Methods of Soil Analysis, Part 2: Microbiological and Biochemical Properties: 12 (SSSA Book Series) [1 ed.] 089118810X, 9780891188100

Citation preview

Published 1994

METHODS OF SOIL ANALYSIS PART 2 Microbiological and Biochemical Properties

Soil Science Society of America Book Series Books in the series are available from the Soil Science Society of America, 677 South Segoe Road, Madison, WI 53711 USA. 1. MINERALS IN SOIL ENVIRONMENTS. Second Edition. 1989. J. B. Dixon and S. B. Weed, editors R. C. Dinauer, managing editor 2. PESTICIDES IN THE SOIL ENVIRONMENT: PROCESSES, IMPACTS, AND MODELING. 1990. H. H. Cheng, editor S. H. Mickelson, managing editor 3. SOIL TESTING AND PLANT ANALYSIS. Third Edition. 1990. R. L. Westerman, editor S. H. Mickelson, managing editor 4. MICRONUTRIENTS IN AGRICULTURE. Second Edition. 1991. J. J. Mortvedt et aI., editors S. H. Mickelson, managing editor 5. METHODS OF SOIL ANALYSIS. Part 2. Microbiological and Biochemical Properties. 1994. R. W. Weaver et aI., editors S. H. Mickelson, managing editor

Methods of Soil Analysis Part 2 Microbiological and Biochemical Properties Editorial Committee: R. W. Weaver, chair Scott Angle Peter Bottomley David Bezdicek Scott Smith Ali Tabatabai Art Wollum

Managing Editor: S. H. Mickelson

Editor-in-Chief SSSA: J. M. Bigham

Number 5 in the Soil Science Society of America Book Series Published by: Soil Science Society of America, Inc.

1994

Copyright © 1994 by the Soil Science Society of America, Inc. ALL RIGHTS RESERVED UNDER THE U.S. COPYRIGHT LAW OF 1976 (P.L. 94-553) Any and all uses beyond the "fair use" provision of the law require written permission from the publishers and/or author(s); not applicable to contributions prepared by officers or employees of the U.S. Government as part of their official duties.

Soil Science Society of America, Inc. 677 South Segoe Road, Madison, Wisconsin 53711 USA

Library of Congress Cataloging-in-Publication Data

Methods of soil analysis. Part 2, Microbiological and biochemical properties / editorial committee, R.W. Weaver, chair ... let al.l. p. cm. - (Soil Science Society of America book series; no. 5)

Includes bibliographical references and index. ISBN 0-89118-810-X 1. Soil microbiology-Methodology. 2. Soils-Analysis. I. Weaver, R. W. (Richard W.), 1944II. Soil Science Society of America. III. Series. QRll1.M34 1994 631.4'17'0287 -dc20 94-20752 CIP

Printed in the USA

CONTENTS FOREWORD ...................................................................... . PREFACE ........................................................................... . CONTRIBUTORS .............................................................. . CONVERSION FACTORS FOR SI AND NON-SI UNITS .............. .

Chapter 1 Soil Sampling for Microbiological Analysis

XV XVll XIX XXlll

1 A. G. WOLLUM. II

1-1 Principles ........................................................................... 1-2 Methods ............................................................................ 1-3 Sources of Error ................................................................... 1-4 Concluding Remarks .............................................................. REFERENCES .........................................................................

Chapter 2 Statistical Treatment of Microbial Data

2 8 12 13 13

15

TIMOTHY B. PARKIN AND JOSEPH A. ROBINSON 2-1 Characteristics of the Lognormal Distribution................................. 2-2 Diagnosing Lognormality ......................................................... 2-3 Estimating Population Parameters from Sample Data...... ............... .. . 2-4 Selecting the Appropriate Location Parameter ................................ 2-5 Hypothesis Testing ................................................................ 2-6 Sample Number Requirements .................................................. 2- 7 Concluding Remarks .............................................................. APPENDIX 1 ........................................................................... APPENDIX 2 ........................................................................... REFERENCES .........................................................................

Chapter 3 Soil Sterilization

16 17 21 25 28 31 32 34 37 38

41 DUANE C. WOLF AND HORACE D. SKIPPPER

3-1 Principles ........................................................................... 3-2 Moist Heat ......................................................................... 3-3 Dry Heat ...... " .......... " ............ "......................................... 3-4 Gamma Irradiation ................................................................ 3-5 Microwave Irradiation ............................................................ 3-6 Gaseous Compounds .............................................................. 3-7 Nongaseous Compounds .......................................................... 3-8 Conclusions ......................................................................... REFERENCES .........................................................................

Chapter 4 Soil Water Potential

41 42 43 44 45 45 47 49 49

53

K. J. McINNES. R. W. WEAVER, AND M. 1. SAVAGE

4-1 Principles ........................................................................... 4-2 Materials............................................................................ 4-3 Procedure........................................................................... 4-4 Comments .......................................................................... REFERENCES ......................................................................... v

54 55 55 57 57

CONTENTS

vi

Chapter 5 Most Probable Number Counts

59 PAUL L. WOOMER

5-1 Principles ........................................................................... 5-2 Methodology.......... ..... .... ........... ...... .......... ........ ...... ........... 5-3 Comments .......................................................................... ACKNOWLEDGMENT .............................................................. REFERENCES ............................ - _ _........................................

Chapter 6 Light Microscopic Methods for Studying Soil Microorganisms

60 65 77 78 78

81

PETER J. BOTTOMLEY 6-1 Sampling of Soil for Microscopic Observation ................................. 6-2 Microscopic Enumeration of Total Bacteria in Soil........................... 6-3 Determining the Proportion of Viable Soil Bacteria Using a Cell Elongation Assay .................................................................... 6-4 Determining the Proportion of Viable Soil Bacteria by Following the Reduction of Tetrazolium Dyes to Formazan ................................ 6-5 Microscopic Determination of the Mycelial Length of Soil Fungi .......... 6-6 Determining the Proportion of Metabolically Active Fungal Mycelia by Following the Hydrolysis of Fluorescein Diacetate .......................... 6-7 Determining the Proportion of Metabolically Active Fungal Mycelia by Following the Reduction of Tetrazolium Dyes to Formazan ............... 6-8 Determining the Weight of Soil Biomass from Microscopic Estimates of Biovolume ......................................................................... 6-9 Microscopic Enumeration of Bacteria with Fluorescently Labeled Oligonucleotides Directed at Specific Regions of 16S Ribosomal RNA ...... ACKNOWLEDGMENT .............................................................. REFERENCES .........................................................................

Chapter 7 Viruses

81 82 92 94 96 98 99 100 101 104 104

107 J. SCOTT ANGLE

7-1 General Principles of Analysis ..... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7-2 Phages .............................................................................. 7-3 Enteric Viruses .................................................................... REFERENCES .........................................................................

Chapter 8 Recovery and Enumeration of Viable Bacteria

109 110 114 115

119

DAVID A. ZUBERER 8-1 Principles of Enumerating Soil Bacteria ........................................ 8-2 Materials and Equipment......................................................... 8-3 Collection and Preparation of Soil Samples.................................... 8-4 Release of Bacteria from Soils ................................................... 8-5 Diluents Used in Recovery and Enumeration of Soil Bacteria .............. 8-6 Preparation of Serial Dilutions .................................................. 8-7 Plating Techniques ......... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8-8 Media for Enumeration of Soil Bacteria ....................................... 8-9 Analysis and Presentation of Plate Count Data ............................... 8-10 Conclusion ... ..................... .... ..... .... ......... ..... .... ........ .... ..... REFERENCES .........................................................................

120 122 123 124 125 127 130 134 139 141 142

CONTENTS

vii

Chapter 9 Coliform Bacteria

145 RONALD F. TURCO

9-1 Recovery and Enumeration of Fecal Coliforms from Soil ................... 9-2 Detection and Enumeration of Total Coliforms ............................... 9-3 Rapid Test for Detection of E. coli in Soil..................................... 9-4 Direct Methods for Detection of E. coli in Soil ............................... ACKNOWLEDGMENT .............................................................. REFERENCES .........................................................................

Chapter 10 Autotrophic Nitrifying Bacteria

147 151 154 156 156 157

159

EDWIN L. SCHMIDT AND L. W. BELSER 10-1 Enumeration by Most Probable Number ..................................... 10-2 Diversity of Nitrifiers ............................................................ 10-3 Immunofluorescence Examination.. .......... .................. ........ ....... 10-4 Isolation of Nitrifiers ............................................................. 10-5 Maintenance of Pure Cultures.................................................. 10-6 Nitrifying Activity in Soils ............................ .. .. .. .. .. .. .. .. .. .. .. .. .. REFERENCES .........................................................................

Chapter 11 Free-living Dinitrogen-fixing Bacteria

160 163 165 167 169 171 176

179

ROGER KNOWLES AND WlLFREDO LASERNA BARRAQUIO 11-1 The Acetylene Reduction Assay.................................... ........... 11-2 Methods for Dinitrogen Fixers in General.................................... 11-3 Methods for Azotobacteraceae ................................................. 11-4 Method for Methanotrophs ..................................................... 11-5 Method for Hydrogen-Using Dinitrogen Fixers.............................. 11-6 Method for Cyanobacteria ...................................................... 11-7 Method for Photosynthetic Purple Nonsulfur Bacteria ...................... 11-8 Method for Clostridia.. ...................................... .................... 11-9 Method for Sulfate-Reducing Bacteria ........................................ REFERENCES . ..... ...... ...... ............. ............................. .......... ..

Chapter 12 Legume Nodule Symbionts

180 181 183 186 188 189 191 192 194 195

199

R.W. WEAVER AND PETER H. GRAHAM 12-1 Nodule Collection and the Isolation of Symbionts ........................... 12-2 Cultivation of Nodule Symbionts .............................................. 12-3 Maintenance of Cultures ........................................................ 12-4 Enumeration of Nodule Symbionts in Soil and Inoculants ................. 12-5 Inoculants for Field Experimentation ......................................... 12-6 Inoculation of Seed .............................................................. 12-7 Field Experimentation Involving Inoculation ................................. 12-8 Growth-Pouch Infection Assays ................................................ REFERENCES .........................................................................

200 203 205 206 210 213 215 216 218

CONTENTS

viii

Chapter 13 Anaerobic Bacteria and Processes

223

HEINRICH F. KASPAR AND JAMES M. TIEDJE 13-1 Principles .......................................................................... 13-2 Methods for Removal of Oxygen .............................................. 13-3 Methods for Reduction of Media .............................................. 13-4 Redox Indicators ................................................................. 13-5 Culture Methods ................................................................. 13-6 Enumeration Methods ........................................................... 13-7 Simple Method to Carry Out Anaerobic Incubations of Soil ............... ACKNOWLEDGEMENTS ........................................................... REFERENCES .........................................................................

Chapter 14 Denitrifiers

224 226 229 231 233 235 240 241 242

245 JAMES M. TIEDJE

14-1 Nitrate Reducing Processes ..................................................... 14-2 Key Physiological and Ecological Features of Respiratory Denitrifiers... 14-3 Enumeration of Denitrifiers .................................................... 14-4 Enumeration of Dissimilatory Nitrate to Ammonium Reducers ........... 14-5 Denitrifier Enzyme Activity .................................................... 14-6 Isolation of Denitrifiers .......................................................... 14-7 Confirmation of Respiratory Denitrification .................................. 14-8 Taxonomic Identification ........................................................ ACKNOWLEDGMENT .............................................................. REFERENCES .........................................................................

