, Pfiffner SM, Barcelona MJ, White DC, Mac-
naughton SJ (1999) Microbial characterization of JP-4 fuel contaminated-site using
a combined lipid biomarker/PCR-DGGE based approach. Environ Microbiol 1:231-241

White DC, Kirkegaard RD, Palmer Jr. RJ, Flemming CA, Chen G, Leung KT, Phiefer CB,
Arrage AA (1999) The biofilm ecology of microbial biofouling, biocide resistance and
corrosion. In: Keevil CW, Godfree A, Holt D, Dow C (eds) Biofilms in the aquatic
environment. Roy Soc Chem, Cambridge, UK, pp 120-130

White DC, Pinkart HC, Ringelberg DB (1996) Biomass measurements: biochemical ap-
proaches. In: Hurst CH, Knudsen GR, Mclnerney MJ, Stetzenbach LD, Walter MV (eds)
Manual of environmental microbiology, 1st ed. ASM Press, Washington, DC, pp 91-101

Zhang CL, Li Y, Wall JD, Larsen L, Sassen R, Huang Y, Wang Y, Peacock A, White DC, Horita J,
Cole DR (2001) Lipid and carbon isotopic evidence of methane-oxidizing and sulfate-
reducing bacteria in association with gas hydrates from the Gulf of Mexico. Geology
30:239-242

Zogg GP, Zak DR, Ringelberg DB, MacDonald NW, Pregitzer KS, White DC (1997) Compo-
sitional and functional shifts in microbial communities related to soil warming. Soil Sci
Soc Amer J 61:475-481

Enumeration of Soil Microorganisms

Julia Foght, Jackie Aislabie

13.1

Sample Preparation and Dilution

■ Introduction

Objectives. Soil is a heterogeneous matrix in which microbes are associated
with organic and inorganic soil particles, forming aggregates. The goals of
sample preparation for conventional enumeration techniques are to release
the microbes from the matrix of a representative soil sample, then disperse
them in a suitable diluent so that individual cells can be enumerated ei-
ther by microscopic visualization or cultivation methods. The basic meth-
ods for soil aggregate disruption and dilution have been in common use
for decades, but individual laboratories often develop variations to create
their own empirical "standard methods." Different soil types may be more
amenable to certain diluents or disruption techniques, so, if examining an
unfamiliar soil type, it is wise to test combinations of methods to empir-
ically optimize enumeration results. The presence of inorganic or organic
contaminants (e.g., crude oil) may require adaptation of the basic methods
to disperse the soil sample adequately or dilute a toxicant (e.g., heavy metal).

Principle. A suitable buffered diluent releases microbial cells from the soil
matrix and is used to dilute the suspension to a cell density suitable for the
enumeration method to be used. The dilution method must not compro-
mise the structural integrity of cells to be enumerated by microscopy, nor
the viability of cells for culture-based enumeration.

Theory. Microbes in soil are distributed heterogeneously in microenviron-
ments of different scales and along depth profiles (Foster 1988; Ranjard and
Richaume 2001). Therefore, representative samples of a suitable size must
be collected for accurate enumeration. The number of individual samples
theoretically required to represent the site can be calculated (Alef and Nan-
nip ieri 1995), but in practical terms the number of samples handled is

Julia Foght: Biological Sciences, University of Alberta, Edmonton AB, Canada T6G 2E9,
E-mail: julia.foght@ualberta.ca

Jackie Aislabie: Landcare Research, Private Bag 3127, Hamilton, New Zealand

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

262 J. Foght, J. Aislabie

dictated by the time and resources available. As a compromise, a composite
sample can be prepared from several samples of equal mass or volume, but
statistical evaluation of the data is relinquished. Commonly, at least 10 g wet
mass of soil is used to prepare the first dilution, although the sample size
maybe adjusted according to the soil type and the organisms to be enumer-
ated. Serial dilutions (commonly ten-fold) of soil suspensions are prepared
with sufficient mixing to disrupt soil aggregates and release occluded mi-
crobes into suspension. Physical disruption of the soil aggregates can be
enhanced by inclusion of small (2-3 mm) sterile glass beads in the diluent,
at least in the first dilution. Suitable sterile diluents, of which many exist,
aid the dispersion of soil aggregates. Diluents are often buffered (Strick-
land et al. 1988) and may contain proteins such as gelatin or tryptone to aid
dispersion, glycerol to aid resuscitation of starved bacterial cells (Trevors
and Cook 1992), or a surface active agent such as 0.1% Tween 80, although
surfactants may reduce counts of sensitive Gram-negative cells (Koch 1994).

■ Equipment

• Top-loading balance capable of weighing to 0.1 g

• 150-mL glass dilution bottles and, optionally, approx. 20 g of 2-3 mm
glass beads per bottle to aid in disruption of soil aggregates

• Spatula or small spoon, sterilized by autoclave or by flaming with ethanol

• Sterile pipettes for serial dilutions: 10-mL wide-mouth glass pipettes are
less likely to plug during initial dilutions

• Optional mixing equipment: reciprocating or gyratory shaker for first
dilution; vortex mixer; Waring blender

■ Reagents

• Suitable sterile, buffered diluent dispensed into dilution bottles, usually
90 or 99 mL each

• Suitable diluents include: 0.1% (w/v) sodium pyrophosphate with or
without 1% glycerol (Trevors and Cook 1992); phosphate-buffered saline
(0.85% (w/v) NaCl, 2.2 mM KH 2 P0 4 ; 4.2 mM Na 2 HP0 4 , pH 7) with or
without 0.01% gelatin or peptone (Koch 1994); 1-10 mM potassium
phosphate (pH 7); or mineral salts medium lacking carbon source (Atlas
1995).

■ Sample Collection

Acceptable aseptic techniques for collection and storage of soil samples are
given in Chapt. 1 in this volume. Soil intended for conventional enumeration

1 3 Enumeration of Soil Microorganisms 263

techniques should not be dried because this can reduce the microbial counts
(Sparling and Cheshire 1979; van Elsas et al. 2002). Analyses should be
conducted as soon as possible after sample collection.

■ Procedure

1. On a top-loading balance use sterile spatula to aseptically dispense 10 g
of soil into the first dilution bottle containing 90 mL of diluent and
record exact wet mass of sample added. This is the 10 _1 dilution. Alef
and Nannipieri (1995) recommend using 20 g soil in 180mL of diluent
to reduce the effects of sample heterogeneity.

2. To express the counts on the basis of soil dry mass, dispense a similar
sample into a tared aluminum pan for determining dry mass (in triplicate
for accuracy). Dry the sample at 105 °C to constant mass overnight, and
record mass.

3. Shake or mix the dilution bottle vigorously manually or mechanically
(using reciprocating shaker or Waring blender) to disrupt soil aggre-
gates; recommended times vary from 1 min to 1 h and can be optimized
empirically for different soils.

4. Perform ten-fold dilutions by transferring a 10.0-mL sample from the
center of the dilution bottle to a fresh 90-mL dilution bottle, or hundred-
fold dilutions with 1.0 mL transferred into 99 mL of diluent. Mixing
between dilutions may be performed by hand by vigorously shaking the
bottle 25 times between each transfer, or with a vortex mixer.

5. Continue with ten-fold serial dilutions appropriate to the enumeration
method to be used, e.g., for aerobic heterotrophs in uncontaminated
agricultural soils dilute to 10" 9 for most probable number (Sect. 13.3)
and 10~ 7 for plate counts (Sect. 13.4).

■ Calculation

1. Dilution factor (reciprocal of dilution) = (1/dilution)

2. Dry-mass correction factor = (wet mass of sample/dry mass of sample)

■ Notes and Points to Watch

• The initial sample(s) must be as representative of the soil as possible and
analysis of replicates is recommended.

• Sample preparation and dilutions must be performed in a standardized
manner that can be replicated, so that results from samples taken at

264 J. Foght, J. Aislabie

different times or from different sample sites can be compared with
confidence.

• Soil dilutions should be used immediately after preparation, as storage
of the cell suspension in buffer may decrease the counts observed (Koch
1994).

• The dilution volumes can be scaled down, using test tubes with 1 g of
soil in 9 mL of diluent and mixing by vortex, but caution should be used
because small sample sizes may not be representative.

• A sonicator bath or probe may be used for initial soil sample disrup-
tion (Strickland et al. 1988), but this equipment is not standard in all
laboratories, and excess sonication will reduce counts.

• Aggregates in hydrocarbon-contaminated soils may be difficult to dis-
perse, yielding inaccurate results. Similarly, microbes with highly hy-
drophobic cell surfaces, such as acid-fast hydrocarbon-degrading bacte-
ria, may themselves aggregate and be difficult to disperse.

• If using sodium pyrophosphate as the diluent, adjust the pH to neutrality,
as it is ca. pH 10 without adjustment (Trevors and Cook 1992).

13.2

Direct (Microscopic) Enumeration

■ Introduction

Objectives. It has long been known that enumeration techniques relying on
cultivation of microbes in environmental samples can underestimate the
total number of cells present by orders of magnitude (Skinner et al. 1952;
Amann et al. 1995). This bias can be overcome in part by using molecular
methods (Chapt. 10) or by using direct microscopic observation of cells
where no cultivation is required. Direct enumeration methods can provide
the total number of cells (live plus dead) or may discriminate between live
and dead cells. Some stains differentiate cells based on phylogeny or the
presence of functional genes, providing information about the types of cells
as well as numbers. Microscopy is suitable for direct enumeration of both
bacteria and fungi.

Principle. A known volume of a soil suspension is filtered through a 0.2 p.m
pore size filter. The microbes on the filter are stained with a fluorescent dye
and counted by using an epifluorescence microscope. At least 20 fields each
containing 20-50 cells are counted and the total count is calculated from
the area observed and the volume of suspension filtered.

1 3 Enumeration of Soil Microorganisms 265

Theory. To reduce the bias inherent in culture-based enumeration meth-
ods, total counts of microbes in soil can be observed directly using mi-
croscopy (Fry 1990; Kepner and Pratt 1994; Bottomley 1994; Bloem 1995).
Traditionally, to aid detection, the cells have been stained with fluores-
cent dyes (reviewed by Bolter et al. 2002) such as acridine orange (AO)
or 4 / ,6-diamino-2-phenylindole (DAPI) which stain DNA- containing cells.
Recently, emphasis has been put on differentiating between actively metab-
olizing cells and resting cells, or on discriminating between live and dead
cells. Hence, new fluorescent dyes have been developed. The redox dye 5-
cyano-2,3-ditolyl tetrazolium chloride (CTC), for example, is used to count
active bacterial cells (Creach et al. 2003). CTC is a colorless membrane-
permeable compound that produces a red-fluorescing precipitate in the
cell wall when reduced by the electron transport system of active bac-
terial cells. Staining with a combination of propidium iodide (PI, which
is excluded from cells with intact membranes) and thiazole orange (TO,
which is taken up by both live and dead cells) provides a method for dis-
criminating between live and dead cells. Numerous commercial stain kits
are available with specific instructions for their use, such as Live/Dead
BacLight kits (Molecular Probes, Invitrogen, Carlsbad, CA, USA). The flu-
orescent in situ hybridization (FISH) method, which detects hybridization
of fluorescently-labeled oligonucleotide probes with target DNA or RNA
sequences, can combine total counts with counts of specific phylogenetic
groups (Amman et al. 1995) by detecting multiple overlapping fluorescent
signals, but, like other microscopic methods, suffers from sensitivity biases
(Bolter et al. 2002).

Potential problems encountered when enumerating microbes in soil
include autofluorescence of soil matrix components, particularly in oil-
contaminated soils, and occlusion of cells by soil particles, particularly
clay-sized particles. In the latter case, methods have been developed to
reduce interference by clays (Boenigk 2004) and confocal laser-scanning
microscopy (CLSM) has been used to overcome problems of limited depth-
of-focus in conventional microscopy.

■ Equipment

• Filter membranes (0.2 p.m pore size) for sterilizing reagents

• Black polycarbonate filter membranes (0.2 p.m pore size, 25 mm diame-
ter, e.g., Millipore; Millipore Corp., Billerica, MA, USA)

• 25-mm filter holder unit consisting of a 15-mL glass reservoir and fritted
glass base (wrapped and heat sterilized), clamp, and vacuum flask

• Blunt-tipped filter forceps for handling filter membranes

266 J. Foght, J. Aislabie

• Vacuum pump with fine control

• Glass microscope slides and coverslips, pre-cleaned

• Epifluorescence microscope with appropriate filters

■ Reagents

• All diluents and reagents sterile and particle-free by filtration through
0.2-|^m pore size membrane filters

• Appropriate diluent for sample (Sect. 13.1)

• Fluorescent stains appropriate to target cells: e.g., DAPI stock solution
( 1 mg/mL) in deionized water, freshly diluted to a working concentration
of 1 jig/mL in filtered deionized water, stains protected from light

• Suitable wash solution: e.g., phosphate wash solution (PWS) containing
10 mM KH 2 P0 4 , 0.85% NaCl and 5 mM MgCl 2 • 6H 2

• Non-fluorescent immersion oil

■ Sample Preparation

Prepare suitable dilutions of soil sample (Sect. 13.1) in sterile, particle-free
diluent.

■ Procedure

1. Prepare dilution series as required in filter-sterilized diluent. Vigorously
mix sample for 5 min and allow suspension to stand for approx. 30 min
to let larger soil particles settle out. If the sample will be kept longer
than 30 min before counting, add a preservative (e.g., filter-sterilized
formaldehyde to final concentration 3.7% or electron-microscopy-grade
glutaraldehyde to final concentration 2.5%).

2. Place black filter membrane in filter unit, add PWS (e.g., 4 ml) to column
reservoir and known volume (e.g., 0.1 mL) of diluted soil suspension,
avoiding settled soil particles. Perform subsequent steps under reduced
lighting for light-sensitive stains like DAPI.

3. Add required volume of stain (e.g., 1 mL DAPI working solution) to
sample in column reservoir and stain in the dark for 7-10 min.

4. Filter slowly through membrane under gentle vacuum. Rinse sides of
column reservoir gently with diluent (two- to three-fold of initial volume)
and allow filter to air dry.

1 3 Enumeration of Soil Microorganisms 267

5. Place a drop of immersion oil on a glass microscope slide, place the
membrane filter on top, and cover with a coverslip. Follow with a drop
of immersion oil and examine under an epifluorescence microscope at
correct wavelength with appropriate filters.

6. Count at least 20 fields of view (FOV) each containing 20-50 cells. Count
randomly located FOV covering a wide area of the filter, avoiding its
edges.

7. Blanks consisting only of reagents should be performed at intervals, or at
least at the beginning and end of sample enumeration. Blanks should be
< 5% of the total cell densities in the samples and should be subtracted
from sample counts before calculation of total numbers.

■ Calculation

Counts are calculated on the basis of wet mass of soil, corrected for back-
ground, and usually expressed on the basis of dry mass of soil.

- Cells/g soil wet mass =

total no. of cells counted total stained area 1

x x

total no. of FOV area of FOV mass of soil on filter

- Cells/g soil dry mass =

(cells/g soil wet mass) x (dry-mass conversion factor)

A specific example is given:

- Area of FOV = 0.01 mm 2

- Stained area of filter = nr 2 = 176.8 mm 2

(diameter of the filter area covered by filtrate = 15 mm)

- Total counts in 20 FOV for 0.1 mL of 10" 3 dilution = 929

- Total counts in 20 FOV for reagent blanks = 40

- Mass of soil on filter = 0.1 mL of 10~ 3 dilution = 10~ 4 g soil wet mass

- Dry mass conversion factor (Sect. 13.1) = 1. 18
Cells/g soil wet mass =

(929 - 40) cells 176.8 mm 2 1 q
x x - 7 9 x 10

20 FOV 0.01mm 2 10" 4 g

- Corrected count = 7.9 x 10 9 x 1. 18 = 9.3 x 10 9 cells/g soil dry mass

268 J. Foght, J. Aislabie

■ Notes and Points to Watch

• An analysis of the sources of variation in the direct count method (Kirch-
man et al. 1982) emphasizes the importance of enumerating replicate
filters to reduce error.

• Starving ("dwarf) cells and ultramicrobacteria (< 0.5 pm diameter)
may not be not retained on the filter membrane or may not be detected
by activity stains (Bolter et al. 2002).

• At low cell densities it is difficult to achieve statistically valid counts, and
efforts must be made to concentrate the sample if possible.

• Hydrocarbon-contaminated samples may suffer from autofluorescence
and poor disruption of aggregates.

13.3

Enumeration by Culture in Liquid Medium

(Most Probable Number Technique)

■ Introduction

Objectives. The Most Probable Number (MPN) method uses statistics to
infer the number of viable organisms in a sample that are able to grow or
metabolize in a liquid medium under given incubation conditions. MPN
tests can be carried out in large volumes in bottles or test tubes, or in
microliter volumes in microtiter well plates, depending on the sample and
the viability assay.

Different media can be used to enumerate both generalist and specialist
microbes in the soil. Total heterotrophs (generalists) can be enumerated in
complex medium, although full-strength medium such as trypticase soy
broth may not be suitable for enumerating microbes in nutrient-poor soils;
for such samples tenth- strength medium maybe appropriate (Alef and Nan-
nipieri 1995). The MPN method can be customized to differentiate among
specialists by providing selective growth substrates. For example, mineral
medium can be supplemented with filter-sterilized crude oil or refined
product (e.g., diesel fuel) to enumerate "total hydrocarbon degraders" or
amended with specific hydrocarbon substrates representing aliphatic and
aromatic components (e.g., n-hexadecane and naphthalene, respectively).
Liquid hydrocarbons can be added directly to broth whereas solid hydro-
carbons can be provided as a fine suspension of crystals or dissolved in
a non-metabolized water-immiscible carrier such as heptamethylnonane
(Efroymson and Alexander 1991). Volatile hydrocarbons may be supplied
in the vapor phase although this can be technically cumbersome.

1 3 Enumeration of Soil Microorganisms 269

Positive tubes maybe identified by various criteria, including: increased
turbidity due to growth; emulsification of crude oil (e.g., "Sheen Screen,"
Brown and Braddock 1990); production of colored metabolites, particularly
from some aromatic substrates (Stieber et al. 1994; Wrenn and Venosa
1996); reduction of an iodonitrotetrazolium (INT) dye after incubation to
indicate metabolism of substrates (Wrenn and Venosa 1996; Johnsen et al.
2002); or evolution of 14 C0 2 from radiolabeled substrates (Carmichael and
Pfaender 1997). It is important that both positive and negative controls be
included with these tests.

Principle. The microorganisms in a soil sample are serially diluted to ex-
tinction, inoculated in replicate into a suitable medium, and incubated
under appropriate conditions to yield a series of cultures that is scored
according to pre-determined criteria. The combination of positive and
negative cultures after incubation is evaluated by statistical methods to
infer the MPN of viable cells in the undiluted sample.

Theory. Culture-based enumeration methods such as MPN and plate count
assay (Sect. 13.4) are biased because only a small proportion of environ-
mental microbes has been cultured (Amann et al. 1995). With improved
culture-based studies (e.g., Connon and Giovannoni 2002), the bias im-
posed by growth-based methods will lessen, but it must be considered
when interpreting results. The advantage to growth-based enumeration
over molecular methods is that the former is technically simpler, usually
easy to interpret, and can yield isolates for further investigation. The ad-
vantage over plate count methods is that MPN is suitable for particulate
samples (such as soil dilutions) that would obscure plate counts at low
dilutions, and can detect microbes that will not grow on solid medium or
are a minor component of a mixed culture. The disadvantages of MPN are
that it yields only a statistical estimate of the viable microbes present and
it requires many tubes and manipulations compared with plate counts.

Typically a decimal dilution series is prepared in suitable diluent and
a fixed volume of each dilution is inoculated into medium in replicate
cultures, usually in multiples of 3, 5, or 10. MPN tests can be conducted in
tubes, vials, or bottles, generally containing 7-10 mL medium per test tube,
or in microtiter plates with 200 yiL per well. After incubation the tubes are
scored qualitatively for criteria such as growth, production of metabolites,
or loss of substrate.

The combination of positive and negative cultures is converted to the
MPN and confidence intervals either by consulting standard probability
tables (e.g., Eaton et al. 1995; Alef and Nannipieri 1995) or using an algo-
rithm (Koch 1994). The method assumes that (1) the microorganisms have
been distributed into the cultures such that the highest dilution positive
tubes were inoculated with a single organism, (2) culture tubes inoculated

270 J. Foght, J. Aislabie

with as few as one viable microbe will produce a positive result, and (3) the
microbes have not been injured or rendered non-viable during sample
handling.

■ Equipment

• Pipettes

• Sterile test tubes or microtiter plates

• Vortex mixer for mixing inoculum into medium (optional)

• Incubation chamber with suitable temperature control and headspace
(e.g., for anaerobes)

• Microtiter plate reader for measuring color changes or optical density
(optional)

• Solvent-resistant filters (e.g., Millex-FG, Millipore Corp.) for filter ster-
ilizing hydrocarbon solutions (optional)

■ Reagents

• Appropriate diluent for sample (Sect. 13.1)

• Sterile liquid or semi-solid medium suitable for growth of target organ-
ism^). For enumeration of generalists, standard or dilute liquid media
(Alef and Nannipieri 1995, Atlas 1995) are appropriate; for enumeration
of specialists, a mineral salts medium amended with selective carbon
sources such as hydrocarbons maybe used (Sect. 13.4).

• Specialty chemicals, depending on criteria for positive cultures, such as
radiolabeled substrates, endpoint reagents, carrier solvents, etc.

• Filter-sterilized liquid hydrocarbons or stock solutions of solid hydro-
carbons dissolved in ethanol or dimethylformamide, for use as selective
carbon sources (optional)

■ Sample Preparation

Perform serial dilutions of a representative soil sample in appropriate dilu-
ent (Sect. 13.1), to exceed the expected viable number of cells by one or two
orders of magnitude.

■ Procedure

1. Dispense replicate volumes of growth medium into suitable receptacles
(e.g., 10 mL in test tubes, 200 pi per well for microtiter plates). Prepare

1 3 Enumeration of Soil Microorganisms 27 1

replicates (typically 3, 5, or 10) for each sample dilution to be tested.
Medium must contain complete nutrients for growth including carbon
source, and may contain indicators such as dyes or radiolabeled sub-
strates.

2. Inoculate replicate tubes with fixed volume of diluted sample (e.g., 1.0 mL
for tubes, 100 pL for microtiter wells) covering at least three decimal
dilutions.

3. Include negative controls (uninoculated medium) and positive controls
(medium inoculated with a culture known to produce a positive result)
for reference.

4. Incubate 7-14 days or longer in the dark under suitable conditions,
taking into account in situ conditions of temperature, 2 levels, etc.

5. Score tubes at intervals for positive results. Continue to incubate until
two successive readings give the same results. Positive indicators include
turbidity (e.g., heterotrophs growing in complex medium), hydrocarbon
emulsification, production of soluble or gaseous metabolic end products
(e.g., 14 C0 2 evolution from radiolabeled substrates, methane, colored
metabolites), and changes in indicators (pH indicators, redox dyes).

6. Identify the highest dilution set with all tubes positive, and the next two
higher dilution sets. Use the pattern of positive and negative tubes with
standard probability tables (e.g., Alef and Nannipieri 1995, Eaton et al.
1995) to calculate the MPN from the dilution factor of the middle set.
When non-standard patterns are encountered, follow the recommended
variations provided with the tables for calculating the MPN.

■ Calculation

Published tables of statistical probability (Alef and Nannipieri 1995; Eaton
et al. 1995; tables are also available on several internet sites such as US Food
& Drug Administration) are used to convert the pattern of positive and
negative tubes into the MPN of viable microbes in the original sample. The
dilution factor and volume of sample used to inoculate the tubes are used in
calculation but the volume of growth medium used in the tubes is not taken
into consideration. Sample volumes reported in standard MPN tables are
designed for water samples and are usually expressed per 1 00 mL of sample;
therefore, they must be corrected for the actual volume of inoculum used
in the test. Values are calculated as the MPN ± 95% confidence intervals
(provided with the tables) and expressed on the basis of soil dry mass by
multiplying the MPN by the dry-mass correction factor (Sect. 13.1).

The simple algorithm below (Eaton et al. 1995) can be used to calcu-
late the MPN without consulting published tables but does not provide

272 J. Foght, J. Aislabie

confidence intervals.
MPN/lOOmL

number of positive tubes x 100
^(mL sample in negative tubes) (mL sample in all tubes)

■ Notes and Points to Watch

• Match the incubation conditions to in situ conditions when feasible.
For example, select an appropriate culture incubation temperature (in-
cluding temperature of the diluent and medium when inoculating), pro-
vide semi-solid medium for enumeration of microaerophiles, anaerobic
medium and headspace for anaerobes, etc.

• If aerobic tubes are sealed, ensure that there is adequate headspace to
maintain aerobic conditions if extended incubation will be required.

• To use a high proportion of sample to growth medium, increase the
strength of the medium (e.g., use double strength medium for 100%
(v/v) inoculum).

• When using turbidity as the criterion for growth, be aware of the tur-
bidity contributed by soil particles at low dilutions, and by particulate
substrates (e.g., suspensions of polycyclic aromatic hydrocarbon crys-
tals).

• If providing low molecular mass hydrocarbons as a carbon source avoid
toxicity to the inoculum by minimizing substrate volumes.

• Multiple MPN tests can be performed on a soil sample to enumerate
different specialist components of the soil microbiota. If generalist MPN
tests or direct counts (Sect. 13.2) are also performed, the specialists can
be expressed as a proportion of the total viable numbers present in the
sample.

• After incubation, the positive MPN tubes may be suitable to use as an
inoculum for subsequent isolation of pure cultures.

13.4

Enumeration by Culture on Solid Medium

(Plate Count Technique)

■ Introduction

Objectives. The plate count technique quantifies the viable microbes in
a sample by counting the number of colonies that form on or in a solid

1 3 Enumeration of Soil Microorganisms 273

growth medium inoculated with dilutions of that sample. Each colony is
assumed to have originated from a single propagule or "colony forming
unit" (CFU), whether that be a bacterial cell, endospore, hyphal fragment,
or spore. Non-selective growth medium may be used to cultivate gener-
alists, or selective medium may be used to enumerate specialists such as
hydrocarbon-degrading bacteria and fungi. Specific enumeration of acti-
nomycetes, filamentous fungi, or yeasts usually requires specialized media
to suppress unwanted soil microbes such as spreading or mucoid colonies
that overgrow the slower-growing colonies on non-selective plates (Labeda
1990). Alternatively, a differential assay can be applied after the colonies
have grown, to distinguish those possessing specific metabolic capabilities
(e.g., production of colored metabolites; Kiyohara et al. 1982). Plates can be
incubated under different atmospheres to enumerate aerobes, anaerobes,
or microaerophiles, or at different temperatures to cultivate psychrotoler-
ant, mesophilic, or thermophilic microbes. Plate counts maybe performed
using several media and incubation conditions to enumerate different sub-
sets of the viable microbes in a soil sample.

Principle. Dilutions of a soil sample, performed in suitable diluent, are
inoculated in replicate onto solid medium for cultivation with or without
selection for specific metabolic types. Plates containing 30-300 colonies are
selected and the colonies counted so that the CFU can be calculated for the
original soil sample using dilution factors and dry-mass correction factors.

Theory. It has long been recognized that the plate count method under-
estimates the actual number of living cells in the sample by one or more
orders of magnitude (Skinner et al. 1952) because soil organisms may not
be viable or are not cultivable under the conditions employed (Amann et
al. 1995). The proportion of viable cells enumerated will depend on the soil
and on the growth medium and incubation conditions. New strategies for
enumerating previously uncultured microbes are being devised to alleviate
the cultivation bias (e.g., Joseph et al. 2003; Stevenson et al. 2004), but selec-
tivity will always be a disadvantage of the plate count method (or any other
cultivation-based enumeration method) compared with direct counts or
molecular methods. The advantages are that the plate count method is rel-
atively rapid and inexpensive and yields well-separated colonies suitable
for subsequent purification and characterization.

The medium and incubation conditions used in the plate count method
determine which metabolic types of microbes will be enumerated, but
the primary assumption for all plate counts is that each colony arises
from a single viable propagule, i.e., a colony forming unit (CFU). After
incubation, colonies are counted only on those plates containing 30-300
colonies, for statistically valid enumeration (Koch 1994). Variations on the
basic method exist, including spread plates appropriate for aerobic bacteria

Vapor

Spray

Overk

274 J. Foght, J. Aislabie

Table 13.1. Recommended methods for providing hydrocarbons on solid medium

Substrate Suitable for:

Spread
Toluene, xylenes
Naphthalene
Phenanthrene
Dibenzothiophene
Pyrene

Octane +J

Hexadecane *J

Jet fuel aJ

Crude oil *J

and yeast, and pour plates suitable for microbes that do not grow well on
surfaces but form subsurface colonies at reduced oxygen tension. Pour
plates also help reduce problems with spreading colonies. Several methods
can be used to supply substrates to mineral salts agar, including overlayer
plates, spray plates and vapor plates (Table 13.1). Plates are incubated in
the dark under suitable conditions of temperature and aeration, often for
extended periods of time (e.g., 2-3 months) to enumerate slow-growing
species (Janssen et al. 2002).

■ Equipment

• Sterile bent glass rod spreaders ("hockey sticks") for inoculating plates

• Flame and beaker of ethanol to surface-sterilize spreaders

• Manual turntable for turning Petri plates while spreading inoculum
(optional)

• Incubators with suitable atmosphere (aerobic or anaerobic) and temper-
ature setting

• Water bath at 50 °C for pour plates or overlayer plates

• Aerosol spray apparatus (e.g., Jet-Pak, Sherwin-Williams Co., Cleveland)
for applying ether solution of substrate to agar surface, or sealed con-
tainers for incubating vapor plates (optional)

■ Reagents

• Solid growth medium in Petri plates, sufficient to inoculate 3 or 5 plates
per dilution. Generalist media for bacteria include Plate Count Agar, Nu-
trient Agar and R2A agar (Difco; Becton, Dickinson & Co., Sparks, MD,

1 3 Enumeration of Soil Microorganisms 275

USA) among many others. General mycological media include Czapek-
Dox Agar (Difco), Malt Extract Agar (Difco) and Mycobiotic ("Mycosel")
Agar (Acumedia, Neogen; Lansing, MI, USA), usually containing antibi-
otics (e.g., oxytetracycline at lOOmg/L and/or streptomycin at 30mg/L)
to suppress bacterial growth. Selective agar for hydrocarbon degraders is
usually a mineral medium such as Bushnell Haas (Difco; Atlas 1995) so-
lidified with 1.5% (w/v) Purified Agar (Oxoid, Basingstoke, UK) or Agar
Noble (Difco) or 0.8% (w/v) gellan gum (Gelrite; Serva, Heidelberg) and
amended with a specific carbon source.

• Overlayer medium is usually prepared with agarose or purified agar
(Bogardt and Hemmingsen 1992).

• An ether or acetone solution of hydrocarbon substrate is used for spray
plates.

■ Sample Preparation

Dilute sample appropriately in suitable diluent (Sect. 13.1) to exceed ex-
pected number by at least one order of magnitude.

■ Procedure

Spread Plates (Bacteria, Yeasts, or Filamentous Fungi)

1. Pipette a fixed volume of inoculum (typically 0.1 mL or 1.0 mL) from
a range of dilutions onto three or five replicate agar plates and spread
evenly using sterile bent glass rod. When inoculum has been absorbed
into agar, invert plates and place in plastic bag to maintain humidity.

2. Incubate at suitable temperature (e.g., ca. 25 °C for temperate soils) and
under appropriate atmosphere, noting the appearance of colonies on
plates with 30-300 colonies. Continue incubating until the number of
colonies is constant. This may take less than a week for fast-growing
bacteria, or more than 2 months for slow-growers or plates incubated
at low temperatures. For extended incubation periods, seal the edges of
plates with laboratory film or tape to prevent drying.

3. Count the colonies on replicate plates having 30-300 colonies, determine
the mean and calculate the CFU per gram dry mass in the original sample
using the dilution factor and dry-mass correction factor.

4. To enumerate colonies that can grow on liquid hydrocarbons spread
a small volume (e.g., 50 p.L per plate) onto the surface of mineral medium
agar either before or after inoculation, leaving small droplets on the agar
surface. For solid hydrocarbons, see spray plate method below.

276 J. Foght, J. Aislabie

Pour Plates (Bacteria or Yeasts)

1. Prepare 20 mL aliquots of molten growth medium agar and bring to ca.
50 °C in a water bath.

2. Add 1.0 mL inoculum to agar, mix briefly, and immediately pour into an
empty sterile Petri plate. Alternatively, add inoculum to an empty sterile
Petri plate, pour molten agar on top and mix by rotating the plate on the
bench top. Allow agar to solidify then incubate as for spread plates.

3. Count both surface and subsurface colonies.

Overlayer Plates (Bogardt and Hemmingsen 1992)

1. Prepare Petri plates containing 20 mL mineral medium with or without
carbon source, depending on whether the overlayer contains a carbon
source.

2. Prepare 5mL sterile molten 1.5% (w/v) agarose or purified agar con-
taining a suspension of particulate, insoluble substrate at a nominal
concentration sufficient to provide an opaque suspension of fine crys-
tals. It may be necessary to dissolve the substrate in a small volume of
ethanol before adding to the agarose. Bring to 50 °C in a water bath.

3. Add inoculum to the molten agarose, mix briefly, and immediately pour
onto the mineral medium base, tipping the plate to distribute the over-
layer evenly. The overlayer should be somewhat opaque. Allow to solidify
then incubate as for spread plates. Alternatively, carefully inoculate the
surface of the overlay as for spread plates.

4. Count colonies that have a surrounding halo of clearing or colored
metabolites (Fig. 13.1).

Spray Plates for Solid Hydrocarbons (Kiyohara et al. 1982)

1. Prepare a solution of crystalline hydrocarbon (e.g., phenanthrene or
dibenzothiophene) in either acetone or anhydrous ethyl ether. CAU-
TION: Ethyl ether is highly flammable and explosive. All procedures
must be carried out in a well-vented fume hood away from any sparks or
flames. Protective clothing and gloves must be worn to prevent exposure
to hydrocarbon mist and precautions, such as spraying inside a card-
board box in the hood, should be taken to prevent contaminating the
fume hood with potentially carcinogenic compounds. The concentration
of the solution does not need to be precise; approx. 10 mg of hydrocarbon
dissolved in 2 mL of solvent should be sufficient to cover one plate.

2. Use an aerosol canister to deliver a fine spray of solution to the surface
of an inoculated plate. The surface should become slightly opaque with

13 Enumeration of Soil Microorganisms

277

*t*> ^i .

Fig. 13.1. Colonies capable of degrading
carbazole form "haloes" of clearing in an
overlayer plate prepared with carbazole.
(From Shotbolt- Brown et al. 1996 with per-
mission)

a thin, even layer of very fine crystals. Seal edges of plates with laboratory
film and place in plastic bags to prevent cross-contamination by vapors.

3. Incubate and score for appearance of colonies. If the substrate can serve as
a carbon source (e.g., phenanthrene), use mineral medium agar lacking
a carbon source and score for colonies that are surrounded by zones of
clearing and are larger than those observed on a parallel control plate
lacking spray. If the substrate does not serve as a carbon source but can
be co-metabolized (e.g., dibenzothiophene), use a low- nutrient agar that
provides a carbon source and score for production of colored metabolites
and/or zones of clearing around the colonies.

Vapor Plates for Volatile Hydrocarbons

1. Inoculate mineral medium agar by the spread plate method.

2. For volatile solid hydrocarbons such as naphthalene, add a few crystals
(< 0.1 g) to the lid of each inverted Petri plate, seal the edges of the
plate with laboratory film, and incubate in sealed plastic bags to prevent
cross-contamination on vapors.

3. Volatile liquid hydrocarbons such as xylenes or jet fuel can be supplied
to individual plates by placing a few drops of hydrocarbon into a plastic
pipette tip stuffed with glass wool and placed on the lid of an inoculated,
inverted mineral salts agar plate. Seal and incubate as above. To supply
vapor to several plates at once, place inoculated plates into a sealable
container with a small beaker containing a "wick" of glass wool or folded
filter paper, and add a small amount of hydrocarbon, just sufficient to

278 J. Foght, J. Aislabie

saturate the headspace for a day or so, to reduce the chance of toxicity.
Replenish hydrocarbon as necessary.

■ Calculation

1. Count the number of colonies arising on the plates, or the number
showing the desired phenotype.

2. Calculate the mean value for replicates, correct for dilution and dry mass,
and express as CFU or phenotype-positive colonies per gram dry mass
of original soil.

■ Notes and Points to Watch

• When spreading inoculum on a plate, make sure that the glass rod is not
too hot. Similarly, do not use pour plates or overlayer plates to enumerate
psychrophiles.

• Incubate plates at temperatures close to those in situ when practical.

• When enumerating specialist populations (e.g., hydrocarbon degraders),
positive and negative controls should be included.

• Antibiotics should be prepared as filter-sterilized concentrated solutions
and added to cooled molten agar immediately before dispensing the agar
into plates. Protect plates from the light before use and during incubation.

• The use of low- nutrient media for enumeration has been recommended
by some researchers, as discussed in Sect. 13.3.

• Davis et al. (2005) suggest that plates with a minimum often colonies per
plate rather than 30-300 colonies should be used for enumeration. This
reduces depression in viable counts due to over-crowding of colonies on
plates leading to inhibition of some species by others, or alternatively
the depletion of nutrients by fast growing colonies that prevents slow
growers from reaching a countable size.

• Problems may arise with long-term incubation of plates for enumeration
of slow-growing bacteria and fungi, including drying of the plates, ap-
pearance of "spreading" or mucoid bacterial colonies or fungal colonies
that obscure other colonies.

• Plate counts over- represent genera that sporulate profusely (e.g., Peni-
cillium, Trichoderma spp., Streptomyces spp.), and under-estimate or
exclude fastidious genera.

• Do not use too much solvent solution for spray plates, as the solvent can
injure the inoculum.

1 3 Enumeration of Soil Microorganisms 279

References

Alef K, Nannipieri P (1995) Methods in applied soil microbiology and biochemistry. Aca-
demic Press, London

Amann RI, Ludwig W, Schleifer, K-H (1995) Phylogenetic identification and in situ detection
of individual microbial cells without cultivation. Microbiol Rev 59:143-169

Atlas RM (1995) Handbook of media for environmental microbiology. ASM Press, Washing-
ton DC

Bloem, J (1995) Fluorescent staining of microbes for total direct counts. In: Akkermans
ADL, van Elsas JD, de Bruijn FJ (eds), Molecular microbial ecology manual. Kluwer
Academic Publ, Dordrecht, The Netherlands, pp 1-12

Boenigk J (2004) A disintegration method for direct counting of bacteria in clay- dominated
sediments: dissolving silicates and subsequent fluorescent staining of bacteria. J Micro-
biol Meth 56:151-159

Bogardt AH, Hemmingsen BB (1992) Enumeration of phenanthrene-degrading bacteria by
the overlayer technique and its use in evaluation of petroleum-contaminated sites. Appl
Environ Microbiol 58:2579-2582

Bolter M, Bloem J, Meiners K, Moller R (2002) Enumeration and biovolume determination
of microbial cells - a methodological review and recommendations for applications in
ecological research. Biol Fertil Soils 36:249-259

Bottomley PJ (1994) Light microscopic methods for studying soil microorganisms. In:
Weaver RW, Angle S, Bottomley P, Bezdicek D, Smith S, Tabatabai A, Wollum A (eds)
Methods of soil analysis, Part 2, Microbiological and biochemical properties. Soil Sci
Soc Am Book Series No. 5, Madison WI, pp 81-105

Brown EJ, Braddock JF (1990) Sheen screen, a miniaturized most-probable-number method
for enumeration of oil-degrading microorganisms. Appl Environ Microbiol 56:3895-
3896

Carmichael LM, Pfaender FK (1997) Polynuclear aromatic hydrocarbon metabolism in soils:
relationship to soil characteristics and preexposure. Environ Toxicol Chem 16:666-
675

Connon SA and Giovannoni SJ (2002) High-throughput methods for culturing microor-
ganisms in very-low-nutrient media yields diverse new marine isolates. Appl Environ
Microbiol 68:3878-3885

Creach V, Baudoux A-C, Betru G, Le Rousiz B (2003) Direct estimate of active bacteria: CTC
use and limitations. J Microbiol Meth 52:19-28

Davis KER, Joseph SJ, Janssen PH (2005) Effects of growth medium, inoculum size, and in-
cubation time on the culturability and isolation of soil bacteria. Appl Environ Microbiol
71:826-834

Eaton AD, Clesceri LS, Greenberg AE (1995) Standard methods for the examination of
water and wastewater, 19 th Edition, American Public Health Association, Washington
DC, Section 9221

Efroymson RA, Alexander M (1991) Biodegradation by an Arthrobacter species of hydro-
carbons partitioned into an organic solvent. Appl Environ Microbiol 57:1441-1447

Foster RC (1988) Microenvironments of soil microorganisms. Biol Fertil Soils 6:189-203

Fry JC (1990) Direct methods and biomass estimation. In: Grigorova R, Norris JF (eds)
Methods in Microbiology, vol 22, Techniques in microbiology. Academic Press, London,
pp 41-85

Janssen PH, Yates PS, Grinton BE, Taylor PM, Sait M (2002) Improved culturability of soil
bacteria and isolation in pure culture of novel members of the divisions Acidobacteria,
Actinobacteria, Proteobacteria, and Verrucomicrobia. Appl Environ Microbiol 68:2391-
2396

280 J. Foght, J. Aislabie

Johnsen AR, Bendixen K, Karlson U (2002). Detection of microbial growth on polycyclic

aromatic hydrocarbons in microtiter plates by using the respiration indicator WST-1.

Appl Environ Microbiol 68:2683-2689
Joseph SJ, Hugenholtz P, Sangwan P, Osborne CA, Janssen PH (2003) Laboratory cultivation

of widespread and previously uncultured soil bacteria. Appl Environ Microbiol 69:7210-

7215
Kepner RL, Pratt JR (1994) Use of fluorochromes for direct enumeration of total bacteria in

environmental samples - past and present. Microbiol Rev 58:603-615
Kirchman D, Sigda J, Kapuscinski R, Mitchell R (1982) Statistical analysis of the direct count

method for enumerating bacteria. Appl Environ Microbiol 44:376-382
Kiyohara H, Nagao K, Yano K (1982) Rapid screen for bacteria degrading water- insoluble,

solid hydrocarbons on agar plates. Appl Environ Microbiol 43:454-457
Koch AL (1994) Growth measurement. In: Gerhardt P, Murray RGE, Wood WA, Krieg NR

(eds) Methods for general and molecular bacteriology. ASM Press, Washington DC,

pp 248-292
Labeda DP (1990) Isolation of biotechnological organisms from nature. The McGraw-Hill

Environmental Biotechnology Series, McGraw-Hill, New York.
Ranjard L, Richaume A (2001) Quantitative and qualitative microscale distribution of bac-
teria in soil. Res Microbiol 152:707-716
Shotbolt-Brown J, Hunter DWF, Aislabie J (1996) Isolation and description of carbazole-

degrading bacteria. Can J Microbiol 42:79-82
Skinner FA, Jones PCT, Mollison JE (1952) A comparison of a direct- and a plate-counting

technique for the quantitative estimation of soil micro-organisms. J Gen Microbiol

6:261-271
Sparling GP, Cheshire MV (1979) Effects of soil drying and storage on subsequent microbial

growth. Soil Biol Biochem 11:317-319
Stevenson BS, Eichorst SA, Wertz JT, Schmidt TM, Breznak JA (2004) New strategies for

cultivation and detection of previously uncultured microbes. Appl Environ Microbiol

70:4748-4755
Stieber M, Haeseler F, Werner P, Frimmel FH (1994) A rapid screening method for microor-
ganisms degrading polycyclic aromatic hydrocarbons in microplates. Appl Microbiol

Biotechnol 40:753-755
Strickland TC, Sollins P, Schimel DS, Kerle EA (1988) Aggregation and aggregate stability

in forest and range soils. Soil Sci Soc Am J 52:829-833
Trevors JT, Cook S (1992) A comparison of plating media and diluents for enumeration of

aerobic bacteria in a loam soil. J Microbiol Meth 14:271-275
Van Elsas JD, Smalla K, Lilley AK, Bailey MJ (2002) Methods for sampling soil microbes In:

Hurst CT, Crawford RL, Knudson GR, Mclnerney MJ, Stetzenbach LD (eds) Manual of

environmental microbiology. 2 nd Edition. ASM Press, Washington DC, pp 505-515
Wrenn BA, Venosa AD (1996) Selective enumeration of aromatic and aliphatic hydrocarbon

degrading bacteria by a most-probable-number procedure. Can J Microbiol 42:252-258

Quantification of Soil Microbial Biomass
by Fumigation-Extraction

Rainer Georg Joergensen, Philip C. Brookes

14.1

General Introduction

The soil microbial biomass responds much more quickly than most other
soil fractions to changing environmental conditions, such as variations
in substrate input (e.g., Powlson et al. 1987) or increases in heavy metal
content (Brookes and McGrath 1984). Much research supports the orig-
inal idea of Powlson and Jenkinson (1976) that the biomass is a much
more sensitive indicator of changing soil conditions than, for example,
the total soil organic matter content. Thus the biomass can serve as an
"early warning" of such changes, long before they are detectable in other
ways. Biomass measurements are certainly useful in studies of soil pro-
tection. They have the advantage that they are relatively cheap and sim-
ple as well as being rapid. There is now a considerable amount of lit-
erature to show that these measurements are useful in determining ef-
fects of stresses on the soil ecosystem. Measurements of the soil microbial
biomass by the fumigation extraction method have been used to estimate
the environmental effects of pesticides (Harden et al. 1993) and antibiotics
(Castro et al. 2002). This method has been repeatedly used to monitor
successfully the bioremediation process of fuel oil contaminated soil (Joer-
gensen et al. 1 994a, 1 994b, 1995,1997; Plante and Voroney 1 998; Franco et al.
2004).

Linked parameters (e.g., biomass-specific respiration or biomass as
a percentage of soil organic C) are also useful because they possess "internal
controls" (see Barajas Aceves et al. 1999 for a discussion). This fact permits
interpretation of measurements in the natural environment, where, unlike
in controlled experiments, there may not be suitable non-contaminated soil
(for example) to provide good "control" or "background" measurements
(Brookes 1995).

Rainer Georg Joergensen: Department of Soil Biology and Plant Nutrition, University of Kas-
sel, Nordbahnhofstr. la, 37213 Witzenhausen, Germany, E-mail: joerge@wiz.uni-kassel.de

Philip C. Brookes: Agriculture and Environment Division, Rothamsted Research, Harpen-
den, Herts., AL5 2JQ, UK

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

282 R.G. Joergensen, RC. Brookes

14.2

Fumigation and Extraction

■ Introduction

Objectives. The biomass of a microbial community can be quantitatively
determined by fumigation and extraction in a large variety of soils devel-
oped under very different environmental conditions, especially in contam-
inated and remediated soils.

Principle. Soils are fumigated with chloroform, incubated for 24 h, and ex-
tracted. Different components can then be measured in the extracts, using
various methods (Sects. 14.3-14.4). Non-fumigated soil is also extracted to
correct for non-biomass soil organic matter.

Theory. Following chloroform fumigation of soil, there is an increase in
the amount of various organic and inorganic components coming from
the cells of soil microorganisms (Jenkinson and Powlson 1976). The mem-
branes of living soil microorganisms are partially lysed by the fumigant
chloroform. After a 24 h incubation period to allow autolysis, a large part
of the soil microbial biomass can be extracted from fumigated soil. The
amount additionally rendered extractable from killed microorganisms is
proportional to the original microbial biomass. Organic C (Vance et al.
1987), total N and NH 4 -N (Brookes et al. 1985), and ninhydrin-reactive N
(Joergensen and Brookes 1990) can be measured in the same 0.5 M K 2 S0 4
extract (Alef and Nannipieri 1995). Organic C (Joergensen 1995) and total
S (Wu et al. 1993) can be measured after extraction with 0.01 M CaCl 2 and
phosphate or total P after extraction with NaHC0 3 (Brookes et al. 1982).

■ Equipment

• Room, incubator, or water bath adjustable to 25 °C

• Implosion-protected desiccator

• Vacuum line (water pump or electric pump)

• Horizontal or overhead shaker

• Deep-freezer at -15 °C

• Folded filter papers (e.g., Whatman 42 or Schleicher & Schuell 595 1/2)

• Glass conical flasks (250 mL)

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 283

■ Reagents

• Ethanol-free chloroform (CHCI3)

• Soda lime

• 0.5MK 2 SO 4

■ Sample Preparation

Use field-moist, sieved (between < 2 and < 5 mm) soil.

■ Procedure

1. Divide a moist soil sample of 50 g into two subsamples of 25 g.

2. Place the non-fumigated control samples in 250 mL conical flasks, extract
immediately with 100 mL 0.5 M K 2 S0 4 (extractant-to-soil ratio of 4:1)
for 30 min by oscillating shaking at 200 rpm (or 45 min overhead shaking
at 40 rpm), filter through a folded filter paper.

3. For the fumigation treatment, place 50-mL glass vials containing the
moist soil into a desiccator containing wet tissue paper and a vial of
soda lime, add a beaker containing 25 mL ethanol-free CHC1 3 and a few
boiling chips and evacuate the desiccator until the CHC1 3 has boiled
vigorously for 2 min.

4. Incubate the desiccator in the dark at 25 °C for 24 h. After fumigation,
remove CHC1 3 by repeated (six- fold) evacuation and extract with 0.5 M
K 2 S0 4 as described above.

5. Store 0.5 M K 2 S0 4 extracts at -15 °C prior to analysis of organic C, total
N, or ninhydrin-reactive N.

■ Notes and Points to Watch

• The desiccator must be kept under vacuum for 24 h to ensure the presence
of a CHCI3 atmosphere, which kills virtually all soil microorganisms.

• Ethanol-free CHC1 3 must be used to measure microbial biomass C be-
cause ethanol cannot be completely removed from the soil after fumiga-
tion. Ethanol-stabilized CHC1 3 can be used if solely microbial biomass N
or ninhydrin-reactive N will be measured (DeLuca and Keeney 1993).

• The soil must be sieved only if homogeneous samples are required (Ocio
and Brookes 1990).

284 R.G. Joergensen, RC. Brookes

• Soil mass can range from 200 mg (Daniel and Anderson 1992) to 200 g
(Ocio and Brookes 1990).

• Soil microbial biomass is extracted by 0.5 M K 2 S0 4 . The high potas-
sium concentration flocculates the soil and prevents adsorption of NHj
released by fumigation. The relatively high salt concentration also in-
hibits decomposition of the microbial material extracted after fumiga-
tion. However, if the extracts have to be stored for a long period, they
must be frozen.

• Upon thawing of frozen K 2 S0 4 soil extracts, a white precipitate of CaS0 4
occurs in near-neutral or alkaline soils. However, this causes no analytical
problems in either method and may be safely ignored (Joergensen and
Olfs 1998).

• Soil water content can fluctuate widely, but must be higher than 30%
water-holding capacity (WHC). Microbial biomass C and biomass N of
soils at 40-50% WHC have been found to be similar to those in saturated
soils (Widmer et al. 1989; Mueller et al. 1992).

• Problems arise for fumigation and extraction in very compressed soils
that cannot be dispersed.

• Young living root cells are also affected by CHC1 3 fumigation. Conse-
quently, in soils containing large amounts of living roots, a pre-extraction
procedure must be carried out (Mueller et al. 1992).

• In substrates containing more than 20% organic matter, e.g., compost,
the ratio extractant-to-soil should be increased to 25:1 or more (Joer-
gensen et al. 1997).

14.3
Biomass C

■ Introduction

Objectives. Very low concentrations of organic C can be measured in 0.5 M
K 2 S0 4 soil extracts of fumigated and non-fumigated soil samples for the
quantitative determination of soil microbial biomass C.

14.3.1

Biomass C by Dichromate Oxidation

Principle. Organic C in the extracts is oxidized by dichromate digestion.
The amount of dichromate left is determined after redox titration by the
change in color from violet to dark green.

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 285

Theory. In the presence of a strong acid and dichromate, organic matter
is oxidized and Cr(+VI) reduced to Cr(+III). The amount of dichromate
left is back-titrated with an iron II ammonium sulfate complex solution
(Kalembasa and Jenkinson 1 973 ) and the amount of C oxidized is calculated.

■ Equipment

• Condenser

• 250-mL round-bottom flask

• Burette

■ Reagents

• K 2 Cr 2 7 solution (66.7 mM = 0.4 N)

• Digestion mixture: Mix two parts cone. H 2 S0 4 with one part cone. H 3 P0 4
(v/v)

• Indicator solution: 0.1% Aldrich (Milwaukee) ferroin solution (1,10-
phenanthroline-iron II sulfate complex)

• Titration solution: 40 mM iron II ammonium sulfate [(NH 4 )2Fe(S0 4 )2 x
6H 2 0] solution dissolved in distilled water, acidified with 20 mL cone.
H 2 S0 4 , and made up to 1 L with distilled water

■ Sample Preparation

Use soil extract prepared as described in Sect. 14.2.

■ Procedure

1. Add 2mL of K 2 Cr 2 7 solution and 15mL of the digestion mixture to
8 mL soil extract in a 250-mL round-bottom flask.

2. Reflux the mixture gently for 30min, allow to cool, and dilute with
20-25 mL water, added through the condenser as a rinse.

3. Back-titrate excess dichromate with titration solution after adding 5
drops of indicator solution to the digested soil extract.

■ Calculation

1. Calculation of extractable organic C

(HB-S)xN xExVDx (VK + SW) x 1000

C (iig/g soil) =

CBxVSx DM

(14.1)

286 R.G. Joergensen, RC. Brookes

HB consumption of titration solution by the hot (refluxed) blank (mL)

S consumption of titration solution by the sample (mL)

N normality of the K 2 Cr 2 7 solution

E 3; conversion of Cr(+VI) to Cr(+III) assuming all organic C on
average as [C(0)]

VD added volume of the K 2 Cr 2 7 solution (mL)

VS added volume of the sample (mL)

VK volume of K 2 S0 4 extractant (mL)

CB consumption of titration solution by the cold (unrefluxed) blank
(mL)

SW total amount of water in the soil sample (mL)

DM total mass of dry soil sample (g)

2. Calculation of microbial biomass C

Biomass C = E c /k^c (14.2)

Eq (organic C extracted from fumigated soils)
- (organic C extracted non-fumigated soils)

k EC 0.38 (Vance et al. 1987)

■ Notes and Points to Watch

• Be careful when working with K 2 Cr 2 7 !