Chapter 15 Actinomycetes

245 250 251 255 256 257 259 265 265 265

269 E. M. H. WELLINGTON AND I. K. TOTH

15-1 Enumeration, Enrichment, and Isolation ..................................... 15-2 Isolation of Physiological Groups .............................................. 15-3 Grouping and Identification of Actinomycetes ............................... REFERENCES .........................................................................

Chapter 16 Frankia and the Actinorhizal Symbiosis

270 279 284 287

291 DAVID D. MYROLD

16-1 Characteristics of Frankia ....................................................... 16-2 Isolation, Culturing, and Maintenance of Frankia Strains ........ .......... 16-3 Quantification and Differentiation of Frankia Strains ....................... 16-4 Characterization of Frankia in Symbiosis ..................................... 16-5 Quantification in Soil ............................................................ 16-6 Conclusion ........................................................................ ACKNOWLEDGMENTS ............................................................. REFERENCES .........................................................................

Chapter 17 Filamentous Fungi

291 294 305 309 316 320 320 322

329 DENNIS PARKINSON

17-1 Qualitative Studies: Isolation Methods.................... .............. ...... 17-2 Quantitative Methods............................................................ REFERENCES .........................................................................

330 342 347

CONTENTS

ix

Chapter 18 Vesicular-Arbuscular Mycorrhizal Fungi

351 DAVID M. SYLVIA

18-1 Quantification of Vesicular-Arbuscular Mycorrhizal Propagules in Soil.. 18-2 Quantification of Vesicular-Arbuscular Mycorrhizal Colonization in Roots ............................................................................... 18-3 Quantification of Vesicular-Arbuscular Mycorrhizal External Hyphae .... 18-4 Recovery of Vesicular-Arbuscular Mycorrhizal Fungal Spores............. 18-5 Identification of Vesicular-Arbuscular Mycorrhizal Fungi .................. 18-6 Assessment of Growth Response and Selection of Effective Isolates ..... 18-7 Production and Use of Vesicular-Arbuscular Mycorrhizal Inocula ........ 18-8 Monoxenic Cultures for Basic Research ...................................... REFERENCES .........................................................................

Chapter 19 Isolation of Microorganisms Producing Antibiotics

352 353 357 360 360 361 366 372 372

379

JEFFRY J. FUHRMANN 19-1 General Principles ........ , ................ .............. ............... .. ..... ... 19-2 Microbiological Media ................... ..... ... ................ ..... ....... .... 19-3 Preparation of Inocula........................................................... 19-4 Dual Culture Detection Methods .............................................. 19-5 Culture Filtrate Methods........................................................ 19-6 Screening Methods ............................................................... 19-7 Methods for Selected Classes of Compounds ................................. 19-8 Concluding Comments .......................................................... REFERENCES .........................................................................

Chapter 20 Microbiological Procedures for Biodegradation Research

380 382 384 385 389 393 396 402 403

407

DENNIS D. FOCHT 20-1 The Enrichment Culture ........................................................ 20-2 Isolation of Pure Cultures....................................................... 20-3 Maintenance of Cultures ........................................................ 20-4 Growth in Liquid Cultures .......... ........ .......... .................... ...... 20-5 Preparation of Washed Cell Suspensions...................................... 20-6 Preparation and Use of Cell-Free Extracts ................................... 20-7 Oxygen Consumption ............................................................ 20-8 Chloride Determination ......................................................... 20-9 Conclusion ........................................................................ REFERENCES .........................................................................

Chapter 21 Algae and Cyanobacteria

409 412 413 415 418 419 421 423 424 424

427 F. BLAINE METTING, JR.

21-1 Identification of Soil Algae and Cyanobacteria .............................. 21-2 Direct Methods for Enumeration.. ............................................ 21-3 Indirect Methods for Enumeration ............................................ 21-4 Methods for Isolation and Purification of Microalgal Cultures ............. 21-5 Methods for Growth and Storage of Microalgal Cultures .................. 21-6 Methods for Estimating Photosynthesis ....................................... 21-7 Methods for Measuring Cyanobacterial Dinitrogen Fixation ............... 21-8 Methods for Studying Endosymbiotic Cyanobacteria in Cycad Roots ... . ACKNOWLEDGMENT .............................................................. REFERENCES .........................................................................

428 432 440 444 447 453 454 454 455 456

CONTENTS

x

459

Chapter 22 Nematodes RUSSELL E. INGHAM 22-1 Nematode Sampling ............................................................. 22-2 Extraction of Nematodes from Soil ............................................ 22-3 Extraction of Nematodes from Plant Material ............................... 22-4 Microscopic Observation and Identification of Nematodes ................. 22-5 Nematode Identification ......................................................... REFERENCES .........................................................................

Chapter 23 Protozoa

461 469 477 479 482 487

491 ELAINE R. INGHAM

23-1 First Consideratons .............................................................. 23-2 Protozoan Ecology ............................................................... 23-3 Methods of Enumeration........................................................ 23-4 Identification ...................................................................... 23-5 Summary .......................................................................... ACKNOWLEDGMENT .............................................................. REFERENCES .........................................................................

(:hapter 24 Arthropods

491 494 498 509 511 511 512

517 ANDREW R. MOLDENKE

24-1 Principles.......................................................................... 24-2 Methods ........................................................................... 24-3 Processing the Extracted Biota Sample ....................................... 24-4 Biota Identification .............................................................. 24-5 Preservation and Archiving ..................................................... 24-6 Archiving .......................................................................... 24-7 Rearing ............................................................................ 24-8 Statistical Methods to Analyze Diversity ...................................... REFERENCES .........................................................................

518 518 528 531 532 533 535 538 539

Chapter 25 Carbon Utilization and Fatty Acid Profiles for Characterization of Bacteria 543 A. C. KENNEDY 25-1 Characterization of Bacteria .................................................... 25-2 Carbon Source Utilization ....................................................... 25-3 Fatty Acid Analysis .............................................................. REFERENCES .........................................................................

Chapter 26 Multilocus Enzyme Electrophoresis Methods for the Analysis of Bacterial Population Genetic Structure

544 544 551 553

557

B.D. EARDLY 26-1 Principles .......................................................................... 26-2 Methods ........................................................................... REFERENCES .........................................................................

559 562 572

CONTENTS

xi

Chapter 27 Spontaneous and Intrinsic Antibiotic Resistance Markers

575

CHARLES HAGEDORN 27-1 Selection of Spontaneous Antibiotic Resistant Strains. .... ... ... ........... 27-2 Selection of Intrinsic Antiobiotic Resistant Strains .......................... 27-3 Evaluation of SAR and IAR Strains .................................. REFERENCES .........................................................................

Chapter 28

Serology and Conjugation of Antibodies

577 581 585 589

593 S. F. WRIGHT

28-1 Antibodies ........................................................................ 28-2 Adsorption of Cross-Reactive Antibodies from Polycional Antiserum.... 28-3 Purification of Antibodies fromAntiserum .................................... 28-4 Conjugation of Antibodies with Fluorescein Isothyocyanate ............... 28-5 Storage of Antibodies and Antibody Conjugates ............................ 28-6 Immunoassays .................................................................... REFERENCES .........................................................................

Chapter 29 Whole-Cell Protein Profiles of Soil Bacteria by Gel Electrophoresis

594 601 602 604 606 606 616

619

DIPANKAR SEN 29-1 Principles .......................................................................... 29-2 Methods ........................................................................... 29-3 General Comments .............................................................. REFERENCES .........................................................................

Chapter 30 Plasmid Profiles

620 621 630 633

635 IAN L. PEPPER

30-1 Plasmid Chromosome Relationships........................................... 30-2 Plasmid Profile Analysis......................................................... 30-3 Applications of Plasmid Profile Analyses for Soil Bacteria ................. REFERENCES .........................................................................

Chapter 31

DNA Fingerprinting and Restriction Fragment Length Polymorphism Analysis

635 638 644 644

647

M. J. SADOWSKY 31-1 DNA Fingerprinting............ ... ..... .......... ........ ... ... ................. 31-2 RFLP Analyses..................... ... ....... ... ............................. .... REFERENCES .........................................................................

Chapter 32 Nucleic Acid Probes

648 656 662

665 A. V. OGRAM AND D. F. BEZDICEK

32-1 Probe Selection ................................................................... 32-2 Isolation and Purification of Fragments to be Used as Probes ............. 32-3 Labeling of Probes ...............................................................

667 669 671

CONTENTS

xii

32-4 Hybridization ..................................................................... 32-5 Detection Systems................................................................ REFERENCES .........................................................................

Chapter 33 Marking Soil Bacteria with lacZY

679 684 686

689

T. E. STALEY AND D. J. DRAHOS 33-1 Principles .......................................................................... 33-2 Materials .......................................................................... 33-3 Procedure ......................................................................... 33-4 Exconjugant-Type Isolate Vigor ............................................... 33-5 Marker Stability .................................................................. 33-6 Recovery from Nonsterile Soil ................................................. 33-7 Attributes and Deficiencies ..................................................... 33-8 Comments ......................................................................... REFERENCES .........................................................................

Chapter 34 Detection of Specific DNA Sequences in Environmental Samples via Polymerase Chain Reaction

691 693 696 700 701 702 703 704 705

707

IAN L. PEPPER AND SURESH D. PILLA! 34-1 Theory ............................................................................. 34-2 Primer Design and Amplification Protocol ................................... 34-3 Optimization of Amplification .................................................. 34-4 Identification of Amplified Products ........................................... 34-5 Quality Control .................................................. ,................ 34-6 Specificity of Amplification ..................................................... 34-7 Sensitivity of Amplification ..................................................... 34-8 Applications in Environmental Microbiology ................................. REFERENCES .........................................................................

Chapter 35 Isolation and Purification of Bacterial DNA from Soil

709 710 714 717 717 719 720 721 725

727

WILLIAM E. HOLBEN 35-1 General Considerations ............................................... . . . .. .. . . . 35-2 Bacterial Fractionation Approach for Recovery of Bacterial Community DNA .................................................................... 35-3 Direct Lysis Approach for the Recovery of Total Bacterial Community DNA .................................................................... 35-4 Bacterial Fractionation Protocol ..................... . .. . . . .. .. . .. .. .. .. . .. .. . . 35-5 Direct Lysis Protocol ............................................................ 35-6 Fractionation of DNA Gradients, Final Purification and Quantitation of Bacterial Community DNA ..................................................... ACKNOWLEDGMENTS ............................................................. REFERENCES .........................................................................

Chapter 36 Microbial Biomass

729 731 733 736 741 743 750 750

753 W. R. HORWATH AND E. A. PAUL

36-1 Soil Sampling, Preparation, and Storage ...................................... 36-2 Physiological Methods ...........................................................

754 754

CONTENTS

xiii

36-3 Chemical Methods ............................................................... 36-4 Comparison of Methods ......................................................... REFERENCES .........................................................................

Chapter 37 Soil Enzymes

763 770 771

775 M. A. TABATABAI

37-1 Principles.......................................................................... 37-2 Factors Affecting Rates of Enzyme Reaction ................................ 37-3 Assay of Enzymes in Soil ....................................................... REFERENCES .........................................................................

Chapter 38 Carbon Mineralization

778 778 790 826

835 L. M. ZIBILSKE

38-1 General Principles ................................................................ 38-2 Experimental Principles ......................................................... 38-3 Field Methods .. ............... .............................. ............... ...... 38-4 Laboratory Methods ............................................................. REFERENCES .........................................................................

Chapter 39 Isotopic Methods for the Study of Soil Organic Matter Dynamics

836 836 841 850 859

865

DUANE C. WOLF, J. O. LEGG, AND THOMAS W. BOUTTON 39-1 Decomposition of 14C-Labeled Organic Matter in Soils .................... 39-2 13C Natural Abundance Technique: Background and Principles ........... 39-3 Decomposition of 15N-Labeled Organic Matter in Soils .................... 39-4 Extraction of Labeled Organic Fractions in Studies of Soil Organic Matter Dynamics ..................................................................... 39-5 Conclusions ....................................................................... REFERENCES .........................................................................

Chapter 40 Practical Considerations in the Use of Nitrogen Tracers in Agricultural and Environmental Research

866 875 887 893 900 901

907

R. D. HAUCK, J. J. MEISINGER, AND R. L. MULVANEY 40-1 Preparing 15N-Labeled Materials............................................... 40-2 Field Study Techniques .......................................................... 40-3 Preparing for and Measuring Nitrogen-Isotope Ratio....................... 40-4 Sources of Nitrogen-15 Supply and Analytical Service ...................... REFERENCES .........................................................................

Chapter 41

Nitrogen Availability Indices

909 919 935 942 943

951

L. G. BUNDY AND J. J. MEISINGER 41-1 Current Status of Nitrogen Availability Indices ................... ..... ...... 41-2 Methods ........................................................................... REFERENCES .........................................................................

951 955 979

CONTENTS

xiv

Chapter 42 Nitrogen Mineralization, Immobilization, and Nitrification

985

STEPHEN C. HART, JOHNM. STARK, ERIC A. DAVIDSON, AND MARY K. FIRESTONE 42-1 Measurement of Gross Nitrogen-Transformation Rates .......... .... ....... 42-2 Field Methods for Estimating Net Rates of Nitrogen Transformations ... 42-3 Laboratory Methods for Estimating Net Nitrogen Transformation Rates ................................................................................... 42-4 Laboratory Methods for Assessing Nitrification .............................. 42-5 Coda ............................................................................... REFERENCES .........................................................................

Chapter 43 Dinitrogen Fixation

987 999 1006 1009 1015 1016

1019 R. W. WEAVER AND SETH K. A. DANSO

43-1 Acetylene Reduction ............................................................ 32-2 Nitrogen Difference.............................................................. 43-3 Nitrogen-15 Isotope Techniques ................................................ 43-4 Use of Dinitrogen-15 Gas ...... .............................. ................... REFERENCES .........................................................................

Chapter 44 Measuring Denitrification in the Field

1019 1025 1030 1038 1043

1047

A. R. MOSIER AND LEIF KLEMEDTSSON

44-1 Methods 44-2 Experimental Protocols ......................................................... . 44-3 Problems with Gas Sampling and Storage Containers ...................... . 44-4 Gas Diffusion Problems ........................................................ . REFERENCES ........................................................................ .

Chapter 45 Sulfur Oxidation and Reduction in Soils

1048 1049 1061 1062 1062

1067 M. A. TABATABAI

45-1 Sulfur Oxidation .................................................................. 45-2 Sulfate Reduction ................................................................ REFERENCES .........................................................................

Chapter 46 Iron and Manganese Oxidation and Reduction

1068 1071 1076

1079

WILLIAM C. GHIORSE 46-1 Iron-Depositing and Manganese-Oxidizing Heterotrophs ................... 46-2 Iron-Oxidizing Autotrophs ...................................................... 46-3 lron- and Manganese-Reducing Heterotrophs ................................ ACKNOWLEDGMENT .............................................................. REFERENCES .........................................................................

1081 1086 1090 1094 1094

Subject Index ................................................................................

1097

FOREWORD

The methods pertinent to soil microbiology were formerly included in Part 2 of the Agronomy Monograph No.9, Methods of Soil Analysis. Since the 2nd edition of this document, the number of biochemical and microbiological methods have expanded greatly. In addition because the clientele of scientists engaged in these efforts are primarily soils based, the ASA Board of Directors in 1993 elected to place this document in the SSSA Book Series. It is most refreshing and encouraging to see this stand-alone contribution specifically dedicated to soil microbiological and biochemical methods. This text will be well received by an ever-expanding spectra of biogeoscientists. It is very timely given the rise in public and private interests in soil and water quality, biodiversity, biodegradation, terrestrial ecology, environmental quality protection, sustainability of the biosphere and issues of global climatic change. For too long soil quality has been defined in terms of soil physical and chemical attributes with little or no regard to biological components. Part of this oversight has been a function of techniques available to accurately identify, define and quantify biological health and diversity. Another aspect is the rather recent explosion of interest and awareness among geoscientists in the functionality and import of soil microbiological and biochemical attributes in nearsurface earth processes. The methods reported herein are at the cutting edge of science. Analytical techniques range in resolution from whole organisms to molecular fragments. In unravelling the identity and behavior of the complex soil biological system, temporal and dynamic diversity are considered in sampling methods. The authors represent a select spectra in biogeoscience expertise and career development. Such a synergistic assemblage of scientists assures that the methodology presented is current and relevant. The document is comprehensive in scope, interdisciplinary in character, and offers a high probability of acceptance among biologists. These methods will serve as the standard bearer for both professional and practicing biological scientists. It is the goal that common methodology will enhance collaboration and interchange among scientists and generate data sets using similar analytical approaches. We commend the authors and editors for their diligence and genius in bringing this new book to fruition in such a timely manner. This addition to the SSSA Book Series, will be well received and widely used by a growing number of biogeoscientist professionals wishing to document soil microbiological-biochemical attributes in near surface earth systems.

Larry P. Wilding, president Soil Science Society of America

xv

PREFACE

The books, Methods of Soil Analysis-Parts 1 and 2, published as Agronomy Monograph No.9 have been the primary references on analytical methods used by soil scientists and persons in other disciplines involved with making measurements on soils. Part 2 of the second edition covered both methods on soil chemistry and soil microbiology. The need for more extensive coverage in both of these areas resulted in necessity of dividing Part 2 into two new books. One covering the topic of soil chemistry and the second covering soil microbiology and soil biochemistry. Revision was so extensive and involved so many new authors that it seemed best to consider this book a new publication rather than a third edition. It is published as one of the Soil Science of America Book Series. Division of some subject matter between the book on chemical methods for soil analysis and this book was not always straightforward because some chemical methods are needed in measuring microbiological and biochemical processes. In such cases, a chemical method is provided within chapters of this book but the depth of coverage on theory is not complete nor are alternative methods presented as is the case for the book on soil chemical methods. Our desire was to make it possible to use the methods in this book independently without having to purchase both books. Early in the book the topics of statistical methods, soil sampling, and measurement of soil moisture tension are covered. These chapters were not covered in the previous editions of Part 2 but are particularly important for investigations in soil microbiology and biochemistry. Several methods are provided on use of molecular techniques that were not in previous editions but are needed in many modern soil microbiology laboratories. The treatment of the material on molecular topics is such that a person would not need extensive training in molecular techniques to take advantage of the methods. It is hoped that many laboratories outside of soil science will take advantage of the methods contained in this book. They will be particularly relevant and useful to laboratories with interest in environmental microbiology or bioremediation. Analytical methods are essential to progress in science and the methods presented in this book are recognized by soil scientists as being among the best currently available. All chapters were reviewed by persons having expertise on particular methods, by an associate editor, and by the editor. The help of the many reviewers,

xvii

xviii

PREFACE

efforts and patience of authors, and advice from the editorial board are all gratefully acknowledged. A book such as this one is very much a team effort and is beyond the capability of any individual or small group of individuals. R. W. Weaver, editor Texas A&M University College Station, Texas

J. Scott Angle, associate editor University of Maryland College Park, Maryland

Peter J. Bottomley, associate editor Oregon State University Corvallis, Oregon

CONTRIBUTORS J. Scott Angle

Professor of Agronomy, Agronomy Department, University of Maryland, College Park, MD 20742

Wilfredo Laserna Barraquio

Associate Professor of Microbiology, Institute of Biology, University of the Philippines, Diliman, QL 1101, Philippines

L. W. Belser

School of Science and Computer Studies, Nelson Polytechnic, Nelson, New Zealand

D. F. Bezdicek

Professor of Soils, Department of Crop and Soil Sciences, Washington State University, Pullman, WA 99164-6420

Peter J. Bottomley

Professor of Microbiology and Soil Science, Department of Microbiology, Oregon State University, Corvallis, OR 97331-3802

Thomas W. Boutton

Associate Professor of Ecology, Department of Rangeland Ecology and Management, Texas A&M University, College Station, TX 77843-2126

L.G. Bundy

Professor of Soil Science, Department of Soil Science, University of Wisconsin, Madison, WI 53706

Seth K. A. Danso

Technical Officer, Soil Fertility and Crop Production Section, Joint FAOIIAEA Division, International Atomic Energy Agency, Wagramerstrasse 5, P.O. Box 100, A-1400 Vienna, Austria

Eric A. Davidson

Associate Research Scientist, The Woods Hole Research Center, P.O. Box 296, Woods Hole, MA 02543

D.J. Drahos

Director of Research and Development and Senior Scientist, SBP Technologies, Inc.; Sybron Chemicals, Inc., Salem, VA 24153

B. D. Eardly

Assistant Professor of Biology, Pennsylvania State University, Berks Campus, Reading, PA 19610

Mary K. Firestone

Professor of Soil Microbial Ecology, Department of Soil Science, University of California, 108 Hilgard Hall, Berkeley, CA 94720

Dennis D. Focht

Professor of Soil Microbiology, Department of Soil and Environmental Sciences, University of California, Riverside, CA 92521

xix

xx

CONTRIBUTORS

Jeffry J. Fuhrmann

Associate Professor of Soil Microbiology, Department of Plant and Soil Sciences, University of Delaware, Newark, DE 19717-1303

William C. Ghiorse

Professor and Chairman, Section of Microbiology, Division of Biological Sciences, Cornell University, Ithaca, NY 14853

Peter H. Graham

Professor, Department of Soil Science, University of Minnesota, St. Paul, MN 55108

Charles Hagedorn

Professor of Soil Microbiology, Department of Crop and Soil Environmental Sciences, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061-0404

Stephen C. Hart

Assistant Professor, School of Forestry, Northern Arizona University, P.O. Box 15018, Flagstaff, AZ 86011-15018

R.D. Hauck

Senior Scientist, Tennessee Valley Authority, NFE-IA, Muscle Shoals, AL 35660

WiUiam E. Holben

Research Assistant Professor, Center for Microbial Ecology and Department of Crop and Soil Sciences, Michigan State University, East Lansing, MI 48824. Currently Research Scientist, Environmental Microbiology, The Agouron Institute, La Jolla, CA 92037-4696; Email bholben@ vaxkiller. agr.org.