• It is impossible to measure organic C with K 2 Cr 2 7 in the presence of
high chloride concentrations.

14.3.2

Biomass C by UV-Persulfate Oxidation

Principle. After removal of inorganic C by acidification, organic C in the
extracts is oxidized by UV light at 210-260 nm in the presence of K 2 S 2 8
to C0 2 , which is measured using an infrared absorption detector.

Theory. The part of the microbial biomass rendered extractable after
CHC1 3 fumigation is easily decomposable. For this reason, it is completely
oxidized to C0 2 by UV-light in the presence of K 2 S 2 8 . Infrared strongly
absorbs C0 2 .

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 287

■ Equipment

• Automatic carbon analyzer with IR-detection (e.g., Dohrman DC 80,
Tekmar-Dohrmann, Cincinnati)

■ Reagents

• Acidification buffer: 50 g sodium hexametaphosphate [((NaP0 4 ) 6 )n] dis-
solved in 900 mL distilled water, acidified to pH 2 with cone. H 3 P0 4 and
made up to 1 L

• Oxidation reagent: 20 g K 2 S 2 8 dissolved in 900 mL distilled water, acid-
ified to pH 2 with cone. H 3 P0 4 and made up to 1 L

■ Sample Preparation

Use soil extract prepared as described in Sect. 14.2.

■ Procedure

For the automated UV-persulfate oxidation method, mix 5 mL K 2 S0 4 soil
extract with 5 mL acidification buffer. Any precipitate of CaS0 4 in the soil
extracts is dissolved by this procedure. The oxidation reagent is automat-
ically fed into the UV oxidation chamber, where the oxidation to C0 2 is
activated by UV light. The resulting C0 2 is measured by IR absorption. The
IR detectors of Dimatec (Essen, Germany), for example, use a wavelength
of 4.45 yarn (80 nm width) with 3.95 yam as reference.

■ Calculation

1. Calculation of extractable organic C

[(SxDS)-(BxDB)]x(VK + SW)

C(ug/gsoil) = (14.3)

r& b DM

S C in sample extract (yig/mL)

B C in blank extract (yig/mL)

DS dilution of sample with the acidification buffer

DB dilution of blank with the acidification buffer

VK volume of K 2 S0 4 extractant (mL)

SW total amount of water in the soil sample (mL)

DM total mass of dry soil sample (g)

288 R.G. Joergensen, RC. Brookes

2. Calculation of microbial biomass C

Biomass C = E c /k EC (14.4)

Eq (organic C extracted from fumigated soils)
- (organic C extracted non-fumigated soils)

k EC 0.45 (Wu et al. 1990; Joergensen 1996a)

■ Notes and Points to Watch

• It is impossible to measure organic C with the UV-persulphate oxidation
method in the presence of high chloride concentrations because chloride
absorbs a large amount of energy in the UV range.

14.3.3

Biomass C by Oven Oxidation

Principle. After removal of inorganic C by acidification, organic C in the
extracts is oxidized at 850 °C in the presence of platinum catalyzer to C0 2 ,
which is measured using an infrared absorption detector.

Theory. Easily decomposable material as the part of the soil microbial
biomass extractable after CHC1 3 fumigation is completely oxidized at
850 °C in the presence of platinum catalyzer. Infrared strongly absorbs
C0 2 . The new auto analyzers with oven systems [Shimadzu 5050 (Shi-
madzu, Kyoto), Dimatoc 100 (Dimatec, Essen), multi N/C 2100 S (Analytik
Jena, Jena, Germany), Maihack Tocor 4, Tocor 200 (SICK, Dusseldorf)]
use small sample volumes so that they are able to measure C in extracts
containing large amounts of salts (Joergensen and Olfs 1998).

■ Equipment

Automatic carbon analyzer with oven systems (for example Shimadzu 5050,
Dimatoc 100, Dimatoc 2000, Analytik Jena multi N/C 2100 S; Analytik Jena
multi N/C 3100), see manuals for detailed description.

■ Sample Preparation

Use soil extract prepared as described in Sect. 14.2.

■ Procedure

Dilute the samples to fit the calibration line and acidify using a few drops
ofHCl.

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 289

■ Calculation

See Sect. 14.3.1.

14.4
Biomass N

■ Introduction

Objectives. Low concentrations of ninhydrin-reactive N or total N can be
measured in 0.5 M K 2 S0 4 soil extracts of fumigated and non-fumigated soil
samples without or with a digestion or oxidation step for the quantitative
determination of soil microbial biomass N.

14.4.1

Ninhydrin-Reactive Nitrogen

Principle. The amount of ninhydrin-reactive compounds released from the
microbial biomass during the CHC1 3 fumigation and extraction by 0.5 M
K 2 S0 4 is closely correlated to the initial soil microbial biomass C and
biomass N content (Joergensen and Brookes 1991).

Theory. Ninhydrin forms a purple complex with molecules containing a-
amino nitrogen and with NHj and other compounds with free a-amino
groups such as amino acids, peptides, and proteins (Moore and Stein 1948).
The presence of reduced ninhydrin (hydrindantin) is essential to obtain
quantitative color development with NHJ .

■ Equipment

• Boiling water bath

• Spectrophotometer

■ Reagents

• Lithium acetate buffer (4 M, pH 5.2): 408 g lithium acetate dihydrate (for
amino acid analysis) dissolved in water (400 mL), adjusted to pH 5.2 with
96% acetic acid, and finally made up to 1 L with water

• Citric acid buffer: citric acid monohydrate (42 g) and NaOH (16 g) dis-
solved in water (900 mL), adjusted to pH 5 with 10 M NaOH if required,
then finally made up to 1 L with water

290 R.G. Joergensen, RC. Brookes

• Ninhydrin reagent: 2 g ninhydrin and 0.3 g hydrindantin dihydrate dis-
solved (for amino acid analysis) in 75 mL dimethylsulfoxide (DMSO),
25 mL of 4 M lithium acetate buffer then added (Moore 1968)

• Ethanol/water mixture (1 + 1, v/v)

• Standard solutions: 10 mM L-leucine prepared in 0.5 M K 2 S0 4 and di-
luted within the range 0-1,000 ]iM

■ Sample Preparation

Use soil extract prepared as described in Sect. 14.2.

■ Procedure

1. Add 0.6 mL of standard solutions, K 2 S0 4 soil extracts or blank, and
1.4 mL of citric acid buffer to 20 mL test tubes (Joergensen and Brookes
1990).

2. Add 1 mL of ninhydrin reagent slowly, mix thoroughly, and close with
loose aluminum lids.

3. Heat the test tubes for 25 min in a vigorously boiling water bath; any
precipitate formed during the addition of the reagents then dissolves.

4. After heating, add 4 mL of the ethanol-to-water mixture, mix the solu-
tions thoroughly, and read the absorbance at 570 nm.

■ Calculation

1. Calculation of extracted ninhydrin-reactive N (N n i n )

xt / / -n (S-B)xNx(VK + SW)

N nin (jig/g soil) = — (14.5)

L x DM

S absorbance of the sample

B absorbance of the blank

JV atomic mass of nitrogen (14)

VK volume of K 2 S0 4 extractant (mL)

SW total amount of water in the soil sample (mL)

L millimolar absorbance coefficient of leucine

DM total mass of dry soil sample (g)

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 291

2. Calculation of microbial ninhydrin-reactive N

Bnin - (N n in extracted from the fumigated soil)

(14.6)
_ (N n in extracted from the non-fumigated soil)

3. Calculation of microbial biomass C

Biomass C = B n i n x 22

(for soils with a pH( H2 o) above 5.0; Joergensen 1996b)

Biomass C = B n i n x 35

(for soils with a pH( H2 o) of or below 5.0; Joergensen 1996b)

■ Notes and Points to Watch

• A reflux digestion is not required for ninhydrin N. This makes it very
suitable for situations with minimal laboratory facilities.

• In both biomass C and N measurements the fraction coming from the
biomass is determined following subtraction of an appropriate "control."
With biomass C this value is often half of the total, while with biomass
ninhydrin N it is commonly about 10% or less. This causes considerably
less error in its determination.

• At 100 °C the reaction with free amino groups of proteins and amino
acids is essentially complete within 1 5 min (e.g., leucine reaches the max-
imum optical density after approximately 5 min). However the reaction
of hydrindantin with NHj requires 25 min.

• The ratio between the volume of the sample and that of citric acid should
not be closer than 0.75:1.75 to avoid the formation of a precipitate after
the addition of the ninhydrin reagent.

• The most common solvent in the ninhydrin method is 2-methoxyethanol
(Amato and Ladd 1998). However, because it is an ether it tends to
form peroxides that destroy ninhydrin and hydrindantin. Dimethylsul-
foxide (DMSO) is peroxide free, has lower toxicity and a higher boil-
ing point (189 °C), and gives a more stable color development than
2-methoxyethanol.

• The ninhydrin method proposed by Amato and Ladd (1988) for 2M
KC1 extracts does not require the use of citric acid buffer. The optimum
reagent-to-sample ratio is 1:2.

292 R.G. Joergensen, RC. Brookes

14.4.2

Total Nitrogen

Principle. Total nitrogen is measured under strong acidic conditions by
Kjeldahl digestion. Ammonium can be measured by distillation (see
Chapt. 16).

Theory. Ammonium is released from amines, peptides and amino acids in
0.5 M K 2 S0 4 soil extracts of fumigated and non-fumigated soil samples. Ni-
trate is additionally reduced to ammonium under strong acidic conditions
in the presence of KCr(S0 4 ) 2 , Zn powder, and CuS0 4 as reducing agents.

■ Equipment

• Digestion block

• Steam distillation apparatus

• Burette or autotitrator

■ Reagents

• Reducing agent: 50 g of chromium(III) potassium sulfate dodecahydrate
(KCr(S0 4 ) 2 x 12H 2 0) dissolved in approx. 700 mLdeionized water, and
after adding 200 mL cone. H 2 S0 4 , cooled and diluted to 1,000 mL

• Zn powder

• CuS0 4 solution (0.1 9 M)

• Cone. H 2 S0 4

• lOMNaOH

• 2% H3BO3

• IO11MHCI

■ Sample Preparation

Use soil extract prepared as described in Sect. 14.2.

■ Procedure

1. Add 10 mL of the reducing agent and approx. 300 mg Zn powder to 30 mL
of the K 2 S0 4 soil extract and leave for at least 2 h at room temperature.

2. Add 0.6 mL of CuS0 4 solution, 8 mL of cone. H 2 S0 4 , heat gently for 2 h
until all the water has disappeared, and then heat for 3 h at the maximum
temperature.

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 293

3. Allow the digest to cool before distillation with 40 mL 10 M NaOH. The
evolved NH 3 is adsorbed in 2% H3BO3.

4. Titrate the resulting solution with 10 pM HC1 to pH 4.8.

■ Calculation

1. Calculation of extractable total N

1X (S-B)xMxNx(VK + SW)

N pg/g soil = 14.7

^ b b Ax DM

S HC1 consumed by sample extract (pL)

B HC1 consumed by blank extract (pL)

M molarity of HC1

N molecular mass of nitrogen (14)

VK volume of K 2 S0 4 extractant (mL)

SW total amount of water in the soil sample (mL)

A sample aliquot (mL)

DM total mass of dry soil sample (g)

2. Calculation of microbial biomass N

Biomass N = En/^en (14.8)

£ N (total N extracted from fumigated soils)
- (total N extracted non-fumigated soils)

fc EN 0.54 (Brookes et al. 1985; Joergensen and Mueller 1996)

■ Notes and Points to Watch

• A method is available in which the extracted total N is oxidized to NO3 ,
which is then determined colorimetrically (Cabrera and Beare 1993).

• If losses of NO3 occur during the fumigation period, they can be cor-
rected by considering the difference between the NO3 extracted initially
and the NO3 extracted at the end of the fumigation period (Brookes et al.
1985).

• If (non-fumigated) soil samples contain large amounts of NO3 or NHJ
in the soil solution, a pre-extraction step should be carried out (Wid-
mer et al. 1989; Mueller et al. 1992; Joergensen et al. 1995).

294 R.G. Joergensen, P.C. Brookes

References

Alef K, Nannipieri P (1995) Methods in Applied Soil Microbiology and Biochemistry. Aca-
demic Press, London
Amato M, Ladd JN (1988) Assay for microbial biomass based on ninhydrin-reactive nitrogen

in extracts of fumigated soils. Soil Biol Biochem 20:107-114
Barajas Aceves M, Grace C, Ansorena J, Dendooven L, Brookes PC (1999) Soil microbial

biomass and organic C in a gradient of zinc concentrations around a spoil tip mine. Soil

Biol Biochem 31:867-876
Brookes PC (1995) The use of microbial parameters in monitoring soil pollution by heavy

metals. Biol Fertil Soils 19:269-279
Brookes PC, Landman A, Pruden G, Jenkinson DS (1985) Chloroform fumigation and the

release of soil nitrogen: A rapid direct extraction method for measuring microbial

biomass nitrogen in soil. Soil Biol Biochem 17:837-842
Brookes PC, McGrath SP (1984) The effects of metal toxicity on the soil microbial biomass.

J Soil Sci 35:341-346
Cabrera ML, Beare MH (1993) Alkaline persulfate oxidation for determining total nitrogen

in microbial biomass extracts. Soil Sci Soc Am J 57:1007-1012
Castro J, Sanchez-Brunete C, Rodriguez JA, Tadeo JL (2002) Persistence of chlorpyrifos and

endosulfan in soil. Fres Environ Bull 11:578-582
Daniel O, Anderson JM (1992) Microbial biomass and activity in contrasting soil materials

after passage through the gut of the earthworm Lumbricus rubellus Hoffmeister. Soil

Biol Biochem 24:465-470
DeLuca TH, Keeney DR (1993) Ethanol-stabilized chloroform as fumigant for estimating

microbial biomass by reaction with ninhydrin. Soil Biol Biochem 25:1297-1298
Franco I, Contin M, Bragato G, De Nobili M (2004) Microbiological resilience of soils

contaminated with crude oil. Geoderma 121:17-30
Harden T, Joergensen RG, Meyer B, Wolters V (1993) Mineralization of straw and formation

of soil microbial biomass in a soil treated with simazine and dinoterb. Soil Biol Biochem

25:1273-1276
Jenkinson DS, Powlson DS (1976) The effects of biocidal treatments on metabolism in soil -

I. Fumigation with chloroform. Soil Biol Biochem 8:167-177
Joergensen RG ( 1 995 ) The fumigation-extraction method to estimate soil microbial biomass:

Extraction with 0.01 m CaCl 2 . Agribiol Res 48:319-324
Joergensen RG (1996a) The fumigation-extraction method to estimate soil microbial

biomass: Calibration of the k^c value. Soil Biol Biochem 28:25-31
Joergensen RG (1996b) Quantification of the microbial biomass by determining ninhydrin-
reactive N. Soil Biol Biochem 28:301-306
Joergensen RG, Brookes PC (1990) Ninhydrin-reactive nitrogen measurements of microbial

biomass in 0.5 M K2SO4 soil extracts. Soil Biol Biochem 22:1023-1027
Joergensen RG, Figge RM, Kupsch L ( 1 997) Microbial decomposition of fuel oil after compost

addition to soil. Z Pflanzenernahr Bodenk 160:21-24
Joergensen RG, Mueller T (1996) The fumigation-extraction method to estimate soil micro-
bial biomass: Calibration of the /cen value. Soil Biol Biochem 28:33-37
Joergensen RG, Olfs HW (1998) The variability between different analytical procedures and

laboratories for measuring soil microbial biomass C and biomass N by the fumigation

extraction method. Z Pflanzenernahr Bodenk 161:51-58
Joergensen, RG, Schmaedeke F, Windhorst K, Meyer B (1994a) Biomasse und Aktivitat

von Mikroorganismen eines mineralolkontaminierten Bodens. In: Alef K, Fiedler H,

Hutzinger O (eds) Band 6: Bodenkontamination, Bodensanierung, Bodeninformation-

ssysteme. Eco-Informa'94, Umweltbundesamt/Wien, pp 225-236

14 Quantification of Soil Microbial Biomass by Fumigation-Extraction 295

Joergensen RG, Schmaedeke F, Windhorst K, Meyer B ( 1 994b) Die Messung der mikrobiellen

Biomasse wahrend der Sanierung eines mit Dieselol kontaminierten Bodens. VDLUFA-

Schriftenr 38:557-560
Joergensen RG, Schmaedeke F, Windhorst K, Meyer B (1995) Biomass and activity of mi-
croorganisms in a fuel oil contaminated soil. Soil Biol Biochem 27:1137-1143
Kalembasa SJ, Jenkinson DS (1973) A comparative study of titrimetric and gravimetric

methods for the determination of organic carbon in soil. J Sci Food Agric 24:1085-1090
Moore S (1968) Amino acid analysis: Aqueous dimethyl sulfoxide as solvent for the ninhydrin

reaction. J Biol Chem 243:6281-6283
Moore S, Stein WH (1948) Photometric ninhydrin method for use in the chromatography

of amino acids. J Biol Chem 176:367-388
Mueller T, Joergensen RG, Meyer B (1992) Estimation of soil microbial biomass C in the

presence of living roots by fumigation-extraction. Soil Biol Biochem 24:179-181
Ocio JA, Brookes PC (1990) Soil microbial biomass measurements in sieved and unsieved

soils. Soil Biol Biochem 22:999-1000
Plante AF, Voroney RP (1998) Decomposition of land applied oily food waste and associated

changes in soil aggregate stability. J Environm Qual 27:395-402
Powlson DS, Brookes PC, Christensen BT (1987) Measurement of soil microbial biomass

provides an early indication of changes in total soil organic matter due to straw incor-
poration. Soil Biol Biochem 19:159-164
Powlson DS, Jenkinson DS (1976) The effects of biocidal treatments on metabolism in

soil. II gamma irradiation, autoclaving, air-drying and fumigation. Soil Biol Biochem

8:179-188
Vance ED, Brookes PC, Jenkinson DS (1987) An extraction method for measuring soil

microbial C. Soil Biol Biochem 19:703-708
Widmer P, Brookes PC, Parry LC (1989) Microbial biomass nitrogen measurements in soils

containing large amounts of inorganic nitrogen. Soil Biol Biochem 21:865-867
Wu J, Joergensen RG, Pommerening B, Chaussod R, Brookes PC (1990) Measurement of soil

microbial biomass C - an automated procedure. Soil Biol Biochem 22:1167-1169
Wu J, O'Donnell AG, Syers JK (1993) Microbial growth and sulphur immobilization following

the incorporation of plant residues into soil. Soil Biol Biochem 25:1567-1573

Determination of Adenylates
and Adenylate Energy Charge

Rainer Georg Joergensen, Markus Raubuch

■ Introduction

Objectives. The determination of adenosine-triphosphate (ATP) extracted
from soil was introduced a long time ago as an estimate of the soil microbial
biomass (Oades and Jenkinson 1979). After a conditioning pre-incubation,
close linear relationships exist between ATP and microbial biomass C de-
termined either by the fumigation incubation technique (Jenkinson 1988)
or by the fumigation extraction method (Chapt. 14; Contin et al. 2001; Dy-
ckmans et al. 2003). A similar close linear relationship exists also between
microbial biomass C and the sum of all three adenylates AMP, ADP, and
ATP (Dyckmans et al. 2003). The determination of adenylates is the quickest
way of estimating microbial biomass, because 24-h incubation periods or
manipulations such as substrate addition are not required as in the fumiga-
tion extraction or the substrate induced respiration methods, respectively.
The measurement of adenylates by high-performance liquid chromatog-
raphy (HPLC) has been repeatedly used to monitor the effects of heavy
metal contamination (Chander et al. 2001) and salinization (Sardinha et al.
2003), but no information is available regarding fuel oil contaminated soil.
However, enzymatic ATP has been successfully used to monitor microbial
activity during fuel oil decomposition, although some quenching of the
bioluminescence by fuel oil residues occurred (Wen et al. 2003).

An important index for the energetic state of the soil microbial commu-
nity is the adenylate energy charge (AEC), which was defined by Atkinson
and Walton (1967) as follows:

(ATP + 0.5 x ADP) /(ATP + ADP + AMP)

High AEC values (> 0. 7) have frequently been described in soils (Brookes
et al. 1987; Brookes 1995; Chander et al. 2001; Dyckmans et al. 2003).
Low AEC values have been demonstrated under drought stress conditions
(Raubuch et al. 2002), but also in Cu contaminated soils (Chander et al.
2001) and in acidic saline soils (Sardinha et al. 2003).

Rainer Georg Joergensen, Markus Raubuch: Department of Soil Biology and Plant Nu-
trition, University of Kassel, Nordbahnhofstr. la, 37213 Witzenhausen, Germany, E-mail:
joerge@wiz.uni-kassel.de

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

298 R.G. Joergensen, M. Raubuch

Principle. Soil adenylates (AMP + ADP + ATP) are extracted with dimethyl-
sulfoxide (DMSO) under strong alkaline conditions in combination with
an ethylene-diamine-tetraacetic acid (EDTA)-containing phosphate buffer.
DMSO destroys microbial cells, the phosphate buffer completely prevents
the adsorption of adenylates under the strong alkaline conditions, and
EDTA promotes the irreversible inactivation of ATP-converting enzymes.

Theory. ATP is rapidly destroyed outside living cells and can be used as
an estimate for the soil microbial biomass assuming a constant ATP-to-
microbial biomass ratio, which is fairly true in the absence of living plant
roots and after a conditioning pre-incubation (Jenkinson 1988). The ATP-
to-microbial biomass C ratio is affected by drought (Raubuch et al. 2002),
temperature (Joergensen and Raubuch 2003), and N limitation (Joergensen
and Raubuch 2002). However, the main problems in measuring ATP in soils
are (1) the enzymatic breakdown of ATP after cell death and (2) adsorp-
tion of ATP to clay minerals during extraction (Martens 2001). The alka-
line DMSO-EDTA-phosphate-buffer extractant solved nearly all method-
ological problems reported earlier (Bai et al. 1988; Martens 1992). This
is especially true in combination with HPLC analysis after derivatization
with chloroacetaldehyde to form the fluorescent l-N 6 -etheno-derivatives
(^-adenylates), which are highly selective for fluorometric determination
(Bai et al. 1989; Dyckmans and Raubuch 1997).

■ Equipment

• Multipoint magnetic stirrer

• Ultrasonic bath

• Evacuation units and filters (0.45-^m cellulose nitrate membrane filters)

• Heating water bath

• Test tube stirrer

• Glassware: 100-mL glass beaker (tall form), 20-mL test tubes

• Pipettes

• HPLC equipment: automatic injector, isocratic precision pump, column
oven, solvent delivery system, fluorescence detector and recording unit

• Analytical column (250 x 4.6 mm; 5 p.m ODS Hypersil, Thermo Electron
Corp., Waltham, MA, USA) with guard column (10 x 4.0 mm, 5 pm ODS
Hypersil)

15 Determination of Adenylates and Adenylate Energy Charge 299

■ Reagents

• DMSO

• Extraction buffer: 20 mM EDTA dissolved in 10 mM Na 3 P0 4 x 12H 2
containing 0.1 M KOH at pH 12

• Tris buffer: 2 mM EDTA dissolved in 10 mM ammonium acetate/20 mM
Tr is (hydroxy methyl) -aminome thane, adjusted to pH 7.75 with acetic
acid (store at 4°C)

• Adenylate releasing reagent: 0.05 mL benzalkonium chloride solution
(ca. 50% in water, Fluka, Fluka AG, Buchs, Switzerland, purum grade)
added to 49.95 mL Tris buffer (store at 4 °C)

• 0.1MKH 2 PO 4

• Chloroacetaldehyde

• TBAHS buffer: 50 mM ammonium acetate, 1 mM EDTA, 0.4 mm tetra-
ft-butylammonium hydrogen sulfate (TBAHS, LiChropur, Merck KGaA,
Darmstadt, Germany)

• Mobile phase for HPLC: TBAHS buffer mixed with methanol at a ratio
of89.5tol0.5(v/v)

• Calibration stock solution I (lOOiig/mL): 14.35 mg AMP-Na 2 x 6H 2 0,
11.59mgADP-K 2 x 2H 2 0, or 11.90 mgATP-Na 2 x 3H 2 0; each dissolved
in 100 mL extraction buffer (store at 4 °C)

• Calibration stock solution II (1 p.g/mL): 1/100 dilution of stock solution I
(store at 4 °C)

• Working standard solutions: a set of four standards each containing 2, 4,
6, 8 ng of AMP, ADP, ATP, respectively, prepared by mixing 100-400 \xL
stock solution II with 0.2 mL chloroacetaldehyde and adding 0.01 M
Na 2 HP0 4 x 2H 2 to give a final volume of 10 mL, heated for 3min at
85 °C, and cooled in an ice bath (store at 4 °C for maximum 7 days)

■ Sample Preparation

Use moist sample equivalent to 1-5 g oven-dry soil, sieved (< 2 mm). The
experimental design reflects the fact that adenylate content responds to
actual conditions, is influenced by mechanical disturbance, water content,
and temperature.

300 R.G. Joergensen, M. Raubuch

■ Procedure

1. Weigh moist soil equivalent to 1-5 g oven-dry soil into a 100-mL glass
beaker (tall form).

2. Add 4 mL DMSO and stir for 2 min on a magnetic stirrer using a mag-
netic stirring bar.

3. Add 16 mL extraction buffer and stir again for 2 min.

4. Sonify for 2 min in an ultrasonic bath.

5. Mix an aliquot of 0.5 mL of soil suspension with 0.5 mL of adenylate
releasing reagent in a 20 mL test tube, mix using a test tube stirrer, and
sonify for another 5 s.

6. Pass the suspension through a membrane filter (0.45 p.m) and wash the
soil residue twice with 1 mL 0.1 M KH 2 P0 4 .

7. Add 0.2 mL chloroacetaldehyde and make up to a final volume of 5 mL
by addition of 0.1 M KH 2 P0 4 .

8. Incubate in a water bath for 30 min at 85 °C to yield the fluorescent
l-N 6 -etheno-derivatives and cool afterward in an ice bath.

9. Store at 4 °C for a maximum 7 days before HPLC measurements.

10. Adjust the column oven to 27 °C.

11. Run HPLC with the mobile phase at 2 mL/min for 3 h for equilibration
of the column.

12. Use a sample loop of 200 p.L.

13. Fluorometric emission is measured at 410 nm with 280 nm as excitation
wavelength.

14. Clean the HPLC after measurement for 30 min at 1 mL/min with a meth-
anol/water (50:50 v/v) solution.

15. Treat calibration standards like soil extractants to prepare calibration
curves.

16. Standard solutions correspond to concentrations 2ng, 4ng, 6ng, 8ng
of AMP, ADP and ATP in 200 ]iL y respectively.

17. There is a linear relationship in adenylate content and signal response
up to 8 ng of each adenylate. The adenylates are detected on the chro-
matogram in the order AMP, ADP, and ATP.

15 Determination of Adenylates and Adenylate Energy Charge 301

■ Calculation

1. Identify AMP, ADP and ATP by retention time according to the retention
time of the standards.

2. Calculate nanograms from areas and linear equation of standards.

3. Take dilution into account (analogous for ADP and AMP).

H x (E + SW) x I

ATP (ng/gsoil) =

A xDM

H ATP in 200 p.L injection volume (ng)

E extractant (4 mL DMSO + 16 mL extraction buffer; mL)

J 25; conversion factor of the injection volume (200 p.L from 5 mL)

SW total amount of water in the soil sample (mL)

A aliquot (0.5 mL)

DM total mass of dry soil sample (g)

4. Molecular masses for conversion into nmol:
AMP = 347.2 g, ADP = 427.2 g, ATP = 507.2 g

5. Total adenylate content (nmol/g soil) = AMP + ADP + ATP

6. Adenylate Energy Charge (AEC)

= (ATP + 0.5 ADP)/(AMP + ADP + ATP)

Notes and Points to Watch

The mobile phase must be degassed in advance. Oxygen disturbs the
measurement, especially of ATP.

The retention time must be checked before the first measurement with
a standard mixture of AMP, ADP, and ATP standards, but do not use
a standard mixture of AMP, ADP and ATP for calibration. ATP contains
impurities of AMP and ADP, ADP contains impurities of AMP.

The column temperature should be constant at 27 °C. The separation of
£-AMP, £-ADP and £-ATP from extracted impurities is improved at 27 °C.
Changing temperatures causes shifts in the retention times.

302 R.G. Joergensen, M. Raubuch

References

Atkinson DE, Walton GM (1967) Adenosine triphosphate conservation in metabolic regu-
lation. Rat liver cleavage enzyme. J Biol Chem 242:3239-3241

Bai QY, Zelles L, Scheunert I, Korte F (1988) A simple procedure for the determination of
adenosine triphosphate in soils. Chemosphere 17:2461-2470

Bai QY, Zelles L, Scheunert I, Korte F (1989) Determination of adenine nucleotides in soil by
ion-paired reverse-phase high-performance liquid chromatography. J Microbiol Meth
9:345-351

Brookes PC (1995) Estimation of the adenylate energy charge in soils. In: Alef K, Nannipieri P
(eds) Methods in Applied Soil Microbiology and Biochemistry. Academic Press, London,
pp 204-213

Brookes PC, Newcombe AD, Jenkinson DS (1987) Adenylate energy charge measurements
in soil. Soil Biol Biochem 19:211-217

Chander KC, Dyckmans J, Joergensen RG, Meyer B, Raubuch M (2001) Different sources of
heavy metals and their long-term effects on soil microbial properties. Biol Fertil Soils
34:241-247

Contin M, Todd A, Brookes PC (2001) The ATP concentration in the soil microbial biomass.
Soil Biol Biochem 33:701-704

Dyckmans J, Chander K, Joergensen RG, Priess J, Raubuch M, Sehy U (2003) Adenylates as an
estimate of microbial biomass C in different soil groups. Soil Biol Biochem 35:1485-1491

Dyckmans J, Raubuch M (1997) A modification of a method to determine adenosine nu-
cleotides in forest organic layers and mineral soils by ion-paired reversed-phase high-
performance liquid chromatography. J Microbiol Meth 30:13-20

Jenkinson DS (1988) The determination of microbial biomass carbon and nitrogen in soil.
In: Wilson JR (ed) Advances in nitrogen cycling in agricultural ecosystems. CABI,
Wallingford, pp 368-386

Joergensen RG, Raubuch M (2002) Adenylate energy charge of a glucose-treated soil without
adding a nitrogen source. Soil Biol Biochem 34:1317-1324

Joergensen RG, Raubuch M (2003) Adenylate in the soil microbial biomass at different
temperatures. Soil Biol Biochem 35:1063-1069

Martens R (1992) A comparison of soil adenine nucleotide measurements by HPLC and
enzymatic analysis. Soil Biol Biochem 24:639-645

Martens R (200 1 ) Estimation of ATP in soil: extraction methods and calculation of extraction
efficiency. Soil Biol Biochem 33:973-982

Oades JM, Jenkinson DS (1979) Adenosine triphosphate content of the soil microbial
biomass. Soil Biol Biochem 11:193-199

Raubuch M, Dyckmans J, Joergensen RG, Kreutzfeldt M (2002) Relation between respi-
ration, ATP content and adenylate energy charge (AEC) after incubation at different
temperatures and after drying and rewetting. J Plant Nutr Soil Sci 165:435-440

Sardinha M, Miiller T, Schmeisky H, Joergensen RG (2003) Microbial performance in a tem-
perate floodplain soil along a salinity gradient. Appl Soil Ecol 23:237-244

Wen G, Voroney RP, McGonigle TP, Inanaga S (2003) Can ATP be measured in soils treated
with industrial oily waste? J Plant Nutr Soil Sci 166:724-730

Determination

of Aerobic N-Mineralization

Rainer Georg Joergensen

■ Introduction

Objectives. N mineralization is the transformation of organic N into inor-
ganic N components (Beck 1983). It is thus an important biological process
of the nitrogen cycle in ecosystems, reflecting the ability of a soil to provide
available nitrogen to plants (Alef 1995). In terrestrial ecosystems, especially
in arable crop production, N is often the most limiting nutrient for plant
growth. The nitrogen availability to soil microorganisms often limits the
decomposition of fuel oil in contaminated soils (Joergensen et al. 1995). N
mineralization mainly depends on temperature, moisture, aeration, type
of organic N, and pH. NHJ is subject to fixation by clays. NO3 can be lost
through denitrification and leaching (Alef 1995).

Principle. A soil is incubated aerobically after removal of plant debris for
two periods. The soil is extracted with 2 M KC1 before and after each of the
two incubation periods. In the soil extracts, NHj, NO3, and, if necessary,
NO2 are measured.

Theory. N mineralization is the catabolic use of N- containing organic com-
ponents, e.g., amino acids, amino sugars, amines, and nucleic acids derived
from plant and animal debris as well as from soil organic matter to meet
the energy demand of the soil microbial biomass. The N mineralization
process can be divided into two steps (Alef 1995): (1) the first step is am-
monification, which is the breakdown of organic NH 2 groups to NHJ.
Except for the hydrolysis of urea by extracellular urease, ammonification
is carried out by proteases bound to cell membranes of all heterotrophic
microorganisms in soil, i.e. more than 95% of the soil microbial com-
munity. (2) The second step is nitrification, which is carried out by het-
erotrophic fungi in acidic soils or obligatory aerobic chemoautotrophic
bacteria [e.g., Nitrosomonas: NHJ —> NO2 (GO = -273.9 kj/mol) andM-
trobacter NO2 —> NO3 (GO = -76.7 kj/mol) under neutral and slightly
alkaline soil conditions. Nitrification is inhibited by fuel oil contamination
in contrast to ammonification (Joergensen et al. 1995). In compacted soils,

Rainer Georg Joergensen: Department of Soil Biology and Plant Nutrition, University of Kas-
sel, Nordbahnhofstr. la, 37213 Witzenhausen, Germany, E-mail: joerge@wiz.uni-kassel.de

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

304 R.G. Joergensen

N mineralization is more strongly affected than C0 2 production (Nieder
etal. 1993;Ahletal. 1998).

■ Equipment

• 100 mL low-density, wide-neck polyethylene (PE) bottles

• Funnels

• 50-mL Erlenmeyer flasks

• Horizontal or overhead shaker

• Folded filter papers (e.g., Whatman 42 or Schleicher & Schuell 595 1/2)

• Room or incubator at 25° C

• Steam distillation apparatus

• Burette or autotitrator

■ Reagents

• 2 M KCl solution

• 2%H 3 B0 3 (extra pure)

• 10}xMHCl

• MgO (extra pure)

• Devarda alloy

■ Sample Preparation

Use field-moist, sieved (between < 2 and < 5 mm) soil at approx. 40-50%
water holding capacity.

■ Procedure

1. Weigh 15 g field moist soil into nine PE bottles.

2. Add 5 mL of water slowly.

3. Incubate at 25 °C in the dark.

4. Remove three replicates after 0, 14, and 28 days.

5. Extract with 60 mL 2 M KCl (extractant-to-soil ratio of 4: 1 ) for 30 min by
oscillating shaking at 200 rpm (or 45 min overhead shaking at 40 rpm).

16 Determination of Aerobic N-Mineralization 305

6. Filter through a folded filter paper.

7. Pipette a 30 mL aliquot into the sample flask of the distillation appara-
tus.

8. Add approx. 200 mg MgO rapidly to volatize NH \ as NH 3 under alkaline
conditions.

9. Stop the first distillation when the distillate reaches the 30 mL mark
on the receiver flask (a 50 mL Erlenmeyer flask containing 5 mL 2%
H3BO3).

10. Add approx. 200 mg Devarda alloy rapidly to reduce NO3 and NO2 to
NHj, which is volatilized under the alkaline conditions of the distilla-
tion flask.

11. Stop the second distillation when the distillate reaches the 30 mL mark
on the receiver flask (a 50-mL Erlenmeyer flask containing 5 mL 2%
H3BO3).

12. Titrate NH+ in each of the two distillates with 10 jim HC1 to pH 4.8.

■ Calculation

1. Calculation of extractable NH+ -N and NO3 -N

xt/ / -n (S-B)xMxNx(VK + SW)

N(ug/gsoil) = (16.1)

^ && Ax DM

S HC1 consumed by sample extract (pL)

B HC1 consumed by blank extract (pL)

M molarity of HC1

N molecular mass of nitrogen (14)

VK volume of K 2 S0 4 extractant (mL)

SW total amount of water in the soil sample (mL)

A sample aliquot (mL)

DM total mass of dry soil sample (g)

2. Calculation of net N mineralized

N [|ig N/(g soil x day)]

_ (NH+-N + NO" -N) t , +1 - (NH+-N + NO" -N) t , (16.2)

306 R.G. Joergensen

td sampling day before the last sampling day (day or day 14)
trf+i last sampling day (day 14 or day 28)
n incubation period (days)

■ Notes and Points to Watch

• If the N mineralization rate of the first incubation period (0-14 days)
does not differ significantly from that of the second incubation period
(14-28 days), the average value of both periods should be used (Beck
1983; Kandeler 1993a). If the N mineralization rate of the first incubation
period is significantly lower than that of the second incubation period,
e.g., due to N immobilization during the decomposition of plant residues,
only the value of the second incubation period should be used. If the N
mineralization rate of the first incubation period is significantly higher
than that of the second incubation period, e.g., due to the increasing
recalcitrance of decomposable soil organic matter, only the value of the
first incubation period should be used.

• The steam distillation method is especially suitable for colored extracts
(Keeney and Nelson 1982; Forster 1995).

• If a soil accumulates NO2 in the soil solution, a colorimetric method
must be used to determine it (Keeney and Nelson 1982; Forster 1995).

• Colorimetric methods are also available for the manual determination
of extractable NO3 (e.g., Kandeler 1993a; Forster 1995), and for auto-
mated segmented flow or flow injection, analyses are also available (e.g.,
Kutscha-Lissberg and Prillinger 1982).

• It is possible to estimate the NO3 content in soil extracts by the decrease
in UV absorbance after reduction of NO3 (Kandeler 1993b).

• Colorimetric methods are also available for the manual determination of
extractable NHJ (e.g., Keeney and Nelson 1982; Kandeler 1993; Forster
1995), and for automated segmented flow or flow injection, analyses are
also available.

• Contamination of chemicals, especially KC1, but also of filter paper,
funnel, extraction bottles, and glassware should be avoided and regularly
checked.

References

Alef K (1995) Nitrogen mineralization in soils. In: Alef K, Nannipieri P (eds) Methods in
Applied Soil Microbiology and Biochemistry. Academic Press, London, pp 234-245

16 Determination of Aerobic N-Mineralization 307

Ahl C, Joergensen RG, Kandeler E, Meyer B, Woehler V (1998) Microbial biomass and
activity in silt and sand loams after long-term shallow tillage in central Germany. Soil
Till Res 49:93-104

Beck T (1983) Die N-Mineralisation von Boden im Brutversuch. Z Pflanzenernahr Bodenk
146:243-252

Forster JC (1995) Soil nitrogen. In: Alef K, Nannipieri P (eds) Methods in Applied Soil
Microbiology and Biochemistry. Academic Press, London, pp 79-87

Joergensen RG, Schmaedeke F, Windhorst K, Meyer B (1995) Biomass and activity of mi-
croorganisms in a fuel oil contaminated soil. Soil Biol Biochem 27:1137-1143

Kandeler E (1993a) Bestimmung der N-Mineralisation im aeroben Brutversuch. In Schin-
ner F, Ohlinger R, Kandeler E, Margesin R (eds) Bodenbiologische Arbeitsmethoden,
2nd ed. Springer, Berlin, pp 158-159

Kandeler E (1993b) Bestimmung von Nitrat. In: Schinner F, Ohlinger R, Kandeler E, Mar-
gesin R (eds) Bodenbiologische Arbeitsmethoden, 2nd ed. Springer, Berlin, pp 369-371

Keeney DR, Nelson DW (1982) Nitrogen - inorganic forms. In: Page AL, Miller, RH, Kee-
ner DR (eds) Methods of Soil Analysis, Part 2. Am Soc Agron, Soil Sci Soc Am, Madison,
pp 643-698

Kutscha-Lissberg P, Prillinger F (1982) Rapid determination of EUF-extractable nitrogen
and boron. Plant Soil 64:63-66

Nieder R, Scheithauer U, Richter J (1993) Dynamics of nitrogen after deeper tillage in arable
loess soils of West Germany. Biol Fertil Soils 16:45-51

Determination of Enzyme Activities
in Contaminated Soil

Rosa Margesin

17.1

General Introduction

Soil biological activities are sensitive to environmental stress; each change
in environmental conditions may result in a shift in the species composition
of the soil microflora and modification of their metabolic rate. Soil enzyme
activities are attractive as indicators for monitoring various impacts on soil
because of their central role in the soil environment. Soil enzymes are the
catalysts of important metabolic processes including the decomposition of
organic inputs and the detoxification of xenobiotics (Schinner et al. 1996;
Dick 1997).

Soil enzyme activities have been used as a biological indicator of pollu-
tion with heavy metals, pesticides, and hydrocarbons (Schinner et al. 1993;
Sparling 1997; van Beelen and Doelman 1997; Margesin et al. 2000a, 2000b).
A number of studies have demonstrated that soil enzymes hold potential for
assessing the impact of hydrocarbons and of fertilization on soil microor-
ganisms and are a useful tool to monitor the early stages of remediation of
contaminated soil (Margesin et al. 2000a, 2000b). The usefulness of various
enzyme parameters depends on the composition and concentration of the
hydrocarbons, as well as on other factors such as the age of contamination
and physico-chemical soil characteristics. While some enzymes activities
are appropriate to monitor the most active phase of biodegradation, oth-
ers are also indicative of low hydrocarbon concentrations (Margesin et al.
2000a).

A broad spectrum of soil enzyme activities should be used to evaluate
the effect of contamination on the different nutrient cycles. In this Chapter,
a small selection of methods for the determination of enzyme activities
in contaminated soil is described. Detailed descriptions of supplementary
methods are given in Schinner et al. (1996). Of course, additional informa-
tion on the soil biological status should be obtained from complementary
methods, such as soil microbial counts (Chap. 13), soil biomass (Chap. 14),
molecular biology (Chap. 10), and fatty acid profiles (Chap. 12).

Rosa Margesin: Institute of Microbiology, Leopold Franzens University, Technikerstrasse
25, 6020 Innsbruck, Austria, E-mail: rosa.margesin@uibk.ac.at

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

310 R. Margesin

17.2

Lipase-Esterase Activity

■ Introduction

Objectives. Soil lipase activity is a valuable tool to monitor the biodegrada-
tion of petroleum hydrocarbons, such as diesel oil, in freshly contaminated
soil (Margesin et al. 1999, 2002a, b). The residual hydrocarbon content
correlates negatively with soil lipase activity in unfertilized as well as in
fertilized soil (Margesin and Schinner 2001). Generally, a correlation be-
tween soil lipase activity and other biological parameters can be found.
Soil lipase activity increases with increasing initial oil loading rates, which
demonstrates the induction of this enzyme activity by the contamina-
tion. This induction is attributed to the appearance of products released
from hydrocarbon biodegradation, which are the substrate for hydrolases
including esterases-lipases. A strong relationship between the ability of mi-
croorganisms to degrade diesel oil and their lipolytic activity was described
by Mills et al. (1978) and Kato et al. (2001). No increase of lipase activity
was observed during the biodegradation of polycyclic aromatic hydrocar-
bons (PAHs) in soil (Margesin et al. 2000a). Assaying lipase activity is also
useful to monitor the biodegradation of carboxyl esters such as lipids and
biodegradable polyesters in soil (Sakai et al. 2002). The described method
can also be applied to determine lipase activity in non-contaminated soil
and could be useful for the screening of lipase-producing soil microorgan-
isms.

Principle. Using p-nitrophenyl butyrate (pNPB) as substrate, soil sam-
ples are incubated at 30 °C and pH 7.25 for lOmin. After cooling on ice
and centrifugation, the released p-nitrophenol (pNP) is determined spec-
trophotometrically at 400 nm. To allow for the adsorption of pNP onto soil,
a calibration curve is prepared in the presence of soil (Margesin et al. 2002).

Theory. A significant proportion of lipids, such as pesticide emulsions,
oils, and lipid conjugates, enter soil in the form of triacylglycerols, the
primary storage fat in plant and animal tissue. The degradation of lipids is
initiated by lipases (glycerol ester hydrolases) acting on the carboxylester
bonds present in acylglycerols to liberate fatty acids and glycerol. Lipases
are produced by a large variety of microorganisms, plants, and animals.

The standard assays to determine the hydrolytic activity of lipase in
soil are based on titration or fluorimetry. Methods that determine the
fatty acids produced from tributyrin (Pokorna 1964; Hankin et al. 1982)
or Tween 20 (Sakai et al. 2002) titrimetrically are easy to use, however,
the disadvantages are the long incubation time (between 18 h and 3 days)
and the possible adsorption of the released fatty acids onto soil colloids.

1 7 Determination of Enzyme Activities in Contaminated Soil 311

Fluorimetric methods (Pancholy and Lynd 1972; Cooper and Morgan 1981)
are more sensitive and specific, but the substrate is relatively expensive.
Colorimetric methods are quick and simple. Chromogenic substrates, such
as p-nitrophenyl esters, are commonly used to assay microbial esterase and
lipase activity (Shirai and Jackson 1982; Plou et al. 1998; Ishimoto et al.
2001; Wei et al. 2003). In soil enzymology, p-nitrophenyl derivatives are
widely used as substrates for measuring phosphatases, arylsulfatase, and
6-glucosidase activity. The described method is based on the use of pNPB as
substrate for the rapid, precise, and simple measurement of lipase activity
in soil.

■ Equipment

• Centrifuge and centrifuge tubes (2,000 g, 2-4 °C)

• Water bath (30 °C)

• Spectrophotometer

■ Reagents

• Phosphate buffer: 100 mM NaH 2 P0 4 -NaOH, pH 7.25 (store at 4 °C)

• Substrate: 100 mM p-nitrophenyl butyrate (pNPB) diluted in 2-propanol
(store aliquots at -20 °C)

• Calibration standards (p-nitrophenol, pNP)

- Stock solution: 1 mg pNP/mL buffer (store at 4 °C)

- Working standard solution: 100 yigpNP/mL buffer (prepare daily
fresh)

- Standards: 0, 25, 50, 75, 100, and 125 jig pNP/5 mL buffer (adjust vol-
umes of 0-1.25 mL of the working standard solution to 5mL with
buffer)

■ Sample Preparation

Use field-moist, sieved (< 5 mm) soil.

■ Procedure

1. Weigh 0.1 g of soil into centrifuge tubes, prepare 3-4 replicates (samples).

2. Add 5 mL of buffer, and prewarm at 30 °C in a water bath for 10 min.

3. Prepare a control (3-4 replicates) without soil.

4. Add 50 p.L of substrate solution to each tube.

312 R. Margesin

5. Mix the contents and incubate the tubes in a water bath at 30 °C for
exactly lOmin.

6. Stop the reaction by cooling the tubes for 10 min on ice.

7. Centrifuge the tubes at 2,000 g and 2-4 °C for 5 min.

8. Pipette the supernatants in test tubes that are held on ice.

9. Immediately afterward, measure the absorbance of the released pNP in
samples and controls at 400 nm against the reagent blank. Dilute the
solution with buffer when absorbance values are too high.

To prepare a calibration curve, treat calibration standards like the soil
samples:

1. Weigh 0.1 g of soil into each of six centrifuge tubes.

2. Add 5 mL of standard solution containing (= reagent blank), 25, 50, 75,
100,andl25}igpNP.

3. Mix and incubate at 30 °C for exactly 10 min.

4. Proceed as described for the soil samples (6-8), and measure the ab-
sorbance of the calibration standards at 400 nm against the reagent blank.

■ Calculation

1. Calculate the pNP concentration from the calibration curve.

2. Subtract the control reading (hydrolysis in absence of soil) from the
sample reading (hydrolysis in presence of soil) and express soil lipase
activity as "|^g of released pNP per gram soil dry mass over 10 min" using
the following formula:

S-C
|ig pNP/(g dry soil x 10 min) =

wm x dm

S pNP concentration of the soil sample (p.g)
C pNP concentration of the control (pg)
wm Soil wet mass (0.1 g)
dm Soil dry mass (g)

Notes and Points to Watch

To measure the chemical pNP release from the substrate, it is necessary
to prepare a control without soil.

17 Determination of Enzyme Activities in Contaminated Soil 313

• To allow for the adsorption of pNP onto soil, the calibration curve is
prepared in the presence of soil.

• It is important to use a neutral pH for this assay since the ester bond in
pNPB is very labile and is fully hydrolyzed at alkaline pH. The degree
of dissociation of pNP (colorless) into p-nitrophenoxide (yellow) is 0%
below pH 5.0, 50% at pH 7.0 and 100% at pH 9.0. Enzyme assays based
on p-nitrophenyl derivatives are usually carried out at a pH of 7.25 to 7.3
(Ishimoto et al. 2001; Wei et al. 2003).

• A reaction temperature of 30 °C is chosen to avoid a high rate of non-
enzymatic substrate hydrolysis, which occurs at higher temperatures.

• Drying and freeze-thawing of soil may affect soil lipase activity, however,
this effect is soil-specific.

• The presence of heavy metals in soil inhibits lipase activity. The metal
sensitivity of soil lipases depends on the soil properties.

• Lipase activity determined with the described assay has been found to
correlate significantly with titrimetric and fluorimetric assays.

17.3

Fluorescein Diacetate Hydrolytic Activity

■ Introduction

Objectives. The rate of fluorescein diacetate (FDA) hydrolysis in soil has
been considered a suitable index of overall enzyme activity (Schmirer and
Rosswall 1982). It is a suitable indicator to indicate the onset of biodegra-
dation of diesel oil and of monoaromatic compounds, such as BTEX (Mar-
gesin et al. 2000a, 2003). The time course of FDA hydrolytic activity during
the bioremediation of soil contaminated with diesel oil is comparable to
that of lipase activity (Sect. 17.2). The biodegradation of PAHs (naphtha-
lene, phenanthrene) in soil also results in an increase of FDA hydrolytic
activity, which, however, is followed by a marked activity decrease (Mar-
gesin et al. 2000a). A short-term, reversible inhibition of FDA hydrolysis
has been noted in BTEX-contaminated soil (Margesin et al. 2003) and
in jet-fuel-contaminated soil (Song and Bartha 1990), being substantially
higher in unfertilized than in fertilized soil. The inhibition terminated after
a significant part of the contamination had disappeared, and a stimulation
of the activity was observed after most of the fuel had been mineralized.
Generally, a correlation between FDA hydrolytic activity and other soil
biological parameters can be found.

314 R. Margesin

Principle. Using FDA as substrate, soil samples are incubated at 25 °C and
pH 7.6 for 2 h. The released fluorescein is extracted with acetone, and
quantified photometrically at 490 nm.

Theory. FDA is hydrolyzed by a number of different enzymes, such as pro-
teases, lipases, and esterases. The ability to hydrolyze FDA is widespread
among soil organisms and has been detected among heterotrophic bacteria,
fungi, algae, and protozoa. The product of this enzyme conversion is flu-
orescein, which can be visualized within cells by fluorescence microscopy.
Fluorescein can also be quantified using fluorometry and spectrophotom-
etry. Schmirer and Rosswall (1982) developed a simple, rapid, and sensitive
spectrophometric method for the measurement of total microbial activity
in soil straw litter.

■ Equipment

• Water bath (25 °C)

• Spectrophotometer

■ Reagents

• Phosphate buffer: 60 mM NaH 2 P0 4 x H 2 0/Na 2 HP0 4 x 2H 2 0, pH 7.6
(store at 4 °C)

• Substrate solution: Fluorescein diacetate (FDA; 2 mg/mL) dissolved in
acetone (store aliquots at -20 °C)

• Acetone (technical grade)

• Calibration standards (fluorescein)

- Stock solution: 1 mg fluorescein/mL acetone (store at 4°C)

- Working standard solution: 100 p.g fluorescein/mL buffer (prepare
daily fresh)

- Standards: 0, 20, 50, 70, 100, and 150p.g fluoresce in/ lOmL buffer
(adjust volumes of 0-1.5 mL of the working standard solution to
10 mL with buffer)

■ Sample Preparation

Use field-moist, sieved (< 5 mm) soil.

■ Procedure

1. Weigh 1 g of soil into 100 mL Erlenmeyer flasks, prepare 3 replicates
(samples).

17 Determination of Enzyme Activities in Contaminated Soil 315

2. Add 10 mL of buffer.

3. Prepare a control (3 replicates) without soil.

4. Add 100 pi of substrate solution to each flask.

5. Mix the contents and incubate the stoppered flasks at 25 °C for 2 h.

6. Stop the reaction by adding 10 mL of acetone.

7. Filter the contents of the flasks.

8. Immediately afterwards, measure the absorbance of the released fluo-
rescein in samples and controls at 490 nm against the reagent blank.

To prepare a calibration curve, treat calibration standards like the soil
samples:

1. Weigh 1 g of soil into each of six flasks.

2. Add 10 mL of standard solution containing (= reagent blank), 20, 50,
70, 100, 120, and 150 pg fluorescein.

3. Proceed as described for the soil samples (5-7), and measure the ab-
sorbance of the calibration standards at 490 nm against the reagent blank.

■ Calculation

3. Calculate the fluorescein concentration from the calibration curve.

4. Subtract the control reading (hydrolysis in absence of soil) from the
sample reading (hydrolysis in presence of soil) and express activity as
"|^g of released fluorescein per gram soil dry mass over 2 h" using the
following formula:

S-C
|ig Fluorescein/(g dry soil x 2 h) =

wm x dm

S pNP concentration of the soil sample (pg)
C pNP concentration of the control (pg)
wm Soil wet mass (1 g)
dm Soil dry mass (g)

Notes and Points to Watch

To measure the chemical fluorescein release from the substrate, it is
necessary to prepare a control without soil.