W. R. Horwath

Faculty Research Associate, 3450 S.W. Campus Way, Crop and Soil Science Department, Oregon State University, Corvallis, OR 97331

Elaine R. Ingham

Associate Professor of Soil Ecology, Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR 97331-2902

Russell E. Ingham

Associate Professor, Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR 973312902

Heinrich F. Kaspar

Cawthron Institute, P.O. Box 175, Nelson, New Zealand

A. C. Kennedy

Soil Scientist, USDA-ARS, Washington State University, Pullman, WA 99164-6421

Leif K1emedtsson

Ph.D., Swedish Environmental Research Institute (IVL), Gothenburg, Sweden

Roger Knowles

Professor of Microbiology, McGill University, Macdonald Campus, Ste. Anne de Bellevue, PQ H9X, Canada

J. O. Legg

Adjunct Professor, Agronomy Department, University of Arkansas, Fayetteville, AR 72701

K. J. McInnes

Assistant Professor of Environmental Physics, Department of Soil and Crop Sciences, Texas A&M University, College Station, TX 77843-2474

CONTRiBUTORS

ni

J. J. Meisinger

Soil Scientist, USDA-ARS, BARC-West, Beltsville, MD 20705

F. Blaine Metting, Jr.

Senior Program Manager, Battelle, Pacific Northwest Laboratories, Richland, WA 99352

Andrew R. Moldenke

Research Professor of Entomology, Oregon State University, Corvallis, OR 97331

A. R. Mosier

Research Chemist, USDA-ARS, Fort Collins, CO 80522

R. L. Mulvaney

Professor, Department of Agronomy, University of Illinois, Urbana, IL 61801

David D. Myrold

Associate Professor, Department of Crop and Soil Sciences, Oregon State University, Corvallis, OR 97331-7306

A. V. Ogram

Assistant Professor of Soils, Department of Crop and Soil Science, Washington State University, Pullman, WA 99164-6420

Timothy B. Parkin

Research Microbiologist, USDA-ARS, National Soil Tilth Laboratory, Ames, IA 50011

Dennis Parkinson

Professor, Department of Biological Sciences, University of Calgary, Calgary, Alberta, Canada TIN 1N4

E. A. Paul

Professor, Crop and Soil Sciences, Michigan State University, East Lansing, MI 48824

Ian L. Pepper

Professor of Environmental Microbiology, Department of Soil and Water Science, University of Arizona, Tucson, AZ 85721

Suresh D. Pillai

Assistant Professor of Environmental Microbiology, Texas A&M University Research Center, EI Paso, TX 79927

Joseph A. Robinson

Associate Director of Biostatistics and Environmental Research, The Upjohn Company, Kalamazoo, MI 49001

M. J. Sadowsky

Associate Professor of Soil Science and Microbiology, Soil Science Department, University of Minnesota, St. Paul, MN 55108

M. J. Savage

Professor of Agrometeorology, Department of Agronomy, University of Natal, Pietermaritzburg 3201, Republic of South Africa

Edwin L. Schmidt

Professor Emeritus of Soil Science, Department of Soil Science, University of Minnesota, St. Paul, MN 55108

Dipankar Sen

Research Scientist, Department of Soil and Crop Sciences, Texas A&M University, College Station, TX 77843

Horace D. Skipper

Professor, Department of Agronomy and Soils, Clemson University, Clemson, SC 29634-0359

xxii

CONTRIBUTORS

T. E. Staley

Research Microbiologist, USDA-ARS, NAA, ASWCRL, Beckley, WV 25813

John M. Stark

Assistant Professor of Microbial Ecology, Department of Biology and the Ecology Center, Utah State University, Logan, UT 84322-5500

David M. Sylvia

Professor of Soil Microbiology, Soil and Water Science Department, University of Florida, Gainesville, FL 326110290

M. A. Tabatabai

Professor of Soil Biochemistry, Department of Agronomy, Iowa State University, Ames, IA 50011

James M. Tiedje

University Distinguished Professor, Center for Microbial Ecology, Michigan State University, East Lansing, MI 48824-1325

I. K. Toth

Research Fellow, Biological Sciences, University of Warwick, Coventry, CV4 7AL U.K.

Ronald F. Turco

Professor, Department of Agronomy, 1150 Lilly Hall of Life Sciences, Purdue University, West Lafayette, IN 47907-1150

R. W. Weaver

Professor of Soil Microbiology, Department of Soil and Crop Sciences, Texas A&M University, College Station, TX 77843-2474

E. M. H. Wellingtot;l

Senior Lecturer, Department of Biological Sciences, University of Warwick, Coventry CV4 7AL U.K.

Duane C. Wolf

Professor, Department of Agronomy, University of Arkansas, Fayetteville, AR 72701

A. G. Wollum, II

Professor of Soil Microbiology, Department of Soil Science, North Carolina State University, Raleigh, NC 27695-7619

Paul L. Woomer

Programme Officer, Tropical Soil Biology and Fertility Programme, P.O. Box 30592, Nairobi, Kenya

S. F. Wright

Research Scientist, USDA-ARS, Soil Microbial Systems Laboratory, BARC-East, Beltsville, MD 20705

L. M. Zibilske

Associate Professor of Soil Microbiology, Department of Plant, Soil and Environmental Sciences, University of Maine, Orono, ME 04469-5722

David A. Zuberer

Professor of Soil Microbiology, Department of Soil and Crop Sciences, Texas A&M University, College Station, TX 77843-2474

Conversion Factors for SI and non-SI Units

xxiii

hectare, ha square kilometer, km 2 (10 3 m)2 square kilometer, km 2 (10 3 m)2 square meter, m 2 square meter, m 2 square millimeter, mm 2 (10- 3 m)2

cubic meter, m 3 cubic meter, m 3 cubic meter, m 3 liter, L (10 -3 m 3 ) liter, L (10 -3 m 3) liter, L (10 -3 m 3 ) liter, L (10 -3 m 3 ) liter, L (10 -3 m 3 ) liter, L (10- 3 m 3 )

2.47 247 0.386 2.47 x 10- 4 10.76 1.55 x 10- 3

9.73 x 35.3 6.10 X 2.84 x 1.057 3.53 x 0.265 33.78 2.11

10- 2

10 4 10- 2

10- 3

kilometer, km (10 3 m) meter, m meter, m micrometer, /Lm (10 -6 m) millimeter, mm (10 -3 m) nanometer, nm (10 -9 m)

Column 1 SI Unit

0.621 1.094 3.28 1.0 3.94 x 10- 2 10

To convert Column 1 into Column 2, mUltiply by

Volume

Area

Length

A

acre-inch cubic foot, ft 3 cubic inch, in 3 bushel, bu quart (liquid), qt cubic foot, ft3 gallon ounce (fluid), oz pint (fluid), pt

acre acre square mile, mi 2 acre square foot, ft 2 square inch, in 2

mile, mi yard, yd foot, ft micron, /L inch, in Angstrom,

Column 2 non-SI Unit

Conversion Factors for SI and non-SI Units

102.8 2.83 x 10- 2 1.64 x 10- 5 35.24 0.946 28.3 3.78 2.96 x 10- 2 0.473

0.405 4.05 x 10- 3 2.590 4.05 x 10 3 9.29 x 10- 2 645

1.609 0.914 0.304 1.0 25.4 0.1

To convert Column 2 into Column I, multiply by

00

c" .... Cj .... ..,Z

0

Z Z

00 .... ~ I::;

0 i:I:l

""1

00

0 i:I:l

(';J

..,~

Z

0

00 ....

i:I:l

tol

~

(';J

0

~.


~

fa

~

~

fa

~

""l

~

~

~

~

~

57.3

35.97

10- 4

5.56 x 10- 3

3.60 x 10- 2

radian, rad

Plane Angle degrees (angle), °

Transpiration and Photosynthesis milligram per square meter second, gram per square decimeter hour, mg m- 2 S-1 gdm- 2 h- 1 micromole (H 20) per square centimilligram (H 20) per s~uare meter second, mg m -2 smeter second, /Lmol cm - 2 S-1 milligram per square meter second, milligram per square centimeter mgm- 2 s- 1 second, mg cm -2 S-1 milligram per square meter second, milligram ~er square decimeter hour, mgm- 2 s- 1 mgdm- h- 1

Energy, Work, Quantity of Heat joule, J British thermal unit, Btu joule, J calorie,cal joule, J erg joule, J foot-pound joule per square meter, J m- 2 calorie per square centimeter (langley) newton, N dyne watt per square meter, W m -2 calorie per square centimeter minute (irradiance), cal cm -2 min-I

9.52 x 10- 4 0.239 10 7 0.735 2.387 x 10- 5 10 5 1.43 x 10- 3

Celsius, °C Fahrenheit, OF

Temperature

Column 2 non-SI Unit

Kelvin, K Celsius, °C

Column 1 SI Unit

1.00 (K - 273) (915°C) + 32

To convert Column 1 into Column 2, multiply by

Conversion Factors for SI and non-SI Units

1.75

2.78

10 4

180

27.8

X

X

10- 2

10- 2

1.05 X 10 3 4.19 10- 7 1.36 4.19 x 10 4 10- 5 698

1.00 (OC + 273) 5/9 (OF - 32)

To convert Column 2 into Column 1, multiply by

00

~

f!3

~ '!-

~

~ if!3

n

;:

I n

;.

gram per kilogram, g kg - I milligram per kilogram, mg kg -I

becquerel, Bq becquerel per kilogram, Bq kg- I gray, Gy (absorbed dosel sievert, Sv (equivalent dosel

0.1

2.7 x 10- 11 2.7 X 10- 2 100 100

2.29 1.20 1.39 1.66

P K Ca Mg

Elemental

centimole per kilogram, cmol kg -I

1

1

cubic meter, m 3 cubic meter per hour, m 3 h- I cubic meter per hour, m 3 h- I hectare-meters, ha-m hectare-meters, ha-m hectare-centimeters, ha-cm

9.73 x 10- 3 9.81 x 10- 3 4.40 8.11 97.28 8.1 x 10- 2

millimho per centimeter, mmho cm -I gauss, G

Plant Nutrient Conversion

Oxide P 20 5 K 20 CaO MgO

curie, Ci picocurie per gram, pCi g-I rad, rd rem (roentgen equivalent manl

Radioactivity

milliequivalents per 100 grams, meq 100 g-I percent, % parts per million, ppm

Concentrations

acre-inches, acre-in cubic feet per second, ft 3 s-I U.S. gallons per minute, gal min -I acre-feet, acre-ft acre-inches, acre-in acre-feet, acre-ft

Water Measurement

siemen per meter, S m -I tesla, T

10 10 4

Electrical Conductivity, Electricity, and Magnetism

0.437 0.830 0.715 0.602

om

3.7 X 10 10 37 0.01

10 1

1

102.8 101.9 0.227 0.123 1.03 x 10- 2 12.33

0.1 10- 4

i -

~:

~ ~

~

~

V:,

~

Z

'"

~

-~

6

'"

~

(dS del)/ko' Thus, (dS dt-l)/k o should always be less than, or at most equal to, the indigenous population. The oxidation rate dS de 1 can be measured easily. Only values for ko are required for the estimate. These values are given in Table 10-2 and were obtained in pure culture (Belser & Schmidt, 1980). Values are given for three genera of NH/ oxidizers, Nitrosomonas, Nitrosospira, and Nitrosolobus, and one genus of N0 2- oxidizer, Nitrobacter.

10-6.2.3 Short-term Nitrification Assay There are several methods by which short-term nitrification activities can be measured. A simple method that appears to be adequate for soils with a significant nitrifier population is recommended here. This method involves preparing soil slurries that are shaken and periodically measured for the accumulation of N0 2-. Sodium chlorate is added to the soil slurries to block the further oxidation of N0 2- (Belser & Mays, 1980). With chlorate present (final concentration 10 mM), the rate at which N0 2- accumulates is equal to the rate of NH/ oxidation and only N0 2- need be measured. Analyses are made quickly and conveniently, with high sensitivity and without problems due to a high N0 3- background.

174

SCHMIDT & BELSER

Table 10-2. Maximum activities per cell determined during exponential growth of nitrifiers in pure culture. Culture Ammonium oxidizers* Nitrosomonas europaea ATCC§ Nitrosomonas sp.§ Nitrosospira briensis Nitrosolobus multi/ormis Nitrite oxidizers~ Nitrobacter winogradskyi Nitrobacter .. agilis"

Activities per cellt 0.011 0.023 0.004 0.023 0.012 0.009

t Picomoles per cell per hour. * Data of Belser and Schmidt (1980). § The ATCC strain is noticeably smaller than many other isolates of Nitrosomonas as typified here by Nitrosomonas sp. (Belser & Schmidt, 1978b, 1980). ~ Data of Rennie and Schmidt (1977). Note: Nitrobacter agilis is now considered to be a strain of N. winogradskyi.

10-6.2.3.1 Materials 1. Sterile 0.5 roM phosphate buffer (section 10-1.1,112 concentration). 2. Ammonium sulfate [(NH4hS04], 0.25 M (sterile). 3. Potassium chlorate (KCI0 3) 1.0 M (sterile). 4. Merthiolate (ethylmercurithiosalicylic acid, sodium salt) 1% (wt/ vol). Store at room temperature in foil-covered bottle. Replace monthly. 5. Reagents and instrumentation for quantitative N02- analysis (see chapter 41 by Bundy and Meisinger). 10-6.2.3.2 Procedure. Prepare soil slurries in duplicate by adding 20 g of moist soil to individual 250-mL cotton-stoppered flasks containing 90 mL of phosphate buffer and 0.2 mL of (NH4hS04 solution (total volume approximately 100 mL depending on the volume weight of the soil). Place flasks on a rotary shaker, and add 1.0 mL of chlorate solution per flask. Let shake for several minutes then take a 5.0-mL aliquot. Add 0.05-mL merthiolate to stop the reaction. Save for nitrite analysis at the end of the assay. Periodically thereafter, collect four to five individual samples spaced at 1- to 2-h intervals, and determine nitrite concentrations when sampling is complete. Calculate the rate at which NOz- accumulates per hour per gram of oven-dry soil. This gives the NH/ oxidation rate. Divide this rate by the appropriate activity per NH4+ oxidizer in Table 10-2 to obtain a rough estimate of the NH4+ oxidizer population per gram of dry weight of soil. Counts can also be made on the suspension to estimate MPN counting effectiveness. Counting efficiency is estimated by dividing the MPN count by the estimated theoretical population.

AUTOTROPHIC NITRIFYING BACTERIA

175

10-6.2.3.3 Comments. The short-term assay is presented as an indicator of the potential activity of the nitrifying population present in a soil at the time of sampling. That population may be fully or partially active at sampling, depending on the interactions of the soil factors that regulate nitrification. The size of that population will be dictated also by the same soil factors. The assay aims to characterize the nitrifying population under conditions such that substrate and oxygen are nonlimiting and incubation time is so short as to essentially avoid increases in the standing population during the assay. The short-term assay provides information on two facets of the nitrifying population. On the one hand the rates of ammonia oxidation obtained by short-term assay reflect the potential overall nitrification rates of a given soil. This is the case since NH/ oxidation is the rate-limiting step in overall nitrification. Hence, comparisons are readily made between different soils as to nitrification potential, and, in addition, the effects of treatment, manipulation, or seasonal change on the nitrifying population of a given soil may be estimated. On the other hand, activity observed by short-term assay is clearly a function of the size of the nitrifying population. Thus, observed activity could be an indicator of population size if it were known how much of the activity could be attributed to a single cell. Such enumeration, however, requires knowledge of the composition of the nitrifying population and of the activity constants of each major component of that population. Unfortunately, neither the nature nor activities of a particular nitrifying population can be defined by current methodologies. Preliminary estimates of nitrifier numbers that have been based on shortterm assay and cell activity constants suggest that actual MPN counts enumerate < 10% of the theoretical population (Sarathchandra, 1978; Belser & Mays, 1982; Berg & Rosswall, 1985). Nitrite oxidation rates can be measured with varying concentrations of N0 2- added to soil slurries along with nitrapyrin [2-chloro-6-(trichloromethyl)pyridine] to inhibit the oxidation of NH/ -N. Nitrite disappearance is followed, and both the maximum activity per gram of soil and the Km are calculated. The Km serves to determine the maximum activity (VmaJ, since the enzyme systems may not be saturated at the N0 2- concentrations used. Various modifications of nitrification potential assays have been proposed. These may supplement or supplant the procedures recommended here. Berg and Rosswall (1985) added chlorate but not ammonium to "undisturbed" soil cores and assayed for nitrite over a 12-h period. The objective was to provide an in situ nitrification rate. The authors predicted that, after further development, the method will become valuable. Killham (1987) proposed a perfusion system for pumping a buffered ammonium solution containing chlorate through a soil column. The method allows for the introduction of various inhibitors such as antibiotics and acetylene to help characterize the nitrification process. Chlorate was also used with apparent success in longer term nitrification incubation assays by Azhar et al. (1989). The equivalent of 2.13 g NaCI0 3 kg- 1 soil allowed continued

SCHMIDT & BELSER

176

nitrate formation for 10 d but not thereafter. Nitrite accumulated during the first 30 d, then declined. Chlorate must be used with considerable caution, however, especially in longer-term incubations. Nitrite-oxidizing bacteria are inhibited because they reduce chlorate to the toxic chlorite form. The presence of nitrite oxidizers and other soil bacteria can lead to the accumulation of chlorite and subsequent inhibition of the ammonia oxidizers as well (Hynes & Knowles, 1983).

REFERENCES Ardakani, M.S., J.T. Rehbock, and A.D. McLaren. 1973. Oxidation of nitrite to nitrate in a soil column. Soil Sci. Soc. Am. Proc. 37:53-56. Azhar, E.S., O. Van Cleemput, and W. Verstraete. 1989. The effect of sodium chlorate and nitrapyrin on the nitrification mediated nitrosation process in soils. Plant Soil 116:133139. Belser, L.W. 1977. Nitrate reduction to nitrite, a possible source of nitrite for growth of nitrite-oxidizing bacteria. Appl. Environ. Microbiol. 34:403-410. Belser, L.W. 1979. Population ecology of nitrifying bacteria. Annu. Rev. Microbiol. 33:309333. Belser, L. W., and E.L. Mays. 1980. The specific inhibition of nitrite oxidation by chlorate and its use in assessing mtrification in soils and sediments. Appl. Environ. Microbiol. 39: 505-510. Belser, L.W., and E.L. Mays. 1982. Use of nitrifier activity measurements to estimate the efficiency of viable nitrifier counts in soils and sediments. Appl. Environ. Microbiol. 43:945-948. Belser, L.W., and E.L. Schmidt. 1978a. Diversity in the ammonia-oxidizing population of a soil. Appl. Environ. Microbiol. 36:584-588. Belser, L.W., and E.L. Schmidt. 1978b. Serological diversity within a terrestrial ammoniaoxidizing population. Appl. Environ. Microbiol. 36:489-593. Belser, L. W., and E.L. Schmidt. 1980. Growth and oxidation of ammonia by three genera of ammonium oxidizers. FEMS Microbiol. Lett. 7:213-216. Berg, P., and T. Rosswall. 1985. Ammonium oxidizer numbers, potential and actual oxidation rates in two Swedish arable soils. BioI. Fert. Soils 1:131-140. Bohlool, B.B., and E.L. Schmidt. 1980. The immunofluorescence approach in microbial ecology. Adv. Microbiol. Ecol. 4:203-241. Boon, B., and H. Laudelout. 1962. Kinetics of nitrite oxidation by Nitrobacter winogradskyi. Biochem. J. 85:440-447. Donaldson, J.M., and G.S. Henderson. 1989. A dilute medium to determine population size of ammonium oxidizers in forest soils. Soil Sci. Soc. Am. J. 53:1608-1611. Hankinson, T.R., and E.L. Schmidt. 1984. Examination of an acid forest soil for ammoniaand nitrite-oxidizing autotrophic bacteria. Can. J. Microbiol. 30:1125-1132. Hankinson, T.R., and E.L. Schmidt. 1988. An acidophilic and a neutrophilic Nitrobacter strain isolated from the numerically predominant nitrite-oxidizing population of an acid forest soil. Appl. Environ. Microbiol. 54:1536-1540. Harms, H., H.P. Koops, and H. Wehrman. 1976. An ammonia-oxidizing bacterium Nitrosovibrio tenus nov. gen. nov. sp. Arch. Mikrobiol. 108:105-111. Hynes, R.K., and R. Knowles. 1983. Inhibition of chemoautotrophic nitrification by sodium chlorate and sodium chlorite: a reexamination. Appl. Environ. Microbiol. 45:11781182. Killham, K. 1987. A new perfusion system for the measurement and characterization of potential rates of soil nitrification. Plant Soil 97:267-272. Knowles, G., A.L. Downing, and M.J. Barrett. 1965. Determination of kinetic constants for nitrifying bacteria in mixed culture with the aid of an electronic computer. J. Gen. Microbiol. 38:263-273. Laudelout, H., R. Lambert, and M.L. Pham. 1976. Influence du pH et de la pression partielle d'oxygene sur la nitrification. Ann. Microbiol. (Paris) 127A:367-382.

AUTOTROPHIC NITRIFYING BACTERIA

177

Matulewich, v'A., P.F. Strom, and M.S. Finstein. 1975. Length of incubation for enumerating nitrifying bacteria present in various environments. App!. Microbio!. 29:265-268. Nishio, M., and C. Furusaka. 1971. Kinetic study of soil percolated with nitrate. Soil Sci. Plant Nutr. 17:61-67. Rennie, R.J., and E.L. Schmidt. 1977. Autecological and kinetic analysis of competition between strains of Nitrobacter in soils. Eco!. Bull. 25:431-441. Sarathchandra, S. U. 1978. Nitrification activities of some New Zealand soils and the effect of some clay types on nitrification. N.Z. J. Agric. Res. 21:615-621. Schmidt, E.L. 1974. Quantitative autecological study of microorganisms in soils by immunofluorescence. Soil Sci. 118:141-149. Smorczewski, W.T., and E.L. Schmidt. 1991. Numbers, activities and diversity of autotrophic ammonia oxidizing bacteria in a freshwater, eutrophic lake sediment. Can. J. Microbio!. 37:828-833. Soriano, S., and N. Walker. 1968. Isolation of ammonia oxidizing autotrophic bacteria. J. App!. Bacterio!' 31:493-497. Soriano, S., and N. Walker. 1973. The nitrifying bacteria in soils from Rothamsted classical fields and elsewhere. J. App!. Bacterio!. 36:523-529. Stanley, P.M., and E.L. Schmidt. 1981. Serological diversity of Nitrobacter from soil and aquatic habitats. App!. Environ. Microbio!. 41:1069-1071. Suzuki, I., V, Dular, and S.C. Kwok. 1974. Ammonia or ammonium ion as substrate for oxidation by Nitrosomonas europaea cells and extracts. J. Bacterio!' 120:556-558. Watson, S.w., E. Bock, H. Harms, H-P. Koops, and A.B. Hooper. 1989. Nitrifying bacteria. p. 1808-1834. In J.T. Staley et a!. (ed.) Bergey's manual of systematic bacterio!' Vol. 3. Williams and Wilkins, Baltimore. Watson, S.W., L.B. Graham, C.C. Remsen, and F.W. Valoi. 1971. A lobular, ammoniaox~dizing bacterium Nitrosolobus multiformis nov. gen. nov. sp. Arch. Mikrobiol. 76.183-203. Woldendorp, V.w., and H.J. Laanbroek. 1989. Activity of nitrifiers in relation to nitrogen nutrition of plants in natural ecosystems. Plant Soil 115:217-228.