316 R. Margesin

• To allow for the adsorption of fluorescein onto soil, the calibration curve
is prepared in the presence of soil. This is especially important in case of
soil containing high amounts of organic matter or clay.

• Depending on the soil to be tested it might be necessary to optimize soil
mass, substrate concentration, and incubation time.

• It is important to use a neutral pH for this enzyme assay since chemical
hydrolysis of FDA occurs under acidic and alkaline pH conditions.

• Chemical hydrolysis of FDA occurs also at higher temperatures, therefore
a reaction temperature of 25 °C is used for this enzyme assay. Store
aliquots of the substrate solution at -20 °C to avoid chemical substrate
hydrolysis, which occurs when the solution is stored for longer periods
at 4 °C or room temperature.

• Acetone is added not only to stop the enzyme reaction, but also to solu-
bilize cell membranes in order to facilitate the extraction of fluorescein
from microbial cells.

• High amounts of heavy metals in soil may interfere with the method.

17.4

Dehydrogenase Activity

■ Introduction

Objectives. Soil dehydrogenase activity is a useful method to monitor the
bioremediation of soil contaminated with petroleum hydrocarbons, such
as diesel oil. The method has been applied to soil containing fresh (Mar-
gesin and Schinner 1997; Margesin et al. 2000a) and aged contamination
(Margesin and Schinner 1999, 2001). A statistically significant positive cor-
relation between this activity and the residual hydrocarbon content has
been repeatedly observed. The substantial increase in soil dehydrogenase
activity after hydrocarbon contamination reflects the adaptation and ex-
ponential growth of hydrocarbon degraders due to the availability of new
carbon sources introduced by the contamination. Soil dehydrogenase activ-
ity declines with decreasing hydrocarbon content due to the loss of available
compounds as a consequence of biodegradation (Margesin et al. 2000a, b).
Measuring dehydrogenase activity is also a useful tool to monitor envi-
ronmental contamination by anionic surfactants (Margesin and Schinner
1998), since this activity is inhibited in presence of high concentrations
of anionic surfactants, such as sodium dodecyl sulfate. However, dehydro-
genase activity is not a suitable parameter to monitor biodegradation of
PAHs in soil (Margesin et al. 2000a).

17 Determination of Enzyme Activities in Contaminated Soil 317

Principle. Soil samples are mixed with [2(p-iodophenyl)-3-(p-nitrophe-
nyl)-5-phenyl tetrazolium chloride] solution (INT), and incubated for 2h
at 40 °C. The reduced iodonitro tetrazolium formazan (INTF) is extracted
with dimethylformamide and ethanol, and quantified photometrically at
464 nm (von Mersi and Schinner 1991; Schinner et al. 1996).

Theory. Dehydrogenases belong to oxidoreductases and catalyze the re-
moval from a substrate of two hydrogens that are taken up by a hydrogen
acceptor or coenzyme. Nicotinamide adenine dinucleotide (NAD) is the
coenzyme used by the majority of dehydrogenases, others use nicotinamide
adenine dinucleotide phosphate (NADP) or flavin adenine dinucleotide
(FAD). Since dehydrogenases are important components of the enzyme
system of all microorganisms, soil dehydrogenase activity reflects a broad
range of microbial oxidative activities, and can be taken as a measure for the
intensity of microbial metabolism in soil (Schinner et al. 1996). Because of
increased sensitivity and reproducibility, the substrate INT has been used
by a number of authors (Trevors 1984; Griffiths 1989; von Mersi and Schin-
ner 1 99 1 ) to determine soil dehydrogenase activity. An alternative substrate
is 2,3,5-triphenyltetrazolium chloride (for details, see Schinner et al. 1996).

■ Equipment

• Water bath or incubator (40 °C)

• Spectrophotometer

■ Reagents

• Buffer: 1 M Tris-HCl, pH 7.0 (store at 4 °C).

• Substrate: dissolve 500 mg of 2(p-iodophenyl)-3-(p-nitrophenyl)-5-phe-
nyl tetrazolium chloride (INT; Serva, Heidelberg) in 2 mL of N,N-dime-
thylformamide by shaking vigorously and using an ultrasonic water
bath. Make up to volume with distilled water in a 100 mL volumetric
flask (prepare daily fresh, and store it in the dark until use).

• Extraction solution: mix 1 part of N,N-dimethylformamide with 1 part
of 96% ethanol.

• Calibration standards (iodonitrotetrazolium formazan (INTF)).

- Stock solution: 100 yig INTF/mL extraction solution (store at 4 °C)

- Standards: 0, 100, 200, 300, and 500 jig INTF/13.5 mL extraction solu-
tion (adjust volumes of 0-5 mL of the stock solution to 13.5 mL with
extraction solution)

318 R. Margesin

■ Sample Preparation

Use field-moist, sieved (< 5 mm) soil. Additionally, prepare autoclaved soil.

■ Procedure

1. Weigh 1 g of soil into 100 mL Erlenmeyer flasks, prepare 3 replicates
(samples).

2. Weigh 1 g of autoclaved soil into 100 mL Erlenmeyer flasks, prepare 2
replicates (controls).

3. Add 1.5 mL of buffer and 2 mL of substrate solution to both samples and
controls.

4. Mix the contents and incubate the stoppered flasks at 40 °C for 2 h.

5. Add lOmL of extraction solution to each flask. For extraction of the
released INTF keep the flasks for 1 h at room temperature in the dark,
and shake vigorously every 20 min.

6. Filter the contents of the flasks.

7. Immediately afterwards, measure the INTF concentration in samples,
controls and calibration standards at 464 nm against the reagent blank.
Dilute solutions with extraction solution when absorbance values are
too high.

■ Calculation

1. Calculate the INTF concentration from the calibration curve.

2. Express soil dehydrogenase activity as "|ig of released INTF per gram
soil dry mass over 2 h" using the following formula:

S-C
|Lig INTF/(g dry soil x 2 h) =

wm x dm

S INTF concentration of the soil sample (p.g)
C INTF concentration of the control (p.g)
wm Soil wet mass (1 g)
dm Soil dry mass (g)

Notes and Points to Watch

INT is very sensitive to light. Both incubation and filtration have to be
carried out in the dark.

17 Determination of Enzyme Activities in Contaminated Soil 319

• The control contains autoclaved soil to estimate the chemical substrate
reduction due to reactive soil components.

• Depending on the soil to be tested, it might be necessary to optimize soil
mass, substrate concentration, and incubation time.

• High amounts of heavy metals (e.g., copper) in soil can interfere with
the method.

• Dehydrogenase activity is significantly reduced in acidic soils.

References

Cooper AB, Morgan HW (1981) Improved fluorimetric method to assay for soil lipase

activity. Soil Biol Biochem 13:307-311
Dick RP (1997) Soil enzyme activities as integrative indicators of soil health. In: Pankhurst

CE, Double BM, Gupta VV (eds) Biological indicators of soil health. CAB Int, Oxon,

pp 121-157
Griffiths BS (1989) Improved extraction of iodonitrotetrazolium-formazan from soil with

dimethylformamide. Soil Biol Biochem 21:179-180
Hankin L, Hill DE, Stephens GR (1982) Effect of mulches on bacterial populations and

enzyme activity in soil and vegetable yields. Plant & Soil 64:193-201
Ishimoto R, Sugimoto M, Kawai F (2001) Screening and characterization of trehalose-oleate

hydrolyzing lipase. FEMS Microbiol Lett 195: 231-235
Kato T, Haruki M, Imanaka T, Morikawa M, Kanaya S (2001) Isolation and characteriza-
tion of psychrotrophic bacteria from oil-reservoir water and oil sands. Appl Microbiol

Biotechnol 55:794-800
Margesin R, Feller G, Hammerle M, Stegner U, Schinner F (2002) A colorimetric method

for the determination of lipase activity in soil. Biotechnol Lett 24:27-33
Margesin R, Schinner F (1997) Bioremediation of diesel-oil-contaminated alpine soils at low

temperatures. Appl Microbiol Biotechnol 47:462-468
Margesin R, Schinner F (1998) Biodegradation of the anionic surfactant sodium dodecyl

sulfate at low temperatures. Int Biodet Biodegradation 41:139-143
Margesin R, Schinner F (1999) A feasibility study for the in situ remediation of a former

tank farm. World J Microbiol Biotechnol 15:615-622
Margesin R, Schinner F (2001) Bioremediation (natural attenuation and biostimulation) of

diesel-oil-contaminated soil in an alpine glacier skiing area. Appl Environ Microbiol

67:3127-3133
Margesin R, Walder G, Schinner F (2000a) The impact of hydrocarbon remediation (diesel oil

and polycyclic aromatic hydrocarbons) on enzyme activities and microbial properties

of soil. Acta Biotechnol 20:313-333
Margesin R, Walder G, Schinner F (2003) Bioremediation assessment of a BTEX-

contaminated soil. Acta Biotechnol 23:29-36
Margesin R, Zimmerbauer A, Schinner F (1999) Soil lipase activity - a useful indicator of

oil biodegradation. Biotechnol Tech 13:859-863
Margesin R, Zimmerbauer A, Schinner F (2000b) Monitoring of bioremediation by soil

biological activities. Chemosphere 40:339-346
Mills AL, Breuil C, Colwell RR (1978) Enumeration of petroleum-degrading marine and

estuarine microorganisms by the most probable number method. Can J Microbiol

24:552-557

320 R. Margesin

Pancholy SK, Lynd JQ (1972) Quantitative fluorescence analysis of soil lipase activity. Soil

Biol Biochem 4: 257-259
Plou FJ, Ferrer M, Nuero OM, Calvo MV, Alcalde M, Reyes F, Ballesteros A (1998) Analysis of

Tween 80 as an esterase/lipase substrate for lipolytic activity. Biotechnol Tech 12: 183- 186
Pokorna V (1964) Method of determining the lipolytic activity of upland and lowland peats

and muds. Soviet Soil Sci 1:85-87
Sakai Y, Hayatsu M, Hayano K (2002) Use of Tween 20 as a substrate for assay of lipase

activity in soils. Soil Sci Plant Nutr 48:729-734
Schinner F, Bayer H, Mitterer M (1993) The influence of herbicides on microbial activity in

soil materials. Austrian J Agric Res 34:22-30
Schinner F, Ohlinger R, Kandeler E, Margesin R (eds; 1996) Methods in Soil Biology. Springer,

Berlin Heidelberg New York
Schniirer J, Rosswall T (1982) Fluorescein diacetate hydrolysis as a measure of total microbial

activity in soil and litter. Appl Environ Microbiol 43:1256-1261
Shirai K, Jackson RL (1982) Lipoprotein lipase-catalyzed hydrolysis of p-nitrophenyl bu-

tyrate. J Biol Chem 257:1253-1258
Song HG, Bartha R (1990) Effects of jet fuel on the microbial community of soil. Appl

Environ Microbiol 56:646-651
Sparling GP (1997) Soil microbial biomass, activity and nutrient cycling as indicators of

soil health. In: Pankhurst CE, Double BM, Gupta VV (eds) Biological indicators of soil

health. CAB Int, Oxon, pp 97-119
Trevors JT (1984) Dehydrogenase activity in soil: A comparison between the INT and TTC

assay. Soil Biol Biochem 16:673-674
van Beelen P, Doelman P (1997) Significance and application of microbial toxicity tests in

assessing ecotoxicological risks of contaminants in soil and sediment. Chemosphere

34:455-499
Von Mersi, Schinner F (1991) An improved and accurate method for determining the

dehydrogenase activity of soils with iodonitrotetrazolium chloride. Biol Fertil Soils

11:216-220
Wei YL, Kurihara T, Suzuki T, Esaki N (2003) A novel esterase from a psychrotrophic

bacterium, Acinetobacter sp. strain no. 6, that belongs to the amidase signature family.

J Mol Cat B-Enzym 23:357-365

^ O Assessment of Ecotoxicity
' ^ of Contaminated Soil Usin

Using Bioassays

Adolf Eisentraeger, Kerstin Hund-Rinke, Joerg Roembke

18.1

General Introduction: Strategy

Bioassays provide important information for the assessment of pollutant
effects of chemicals or environmental samples. In contrast to chemical
analyses, they also detect effects of multiple contaminants and metabo-
lites. Standardized bioassays can be used for the path-related, toxicological
characterization of soils and soil materials, taking into account possible
transfer of pollutants to the groundwater and potential effects on soil mi-
croorganisms, earthworms, and plants. A large number of bioassays have
been applied for the characterization of contaminated soil or soil materials
(Spurgeon et al. 2002). Most of them have been developed for the testing
of chemicals and then have been adapted for testing of contaminated soil
samples. In the first case, uncontaminated soil is spiked with chemicals in
defined concentrations and dose-response relationships are obtained and
evaluated further. While this is a straightforward and often standardized
approach, testing and assessing contaminated soils is more difficult, since
many soil samples are contaminated with different kinds of known and un-
known chemicals that can not be quantified comprehensively. In addition,
the soils have different properties (e.g., pH, texture), which themselves can
affect organisms. Whereas the uncontaminated soil sample can be used
as reference sample for the testing of chemicals, it is very difficult to se-
lect uncontaminated reference samples for contaminated soils. Therefore,
bioassays can not be transferred easily to the testing of contaminated soils
and the evaluation of test results is completely different.

The results of bioassays using soil are affected by mobilization, bioavail-
ability, and pathways of transfer of contaminants. The latter ones can be
varied by the kind of organisms and the test design chosen. In addition,
by using a battery of different test systems the effects of contaminants can
be assessed as a whole (i.e., whether they are known or not), thus covering

Adolf Eisentraeger: Institute of Hygiene and Environmental Medicine, Aachen University of

Technology, Pauwelsstr. 30, 52074 Aachen, Germany, E-mail: adolf.eisentraeger@post.rwth-

aachen.de

Kerstin Hund-Rinke: Fraunhofer Institute for Molecular Biology and Applied Ecology, P.O.

Box 1260, 57377 Schmallenberg, Germany

Joerg Roembke: ECT Oekotoxikologie GmbH, Boettgerstr. 2-14, 65439 Floersheim, Germany

Soil Biology, Volume 5

Manual for Soil Analysis

R. Margesin, F. Schinner (Eds.)

322 A. Eisentraeger et al.

potential synergistic or antagonistic effects. Therefore, bioassays are useful
tools for complementing chemical analysis.

Several joint research projects were carried out in the recent years in
order to optimize bioassays with respect to sample treatment, test perfor-
mance, and evaluation of data (Hund-Rinke et al. 2002a, 2002b). After per-
forming a round-robin test and a laboratory intercomparison test, a testing
strategy was proposed by researchers involved in these projects (Dechema
2001; Eisentraeger et al. 2004). The strategy allows a cost efficient assess-
ment of soil samples in addition to chemical analysis in a stepwise approach
using a maximum of nine standardized bioassays. Detailed advice both on
sample treatment and on the interpretation of test data is given. Depending
on the results, recommendations are given whether remediated soils can
be incorporated at unsealed or sealed sites and whether they can be used as
upper or lower soils. The proposed strategy can be expected to contribute
to the discussion on the standardization of soil testing and to promote the
incorporation of biological test methods into soil protection legislation.

As mentioned above, nine different bioassays are applied in a stepwise
approach. The four pathways from soil to (ground)water, to soil microflora,
to soil fauna and higher plants are assessed using several test systems.

The water-extractable ecotoxicological potential of soils and soil mate-
rials is examined to assess whether undesirable effects in the groundwater
or surface water might occur (i.e., after toxic substances have been set free)
by using two aquatic ecotoxicity tests with bacteria and algae (Table 18.1).
Since only the potential is of significance in this context, the selected test
systems are not considered to be ecologically relevant.

Genotoxic substances in contaminated soils may be hazardous both for
soil organisms and human beings. The latter may be exposed via the path
soil - groundwater - since it is one of the major sources for drinking water.
Therefore, the water extractable genotoxic potential is assessed by testing
water extracts using a test of genotoxicity known as the umu test (because
of its dependence on umuC gene induction) according to ISO 13829 (2000;
Table 18.1). The Salmonella! 'microsome test (Ames test) according to DIN
38415 T4 (1999) should be carried out additionally, but only if the umu test
is negative and there are strong hints from chemical analysis or site history
that mutagenic compounds are present (Table 18.1). It should be noted that
the approach presented here is a screening method to identify substances
that can cause gene mutations; it cannot be used to identify clastogenic
substances.

Several bioassays are employed to ascertain different aspects of the habi-
tat function of soils (Table 18.2): Microorganisms are chosen for these that
differ in trophic levels, exposition, and habitat (e.g., bulk soil, air- filled soil
pores; water film of soil pores). Effects on the soil microflora are quantified
via respiration and ammonium oxidation activity. The combined earth-

18 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays

323

Table 18.1. Biological test systems for the ecotoxicological assessment of the water-
extractable ecotoxic and genotoxic potential of contaminated soil or soil materials. (Eisen-
traeger et al. 2004)

Test system

Standard

Comments/modifications

Water quality - determination
of the inhibitory effect of water
samples on the light emission
of Vibrio fischeri luminescent
bacteria test - parts 1-3

Water quality - fresh water al-
gal growth inhibition test with
Scenedesmus subspicatus and
Selenastrum capricornutum

Water quality - determination
of the genotoxicity of water
and waste water using the umu
test

ISO 11348 (1998)

ISO 8692 (1989)

ISO 13829 (2000)

Alternatively testing on

microplates possible

(Eisentraeger et al. 2003;

Rilaetal.2003)

Water quality - determina-
tion of the genotoxicity of wa-
ter and waste water using the
Salmonella/ microsome test
(Ames test)

DIN 38415 T4 (1999)

worm mortality/reproduction test and the Collembola reproduction test
are used to assess effects on soil fauna. Further, the emergence and growth
of Brassica rapa and Avena sativa are used to assess a soil's capacity to
function as a habitat for higher plants.

18.2

Sample Preparation

■ Introduction

Objectives and Principles. Suitable sample preparation is a prerequisite for
obtaining reliable results (ISO 15799 2003). The preparation of soil sam-
ples includes transport, sieving, determination of maximum water-holding
capacity, and water content, as well as adjustment of water content and stor-
age.

Theory. Tests should be performed as soon as possible after sampling. The
period of storage should be minimized, at least for soils containing degrad-

324

A. Eisentraeger et al.

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1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 325

able contaminants. Soil sampling may cause an alteration of the soil con-
ditions, especially of the redox potential. This may result in a degradation
of the contaminants and lead to an inaccurate evaluation of toxic potential.
Due to altered environmental conditions, for example, it was found at a mu-
nition site that the TNT concentration after excavation quickly decreased
by a factor of about 10.

Microbial activity may diminish in storage, leading to erroneous results
with respect to microbial respiration and nitrification rates (see below).
The extent and time period of decrease varies for different soils. If no
degradation is expected, the maximum storage period for uncontaminated
soil samples should not exceed 3 months at 4 °C in the dark. If soil samples
have to be stored for a longer period, -20 °C may be best. Certain aspects
of these procedures, however, remain controversial, especially regarding
potential ammonium oxidation activity (Sect. 18.5.2).

Samples should be stored in a way that changes in the soil water content
are minimized. The vessels used should not influence the composition of the
samples. For soil samples contaminated with organic pollutants, stainless
steel, aluminum, or glass vessels should be used. Container materials of
lower quality may be used for large amounts of soil, in which case it must
be assured that the ratio of soil mass to vessel mass is appropriate. The
decrease of contaminant concentrations (e.g., by sorption to the wall of the
vessel) also must be negligible. Suitable containers for samples containing
inorganic contaminants (heavy metals) usually made of plastic and or other
materials free of heavy metals.

A draft version of a sequential approach to estimate the storage capacity
of soil samples contaminated with volatile organic compounds (VOC) was
set up by Rila and Eisentraeger (2003). This approach is based on the quan-
tification of the VOCs, on the one hand, and of the microbial respiratory
activity, on the other, under the assumption that toxic characteristics of
samples with a high microbial activity and a high VOC concentration are
altered during storage.

The procedure of sample preparation is summarized in Fig. 18.1. Ac-
cording to ISO/DIS 21268 (2004), for the tests with soil eluate, and for those
using terrestrial test organisms other than microorganisms, the soil sam-
ples are sieved to < 4 mm. For microbial tests the soil fraction < 2 mm
is needed. The microbial tests are performed using the indigenous soil
microflora, whereas the other tests are performed with introduced organ-
isms. These latter tests usually require huge amounts of soil. Especially for
highly silty and loamy soils, sieving of large soil volumes to smaller soil
fractions may be difficult with an acceptable expenditure of work, as the
holes of the sieves may plug up within several minutes. This makes frequent
cleaning necessary. Therefore, depending on the organisms introduced, it
was decided to apply different procedures.

326

A. Eisentraeger et al.

SOIL SAMPLING, transportation, storage,
documentation of soil treatment

MAIN SIEVING

(< 4 mm nominal screen size; soil sample

of at least 2 kg), quantification of water

content and water holding capacity

WATER EXTRACTION

using a glass flask (500 ml) with defined

amount of soil (175 g ± 5 g), addition of

water (soil / water 1+2)

AGITATION for 24 h:

end-over-end tumbler: 5-10 rpm or

roller table rotating at about 10 rpm

SETTLING

of suspended soils (15 ± 5 min)

CENTRIFUGATION

at 2,000 - 3,000 g for 20 min

FILTRATION a >

(0.45 um glass fibre membrane filter

without organic glue or regenerated

cellulose)

BIOASSAYS:

water extractable ecotoxicity and
genotoxicity

Adjustment of the water content to
50 ± 10 % WHCmax

INVESTIGATION OF PATHWAYS
SOIL - SOIL FAUNA and PLANTS

Sieving < 2 mm; adjustment of the water
content to 50 ± 10 % WHCmax

INVESTIGATION OF PATHWAY
SOIL - SOIL MICRO-ORGANISMS

Centrifugation at 10,000 - 20,000 g b >

if no genotoxic potential:
solid phase extraction of water extract

BIOASSAYS:

water extractable genotoxicity

a ) As the standard can be used for
the determination of organic pol-
lutants in the eluate and for its
ecotoxicological characterization,
it must be kept in mind that the
use of 0.45 yam filters may reduce
both the concentration of organic
pollutants and the toxicity because
pollutants adsorbed to soil parti-
cles will be blocked by the filter.

b ) If the concentration of bound
pollutants or the global toxicity
of the eluate are of interest other
techniques such as settling or cen-
trifugation are recommended.

Fig. 18.1. Procedure proposed for the preparation of soil samples for ecotoxicological testing
(modified according to Pfeifer et al. 2000; Dechema 2001; Rila and Eisentraeger 2003;
Eisentraeger et al. 2004, ISO/DIS 21268-1 2004)

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 327

Sample Preparation for Investigation of the Pathways: Soil to-Soil Organisms, and Soil
to Plants. It has to be considered that biological determinations require
optimal water content. The water content also has an influence on the
oxygen supply; water saturation, for example, results in a limited oxy-
gen supply. As the demands of organisms regarding humidity and oxygen
supply differ, the optimal water content depends on the planned investi-
gations. For microbiological assessments a water content of 50 ± 10% of
the maximum water holding capacity (WHC max ) is recommended. Soils
with this water content provide the microorganisms living in the soil pore
water with sufficient water as well as oxygen. Similar conditions are rec-
ommended for collembola that live in air-filled soil pores. For tests with
earthworms the appropriate water content is higher, since their surface is
rather sensitive to desiccation. Due to their large size, the thin water film in
soil pores, sufficient for microorganisms, is insufficient for them. Further-
more, the earthworm species used in ecotoxicological tests is a compost
worm, adapted to higher humidity.

The soil has to be dried if the water content is too high for the planned
investigations or for sieving. During the drying process local complete
drying should be avoided. This is essential for microbial investigations,
since complete drying causes a reduction of the microbial population.
Moreover, the aggregates formed in silty and loamy soils will be difficult to
destroy. Localized drying can be prevented by turning the soil periodically.
In addition, structural changes of the soil due to drying can cause problems
in plant tests since roots cannot permeate hard soil aggregates. In such
cases, the soil samples must be re-wetted carefully by hand in order to
reach a moisture suitable for plants, i.e., automatic moistening via wicks
(Sect. 18.5.5) will not be sufficient.

Water Extraction for Ecotoxicological and Genotoxicological Testing. In order to
make a pragmatic estimation of the fraction of contaminants that might
migrate to the groundwater, soil samples are extracted with water in a simple
batch assay. By choosing a dry soil-to-water ratio of 1:2 it is guaranteed,
on the one hand, that enough water sample is available for biotesting and,
on the other, that concentrations of the water-extractable contaminants
remain high. Meanwhile, this approach is accepted, has been successfully
validated in a ring test, and its standardization is in progress (ISO/DIS
21268-1 2004).

Preparation of Solid-Phase Extracts from the Water Extracts for Genotoxicological
Testing. As stated earlier, groundwater contaminated with genotoxic sub-
stances can be hazardous. The water-extractable genotoxic potential is
assessed in order to roughly estimate whether genotoxic compounds might
be mobilized by water. In the first step of the procedure, the same water
extract is tested as is used for the assessment of the water-extractable eco-

328 A. Eisentraeger et al.

toxicological potential. If there is a genotoxic effect in the umu test, with
or without metabolic activation, a high risk of transfer of genotoxic sub-
stances from soil to groundwater exists. If there is no genotoxic effect, the
water extract should be concentrated by a (low) factor of 15 using Serdolit
PAD-1 resin (Boehringer Ingelheim Pharma GmbH & Co. KG, Ingelheim,
Germany) as described below.

■ Equipment and Reagents

• 2 mm sieve (in exceptional cases 4 mm, see Sect. 18.1)

• Cylinders of metal, glass, or plastic (diameter 5-8 cm), sealed at one end
with a finely meshed fabric for determining water-holding capacity

• Sand bath fitted out at the bottom with a discharge valve, filled with fine
sand (grain size 0.1-0.7 mm); to about 10 cm, then saturated with water
before starting the test by closing the valve while letting in the water and
opening it afterward (so that the surplus water can run off), and the sand
then covered with a moist fabric

• Analytical balance

• Heating apparatus for determining water content or drying cabinet and
exsiccator

• Shakers for water extraction

• Centrifuge

• Glass microfiber filters

• Pentane, acetone, dimethylsulfoxide (DMSO), methanol, dichlorome-
thane, cone. HC1, cone. NaOH (of analytical grade)

• Resin (e.g., Serdolit PAD-1 resin; Boehringer Ingelheim, No. 42442)

■ Sample Preparation

The preparation of the soil consists of the following steps:

• Sieving

• Water extraction for aquatic test systems

• Solid phase extraction of the water extract for aquatic genotoxicity test
systems

• Determination of the WHC max (Chap. 2)

• Determination of the water content of the sieved soil (Chap. 2)

• Adjustment of the water content to a specific percentage of WHC
(Chap. 2)

max

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 329

■ Procedure

In the guideline ISO 10381-6 (1993) collection, handling, and storage of
soil for the assessment of aerobic microbial processes in the laboratory is
described. For testing contaminated soils it has to be considered that some
contaminants may interact with vessel material (see Sect. 18.1). Moreover,
alteration of the redox potential during storage should be minimized for
anaerobic soils for which only investigation by aquatic ecotoxicological and
genotoxicological tests is relevant.