Published 1994

Chapter 11 Free-living Dinitrogen-fixing Bacteria ROGER KNOWLES

WILFREDO LASERNA BARRAQUlO, McGill University, Macdonald Campus, Ste. Anne de Bellevue, Quebec, Canada AND

Biological nitrogen (N2) fixation assumes great significance in natural and agricultural systems in view of the impending scarcity of inorganic fertilizers. Free-living N2-fixing microorganisms are widely distributed and found in almost every ecological niche-in soils, associated with plants, in aquatic systems and sediments (Knowles, 1978). This distribution is a function of their great biochemical, taxonomic, and ecological diversity. In the first edition of this monograph, the only free-living N2-fixing bacteria specifically considered were the azotobacters (Clark, 1965). Of the others not included, some, like the clostridia, had been fully documented N2 fixers for many years. A few eukaryotes (e.g., Aureobasidium) could not be confirmed and are not now considered to be N2 fixers. The present list of N2-fixing microorganisms includes at least some species or strains of the aerobic or microaerophilic Azotobacter, Azomonas, Beggiatoa, Beijerinckia, Campylobacter, Derxia, Acetobacter, Aquaspirillum, Azospirillum, Herbaspirillum, Corynebacterium (Xanthobacter), Lignobacter, methanotrophs, Mycobacterium (Xanthobacter), Pseudomonas, and Thiobacillus; the facultatively anaerobic Enterobacteriaceae, Alcaligenes, Bacillus, Vibrio, and purple photosynthetic nonsulfur bacteria; the obligately anaerobic Clostridium, Desulfovibrio, Desulfotomaculum, Propionispira, methanogens and purple and green photosynthetic S bacteria; and finally, many cyanobacteria (blue-green algae). For a fuller discussion of this list with appropriate references, see Knowles (1978), Salkinoja-Salonen et al. (1979), and Sprent and Sprent (1990). For many of these groups of N2fixing microorganisms, no well-established enrichment or enumeration methods have been thoroughly tested in different laboratories. In this chapter, the emphasis is on enumeration techniques, and for this reason, some of the methods suggested here must be taken as tentative and subject to modification as may be found necessary. Most of the groups mentioned above are included except for the N2-fixing bacilli and photosynthetic S bacteria for which simple one- or two-step enumeration procedures have not been reported. Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 179

180

KNOWLES & BARRAQUIO

Enrichment procedures (Veldkamp, 1970) permit isolation of specific bacteria, and rapid routine testing of N2 -fixing ability is relatively easy using the catalysis of the reduction of C2H 2 to C2H 4 (by the versatile nitrogenase system) as outlined in chapter 43 by Weaver and Danso in this book and in section 11-1 (Hardy et aI., 1968). Many enrichment culture techniques can be easily adapted to quantitative enumeration by an extinction dilution or most probable number (MPN) method (see chapter 5 by Woomer in this book), but there are some problems. The validity of such methods depends on the ease with which cultures of the desired organism develop from small inocula (and ideally from a single cell). For some bacteria, establishment of a culture from a single cell does not occur readily. One of the best criteria for use in MPN counts is the positive C2H 2 reduction test (Okon et aI., 1977; Patriquin & Knowles, 1972; Villemin et aI., 1974), but the numerous C2H 4 analyses involved make this tedious for other than confirmatory purposes. The C2H 2 reduction assay cannot be used to test for CH4-supported N2 fixers since C 2H 2 inhibits the initial oxidation of CH4 , thus depriving the methanotroph of its source of C and energy (De Bont & Mulder, 1976), as discussed later in section 11-4. The selective culture of N 2-fixing aerobes is difficult because it is virtually impossible to exclude the development of anaerobic microsites that permit growth and activity of facultative and even obligately anaerobic bacteria. For example, Line and Loutit (1973) demonstrated that C2 H 2 was reduced to C2H 4 by a Clostridium growing along with a Pseudomonas culture on an aerobic agar slant. The microaerophilic fixers are particularly troublesome in this respect. For this reason, MPN techniques are here described employing aerobic semisolid cultures in which subsurface plates of microaerophiles develop at appropriate O 2 concentrations within the agar. Inconsistencies in the compositions of the media described are a reflection of the varied published sources from which the methods are derived. Fluid for serial dilutions should maintain the integrity of bacterial cells (Billson et aI., 1970; Ridge, 1970), but it should not contain combined N. Thus, the use of 0.1 % peptone solution, for example, is clearly undesirable. Iron may be supplied as FeCl3 or FeS04 ·7H20, or in chelated form as ferric monosodium ethylenediaminetetraacetic acid (FeEDTA) or ethylenediaminetetraacetic acid tetrasodium salt (Sequestrene, Ciba-Geigy Corp., Greensboro, NC). Other trace element supplements that are frequently necessary for successive transfer of pure cultures mayor may not be necessary for the initial enrichment cultures involved in MPN methods. 11-1 THE ACETYLENE REDUCTION ASSAY The versatile nitrogenase system reduces several low molecular weight substrates such as N 2, C2H 2 , N20, N3 -, and CN-. The sensitive methods available for the detection and measurement of C2 H 2 and its reduction product C2H 4 , permit the use of this reaction as an indirect assay of nitrogenase activity (Hardy et aI., 1968). The C2H 2 reduction (or C2H 2-C2H 4)

DINITROGEN·FIXING BACTERIA

181

assay is about 1000 times more sensitive than the 15N2 method, and fur· thermore, it is cheap and simple in its application (Hardy et aI., 1968). Acetylene is introduced into the system at a concentration sufficient to saturate the nitrogenase sites (usually 0.05-0.1 atm) and completely to inhibit the reduction of air N2. After an appropriate incubation time the amount of C2H 4 formed is determined by gas chromatography. The assay is described in chapter 43 in this book. The great sensitivity permits assays of pure cultures, enrichments, some MPN tubes, and active natural systems, for example, root nodules, to be completed in < 1 h (see chapter 43). However, the low activities of many other natural soil, sediment, and rhizosphere samples require longer assays, which bring a number of potential problems: 1. Acetylene is not the physiological substrate of nitrogenase, and its presence may impose N limitation on the N2-fixing microorganisms and may cause superinduction (and therefore overestimation) of nitrogenase (David & Fay, 1977). 2. It is extremely difficult to match, in a closed system assay (C 2H 2 or 15N2), such conditions as O 2 concentration, nutrient and moisture avail· ability, and light intensity that exist in situ. This is particularly true in long assays where changes can occur in microbial populations (Okon et aI., 1977) as well as in the other parameters just mentioned. 3. Acetylene has side effects not shown by N2. These include the inhibition of (i) nitrogenase-dependent H2 evolution (Hardy et aI., 1968), (ii) conventional hydrogenase activity (Smith et aI., 1976), (iii) uptakehydrogenase activity (Smith et aI., 1976), (iv) cell proliferation in clostridia (Brouzes & Knowles, 1971), (v) N2 reduction by denitrifiers (Fedorova et aI., 1973), (vi) oxidation of NH4 + to NH 20H by Nitrosomonas (Hynes & Knowles, 1978) (vii) methanogenesis (Raimbault, 1975), and (viii) oxida· tion of CH 4 to CH3 0H by methanotrophic bacteria (De Bont & Mulder, 1976). 4. Another side effect of C2H 2 is that it inhibits the further metabolism of C2H 4. Thus, the actual net accumulation of "endogenous" C2H 4 could be greater in the presence of C2H 2 than in its absence, and this C 2H 4 would be measured as part of the C2H 4 produced from the C2H 2 (Witty, 1979). A control for this employing CO is described by Nohrsted (1983). The reader is referred to chapter 43, Turner and Gibson (1980), and Knowles (1980) for a fuller discussion of these and other aspects of the C2H 2 reduction assay. 11-2 METHODS FOR DINITROGEN FIXERS IN GENERAL 11-2.1 Principles The great biochemical and physiological diversity of free-living N2fixers make the estimation of their total number at one sampling time difficult. Enumeration procedures usually require the use of specific media

182

KNOWLES & BARRAQUIO

and incubation conditions to demonstrate the presence of at least a generic group of N2 fixers. The development of a single isolation medium to accommodate most of the commonly occurring free-living heterotrophic N2 fixers in soil has been most helpful. The enumeration procedure described here uses a combined C medium on which at least nine genera of N2 fixers can be isolated (Rennie, 1981). It requires plating the sample on the combined C agar medium, incubating the plates aerobically and anaerobically, and then testing the individual colonies for acetylene-reducing activity. Classification of the acetylene-reducing isolates into major genera is achieved through a series of biochemical tests supplemented with some physiological and cultural tests (Rennie, 1980). 11-2.1.1 Prepared Materials 1. Sterile dilution blanks (see chapter 8 by Zuberer in this book). 2. Nitrogen-deficient combined C medium (Rennie, 1981): a. Solution A: To 900 mL of distilled water, add 0.8 g of potassium monohydrogen phosphate (K2HP0 4 ), 0.2 g of potassium dihydrogen phosphate (KH2P0 4 ), 0.1 g of sodium chloride (NaCl), 28 mg of ethylenediaminetetraacetic acid disodium salt Fe (Na2FeEDTA), 25 mg of sodium molybdate dihydrate (Na2Mo0 4 ·2H20), 100 mg of yeast extract, 5 g of mannitol, 5 g of sucrose, and 0.5 mL of sodium lactate (60%, v/v). For the agar medium, add 12 g of purified agar (Difco Noble Agar or Oxoid Ionagar)1. b. Solution B: To 100 mL of distilled water, add 0.2 g of magnesium sulfate heptahydrate (MgS0 4 ·7H20), and 0.06 g of calcium chloride (CaCI2). Sterilize the solutions separately at 121°C for 15 min, cool, and mix. Add filter-sterilized biotin (5 !-lg L -1) and para-aminobenzoic acid (10 !-lg L -1). Adjust the final pH to 7.0 using sterile acid or alkali. 3. Sterile glass or disposable plastic petri dishes. 4. Anaerobic jar with H 2-C02 GasPak kits and catalyst (Becton Dickinson and Co., BBL Microbiology Systems, Cockeysville, MD). Alternatively, a jar with gassing ports or a vacuum desiccator. 5. Supply of N2. 6. A rotating petri dish turntable or glass hockey stick spreader. 7. 7-mL bijou vials (Sterillin Co., Ltd., Teddington, Middlesex, UK) or serum bottles (of 6- to lO-mL capacity) containing 3 mL of the liquid combined carbon medium. 8. Sterile serum stoppers (Suba seal from W. Freeman and Co., Barnsley, Yorkshire, UK) for the above vials of medium. lDifco Noble Agar, Difco Laboratories, Detroit, MI; or Oxoid Ionagar, Oxoid, Division of Oxo Limited, London.