Sieving (According to ISO 10381-6 1993)

If the soil is too wet for sieving, it should be spread out, where possible in
a gentle air stream, to facilitate uniform drying. The soil should be finger
crumbled and turned over frequently to avoid excessive surface drying.
Normally this procedure should be performed at ambient temperature.
The soil should not be dried more than necessary to facilitate sieving.

Water Extraction (According to ISO/DIS 21268-2 2004)

The soil samples are extracted by a ratio of 1 part soil dry mass to 2 parts of
water with a minimum amount of 100 g soil dry mass. The water content in
the soil has to be considered. The samples are shaken intensively to simulate
worst-case conditions for 24 h and then centrifuged. The supernatant is
filtered with a glass microfiber filter and stored at 4°C in Duran (Schott
AG, Mainz) glass bottles in the dark. The pH of the elutriates is adjusted
to 7 ± 1 with cone. HC1 or NaOH. Ecotoxicological and genotoxicological
testing should be performed within 8 days.

Preparation of Solid-Phase Extracts from the Water Extracts
for Genotoxicological Testing

The solid-phase extraction of the water extract is performed with Serdo-
lit PAD-1 resin, an ethylstyrene-DVB-copolymer with a particle size of
0.3-1.0 mm and a pore diameter of ca. 25 nm with a specific surface of
ca. 250m 2 /g. The PAD-1 beads are pretreated by rinsing for 2h in warm
10% (v/v) HC1, Millipore water, 10% (v/v) NaOH, and Millipore water
successively followed by 8h Soxhlet extraction with pentane/acetone in
a ratio of 1:2. The beads are dried at a temperature of 1 10 °C. Shortly before
solid phase extraction 10 g PAD-1 beads are preconditioned by shaking
them with 25 mL methanol.

The water extract should be concentrated by a factor of 15 by mixing
375 mL with 10 g Serdolit PAD-1 beads. This suspension is placed on an
overhead shaker for 2.5 h. The beads are removed from the water extract
and dried under nitrogen atmosphere in a Baker-spe-10 system (J.T. Baker,

330 A. Eisentraeger et al.

Phillipsburg, New Jersey, USA). The dried beads are then extracted with
a mixture of 9 parts dichloromethane and 1 part methanol. One mL of
DMSO is added to the solvent, which is then evaporated under nitrogen
atmosphere to a final volume of 1 mL. The concentrated sample is stored
for less than 8 days at 4°C. The sample is adjusted with distilled water to
a volume of 25 mL for the genotoxicity tests. The final DMSO concentration
is 4%. Therefore, the concentration factor for the water soil extract is 15.

■ Notes and Points to Watch

• As already mentioned in Sect. 18.1, localized drying of the soil has to be
avoided.

• The soil should be processed as soon as possible after sampling. Any
delays due to transportation should be minimized.

• Microbial tests: if storage is unavoidable, this should not exceed 3 months,
unless evidence of continued microbial activity is provided. Even at low
temperatures the active soil microflora decreases with increasing storage
time; the rate of decrease depends on the composition of the soil and the
microflora.

• Soil fauna tests and tests using higher plants: there are no specific recom-
mendations for soil storage with respect to soil fauna and higher plants
in ISO standards. Therefore it is recommended to store the soil sam-
ples under the same conditions as for testing of microbes and microbial
processes.

• Aquatic tests: for testing the leaching potential, water extracts for aquatic
tests should be prepared immediately after sieving. If the tests cannot be
performed within 8 days (storage of the extracts at 4 ± 2 °C in the dark),
extracts should be stored at -20 °C.

• An ISO guidance paper on the long and short term storage of soil samples
is in process.

18.3

Water-Extractable Ecotoxicity

18.3.1

Vibrio fischeri Luminescence-Inhibition Assay

■ Introduction

Objectives. This test is an acute toxicity test with the marine lumines-
cent bacterium Vibrio fischeri NRRL B-11177 (formerly known as Photo-

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 331

bacterium phosphoreum). It is standardized for the determination of the
inhibitory effect of water samples in the ISO guideline 11348 parts 1-3
(1998). In the strategy presented here, it is used to determine whether toxic
substances are present in the aqueous soil extracts.

Principle. The test system measures the light output of the luminescent
bacteria after they have been challenged by a sample and compares it to
the light output of a blank control sample. The difference in light output
(between the sample and the control) is attributed to the effect of the sample
on the organisms. The test is based on the fact that the light output of the
bacteria is reduced when it is introduced to toxic chemicals.

Theory. V.fischeri emits a part of its metabolic energy as blue-green light
(490 nm). Biochemically luminescence is a byway of the respiratory chain.
Reduction equivalents are separated and transmitted to a special acceptor
(flavin mononucleotide, FMN; Engebrecht et al. 1983). During the oxidation
of substrates by dehydrogenase hydrogen is transferred to nicotinamide
adenine dinucleotide (NAD). The reduced NAD (NADH 2 ) transfers the hy-
drogen normally to the electron transport chain. To get bacterial lumines-
cence, a part of the NADH 2 is used to build reduced flavin mononucleotide
(FMNH 2 ). FMNH 2 builds a complex with luciferase which involves the
oxidation of a long-chain aliphatic aldehyde, developing an excited energy
state. The complex decomposes and emits a photon. The oxidation prod-
ucts FMN and the long chain fatty acid are reduced in the next reaction
cycle by NADPH 2 .

FMNH 2 + RCHO + 2 -> Luciferase -> FMN + RCOOH + H 2 + hv

This luminescence is inhibited in the presence of hazardous substances.
Since it is dependent on reduction equivalents, the luminescence inhibitory
test is a physiological test belonging to the electron-transport-chain-activi-
ty group.

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculations are
described in detail in ISO 1 1348 (1998).

18.3.2

Desmodesmus subspicatus Growth-Inhibition Assay

■ Introduction

Objectives. This fresh water algal growth inhibition assay is performed
according to the standard ISO 8692 (1989). It is applicable both for the

332 A. Eisentraeger et al.

characterization of chemicals and aquatic environmental samples. While
the standard allows the testing with two strains (Desmodesmus subspica-
tus, formerly Scenedesmus subspicatus, and Selenastrum capricornutum),
the strategy for soil characterization presented here has been set up and
validated using the strain D. subspicatus. The algal growth inhibition test
complements the acute bacterial luminescence test with V.fischeri.

Principle. The growth of D. subspicatus in batch cultivation in a defined
medium over 72 ± 2 h is quantified both in the presence and the absence
of a sample. The cell density is measured at least every 24 h using direct
methods like cell counting or indirect methods correlating with the di-
rect methods, such as in vivo chlorophyll fluorescence measurement. The
inhibition is measured as a reduction in growth rate.

Theory. D. subspicatus is a fresh water algae that can be easily cultivated
under defined conditions at 23 ± 2 °C with a light intensity in the range of
35 x 10 18 to70 x 10 18 photons/m 2 /s. Since it is based on growth inhibition,
all specific or nonspecific toxic effects relevant to reproduction of these
algae are assessed with this test system.

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculations are
described in detail in ISO 8692 (1989).

18.4

Water-Extractable Genotoxicity

18.4.1

The umu Test

■ Introduction

Objectives. The umu test is a short-term genotoxicity assay carried out on
microplates within less than 8 h. It is standardized for the examination of
water and waste water (ISO 13829 2000). The water-extractable potential of
soil samples is assessed by testing the water extract and (if the water extract
is not genotoxic) the 15-fold concentrated water extract. The results give
hints as to whether genotoxic substances might migrate to the groundwater.
The umu test was chosen since it is widely applied for the examination of
aquatic environmental samples and since both costs and time needed are
reasonable. The procedure has been optimized and validated by charac-
terizing large numbers of contaminated and uncontaminated soil samples
(Ehrlichmann et al. 2000; Rila et al. 2002; Rila and Eisentraeger 2003).

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 333

Principle. The bioassay is performed with the genetically engineered bac-
terium Salmonella choleraesuis subsp. choleraesuis TA1535/pSK1002 (for-
merly Salmonella typhimurium). This strain is exposed to different con-
centrations of the samples. Different kinds of genotoxic substances can be
detected using this test since the strain responds with different types of
genotoxic lesions, depending on the toxin.

Theory. The test is based on the capability of genotoxic agents to in-
duce the umuC gene which is a part of the SOS repair system in re-
sponse to genotoxic substances. The umuC gene is fused with the lacZ
gene for /J-galactosidase activity. The /?-galactosidase converts ONPG (o-
nitrophenol-/?-D-galactopyranoside) to galactose, and the yellow substance
o-nitrophenol is quantified photometrically at 420 ± 20 nm. The tests are
preformed both with and without metabolic activation by S9-mixture (liver
enzymes). Cytotoxic characteristics of the samples are quantified photo-
metrically in parallel.

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculations are
described in detail in ISO 8692 (1989).

18.4.2

Salmonella/Microsome Assay (Ames Test)

■ Introduction

Objectives. The Salmonella/microsome assay (Ames test) is a bacterial mu-
tagenicity assay that is standardized according to DIN 38415 T4 (1999) for
the determination of the genotoxic potential of water and waste water
(Ames et al. 1975). In the strategy presented here, it is recommended if the
umu test is negative and if there are strong hints from chemical analysis or
site history that mutagenic compounds are present. Thus it complements
the umu test in some cases.

This method includes sterile filtration of the aquatic sample prior to the
test. Due to this filtration, solid particles will be separated from the test
sample. It may be possible that genotoxic substances are adsorbed by these
particles. If so, they will not be detected.

Principle. The bacterial strains Salmonella typhimurium TA 100 and TA 98
should be used. The possible mutagenic activity of the sample is detected by
comparing, for the bacterial strain and its activation condition, the number
of mutant colonies on plates treated with the negative control and on plates
treated with undiluted and diluted test samples.

334 A. Eisentraeger et al.

The bacteria will be exposed under defined conditions to various doses of
the test sample and incubated for 48-72 h at 37 ± 1 °C. Under this exposure,
genotoxic agents contained in water or waste water may be able to induce
mutations in one or both marker genes (hisG46 for TA 100 and hisD3052
for TA 98) in correlation with the dosage. Such induction of mutations will
cause a dose-related increase of the numbers of mutant colonies of one or
both strains to a biologically relevant degree above that in the control.

Theory. Bacteria that are not able to synthesize histidine are exposed to
mutagenous substances inducing a reversion to the wild type growing in the
absence of histidine. Histidine auxotrophy is caused by different mutations
in the histidine operon: S. typhimurium TA 98 contains the frameshift
mutation hisD3052 reverting to histidine independency by addition or
loss of base pairs. S. typhimurium TA 100 bears the base pair substitution
hisG46 which can be reverted via base pair substitutions (transition or
transversion).

The tester strains are deep rough enabling larger molecules also to pen-
etrate the bacterial cell wall and produce mutations (rfa mutation). The
excision repair system is disabled (Z\uvrB), increasing the sensitivity by
reducing the capability to repair DNA damage. Furthermore, they contain
the plasmid pKMlOl coding for an ampicillin resistance.

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculations are
described in detail in DIN 38415 T4 (1999). An ISO standard is in prepara-
tion.

18.5

Habitat Function:

Soil/Microorganisms, Soil/Soil Fauna, Soil/Higher Plants

18.5.1

Respiration Curve Test

■ Introduction

Objectives. The determination of respiration curves provides information
on the microbial biomass in soils and its activity. The method is suitable
for monitoring soil quality and evaluating the ecotoxicological potential of
soils. It can be used for soils sampled in the field or during remediation
processes. The method is also suitable for soils that are contaminated
experimentally either in the field or in the laboratory (chemical testing).

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 335

Principle. The C0 2 production or 2 consumption (respiration rate) from
unamended soils as well as the decomposition of an easily biodegradable
substrate (glucose + ammonium + phosphate) is monitored regularly (e.g.,
every hour). From the C0 2 -production or 2 -consumption data the dif-
ferent microbial parameters, such as basal respiration, substrate-induced
respiration, lag time, are calculated.

Theory. Basal respiration and substrate-induced respiration (SIR) are wide-
ly used physiological methods for the characterization of soil microbial
activity and biomass. Basal respiration gives information on the actual
state of microbial activity in the soil. After addition of an easily biodegrad-
able carbon source respiration activity increases. At the time of substrate
addition the activity can be described by

SIR = r + K

where r is the initial respiration rate of growing microorganisms.

In the course of an incubation period the respiration rate increases and
can be described by

dp/dt = rept + K

This equation is based on the assumption that the increase of the respi-
ration rate dp/dt after substrate addition in the SIR method represents
the sum of the respiration rates of growing (rept) and non-growing (K)
microorganisms (Stenstrom et al. 1998).

The microbial respiration activity is affected by several parameters. Wa-
ter content, temperature (Blagodatskaya et al. 1996), the quality of the soil
organic matter (Wander 2004), as well as contaminants (e.g., Blagodatskaya
and Anan'eva 1996; Kandeler et al. 1996) show an influence.

■ Procedure

Sample preparation, equipment, reagents, procedure, and calculations are
described in detail in ISO 17155 (2002). A prerequisite is equipment that
allows the determination of C0 2 release or 2 uptake at short time in-
tervals. Basal respiration is measured first. The respiration rates should
be measured until constant rates are obtained. After measuring the basal
respiration, a defined substrate mixture containing glucose, potassium di-
hydrogen phosphate, and diammonium sulfate is added. The mixture is
made up of: 80 g glucose, 13 g KH 2 P0 4) and 2 g (NH 4 ) 2 S0 4 . In testing, 0.2 g
mixture is used per gram of soil in which at least 1 g organic matter is
found in 100 g soil dry mass. The measurement of C0 2 evolution or 2
consumption has to be continued until the respiration curve reaches its
peak and the respiration rates are declining.

336 A. Eisentraeger et al.

The ecotoxicological potential of soils is described by several parameters:

• Respiratory activation quotient: basal respiration rate divided by sub-
strate-induced respiration rate (Qr = Rb/Rs)

• Lag time (ti ag ): the time from addition of a growth substrate until ex-
ponential growth starts, - a reflection of the vitality of the growing
organisms

• Time to the peak maximum (t pea kmax): the time from addition of growth
substrate to the maximum respiration rate - another reflection of the
vitality of the growing organisms

According to the guideline, Q R > 0.3, ti ag > 20 h, and t pea kmax > 50h
indicate polluted materials.

■ Notes and Points to Watch

• Increased respiratory activation quotients may occur for two reasons.
On one hand, they are an indicator of bioavailable carbon sources. These
may be of biological origin, as for example compost, or biodegradable
organic contaminants (e.g., mineral oil, anthracene oil, phenanthrene)
that have the same effect (Hund and Schenk 1994). Sufficient amounts
of biodegradable carbon sources always result in increased respiration
activities when a sufficient amount of further nutrients (e.g., nitrogen,
phosphate) is available. On the other hand increased Q R s may be an
indicator of contaminants that are not biodegradable, e.g., heavy metals
(Nordgren et al. 1988). Up to now, it is not known how to distinguish
which parameters are responsible for a stress-induced respiration caus-
ing increased quotients.

• It has to be considered for the assessment that increased values indi-
cate amended/contaminated soils, whereas not all contaminated soils
show higher values. Accordingly, it cannot be concluded that the habitat
function of a soil is intact when the respiration values are in a normal
range.

• In the literature, the derivation of a metabolic quotient (basal respira-
tion divided by microbial biomass) as an indicator for an ecosystem is
described (Insam and Domsch 1988; Anderson and Domsch 1990). In
soils with a recent input of easily biodegradable substrates, mainly r-
strategists occur. They usually respire more C0 2 per unit degradable C
than k-strategists, which prevail in soils that have not received fresh or-
ganic matter and have evolved a more complex detritus food web (Insam
1990). Since the substrate-induced respiration can be used to calculate
the microbial biomass, it could be concluded that the metabolic quotient

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 337

and the respiration activation quotient are comparable. In this context it
should be noted that the estimation of the microbial biomass by Ander-
son and Domsch (1978) is based on a linear regression between SIR and
the microbial biomass according to the fumigation- incubation method.
The conversion factor was elaborated on the basis of a range of soils.
However, in other soils the population may differ from the originally
investigated soils (e.g., forest soils vs. contaminated soils) and different
conversion factors may be necessary (Hintze et al. 1994). One should,
therefore, avoid calculating the microbial biomass of soils on the basis
of the substrate-induced respiration for which the conversion factor is
unknown.

18.5.2

Ammonium Oxidation Test

■ Introduction

Objectives. This test is a rapid procedure for determining the potential
rate of ammonium oxidation in soils. The method is suitable for all soils
containing a population of nitrifying organisms. It can be used as a rapid
screening test for monitoring the quality of soils and wastes, and it is
suitable for testing the effects of cultivation methods, chemical substances,
and pollution in soils.

Principle. Ammonium oxidation, the first step in autotrophic nitrification
in soil, is used to assess the potential activity of microbial nitrifying pop-
ulations. Autotrophic ammonium-oxidizing bacteria are exposed to am-
monium sulfate in a soil slurry. Oxidation of the nitrite formed by nitrite-
oxidizing bacteria in the slurry is inhibited by the addition of sodium
chlorate. The subsequent accumulation of nitrite is measured over a 6-h
incubation period and is taken as an estimate of the potential activity of
ammonium oxidizing bacteria. For the assessment of soil quality the nitri-
fication activity in a test soil, in a control soil, and in a mixture of both soils
is determined.

Theory. In soils with pH > 5. 5 nitrification is performed by chemoau-
totrophic nitrifiers (Focht and Verstraete 1977). The procedure consists of
two steps. Ammonium is oxidized to nitrite by one group of nitrifiers, while
nitrite is oxidized to nitrate by a second group. Since nitrite is oxidized as
it is produced, the rate at which ammonium is oxidized is equal to that at
which nitrite plus nitrate accumulate. To avoid the application of two meth-
ods - one for the determination of nitrite and one for determining nitrate -
a procedure was developed to completely and specifically block the oxida-
tion of nitrite. With this method it is possible to get information on the

338 A. Eisentraeger et al.

nitrification process by using only one analytical method, since the rate at
which nitrite alone accumulates equals the rate of ammonium oxidation. In
soils with a high background of nitrate this method is much more sensitive,
since nitrite normally is undetectable at the beginning of the incubation.
A prerequisite for a correct measurement is (1) that the inhibitor does not
inhibit ammonium oxidation, and (2) that the inhibitor completely blocks
nitrite oxidation. Chlorate has proved to be an appropriate inhibitor. At
suitable concentrations an inhibition of ammonium oxidation seems to be
negligible. Although, in some cases, the inhibition of nitrite oxidation can
be incomplete, this does not seem to be a real problem. It is negligible when
Vmax f° r nitrite oxidation is lower than the rate of ammonium oxidation. It
might be a problem, if V max is larger. Since chlorate mainly influence the K m
of the reaction, the initial rate of the reaction is the best estimate of the am-
monium oxidation rate. Leakage will be lowest at low nitrite concentrations
(Belser and Mays 1980).

The results present a potential activity, since several test parameters are
different from natural conditions: Ammonium is added in surplus, aeration
is probably more intensive by shaking in the laboratory than under field
conditions, and the incubation temperature of 25 °C usually far exceeds
real soil conditions.

Several methods exist to get information on nitrification in soil. Some
of these are characterized by incubation periods of several weeks (e.g., ISO
14238 1997). For soil assessments the determination of the ammonium
oxidation activity was selected since this procedure has several advantages,
especially for investigation of contaminated soils and for soil remediation
procedures. These applications frequently require results within a short
period of time, as they contribute to decisions whether a soil has to be
remediated, whether a remediation has to be continued, or whether the
habitat function of the soil (at least with respect to microorganisms) is intact
so that the soil can leave the remediation plant. This is important in avoiding
unneeded and expensive retention of soil in the remediation plants. As the
potential ammonium oxidation method yields results in a short period
of time, and furthermore is suitable for soils with high nitrate contents
(during bioremediation nitrogen has to be added to achieve degradation
of contaminants), this method was selected for the ecotoxicological soil
assessment.

■ Procedure

Sample preparation, equipment, reagents, procedure, and calculations are
described in detail in ISO 15685 (2004). For soil assessments three different
test designs are applied:

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 339

1. Test soil

2. Reference soil (uncontaminated soil with a nitrification activity of about
200-800 ng N/g dry mass of soil/h)

3. Mixture of test soil and reference soil (1:1 with regard to soil dry mass)

The soils are adjusted to 60% of WHC max and incubated for 2 days at 20 ° C.
The mixture is prepared immediately before testing. The mixture and the
two soils are incubated again for 1 day at 20 °C, after which the nitrification
activity is determined. Soil samples are mixed with test medium containing
phosphate, sodium chlorate and diammonium sulfate. The slurries are
incubated for 6h at 25 ± 2°C on an orbital shaking incubator (about
175 rpm). 2-mL samples are taken after 2 and 6 h, and the nitrite content is
determined. The mentioned time interval is a recommendation.

■ Calculation

The rate of ammonium oxidation (ng NO2 -N /g dry mass of soil/h) is
calculated from the difference of NO2 -N concentrations at the different
measuring times. The following formula is applied for the assessment of
the test soil:

M m + SD m < 0. 9 x (M c - M P )/2 (18.1)

M m mean ammonium oxidation activity in soil mixture

SD m standard deviation of ammonium activity in replicate test vessels with
soil mixture

M c mean ammonium oxidation activity in control soil

M P mean ammonium oxidation activity in polluted soil

The polluted soil is considered to be toxic if the mixture has an am-
monium oxidation activity significantly slower than 90% of the calculated
mean activity of the two single soils.

■ Notes and Points to Watch

• The suitability of storing soil samples at -20 °C is discussed controver-
sially. The investigation of 12 soils differing in their physico-chemical
properties has revealed that storage at -20 °C for 13 months does not
affect the nitrifiers in annually frozen soils in any decisive way (Sten-
berg et al. 1998). As the procedure, however, does not seem to be suitable
for every soil, in the guideline ISO 15685 (2004) storage at -20 °C is not
generally recommended. The different results found in the literature on
the effects of freezing as a storage method can be explained in a number

340 A. Eisentraeger et al.

of ways: The populations in soils annually subjected to several freeze and
thaw cycles seem to be adapted and more resistant to freezing than the
microflora in soils where freeze and thaw cycles are not a regular occur-
rence. Furthermore, the growth status of the microorganisms at the time
of sampling may play a role. Active cells seem to be more sensitive to
freezing and thawing than less active cells. Therefore, samples collected
shortly after managing processes such as fertilizing or tilling may show
cell depletion. Furthermore, the selected procedure of freezing and thaw-
ing may influence the results. Slow rates of temperature change seem to
result in greater microbial losses. Storage in small portions and rapid
temperature flux maybe preferable (Stenberg et al. 1998). In conclusion,
soils should only be stored if the effect is known and acceptable.

18.5.3

Combined Earthworm Mortality/Reproduction Test

■ Introduction

Objectives. The determination of the survival and the reproductive success
of earthworms as representatives of soil macrofauna provide information
on these saprophagous soft-bodied invertebrates that in many soils play an
important role as ecosystem engineers. The method is suitable for moni-
toring soil quality and the evaluation of the ecotoxicological potential of
soils. It can be used for soils sampled in the field or during remediation pro-
cesses. Furthermore the method is suitable for soils that are contaminated
experimentally in the field or in the laboratory (e.g., chemical testing, in
particular pesticide testing).

Principle. Adult earthworms are either exposed to potentially contami-
nated soil samples or to a range of concentrations of a test substance mixed
in an artificial or natural control soil. The mortality and the biomass of the
adult worms are measured after 28 days. The effect on the reproduction is
determined by counting the number of juveniles hatched from the cocoons
after an additional period of 4 weeks. Based on these measurements, the
ecotoxicological potential of the test soil is determined.

Theory. Earthworms are important members of the soil community due
to their ability to change or create their habitat through various activities,
thus correctly considered to be "ecosystem engineers" (Lavelle et al. 1997):

• Penetrating the soil and building burrows, as well as increasing pore
space

• Transporting soil and organic matter by casting

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 341

• Comminuting organic material (including cattle feces in meadows) as
a first step in its breakdown

• Providing nutrients to plants (e.g., by concentrating them in burrow
linings or by increasing the availability of nutrients like phosphorus)

• Relocating seeds in the soil profile

• Changing the diversity and improving the activity of the microbial com-
munity by selective feeding and providing feces rich in nutrients

Finally, earthworms are closely exposed to all contaminants occurring
in the soil solution but also - by feeding - to all chemicals adsorbed to soil
particles.

These activities thus finally lead to an improved soil structure, i.e. to
stabilization of soil aggregates, to increase in water infiltration (partly by
higher water-holding capacity; Urbanek and Dolezak 1992; Edwards and
Shipitalo 1998), often to the formation of a humic layer close to the soil
surface (mainly in forest ecosystems; Doube and Brown 1998), and to an
increased yield in orchards or grassland (Blakemore 1997). The activities
described above are performed by various earthworm species to a very dif-
ferent extent. Still, large, deep -burrowing worms like Lumbricus terrestris
are involved in several of these activities, especially concerning soil struc-
ture and organic matter breakdown (Swift et al. 1979). In the light of this
knowledge, it is difficult to understand why the main earthworm species
used in tests are the two closely related compost worms Eisenia fetida or
Eisenia andrei. Ecologically, these species are less important than the deep-
burrowing worms (L0kke and van Gestel 1998). On the other hand, from
a practical point of view the compost worms are more suitable than any
other lumbricid species because they reproduce very quickly and easily in
the laboratory, and mass cultures can be obtained. In addition, the sensi-
tivity of these species is in the same general order of magnitude as other
earthworm species. In most cases the differences between species are, de-
pending on the chemical or contaminant mixture tested, not larger than by
a factor of 10 (Roembke 1997; Jones and Hart 1998).

Concerning the test endpoints, the determination of mortality covers
strong acute effects. However, from an ecological point of view such effects
are clearly less important than long-term, chronic effects usually occurring
at relatively low and thus more realistic concentrations (see "Notes and
Points to Watch"). For this reason, reproduction is the test variable of
highest relevance.

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculation of
the test results are described in detail in the ISO guidelines 1 1268-1 (1993)

342 A. Eisentraeger et al.

and 11268-2 (1998). In deviation from these guidelines in which the acute
and chronic endpoints are determined in individual test runs, it is recom-
mended to use a combined test method for the assessment of contaminated
soils. For the assessment of single chemicals, separate tests should still
be used in order to be in agreement with legal requirements concerning
the risk assessment of chemicals (e.g., the EU guideline describing the
registration of pesticides; European Union 1991).

Ten adult earthworms of the species E.fetida or E. andrei per test vessel
are exposed to a series of mixtures of the potentially contaminated test soil
and an uncontaminated control or reference soil at 20 ± 2 °C for 4 weeks.
If the mortality in the contaminated test soil is higher than 20%, the test is
stopped. Otherwise, at the end of this period, the adult worms are removed
from the vessels and the surviving animals are counted and weighed. After-
wards, the test soil remains in the same vessels for another 4 weeks. After
56 days the juveniles are extracted from test and control soils and counted.
For the endpoint reproduction the data of the test soil vessels are compared
with those from the controls. An inhibition of reproduction of 50% com-
pared to the control is indicative of a contaminated soil sample. A soil that
causes mortality higher than 20% is also classified as contaminated.

■ Notes and Points to Watch

• As already mentioned, the acute test endpoint mortality is ecologically
not relevant due to the following reasons: Lumbricid worms die slowly
and only at high concentrations of soil contaminants. In real field situa-
tions (with the exception of relatively small areas like mining deposits)
the concentrations of chemicals are low but these substances, in particu-
lar metals, are often persistent. Such effects are much better determined
by using chronic sensitive endpoints like reproduction. Ecologically, in
many populations of earthworms any impact more strongly affects the
reproductive rate than it does mortality rate. A short-term decrease in
the number of individuals is easier to compensate than a long-term re-
duction in the number of juveniles. For this reason, the assessment of
the biological quality of soil should be based on the chronic endpoint
reproduction.

18.5.4

Collembola Reproduction Test

■ Introduction

Objectives. The determination of the survival and the reproductive success
of collembolans as representatives of soil mesofauna provides information

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 343

on these saprophagous hard-bodied invertebrates, an important part of
the soil food web in many soils. The method is suitable for monitoring soil
quality and evaluating the ecotoxicological potential of soils. It can be used
for soils sampled in the field or during remediation processes. Furthermore,
the method is suitable for soils contaminated experimentally in the field or
in the laboratory (e.g., chemical testing, in particular pesticide testing).

Principle. Juvenile collembolans are either exposed to potentially contam-
inated soil samples or to a range of concentrations of the test substance
mixed in artificial soil. The mortality of the adult springtails as well as the
reproduction (= number of juveniles) are measured at the end of the expo-
sure period of 28 days. Based on these measurements, the ecotoxicological
potential of the test soil is determined.

Theory. The species Folsomia Candida (Collembola) is tested as a repre-
sentative of hard-bodied soil invertebrates, in particular arthropods (Ac-
hazi et al. 2000). These organisms, mainly consisting of springtails (Collem-
bola) and mites (Acari), are among the most numerous invertebrates in
a wide range of soil types, especially of the Northern hemisphere. Due
to their high numbers they are an important part of the soil food web
(Weigmann 1993). In addition, the springtails control by their feeding ac-
tivity the population cycles of microorganisms, which in turn are extremely
important as mineralizers of organic matter (Swift et al. 1979). To a lesser
extent, springtails can also influence the numbers of nematodes (Hopkin
1997). Finally, they are exposed to contaminants via pore water and air
space.

The species F. Candida is distributed worldwide (mainly by anthro-
pogenic activities). It prefers soils with an elevated content of organic
matter but is not only a compost inhabitant (e.g., it occurs in comparatively
low numbers in agricultural soils; Petersen 1994; Hopkin 1997). Its use is
criticized for the same reasons discussed for compost worms. However,
the response is similar: F. Candida is easily cultured and its sensitivity, as
far as known, is not considerably different from other collembolans (Ac-
hazi et al. 2000). As in the case of earthworms, the endpoint reproduction
is ecologically highly important (see Sect. 18.5.5).

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculation of the
test results are described in detail in the ISO guideline 11267 (1999). Ten
juvenile springtails of the species F. Candida per test vessel are exposed to
a potentially contaminated soil sample or a series of mixtures between the
test soil and an uncontaminated control or reference soil (plus a control) at
20±2 ° C for 4 weeks. At the end of this period, the collembolans are removed

344 A. Eisentraeger et al.

from the vessels and the surviving animals are counted (juveniles and
adults separately) by using photographs or an automatic image processing
system. For the endpoint reproduction, the data from the test soil vessels
are compared with the controls. An inhibition of reproduction of 50%
compared to the control is indicative of a contaminated soil sample.

■ Notes and Points to Watch

• The common test species F. Candida is difficult to distinguish from other
species of the same genus, in particular F. fimetaria (Wiles and Krogh
1998). This species has also been proposed for ecotoxicological testing,
but it reproduces sexually and is, as such, more difficult to handle. Due to
such practical problems and since it is not known whether the two species
are equally sensitive to chemicals, any mixing of them must be carefully
avoided. In cases of doubt a taxonomist specialized in collembolans
should be consulted.

18.5.5

Plant Growth Test

■ Introduction

Objectives. The determination of the emergence and growth of different
plant species allows assessment of the quality of a certain soil as a habitat
for terrestrial primary producers (i.e., in terms of nutrient cycling, the basis
of the whole ecosystem). The method is suitable for monitoring soil quality
and evaluating the ecotoxicological potential of soils. It can be used for
soils sampled in the field or during remediation processes. Furthermore,
the method is suitable for soils that are contaminated experimentally in the
field or in the laboratory (chemical testing, in particular pesticide testing).

Principle. This phytotoxicity test is based on the emergence and early
growth response of a variety of terrestrial plant species to potentially
contaminated soil. Seeds of selected species of plants are planted in pots
containing the test soil and in control pots. They are kept under growing
conditions for the chosen plants and the emergence and mass of the test
plants are compared against those of control plants.

Theory. The importance of plants as the basis of ecosystem performance,
but also for the production of food and forage, cannot be overestimated
(Riepert et al. 2000). In 1984, plants were added to the list of terrestrial
test species by the OECD. These selected species still represent agricultural
plants, while wild herbs, trees, etc., are usually not tested (Boutin et al. 1 995).

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 345

For the testing of chemicals, often two exposure pathways are distinguished:
airborne via aboveground plant parts (e.g., after the spraying of pesticides)
or via soil mixtures. Obviously, in the case of contaminated soil only the
latter test version is used.

Concerning the measurement endpoints, the fresh biomass of the above-
ground parts has been selected due to the practicability of evaluating it and
its high sensitivity. However, one must be aware that this selection has been
done for an acute test with a duration of 1 4 days. Further research will clarify
whether long-lasting chronic tests (e.g., using the endpoint reproduction)
will be more sensitive (ISO 22030 2004).

■ Procedure

Equipment, reagents, sample preparation, procedure, and calculation of
the test results are described in detail in the ISO guideline 11269-2 (1995).
In supplementing the guideline the test was changed in two ways:

1 . In addition to the pure test soils, mixtures of the potentially contaminated
soils with a suitable control or reference soil are made in a ratio of
50:50.

2. While the ISO lists 15 potential test species, it is recommended to use
only the monocotyledonous species Avena sativa (oat) and one of the two
named dicotyledonous species, either Brassica rapa (turnip) or Lepidum
sativum (cress), for soil quality assessment. Each treatment is tested in
four replicates (10 seeds per replicate (= test vessel)). Watering is done
by using a semi-automated wick method (Stalder and Pestemer 1980).
After emergence, the seedlings are thinned to a final number of five per
vessel. Fourteen days later the aboveground parts of the plants (fresh
mass) are harvested and weighed.

■ Evaluation

The evaluation is done according to the following formula (Winkel and
Wilke 2000):

M g + SD Mg < 0. 9 x M b (18.2)

M g Biomass measured in the vessels with the 50:50 mixture of test

and control soil

SD Mg Standard deviation of the 50:50 mixture between test and control

soil

346 A. Eisentraeger et al.

0. 9 x Mb The calculated mean between the test and the control soil (bio-

mass t est soil + biomass C ontroisoii) divided by 2 minus a tolerance
value of 10%.

A soil is classified as toxic if the biomass measured in the vessels with
the 50:50 mixture of test and control soil is > 10% lower than the mean
biomass determined in the test and control soils.

Notes and Points to Watch

In addition to storage problems already mentioned in the context of other
terrestrial tests, it must be pointed out that in the case of plant testing the
amount of plant- available nitrogen is very important for the growth of
the test organisms, including the controls. If the plants grow badly in the
controls it is difficult to identify effects occurring in the test vessels with
test soils. For this reason, Riepert and Felgentreu (2000) recommended
to avoid the use of fresh field soils because they don't contain enough
available nitrogen due to high microbial activity. In order to solve this
general problem fertilizer could be added to the water reservoirs used in
the plant tests. Since all plants (both in the test as well as in the control
vessels) are on the same nutrient level any effect caused by nitrogen
availability would be eliminated. However, one must be cautious since
some soils might be already so rich in nutrients that over-fertilization
could occur.

Another problem in testing potentially contaminated soils with plants
is the fact that structural properties of the soil can affect the plants
too. If the habitat function of the soil has to be assessed in general, the
distinction between chemical and physical properties is not necessary.
However, there are many field soils which are not suitable for the growth
of crop species (e.g., acid soils). In order to avoid false positive results,
the ecological requirements of the common test species (oat, turnip)
are currently being studied (Jessen-Hesse et al. 2003). These data will
allow the determination of which soils can be tested with the current test
species and which cannot.

18.5.6

Test Performance for the Derivation of Threshold Values

■ Introduction

Objectives. The described terrestrial ecotoxicological tests are also suitable
for the derivation of threshold values to protect the habitat function of

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 347

soils for soil organisms. The protection of this soil function is required in
the German Soil Protection Act (BBodSchG 1998). The threshold values
indicate the contamination pathway soil to soil organisms.

Principle. Soils are contaminated experimentally and the biological effect
is investigated (chemical testing). Several concentrations are tested and the
ecotoxicological potential is determined. Based on these measurements
LC 50 (lethal concentration) or EC 50 (effective concentration) values for the
different endpoints are calculated, using appropriate statistical methods.

Theory. Ecotoxicological tests provide information on the toxicity of pri-
ority contaminants. For the derivation of trigger values it has to be kept
in mind that only a limited number of species and organisms have been
tested. To protect the "whole" ecosystem, extrapolation methods have to
be applied. Depending on the amount of available data the extrapolation
method DIBAEX (distribution based extrapolation; Wagner and L0kke
1991) or FAME (factorial application method; European Union 1996) may
be suitable. For the derivation of trigger values concerning the pathway
soil to soil organisms, this procedure was successfully applied in Germany
(Wilke et al. 2001; Wilke et al. 2004). In Germany trigger values are those
which, if exceeded, indicate a harmful soil change or site contamination.
If such cases occur, investigations of the site have to be performed. Since
they indicate a potential effect, EC 50 and LC 50 values instead of NOEC
(no-observed-effect concentration) or LOEC (lowest-observed-effect con-
centration) values are applied for the derivation.

■ Procedure

Chemical testing is described in detail in the different guidelines (mainly
from OECD) mentioned in the pertinent sections.

■ Notes and Points to Watch

• Control soils have to be selected carefully (ISO 15799 2004). For the
derivation of trigger values, natural soils are recommended, but at least
a sandy soil with low sorption capacity should be used. For higher
environmental relevance, loamy and silty soils should be employed. If
artificial soil is used and if the test chemical has a high logK ow value
(octanol-water partitioning coefficient; e.g., > 2; European Plant Protec-
tion Organization 2003) this test substrate should contain only 5% peat
instead of 10% in order to test a more field-relevant situation concerning
the bioavailability of the test substance.

348

A. Eisentraeger et al.

18.6

Combined Performance of Bioassays

and Assessment of the Results

18.6.1

Water-Extractable Ecotoxic Potential

The procedure proposed here, and based on only two bioassays, is a qual-
itative one that offers a way to quickly obtain results and keep costs down
(Fig. 18.2). Dilution values are defined to indicate ecotoxicological poten-
tial if exceeded. The water extracts should be diluted using a factor of 2,
and lowest ineffective dilution values (LID) should be assessed. The LID
is defined as the lowest dilution with less than 20% inhibition in the lu-
minescence algae test. For the qualitative evaluation it is not necessary to
determine EC values by data transformation from dose response curves. Of
course, it might be useful to determine EC values if toxic potentials of soil
samples (e.g., from the same site during remediation) have to be compared.
The V. fischeri luminescence inhibition assay (ISO 11348 1998; Sect.
18.3.1) should be performed at first. If the LID value exceeds 8, a risk of
pollutant leakage exists and it is recommended that the remediated soils
should not be incorporated at unsealed sites. If the luminescence inhibi-
tion assay is negative, the 72 h algae growth inhibition assay (ISO 8692

Vibrio fischeri luminescence

inhibition test

(ISO 11348)

Desmodesmus subspicatus

growth inhibition test (ISO 8692)

LID>8

LID>4

Risk of pollutant leakage is
given

No incorporation of

remediated soil at unsealed

sites

Vibrio fischeri luminescence

inhibition test

(ISO 11348)

and

Desmodesmus subspicatus

growth inhibition test (ISO 8692)

LID<8

LID<4

■=>

Risk of pollutant leakage is
low

Fig. 18.2. Procedure proposed for the assessment of the water extractable ecotoxicological
potential of soils and soil materials. The assessment based on LID values is allowed if
(1) a dose response relationship is obtained, or (2) nearly 100% inhibition is obtained in
several tested dilutions, or (3) no significant inhibition is obtained in the dilutions up to the
threshold value. (Maxam et al. 2000; Pfeifer et al. 2000; Dechema 2001; Rila and Eisentraeger
2003; Eisentraeger et al. 2004)

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 349

1989; Sect. 18.6.2) should be performed additionally. The risk of pollutant
leakage is low if this LID value is < 4 (or the LID value of the lumines-
cence inhibition assay is < 8). These threshold values are derived from
the experiences gained during the earlier-mentioned research projects
(Rila and Eisentraeger 2003) and from the results of a ring test (Hund-
Rinke et al. 2002a, b). Low inhibitions are obtained with uncontaminated
soil samples such as the natural standard soils LUFA 2.1, 2.2 and 2.3 (land-
wirtschaftliche Untersuchungs- und Forschungsanstalt, Speyer, Germany).
This "background toxicity" might be caused by humic substances.

Ecotoxic effects of a wide range of water-extractable contaminants can
be detected by using these two test systems. In a round robin test eight
contaminated soils were investigated using four aquatic test systems (lumi-
nescence and growth test with V.fischeri, tests with algae and daphnids). It
was shown that daphnids are mostly less sensitive than the tests with algae
and the luminescence test with V.fischeri. The daphnids test was more sensi-
tive, however, for soils contaminated with heavy metals (Hund-Rinke et al.
2002c). As heavy metals are routinely measured by chemical analyses, it
was decided to exclude the test with daphnids from the base set of aquatic
ecotoxicological test systems for soil assessment. The approach presented
here is cost effective: No range-finding test has to be carried out and the
algae growth inhibition test can be performed in microplates, so long as
the validity criteria of ISO 8692 are fulfilled (Eisentraeger et al. 2003). If
other or further testing is regarded as necessary, ecological relevance and
practicability should be considered.

18.6.2

Water-Extractable Genotoxicity

Cost effectiveness and speed are also major aspects of the assessment
scheme for water-extractable genotoxic potential. It should thus be noted
that this is a screening method that cannot be used to identify clastogenic
substances, but is able to roughly estimate whether genotoxic compounds
can be mobilized by water. The procedure is mainly based on the assessment
of the genotoxic potential of water extracts using the umu test according to
ISO 13829 (2000; Sect. 18.4.1). The umu test can be performed in less than
a day with and without metabolic activation. The Salmonella/microsome
test (Ames test) according to DIN 38415 T4 (Sect. 18.4.2) should be carried
out additionally if the umu test is negative and if there are strong hints from
chemical analysis or site history that mutagenic compounds are present.

In the first step of the procedure (Fig. 18.3) the same water extract is
tested as used for the assessment of the water-extractable ecotoxicological
potential. If there is a genotoxic effect in the umu test, with or without

350

A. Eisentraeger et al.

First step:

umu-test
optional:
Ames-test

D Li >3
LID>6

umu-test
optional:
Ames-test

D LI =1.5
LID = 3

Second step:

Investigation of coi

umu-test
optional:
Ames-test

D Li >3
LID>6

umu-test
optional:
Ames-test

Dii=1.5
LID = 3

Investigation of the soil elutriates

genotoxic

acute danger for path soil-
groundwater

not
genotoxic

Risk of acute danger for

path soil-groundwater

is low

genotoxic

acute danger for path
soil-groundwater

not
genotoxic

no danger for path
soil-groundwater

Fig. 18.3. Assessment of the water-extractable genotoxic potential of soils and soil materials
using the umu test according to ISO 13829. The Salmonella/microsome test (Ames test)
according to DIN 38415 T4 should be carried out if the umu test is negative and there are
strong hints from chemical analysis or site history that mutagenic compounds are present.
(Eisentraeger et al. 2000; Dechema 2001; Eisentraeger et al. 2001; Rila and Eisentraeger 2003;
Eisentraeger et al. 2004; modified according to Ehrlichmann et al. 2000)

metabolic activation, a high risk of transfer of genotoxic substances from
soil to the ground water exists. If there is no genotoxic effect, the water
extract should be concentrated by a (low) factor of 15 using Serdolit PAD-
1 resin. During the ring test mentioned above (Hund-Rinke et al. 2002a)
the water extracts were concentrated by a factor of 30, as performed by
Ehrlichmann et al. (2000). The factor was reduced to 15 on the basis of
results obtained during this test and further studies (Rila and Eisentraeger
2003), since several obviously uncontaminated soil samples (e.g., LUFA 2.1
and LUFA 2.2) tested positive after 30-fold concentration.

18.6.3

Assessment of the Habitat Function

Criteria for the combined assessment of the pathways from soil to soil
microorganisms, fauna, and higher plants were elaborated (Fig. 18.4). The

18 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays

351

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352 A. Eisentraeger et al.

test design was approved in a round robin test. For special requirements
it is possible to complement the basic test set by further tests. Test results
are interpreted using different strategies selected, depending on the test
system employed.

Soil Microflora - Respiration Activity. The respiration activity is assessed by
the ratio basal respiration:SIR, and by considering the 2 uptake or C0 2
production over time.

Soil Microflora - Ammonium Oxidation Activity. The nitrification activity of the
test soil is assessed by comparison with a control soil and a 1:1 mixture of
test soil and control soil. If the nitrification activity in the mixture is below
90% of the mean value of the activity in the test and control soil, the habitat
function is assessed as "disturbed" for this criterion.

Soil Fauna. Regarding soil fauna, a minimal habitat function is demanded.
The assessment is based on the comparison between the mortality rate
and the reproduction in the test soil and in the control. The habitat func-
tion is considered disturbed if the mortality rate surpasses 20% and the
reproduction rate falls below 50% compared to the control.

Soil Flora. To evaluate potential effects on the soil flora two test strategies
have been elaborated. For both strategies a control soil is needed. The first
strategy directly compares the biomass production in the test soil and in
the control soil. A second possibility is to compare the biomass production
in (1) the test soil, (2) a control soil, and (3) a 1:1 mixture of the test and
control soils. A biomass determined to be less than 70% in the test soil as
compared to the control or less than 90% in comparison to the mean value
of the mixed test and control soils is regarded as insufficient and the sample
is assessed "toxic".

Preferably, a control soil from the site should have the same physico-
chemical soil properties as the contaminated soil but no contamination.
However, in many cases such a soil is not available and it is then recom-
mended to use a sandy soil (e.g., LUFA standard soil 2.2) to avoid a high
sorption of contaminants (for more details see ISO 15799 2003). In cases
where the geographical or pedological typicality of the selected soil is
important, approaches like the EURO Soil concept (Kuhnt and Muntau
1992) or the German Refesol proposal can help to find appropriate control
soils.

The terrestrial tests were selected to give information on the habitat
function of the soil. If the habitat function of a soil is reduced, this may
result from anthropogenic contaminants (e.g., heavy metals, PAHs, TNT),
a high salt content caused by the addition of large amounts of organic
material (e.g., compost), or alow pH. Therefore, expert knowledge is needed
to decide whether a test is suitable for a specific soil or soil material.

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 353

Moreover, results indicating a toxic potential have to be critically examined
with respect to further decisions regarding the use of the test material.
If there seems to be a need to replace a test or to perform further tests,
ecological relevance and practicability should be considered. Under certain
circumstances, field monitoring approaches at the assessment site may be
appropriate (Roembke and Notenboom 2002).

18.6.4

Overall Assessment - Combined Strategy

In Fig. 18.5, a stepwise procedure for the combined evaluation of remediated
soil samples is given as an example for the cost effective application of these
bioassays.

1. In the first step it is determined by chemical analyses whether thresh-
old values for single contaminants are exceeded; these values are laid
down in national laws, decrees, or guidance papers (e.g., in Germany:
BBodSchV 1999). If a threshold value is exceeded, different possibilities
exist. Risk-reduction measures to decrease the contaminant levels may
be necessary, and/or the further use of this soil is restricted, because
the remediation goal for this soil has not been reached. It should be
evident that those soils which are clearly contaminated, where thresh-
old values are exceeded, do not have to be tested biologically at all. For
the other soils, in which such thresholds have not been exceeded, the
water- extractable ecotoxicological and genotoxic potential is tested. If
the threshold values of at least one bioassay are exceeded, the source
of the toxicity should be identified and appropriate measures taken.
If the test results do not indicate a risk for groundwater or surface
water, the remediated soil can, for example, be incorporated as sub-
soil.

2. If, depending on the envisaged use of the soil, the habitat function
of the soil has to be assessed, terrestrial ecotoxicological tests have
to be performed in a second step. Again, if the assessment criteria
are exceeded, the source of the toxicity should be identified and ap-
propriate measures taken. If the values are not exceeded, the habitat
function is substantiated and the remediated soil can be used as top-
soil.

In the overview thus far presented it has been shown that ecotoxicological
test systems are available for the assessment of the retention function and
for the habitat function of soils. In addition, the results of these tests
can be evaluated to determine whether the soil might cause a risk to the
environment. Finally, it should be noted that it may be necessary to modify

354

A. Eisentraeger et al.

Stepl

Threshold values for chemical

analysis of contaminants are

exceeded

yes

Limited use of remediated soil -
remediation target not reached

Determination of water extractable

ecotoxicity and genotoxicity:

threshold values exceeded

yes

Toxicity identification

Low danger of pollutant discharge or

hazard for groundwater - remediated

soil can be incorporated as subsoil

Step 2: (if testing of habitat function is necessary)

Ecotoxicological testing of habitat

function of soil: threshold values

exceeded

yes

Toxicity identification

Habitat function is given

Remediated soil can be incorporated
as topsoil

Fig. 18.5. Stepwise procedure for the examination of soils or soil materials using the test
systems of Tables 18.1-18.2 and Figs. 18.2-18.4 for remediated soil. (Dechema 2001; Eisen-
traeger et al. 2004)

the stepwise procedure presented here in specific cases. These modifications
might depend on the kind of sample to be tested, the kind of site, the
kind of contamination, the overall aim of the investigation (precautionary,
complementary to remediation, on-site analytics), and of course on the
money available.

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 355

References

Achazi R, Roembke J, Riepert F (2000) Collembolen als Testorganismen. In: Heiden S, Erb R,
Dott W, Eisentraeger A (eds) Toxikologische Beurteilung von Boden. Leistungsfahigkeit
biologischer Testverfahren. Spektrum Akademischer Verlag, Heidelberg, pp 83-103

Ames BN, McCann J, Yamasaki E (1975) Methods for detecting carcinogens and mutagens
with the Salmonella/mammalian-microsome mutagenicity test. Mutation Res 31:347-
364

Anderson JPE, Domsch KH (1978) A physiological method for the quantitative measurement
of microbial biomass in soils. Soil Biol Biochem 10:215-221

Anderson T-H, Domsch KH (1990) Metabolic process and soil microbial biomass. Soil Biol
Biochem 22:251-255

BBodSchG (Bundes-Bodenschutzgesetz) (1998) Gesetz zum Schutz vor schadlichen Boden-
veranderungen und zur Sanierung von Altlasten. Bundesgesetzblatt I, p 502 vom 17.
Marz 1998

BBodSchV (Bundes-Bodenschutzverordnung) (1999) Bundes-Bodenschutz- und Altlasten-
verordnung. Verordnung zur Durchfiihrung des Bundes-Bodenschutzgesetzes. BGBl I,
36, pp 1554- 582 vom 16.06.1999

Belser LW, Mays EL (1980) Specific inhibition of nitrite oxidation by chlorate and its use in
assessing nitrification in soils and sediments. Appl Environ Microbiol 39:505-510

Blagodatskaya EV, Anan'eva ND (1996) Assessment of the resistance of soil microbial com-
munities to pollutants. Eurasian Soil Sci 29:1251-1255

Blagodatskaya EV, Anan'eva ND, Myakshina TN (1996) Description of a soil microbe com-
munity in terms of metabolic quotient. Eurasian Soil Sci 28:86-95

Blakemore RJ (1997) Agronomic potential of earthworms in brigalow soils of south-east
Queensland. Soil Biol Biochem 29:603-608

Boutin C, Freemark KE, Keddy CJ (1995) Overview and rationale for developing regulatory
guidelines for nontarget plant testing with chemical pesticides. Environ Toxicol Chem
14:1465-1475

DECHEM A (200 1 ) Methods for toxicological/ecotoxicological Assessment of Soil. 7th Report
of the interdisciplinary DECHEMA committee "Environmental Biotechnology - Soil".
DECHEMA Deutsche Gesellschaft fuer Chemisches Apparatewesen, Chemische Technik
und Biotechnologie eV, Frankfurt am Main, Germany

DIN 38415 T4 (1998) Deutsche Einheitsverfahren zur Wasser-, Abwasser- und Schlam-
muntersuchung - Suborganismische Testverfahren (Gruppe T) - Teil 4: Bestimmung des
erbgutveraendernden Potentials mit dem Salmonella-Mikrosomen-Test (Ames-Test)
(T 4). VCH, Weinheim, Germany

Doube BM, Brown GC (1998) Life in a complex community: Functional interactions be-
tween earthworms, organic matter, microorganisms and plants. In: Edwards CA (ed)
Earthworm ecology. CRC Press, Boca Raton, pp 179-212

Edwards WM, Shipitalo MJ (1998) Consequences of earthworms in agricultural soils: Aggre-
gation and porosity. In: Edwards CA (ed) Earthworm ecology. CRC Press, Boca Raton,
pp 147-161

Ehrlichmann H, Dott W, Eisentraeger A (2000) Assessment of the water-extractable geno-
toxic potential of soil samples from contaminated sites. Ecotoxicol Environ Safety 46:73-
80

Eisentraeger A, Dott W, Klein J, Hahn S (2003) Comparative studies on algal toxicity testing
using fluorometric microplate and Erlenmeyer flask growth inhibition assays. Ecotoxicol
Environ Safety 54:346-354

Eisentraeger A, Pfeifer F, Dott W (2000): Erfassung des waeErig extrahierbaren genotox-
ischen Potentials kontaminierter Boeden. In: Heiden S, Erb R, Dott W, Eisentraeger A

356 A. Eisentraeger et al.

(Hrsg.): Toxikologische Beurteilung von Boeden - Leistungsfaehigkeit biologischer
Testverfahren. Spektrum Akademischer Verlag, Heidelberg, Berlin, pp 131-146

Eisentraeger A, Rila J-P, Ehrlichmann H, Dott W (2001): Erfassung des waeErig extrahier-
baren genotoxischen Potentials von Boeden und Bodenmaterialien mit optimierten
bakteriellen Genotoxizitaetstests. In: Neumann-Hensel H, Ahlf W, Wachendoerfer V
(eds) Nachweis von Umweltchemikalien - Auswerte- und Interpretationsmethoden
fuer Toxizitaetsdaten aus einer oekotoxikologischen Testkombination. (Initiativen zum
Umweltschutz Band 29). Erich Schmidt Verlag, Berlin, pp 47-60