DINITROGEN·FIXING BACTERIA

183

11-2.1.2 Procedure Collect and prepare the sample as described in chapter 1 and prepare serial dilutions as described in chapter 8. Dispense the melted agar medium into sterile petri dishes, and dry the agar surface by incubating at 30 to 37°C for a few hours. Then, from the highest serial dilution prepared (10- 4 is usually adequate), transfer an aliquot of 0.2 to 0.5 mL to each of five replicate plates. Make similar transfers from the successive lower dilutions, including the 10- 1 . Spread the aliquot over the agar surface as it is released from the pipette by rotating the plate on a turntable, or spread it using a glass rod hockey stick sterilized with 70% alcohol and flaming. Then again dry the agar surface. Incubate the plates aerobically or anaerobically at 25 to 30°C for 2 to 3 d. For anaerobic incubation, place the plates in the anaerobic jar (or desiccator), and generate the appropriate H 2-C02-N2 atmosphere using the GasPak kit. Alternatively, flush the chamber with N2, or evacuate and backfill it with N2. Record the number and cultural characteristics of colonies. Transfer individual colonies to sterile bijou vials or serum bottles each containing 3 mL of the liquid combined C medium. Seal the vials with sterile Suba seals and incubate for 24 h at 25 to 30°C statically. If an anaerobic atmosphere is required, evacuate and backfill the vials with 02-free N2 or He using a manifold equipped with sterilized needles and cotton wool filters. Replace about 10% of the gas phase in the vial with C2H2> and continue incubation for up to 24 h. Determine the C2H 4 produced by GC analysis of a 0.2- to 0.5-mL sample of the gas phase (see chapter 43). Record as positive those isolates that exhibit two or three times more C2H 4 than is present in an uninoculated control. 11-2.2 Classification into Major Genera The nine genera of N2-fixing bacteria that can be isolated using the combined C medium include Azotobacter, Azospirillum, Enterobacter, Klebsiella, Erwinia, Bacillus, Clostridium, Derxia, and Rhodospirillum (Rennie, 1981). A computer program designed specifically for the identification of these nine genera of N2-fixing microorganisms has been developed and tested (Rennie, 1980). It is based on interpretation of the 70 biochemical tests of the API 20E and 50E, supplemented with tests for acetylene reduction, nitrate and nitrite reduction, catalase, oxidase, motility, and growth on MacConkey's bile salt medium. Identification can be to the genus and often to species level. 11-3 METHODS FOR AZOTOBACTERACEAE 11-3.1 Principles In general, it is not possible to use the MPN tube method for enumeration of N2-fixing obligate aerobes because of the ease with which

184

KNOWLES & BARRAQUIO

anaerobic and facultative N2 fixers can show activity in 02-deficient microsites (Line & Loutit, 1973). Therefore, for members of the Azotobacteraceae, various modifications of surface plating or MPN plate procedures are used with a N-free solution solidified with purified agar or silica gel and containing an appropriate carbohydrate. Glucose is used by all species; mannitol and benzoate are used by only some species. Of the four genera that now comprise the Azotobacteraceae (Azotobacter, Azomonas, Beijerinckia, and Derxia) (Tchan, 1984), reasonably good enumeration procedures exist for the first three, and two methods based on Brown et al. (1962) for Azotobacter and Strijdom (1966) for Beijerinckia are described below. Enumeration of Derxia in natural samples is difficult, and a method is not included here. 11-3.2 Method for Azotobacter and Azomonas 11-3.2.1 Prepared Materials 1. Sterile dilution blanks (see chapter 8 in this book). 2. Medium for Azotobacter (Brown et aI., 1962): To 1000 mL of distilled water, add 5 g of glucose, 0.2 g of magnesium sulfate heptahydrate (MgS0 4·7H20), 0.04 of ferrous sulfate heptahydrate (FeS0 4·7H20), 0.005 g of sodium molybdate dihydrate (Na2Mo04·2H20), 0.15 g of anhydrous calcium chloride (CaCI2), and 15 g of agar. Purified agar (e.g., Difco Noble Agar or Oxoid Ionagar) is preferable, and only about 12 g of such preparations is required. Sterilize at 121°C for 15 min. Then to the melted and cooled agar, add separately sterilized potassium monohydrogen phosphate (K2HP0 4) solution to give a final concentration of 0.8 g L -1. The final pH is 6.8 to 7.0. 3. Sterile glass or disposable plastic petri dishes. 4. Rotating petri dish turntable or glass hockey stick spreader.

11-3.2.2 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. Dispense the melted cooled agar medium containing the K2HP0 4 into sterile petri dishes. Incubate the plates partially open at 30 to 37°C under a clean air hood or in a sterile room until the surface is quite dry. Then, from the highest serial dilution prepared (10- 4 is usually adequate), transfer an aliquot of 0.2 to 0.5 mL to each of five replicate plates. Make similar transfers from the successive lower dilutions, including the 10- 1 . Spread the aliquot over the agar surface as it is released from the pipette by rotating the plate on a turntable, or spread it using a glass rod hockey stick

DINITROGEN-FIXING BACTERIA

185

sterilized with 70% alcohol and flaming. The agar surface should again be dried before incubation. Incubate at 25 to 30°C for 3 to 5 d. Many so-called oligonitrophiles (combined N scavengers) produce small colonies on plates prepared as described. Their colonies are usually < 1 mm in diameter but occasionally may be up to 3 mm diam. and clearly transparent. These generally are not N2 fixers. Azotobacter (and possibly certain species of Azomonas) produce colonies 2 or more millimeters in diameter having a creamy mucilaginous appearance. Azotobacter chroocaecum, probably the most commonly occurring species, can produce colonies up to 10 mm in diameter that gradually turn brown to black with age. The blackening of such colonies is intensified if sodium benzoate is used in the medium instead of glucose (Aleem, 1953). 11-3.3 Method for Beijerinckia 11-3.3.1 Prepared Materials 1. Sterile dilution blanks (see chapter 8 in this book). 2. Medium for Beijerinckia (Becking, 1961): a. Solution A: To 500 mL of distilled water, add 20 g of glucose, and sterilize at 121°C for 15 min. b. Solution B: To 500 mL of distilled water add 1.0 g of potassium dihydrogen phosphate (KH2P0 4 ), 0.5 g of magnesium sulfate heptahydrate (MgS0 4 ·7H20) and 0.02 g of sodium molybdate dihydrate (Na 2Mo04 ·2H20). Adjust to pH 5.0, and sterilize at 121°C for 15 min. Mix solutions A and B after cooling. 3. Petri dishes, each containing 1 g of finely powdered soil (Strijdom, 1966). Sterilize the soil by autoclaving (121°C) for 1 h on each of two successive days. 11-3.3.2 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. To each of the petri dishes containing soil, add 5 mL of the liquid medium. Then, from the highest serial dilution prepared (10- 3 or 10- 4), transfer an aliquot of 1.0 mL to each of five replicate plates. Make similar transfers from the successive lower dilutions, distributing each aliquot over the surface of the medium-saturated soil. Incubate at 30°C for up to 3 wk, adding fresh medium if necessary to keep the soil surface moist (Strijdom, 1966). Then examine each petri dish for the presence or absence of raised mucilaginous viscous growth typical of Beijerinckia spp. If necessary, check growth microscopically. Record all petri dishes as positive or negative for Beijerinckia, and calculate the MPN by reference to appropriate tables found in chapter 5.

186

KNOWLES & BARRAQUIO

11-4 METHOD FOR METHANOTROPHS 11-4.1 Principles Methanotrophic bacteria (methane-using methylotrophs) of the genera Methylomonas, Methylobacter, Methylocystis, Methylosinus, and Methylococcus can fix N2 (Oakley & Murrell, 1988) and are obligately microaerophilic when doing so (De Bont & Mulder, 1974). Acetylene inhibits the oxidation of methane but not of methanol by these organisms (De Bont & Mulder, 1976). Thus, it is not possible to test C2H 2 reduction by enrichment cultures using an N-deficient mineral salts medium incubated under CH4. Methods have been described for the enumeration of methanotrophs (e.g., Megraw & Knowles, 1987) but not for N2-fixing methanotrophs. We, therefore, suggest that the following methods be used on a trial basis only. 11-4.2 Plate Count Method for Methanotrophs 11-4.2.1 Prepared Materials 1. Sterile dilution blanks (see chapter 8 in this book). 2. Nitrogen- and C-deficient medium for methanotrophs (after De Bont & Mulder, 1974; Whittenbury et aI., 1970): To 1000 mL of distilled water, add 0.5 g of potassium monohydrogen phosphate (K2HP0 4), 0.5 g of potassium dihydrogen phosphate (KH2P0 4), 0.2 g of magnesium sulfate heptahydrate (MgS0 4·7H20), 0.015 g of calcium chloride (CaCI2), 0.001 g of ferrous sulfate heptahydrate (FeS04·7H20) or 0.004 g of Sequestrene Fe (Ciba-Geigy), 0.001 g of sodium molybdate dihydrate (Na2Mo04·2H20), and 10 mL of soil extract (see chapter 8 in this book). Adjust the pH to 6.8 to 7.0. Sterilize at 121°C for 20 min (the phosphates should be sterilized separately). 3. Sterile glass or disposable plastic petri dishes. 4. Sterile membrane filters (47 mm, 0.45 ~m, e.g., Millipore HAWG) , glass fiber pads, and filtration unit. 5. Anaerobic jar with gassing ports, desiccator, or plastic bags (of impermeable Saran-type material) with rubber hose connections. 6. Supplies of CH4 and of N2 (or a mixture of 2-4% O 2 in N2). 11-4.2.2 Procedure Collect and prepare the samples as described in chapter 1, and prepare serial dilutions as described in chapter 8. Filter aliquots (5 mL) of the dilutions through membrane filters and place the membranes onto glass-fiber pads soaked with 2 mL of the N- and C-deficient medium (Megraw & Knowles, 1987). From each dilution prepare duplicate filters for incubation in the presence and absence of CH4 as described below.

DINITROGEN-FIXING BACTERIA

187

Place the plates in a chamber (anaerobic jar or desiccator) connected to cylinders of N2 and CH4 , a vacuum pump, and manometer or vacuum gauge. Evacuate to a residual pressure of 76 to 152 mm of Hg (608-684 mm vacuum), and partially backfill the chamber to a residual air pressure of 608 (152-mm vacuum) with N2. Introduce CH4 to one atmosphere. Alternatively place the plates in plastic bags, and flush with a mixture of 2 to 4% O2 and;::: 20% CH4 in N2 (Hill, 1973). Note that 5 to 15% CH4 is the explosive range. Incubate a duplicate control set of membranes in the absence of CH4 • Incubate at 25°C for 2 to 3 or more weeks, and examine for colonies appearing on the CH4 series but not on the control series. Representative colonies may need to be checked for ability to use CH4 (Whittenbury et aI., 1970). 11-4.3 Most Probable Number Method for Methanotrophs 11-4.3.1 Prepared Materials 1. Sterile dilution blanks (see chapter 8 in this book). 2. Nitrogen- and C-deficient semisolid medium for methanotrophs, prepared exactly as in section 11-4.2.1 but adding 1.75 g of purified agar (Difco Noble Agar or Oxoid Ionagar)l per liter of medium. Dispense about 3 to 5 mL of the medium in each of a series of vials or serum bottles (of 6- to lO-mL capacity), plug with cotton or foam, and sterilize at 121°C for 15 min. 3. Sterile serum stoppers for the above vials of medium.