Eisentraeger A, Rila J-P, Hund-Rinke K, Roembke J (2004) Proposal of a testing strategy and
assessment criteria for the ecotoxicological assessment of soil or soil materials. J Soil
Sediment 4:123-128

Engebrecht J, Nealson KH, Silverman M (1983) Bacterial bioluminescence: Isolation and
genetic analysis of functions from Vibrio fischeri. Cell 32:773-781

European Plant Protection Organization (2003) EPPO Standards. Environmental Risk As-
sessment scheme for plant protection products. Bull OEPP/EPPO 33:195-208

European Union (1991) Council directive concerning the placing of plant protection prod-
ucts on the market No. 91/414/EEC. Brussels, Belgium

European Union (1996) Technical guidance documents in support of Directive 93/67/EEC
on risk assessment of new notified substances and Regulation (EC) No. 1488/94 on risk
assessment of existing substances. Brussels, Belgium

Focht DD, Verstraete W (1977) Biochemical ecology of nitrification and denitrification. Adv
Microbiol Ecol 1:135-214

Hintze TH, Gehlen P, Schroder D (1994) Are microbial biomass estimations equally valid
with arable soils and forest soils? Soil Biol Biochem 26:1207-1211

Hopkin SP (1997) Biology of the springtails (Insecta: Collembola). Oxford Univ Press,
Oxford, pp 1-330

Hund K, Schenk B (1994) The microbial respiration quotient as indicator for bioremediation
processes. Chemosphere 28:477-490

Hund-Rinke K, Koerdel W, Hennecke D, Eisentraeger A, Heiden S (2002a) Bioassays for
the ecotoxicological and genotoxicological assessment of contaminated soils (results
of a round robin test). Part I Assessment of a possible groundwater contamination:
Ecotoxicological and genotoxicological tests with aqueous soil extracts. J Soil Sediment
1:43-50

Hund-Rinke K, Koerdel W, Hennecke D, Achazi R, Warnecke D, Wilke B-M, Winkel B,
Heiden S (2002b) Bioassays for the ecotoxicological and genotoxicological assessment
of contaminated soils (results of a round robin test). Part II Assessment of the habitat
function of soils. J Soil Sediment 2:83-90

Hund-Rinke K, Koerdel W, Heiden S, Erb R (eds; 2002c) Oekotoxikologische Testbatterien -
Ergebnisse eines DBU-gefoerderten Ringtests. Erich Schmidt Verlag, Berlin,

Insam H (1990) Are the soil microbial biomass and basal respiration governed by the
climatic regime? Soil Biol Biochem 22:525-532

Insam H, Domsch KH (1988) Relationship between soil organic carbon and microbial
biomass on chronosequences of reclamation sites. Microb Ecol 15:177-188

ISO 10381-6 (1993) Soil quality - Sampling - Part 6: Guidance on the collection, handling
and storage of soil for the assessment of aerobic microbial processes in the laboratory

ISO 11267 (1999) Soil quality- Inhibition of reproduction of collembola (Folsomi a Candida)
by soil pollutants

ISO 1 1268-1 (1993) Soil quality - Effects of pollutants on earthworms (Eisenia fetida) - Part
1: Determination of acute toxicity using artificial soil substrate

ISO 1 1268-2 (1998) Soil quality - Effects of pollutants on earthworms (Eisenia fetida) - Part
2: Determination of effects on reproduction

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 357

ISO 11269-2 (1995) Soil quality - Determination of the effects of pollutants on soil flora -

Part 2: Effects of chemicals on the emergence and growth of higher plants
ISO 11348 parts 1-3 (1998) Water quality - Determination of the inhibitory effect of water

samples on the light emission of Vibrio fischeri (Luminescent bacteria test)
ISO 11465 (1993) Soil quality - Determination of dry matter and water content on a mass

basis - gravimetric method
ISO 13829 (2000) Water quality - Determination of the genotoxicity of water and waste

water using the umu-test
ISO 14238 (1997) Soil quality - Biological methods - Determination of nitrogen mineral-
ization and nitrification in soils and the influence of chemicals on these processes
ISO 15685 (2004) Soil quality - Determination of potential nitrification and inhibition of

nitrification - Rapid test by ammonium oxidation
ISO 15799 (2003) Soil quality - Guidance on the Ecotoxicological characterization of soils

and soil materials
ISO 17155 (2002) Soil quality - determination of the activity of the soil microflora using

respiration curves
ISO 22030 (2004) Final Draft: Soil quality - Biological methods - Chronic toxicity in higher

plants
ISO 8692 (1989) Water quality - Fresh water algal growth inhibition test with Scenedesmus

subspicatus and Selenastrum capricornutum
ISO/DIS 21268-1 (2004) Soil quality - Leaching procedures for subsequent chemical and

ecotoxicological testing - Part 1: Batch test using a liquid to solid ration of 21 to

lkg
ISO/DIS 21268-2 (2004) Soil quality - Leaching procedures for subsequent chemical and

ecotoxicological testing - Part 2: Batch test using a liquid to solid ration of 10 1/kg dry

matter
Jessen-Hesse V, Terytze K, Roembke J, Jansch S, Eisentraeger A, Filzek P, Grabert E, Hiittner S,

Hund-Rinke K, Jander JP, Neumann-Hensel H (2003) Preparation and performance of

ecotoxicological test methods under practical conditions - a project overview and first

results. Mitt Deut Bodenkundl Ges 102:773-774
Jones A, Hart ADM (1998) Comparison of laboratory toxicity tests for pesticides with field

effects on earthworm populations: a review. In: Sheppard SC, Bembridge JD, Holm-

strup M, Posthuma L (eds) Advances in earthworm ecotoxicology. SETAC Press, Pen-

sacola, pp 247-267
Kandeler E, Kampichler C, Horak O (1996) Influence of heavy metals on the functional

diversity of soil microbial communities. Biol Fertil Soils 23:299-306
Kuhnt G, Muntau H (1992) EURO-Soils: Identification, collection, treatment, characteriza-
tion. Joint Research Center, European Commission, Ispra
Lavelle P, Bignell D, Lepage M (1997) Soil function in a changing world: The role of inver-
tebrate ecosystem engineers. Eur J Soil Biol 33:159-193
Lokke H, Gestel CAM (1998) Handbook of soil invertebrate toxicity tests. Wiley & Sons,

Chichester
Nordgren A, Baath E, Soderstrom B (1988) Evaluation of soil respiration characteristics to

assess heavy metal effects on soil microorganisms using glutamic acid as a substrate.

Soil Biol Biochem 220:949-954
OECD (Organisation for Economic Co-operation and Development) (1984) Guidelines for

testing chemicals No. 208: Terrestrial plants, growth test
Petersen H (1994) A review of collembolan ecology in ecosystem content. Acta Zool Fennica

195:11-118
Pfeifer F, Haake F, Koerdel W, Eisentraeger A (2000) Untersuchung der Rueckhaltefunk-

tion von Boeden mit aquatischen Testsystemen. In: Heiden S, Erb R, Dott W, Eisen-

358 A. Eisentraeger et al.

traeger A (eds) Toxikologische Beurteilung von Boeden - Leistungsfaehigkeit biolog-

ischer Testverfahren. Spektrum Akademischer Verlag, Heidelberg, pp 1-18
Riepert F, Felgentreu D (2000) Relevance of the soil storage to biomass development, N-

mineralization and microbial activity using the higher plant growth test. Appl Soil Ecol

20:57-68
Riepert F, Wilke B-M, Kalsch W, Winkel B (2000) Pflanzen als Testorganismen zur Charakter-

isierung der Lebensraumfunktion des Bodens als Pflanzensstandort. In: Heiden S, Erb R,

Dott W, Eisentraeger A (eds) Toxikologische Beurteilung von Boden. Leistungsfahigkeit

biologischer Testverfahren. Spektrum Akademischer Verlag, Heidelberg, pp 19-42
Rila J-P, Eisentraeger A (2003) Application of bioassays for risk characterization and re-
mediation control of soils polluted with nitroaromatics and PAHs. Water Air Soil Poll

148:223-242
Rila J-P, Hund-Rinke K, Pfeifer F, Dott W, Eisentraeger A (2003) Validation of mi-

croplate bioassays for the assessment of contaminated and remediated sites (laboratory-

intercomparison study). J Soil Sediments 3:273-283
Rila J-P, Maxam G, Eisentraeger A (2002) Kap. IV: Genotoxizitatstests. In: Hund-Rinke K,

Koerdel W, Heiden S, Erb R (eds) Okotoxikologische Testbatterien - Ergebnisse eines

DBU-geforderten Ringtests. Erich Schmidt Verlag, Berlin, pp 131-156
Roembke J (1997) Taxonomische Probleme in der terrestrischen Okotoxikologie (La-

borebene). Abh Ber Naturkundemus Gorlitz 69:33-44
Roembke J, Notenboom J (2002) Ecotoxicological approaches in the field. In: Sunahara G,

Renoux AY, Thellen C, Gaudet CL, Pilon A (eds) Environmental analysis of contaminated

sites. Wiley, Chichester, pp 181-195
Spurgeon, D, Svendsen C, Hankard P, Weeks J, Kille P, Fishwick S (2002) Review of sub-
lethal ecotoxicological tests for measuring harm in terrestrial ecosystems. Environment

Agency, Bristol
Stalder L, Pestemer W (1980) Availability to plants of herbicide residues in soil. Part I:

A rapid method for estimating potentially available residues of herbicides. Weed Res

20:341-347
Stenberg B, Johannson M, Pell M, Sjodahl-Svensson K, Stenstrom J, Torstensson L (1998)

Microbial biomass and activities in soil as affected by frozen and cold storage. Soil Biol

Biochem 30:393-402
Stenstrom J, Stenberg B, Johannson M (1998) Kinetics of substrate-induced respiration

(SIR). Theory. Ambio 27:35-39
Swift MJ, Heal OW, Anderson JM (1979) Decomposition in terrestrial ecosystems. Studies

in ecology, vol 5. Blackwell Sci Publ, pp 1-372
Urbanek J, Dolezal F (1992) Review of some case studies on the abundance and on the

hydraulic efficiency of earthworm channels in Czechoslovak soils, with reference to the

subsurface pipe drainage. Soil Biol Biochem 24:1563-1572
Wagner C, Lokke H (1991) Estimation of ecotoxicological protection levels from NOEC

toxicity data. Water Res 25:1237-1242
Wander M (2004) Soil organic matter fractions and their relevance to soil function. In:

Magdoff F, Weil R (eds) Advances in agroecology. CRC-Press, Boca Raton, pp 67-102
Weigmann G (1993) Zur Bedeutung von Bodenarthropoden fur die Funktion und die

Kennzeichnung von von Okosystemen. Mitt Deut Ges Allg Angew Entomol 8:479-490
Wiles JA, Krogh PH (1998) Tests with the collembolans Isotoma viridis, Folsomia Candida and

Folsomia fimetaria. In: Lokke H, van Gestel CAM (eds) Handbook of soil invertebrate

toxicity tests. Wiley, Chichester, pp 131-156
Wilke B-M, Hund-Rinke K, Pieper S, Roembke J, Marschner A (2004) Entwicklung von

Prufwertempfehlungen fur ausgewahlte Schadstoffe zum Schutz des Bodens als Leben-

sraum fur Bodenorganismen. UWSF - Z Umweltchem Okotox 16:145-214

1 8 Assessment of Ecotoxicity of Contaminated Soil Using Bioassays 359

Wilke B-M, Pieper S, Roembke J. (2001) Ableitung von Priifwerten fur den Wirkungspfad

Boden - Bodenorganismus. Bodenschutz 3:93-100
Winkel B, Wilke B-M (2000) Abschluftbericht des BMBF-Verbundvorhabens: Biologische

Verfahren zur Bodensanierung, Teilverbund 4.1: Praxiserprobung biologischer Testver-

fahren

Subject Index

AAS (atomic absorption spectrometry)

156, 166
Abiotic processes 142
Acclimation 189-197
Acenaphthene 110
Acenaphthylen 110
Acidity 69

Adenosine diphosphate (ADP) 297
Adenosine monophosphate (AMP) 297
Adenosine triphosphate (ATP) 297
Adenylate energy charge (AEC)

297-302
Adenylates 297

ADP (adenosine diphosphate) 297
AEC (adenylate energy charge)

297-302
Algal growth inhibition test 323, 331
Aliphatic compounds 100
Alkanes 103, 189, 274
Alkyl benzenes 104
Alkyl naphthalenes 104
Aluminium (Al) 116
Ames test 333
Ammonium 82, 86, 240, 337
Ammonium oxidation activity 322, 337
AMP (adenosine monophosphate) 297
Antagonistic effect 322
Anthracene 110
Antibodies 124
Antimony (Sb) 117
Apatite 87

Aromatic hydrocarbons 123, 184
Arsenic (As) 116
Atomic absorption spectrometry (AAS)

156, 166
ATP (adenosine triphosphate) 297
Auger 30

Autochthonous microorganisms 157
Avena sativa 344

Barium (Ba) 116

Benz(a)anthracene 110

Benzene 101

Benzene, toluene, ethylbenzene, xylene

(BTEX) 99, 123, 127, 182, 240, 313
Benzo(a)pyrene 110
Benzo(b)fluoranthene 110, 113
Benzo(ghi)perylene 110
Benzo(k)fluoranthene 110,113
Beryllium (Be) 116
Bioaugmentation 143, 144, 157
Biodegradation potential 132,189
Biodegradation rate 98
Bioleaching 155
Biological parameters 42, 189, 201, 261,

281,297,303,309,321
Bioluminescence 244, 330
Biomarker 256
BiomassC 281-288
Biomass N 289-293
Biomass, microbial 252,281-295
Bioreactor experiment 138
Bioremediation, hydrocarbons 131-153
Bioremediation, metals

155-159,161-177
Bioreporter 233-250

- systems 235-241

- MPN analysis 247

- single point measurement 241

- vapor phase sensing 244
Biosurfactant production 145
Bismuth (Bi) 116

Blastn 212

Boron (Bo) 116

Brassica juncea 168, 173

Brassica rapa 344

BTEX (benzene, toluene, ethylbenzene,

xylene) 99, 123, 127, 182, 240, 313
Bulk density 52

362

Subject Index

C:N ratio 149

C:P ratio 149

Cadmium (Cd) 116,240

Caesium (Cs) 116

Calcium (Ca) 116

Capillary GC-MS 251

Carbon allocation 190

Carbon analyzer 288

Carbon tetrachloride 101

Catabolic genotypes 208-214

Catechol-2,3-dioxygenase (xylE) 235

Cerium (Ce) 116

Chemical soil properties 71-93

Chemoautotrophic bacteria 303

Chlorobenzene 101

Chloroform 101,240

Chloroform fumigation 282

Chromium (Cr) 116,240

Chrysene 110

Clod method 57

Cloning 211-215,226,227

C0 2 evolution 334

14 C0 2 evolution 190

Cobalt (Co) 116,240

Collembola reproduction test 342

Colony forming unit (CFU) 273, 278

Cometabolic degradation 150

Community structure 209, 218, 254, 255

Composting 140

Composting bins 140

Copper (Cu) 116,240

Core method 52

Creosote 109

Crude oil 179-188,269,274

CTC (5-cyano-2,3-ditolyl tetrazolium

chloride) 265
Culture-dependent methods 201
Culture-independent methods 201
Cyano ditolyl tetrazolium chloride (CTC)

265
Cycloalkanes 104
Cyclone fermenters 135

2,4-D 240

DAPI (4,6-diamino-2-phenylindole) 265
Decane 105, 106
Dehydrogenase activity 316
Denaturing gradient gel electrophoresis
(DGGE) 209-224

Deschampsia caespitosa 173-175
Desmodesmus subspicatus 331
DGGE (denaturing gradient gel

electrophoresis) 209-224
Diamino phenylindole (DAPI) 265
Dibenzo(ah)anthracene 110
Dichlorobenzene 101
Dichloroethane 101
Dichloroethene 101
Dichloromethane 101
Dichlorophenol 240
Dichromate oxidation 284
Diesel 103, 179-188, 310, 313, 316
Digestion 116
Dilution 262, 269
DNA

- amplification 208
-polymerase 209,213

- purification 202, 206

- quality 207

- quantity 207

- sequence identification 216

- total community 202
16S rDNA

- amplification 210

- cloning and sequencing 211,214

- genotypes 208
Drilling 29

Dry combustion, carbon 72
Dry combustion, nitrogen 77
Dry mass 47

Earthworm mortality 340

Earthworm reproduction 340

Ecotoxicity 321-359

Eicosane 105

Eisenia andrei 341

Eisenia fetida 341

ELISA 121

EnSys immunoassay 123

Enumeration, soil microorganisms 261,

264,268,281-296
EnviroGard immunoassay 127-130
Enzyme activity 309-320
Epifluorescence 264
Esterase activity 310
Ethylbenzene 101
Excavation 32, 54

Subject Index

363

FastA 212,216
Fatty acids

- branched unsaturated 254

- mid-chain branched 254

- monounsaturated 254

- naming 257

- polyunsaturated 254

- profiles 251-259

- saturated 254

- terminally branched 254
Feasibilibility study

- bioleaching 155-159

- hydrocarbons 131-153

- phy to remediation 161-177
Fertilization 167

FID (flame ionization detector) 104

Field capacity 60

Field moist 48

FISH (fluorescent in situ hybridization)

265
Flame ionization detector (FID) 104
Florisil 104
Fluoranthene 110
Fluorene 110

Fluorescein diacetate hydrolysis 313
Folsomia Candida 343
Fumigation-extraction 281, 297
Functional genes 227
Fungi 155,156,275

/?-Galactosidase (lacZ) 235
Gallium (Ga) 116

Gas chromatography (GC) 100-111, 251
Gasoline 104, 179-188
Gasoline-specific compounds 182, 184
GC (gas chromatography) 100,107,111,

233
GC-MS 183

Gel electrophoresis 204, 206, 207
Genomics 226

Genotoxicity 322, 332-334, 349
Germanium (Ge) 116
Green fluorescent protein (GFP) 236
Growth-inhibition assay 331

Habitat function 322, 334, 350
Halogenated hydrocarbons 101
Headspace analyzer 100
Heating oil 103

Heavy metal: see metal
Heterotrophic leaching 155
High-performance liquid chromatography

(HPLC) 297
Hopane 179-188
HPLC (high-performance liquid

chromatography) 297
Humic acids 71, 203
Humification 72
Humus 71

Hydrindantin (ninhydrin) 289
Hydrocarbons

- biodegradation 179-199

- 14 C-labeled 189

- degrading bacteria 142

- mineralization 196

- monoaromatic 100, 189

- polycyclic aromatic 106, 109, 123, 127,
189,310,313,316

- substrates 191,268,275-277

- volatile 99, 184, 195, 268, 277
Hydrolases, hydrolytic activity 310,313

ICP-AES (inductively-coupled

plasma-atomic emission spectrometry)

156, 166
Immobilization 161, 171
Immunoassays 121-130

- detection limits 123

- ELISA 121

- EnSys 123

- EnviroGard 127-130

- RaPID 121-126
Indeno(l,2,3-cd)pyrene 110
Indium (In) 116
Inoculum 134, 157

INT (iodonitrotetrazolium chloride) 317

Internal markers 179

Internal standards 101, 105, 112, 116, 182

Iodonitrotetrazolium chloride (INT) 317

Iron(Fe) 116,240

Isoalkanes 104

Isoparaffins 184

Jet fuel 103,274

Kerosene 99
Kjeldahl method 79

364

Subject Index

fi-Lactamase {bid) 236

Land treatment 142

Lead(Pb) 116,240

Lipase activity 310

Liquid scintillation counter 191

Lithium (Li) 116

Loss on ignition (LOI) 72, 74

Lubrication oil 103

Luciferase AB (luxAB) 239

Luciferase CDABE (luxCDABE) 239

Luciferase, bacterial (lux) 238

Luciferase, insect (luc) 238

Luminescence inhibition assay 330

Magnesium (Mg) 116

Mangan (Mn) 116

Mass selective detector (MSD) 102, 1 1 1

Matric pressure 60

Maximum water-holding capacity 5 1

Membrane filter 265

Mercury (Hg) 116,240

Metagenomic libraries 226

Metal 115-118, 155-159, 161-177

- accumulation 168

- immobilization 155, 161

- leaching 155

- mobilization 155

- speciation 115
Metal-contaminated soil 155, 161
Methyl tert-butyl ether (MTBE) 99
Microarrays 227

Microbial activity 1 94, 309
Microbial community 209, 218, 254, 255
Microbial enrichment, selective 134
Microbial remediation 131-153, 155-159
Microscope, enumeration 264
Microtiter plate 270
Microtox 239
Microwave oven 116
Mineralization 72, 189
Mineralization potential 193
Mini-transposon 239
Molecular techniques 202-231
Monitoring 98, 233, 309

- biodegradation of hydrocarbons 142,
310,313,316

- biodegradation of carboxyl esters 310

- biodegradation of surfactants 316

- impact of contamination 321

- impact of fertilization 197,313

- of process 142

Monoaromatic hydrocarbons 100, 189
Most probable number (MPN) 268, 247
MPN (most probable number) 268, 247
MTBE (methyl tert-butyl ether) 99

Naphthalene 100, 110, 182, 240, 274
Nickel (Ni) 116,240
Ninhydrin (hydrintantin) 289
Ninhydrin-reactive nitrogen

289-291
Niobium (Nb) 116
Nitrate 82, 84, 240, 337
Nitrification 303, 337
Nitrite 82, 337
Nitrobacter 303
Nitrogen (N)

- immobilization 306

- inorganic 82

- mineralization 303-308

- ninhydrin-reactive 289-291

- organic 303

- total 76,292
Nitrosomonas 303
Nutrient sources 149, 156
Nutritional factors 148

O2 consumption 335
Oligonucleotide sequences 212
Organic acids 156, 158
Organic carbon 71
Organic matter 71
Organic nitrogen 303
Oven oxidation 288
Overlayer plate 274, 276
Oxido reductases 317

PAHs (polycyclic aromatic hydrocarbons)
106, 109, 123, 127, 189, 310, 313, 316

- alkylated 184

- deuterated 110

- native 110
Paraffins 184

PCBs (polychlorinated biphenyls) 127,
208, 240

Subject Index

365

PCR amplification 208-22 1

- catabolic genotypes 210,213

- control 214

- 16SrDNA 210

PCR cloning kits 210,215
PCR primers

- catabolic genes 212

- 16SrDNA 214

- universal 220

PCR thermocycler 210

Pentatriacontane 105

Percolation 156, 157

Percussion boring 28

Permeability 48, 52

Petroleum hydrocarbons 98, 123, 127,

137,139,142,184,189,310,316
pH value 68
Phenanthrene 1 10, 274
Phenol 240
Phosphorus (P)

- labile 90

- quantification 91, 116

- total 88

Physical soil properties 47-71

Phytoextraction 161, 163, 168, 172

Phytoremediation 161

Phytostabilization 161, 171-175

Plant growth test 344

Plants 161

Plate count 272

Platinum catalyzer 288

PLFA (polar lipid fatty acids) 251, 253

Polar lipid fatty acids (PLFA) 251, 253

Polychlorinated biphenyls (PCBs) 127,

208, 240
Polycyclic aromatic hydrocarbons (PAHs)

106, 109, 123, 127, 189, 310, 313, 316
Polyvinylpolypyrrolidone (PVPP) 206
Pore size distribution 59
Pore water pressure 50
Porosity 52
Potassium (K) 116
Pour plate 276
Pre-extraction 284, 293
Pre-incubation 298
Pressure plate extractor 65
Primer design 212

Priority pollutants 181

Proton activity 68

Purge and trap 182

PVPP (polyvinylpolypyrrolidone) 206

Pyrene 110

Radioactivity 198

Radiolabeled substrates 269

Radiorespirometry 189

RaPID Immunoassay 121-126

Redox titration 284

Reporter systems 235

Respiration activity 193

Respiration curve test 334

Respirometer 136

Retention time 102

Rhenium (Re) 116

Ribosomal Database Project (RDP) 217

Rubidium (Rb) 116

Salmonella typhimurium 333
Salmonella/microsome test (Ames test)

323, 333
Scale up 141
Scintillation cocktail 192
Segmented flow analysis (SFA) 85
Selected ion monitoring (SIM) 184
Selective microbial enrichment 134
Selenium (Se) 116
Sequencing 211-215
Serial dilution 262, 263, 270
Sewage sludge 140
SFA (segmented flow analysis) 85
Shotgun cloning 226, 227
Silicon (Si) 116
Silver (Ag) 116,240
SIM (selected ion monitoring) 184
Slurry bioreactors 137
Sodium (Na) 116
Soil

- aeration 52

- aggregates 264

- biological activity 309

- column 134

- DNA: see DNA

- enzymes 309

- microcosm 136

- nucleic acid extraction 202, 205

366

Subject Index

- nutrients 76

- pores 59

- quality 2

Soil sampling 1-37, 132-134

- along a linear source 21

- circular grids 15

- container 34

- equipment 133

- irregular sampling 12

- methods 25-37

- non- systematic patterns 12

- pretreatment 37-40

- quantity 24

- random sampling 17

- rectangular grid 20

- regular grids 16

- sample type 25

- sampler 32

- sampling points 10

- sampling site 22

- strategy 7-25

- stratified random sampling 17

- systematic sampling 16

- unaligned random sampling 19

- undisturbed samples 27
Soil storage 41-44

Soil water characteristics 62, 65
Solid phase extraction 329
Solid-phase microextraction (SPME)

143
Solvent extraction 183
SOS Chromotest 235
SPME (solid-phase microextraction)
Spore suspension 158
Spray plate 276
Spread plate 275
Sterile control 135
Streamline test 166
Stripping apparatus 192
Strontium (Sr) 116
Styrene 101, 182
Suction table 62
Sulfur (S) 116
Surfactants 144-147
Synergistic effect 322

143

Tellurium (Te) 116

Tert-amyl methyl ether (TAME) 99

Tetrachloroethene 101

Tetracontane 105, 106

Tetramethylbutane 179

Threshold values 346

Time domain reflectometry (TDR) 48

Titanium (Ti) 116

Toluene 101,274

Toxicity 321-359

TPHs (total petroleum hydrocarbons) 98,

127, 137, 139, 142, 184, 189, 310, 316
Transposon 239
Treatability study 162-166

- bioleaching 155-159

- hydrocarbons 131-153

- phy to remediation 161-177
Triacontane 105
Trichlorobenzene 101
Trichloroethane 101
Trichlorophenol 240
Trimethyl cyclopentane 179
Trimethyl phenanthrene 179
Tungsten (W) 116

Umu test 322, 332

Uranium (U) 116

Urogen III methyltransferase (UMT)

UV-persulfate oxidation 286

237

Vanadium (V) 116
Vapor phase contaminants 244
Vapor plate 277
Vibrio fischeri 323, 330
Vitotox 239

Volatile hydrocarbons 99, 184, 195, 268,
277

Water content 47
Water content adjustment
Water extraction 329
Water retention 59
Water-holding capacity 50
Wilting point 60

Xylene 101,182,240,274

TAME (tert-amyl methyl ether) 99
Tantalum (Ta) 116

Zinc(Zn) 116,240
Zirconium (Zr) 116