11-4.3.2 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. From the highest dilution prepared, transfer a O.l-mL aliquot to each of five vials of the semisolid medium. Then make similar transfers from the successive lower dilutions as appropriate. Do not insert serum stoppers at this stage. Incubate at 30 to 35°C in a desiccator or other chamber filled with an atmosphere of 20% CH4 in air until well-defined pellicle formation has occurred. Do not disturb the pellicle by shaking the tubes. Add 0.1 mL of a 3 to 5% methanol solution to give a final concentration in the medium of 0.1% methanol (De Bont & Mulder, 1974). Replace the aerobic plug with a sterile serum stopper, and replace about 10% of the gas phase with C2H 2 • Continue the incubation for a further 6 to 24 h, and determine the C2H 4 produced by GC analysis of a 0.1- to 0.5-mL sample of the gas phase (see chapter 43 in this book). Compare with the C2H 4 observed in uninoculated vials treated in exactly the same way. Record as positive those vials showing pellicle formation and significantly more C2H 4 than is present in an uninoculated control. Calculate the MPN of N 2 -fixing methylotrophs by referring to the statistical tables found in chapter 5.

188

KNOWLES & BARRAQUIO

11-5 METHOD FOR HYDROGEN·USING DINITROGEN FIXERS 11-5.1 Principles "All strains previously assigned to Corynebacterium autotrophicum for taxonomic reasons proved to be able to fix nitrogen under autotrophic conditions, and all strains isolated as N2-fixing hydrogen bacteria so far have been found to belong to C. autotrophicum" (Wiegel & Schlegel, 1976). These bacteria, now classified as Xanthobacter, can couple the oxidation of H2 to the fixation of N2 and CO2, Dinitrogen-fixing cultures are highly microaerophilic with an optimum O 2 concentration in the region of 3 to 5% (De Bont & Leijten, 1976). The efficient enrichment and enumeration of this organism has been reported using an N-deficient liquid medium free of organic C incubated under an atmosphere of 10% O 2, 10% CO2, 20% H 2, and 60% N2 (De Bont & Leijten, 1976). However, some H 2-using N2 fixers such as Azospirillum brasilense, Azomonas agiUs, Bacillus polymyxa, Beijerinckia indica, and Azotobacter vinelandii do not grow chemolithotrophically (Bowien & Schlegel, 1981; Malik & Schlegel, 1981; Pedrosa et aI., 1980; Wong & Maier, 1985). Except for the cyanobacteria and other phototrophic bacteria, all known aerobic H 2-using N2 fixers are basically heterotrophs (Bowien & Schlegel, 1981). The method described here is an MPN technique using a semisolid medium for heterotrophs incubated under aerobic conditions (Barraquio et aI., 1988). The semisolid medium allows expression of 02-sensitive nitrogenase, and hydrogenase that is often equally O 2 sensitive. 11-5.2 Prepared Materials 1. Sterile dilution blanks (see chapter 8 in this book). 2. Combined C medium described by Rennie (1981) (section 11-2.1.1) and modified by Barraquio et al. (1988) as follows: To 1000 mL of distilled water, add 2.0 g of Difco Noble agar, 0.06 g of sequestrene NaFe (13% Fe, CIBA-Geigy Corp., Greensboro, NC) to replace sodium ferrous EDTA, 5.9 mg (instead of 25 mg) of sodium molybdate dihydrate (NaMo0 4 ·2H20), and 2.5 g of sodium malate. Omit the vitamins. Nickel chloride (NiC1 2) at a final concentration of 1 to 5 !lM may be added to enhance H 2-uptake activity. Prepare the medium as described in section 11-2.1.1. Dispense 4 mL of the melted semisolid medium into each of a series of pre-sterilized cotton-plugged 14-mL serum bottles (Wheaton, Millville, NJ). This volume of the medium will give a depth of about 13 mm. 3. Sterile butyl rubber stoppers and aluminum crimps for the above serum bottles.

DINITROGEN-FIXING BACTERIA

189

4. A supply of gaseous tritium with a specific activity of about 20 !lCi (740 kBq) per cc. Appropriate license and laboratory facilities are required. 11-5.3 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. From the highest serial dilution prepared, transfer a 0.1-mL aliquot to each of five serum bottles of semisolid medium. Then make similar transfers from the successive lower dilutions as deemed appropriate. Do not insert butyl stoppers at this stage. Incubate statically at 30 to 35°C for 3 d or more, and record the presence and absence of growth as determined visually. Replace the aerobic plug with a sterile butyl stopper and apply an aluminum seal. Replace about 1% (relatively low concentration so as not to inhibit greatly H 2-uptake activity) of the gas phase in the bottle with C2H 2. Continue incubation for a further 24 h and determine the C2H 4 produced by GC analysis of a 0.2- to 0.5-mL sample of the gas phase (see chapter 43). Record as positive those bottles that contain two or three times more CzH4 than is present in an uninoculated control. Use the same cultures for the tritium (H3H) uptake assay. Inject 0.3 mL of H3H into the bottle, and then continue incubation for another 24 h. Vortex the bottles vigorously, and then transfer 0.1-mL aliquots into 6-mL scintillation vials each containing 4-mL scintillation fluid (Ready-Solv MP, Beckman Instruments, Fullerton, CA). Determine radioactivity of the samples by liquid scintillation (Beckman LS 7500 counter) with 3H-galactose or other tritiated compound as the standard. An alternative method of determining H 2-using activity that can be tried in laboratories not allowed to handle isotopes is to inject 1% H2 into each serum bottle and then determine the H2 concentration at the start and after 24 to 48 h of incubation by GC analysis (Chan et aI., 1980). Record as positive those cultures that exhibit significantly more radioactivity (or less H2 if done by the alternative method) than is present in an uninoculated control. Calculate the MPN of H 2-using N2 fixers by referring to the statistical tables found in chapter 5 by Woomer in this book. The MPN of H 2-using non-N2-fixers and N2-fixing non-H 2-utilizers can also be determined from the data. 11-6 METHOD FOR CYANOBACTERIA 11-6.1 Principles Cyanobacteria such as Anabaena and Nostoc can compartmentalize their nitrogenase in specialized heterocysts in which a sufficiently reducing environment is maintained. They thus can fix N2 under highly aerobic

KNOWLES & BARRAQUIO

190

conditions (Stewart, 1973). Other cyanobacteria, e.g., the filamentous Plectonema and Oscillatoria and the unicellular Gloeocapsa, do not produce heterocysts and generally synthesize nitrogenase under micro aerobic to anaerobic conditions (Stewart, 1973). At high light intensities (> 9.8 !-tmol m- 2 S-1 (> 500 Ix) for Gloeocapsa) the production of O2 is stimulated to such an extent that N2 fixation may be inhibited (Stewart, 1973). Although studies of soils have revealed mainly the heterocystous N2-fixing cyanobacteria, this could be due to the inadequacy of existing methods to select for the nonheterocystous N2 fixers. Most studies on the distribution of the cyanobacteria use enrichment culture in C- and N-free mineral salts solutions. The enumeration method described here uses such a medium solidified with agar for a plate count and is slightly modified from that of Jurgensen and Davey (1968). 11-6.2 Prepared Materials 1. Sterile dilution blanks (see chapter 8 in this book). 2. Nitrogen- and organic C-deficient medium for cyanobacteria (after Cameron & Fuller, 1960; Wieringa, 1968): To 1000 mL of distilled water, add 0.35 g of potassium monohydrogen phosphate (K2HP0 4), 0.2 g of magnesium sulfate heptahydrate (MgS0 4·7H20), 0.15 g of calcium chloride dihydrate (CaCI2·2H20), 0.1 g of sodium chloride (NaCl), 0.005 g of ferrous sulfate heptahydrate (FeS04·7H20), 100 !-tg of sodium molybdate dihydrate (Na2Mo04·2H20), 5 !-tg of cupric sulfate pentahydrate (CuS04·5H20), 10 !-tg of boric acid (H3B03), 70 !-tg of zinc sulfate heptahydrate (ZnS0 4·7H20), 10 !-tg of manganous sulfate monohydrate (MnS04·H20) , and 15 g of purified agar (Difco Noble Agar or Oxoid Ionagar).1 Autoclave at 121°C for 15 min. 3. Sterile glass or plastic disposable petri dishes. 11-6.3 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. From the highest serial dilution prepared, transfer a 1.0-mL aliquot to each of five replicate petri dishes. Make similar transfers from the successive lower dilutions as necessary. Pour at least 25 mL of the melted and cooled (45°C) agar medium into each dish, swirl, and allow to solidify. Incubate under illumination of no more than about 49 !-tmol m- 2 S-1 (2500 Ix) from cool-white fluorescent lamps, with a 16-h light period alternating with an 8-h dark period at 20 to 30°C. Count the colonies that develop after about 6 wk of incubation. Such a count may include organisms that are unable to fix N2 but are efficient scavengers of fixed N.

DINITROGEN-FIXING BACTERIA

191

11-7 METHOD FOR PHOTOSYNTHETIC PURPLE NONSULFUR BACTERIA 11-7 _1 Principles The genera Rhodobacter, Rhodospirillum, Rhodopseudomonas, and Rhodomicrobium, all members of the Rhodospirillaceae (Imhoff & Truper, 1989) are photoorganotrophs that carry out photo assimilation of simple organic compounds such as malate and succinate. Organic compounds or H2 are used as photosynthetic electron donors, and under anaerobic conditions, many species fix N2. There are no well-documented accounts of the enumeration of these N2 fixers from natural environments. The method that follows is modified from the work of Gibson (1975), Weaver et al. (1975), Finke and Seeley (1978), and Siefert et al. (1978) and employs an illuminated anaerobic plate count on aN-deficient malatecontaining medium. 11-7.2 Prepared Materials 1. Sterile dilution blanks (chapter 8). 2. Nitrogen-deficient malate agar medium (after Gibson, 1975; Siefert et al., 1978; Weaver et al., 1975): a. Solution A: To 250 mL of distilled water, add 4.0 g of ethylenediaminetetraacetic acid disodium salt (EDTA disodium), 0.2 g of ferrous sulfate heptahydrate (FeS04·7H20), 0.7 g of boric acid (H3 B0 3 ), 0.39 g of manganous sulfate monohydrate (MnS04·H20), 0.06 g of zinc sulfate heptahydrate (ZnS0 4·7H20), 0.025 g of sodium molybdate dihydrate (NazMo0 4·2HzO), and 0.01 g of cupric nitrate trihydrate [Cu(N0 3 h·3H zO]. Autoclave at 121°C for 15 min. b. Solution B: To 1000 mL of distilled water, add 100 mg of nicotinic acid, 50 mg of thiamine hydrochloride, 10 mg of biotin, and 10 mg of p-aminobenzoic acid. Sterilize by membrane filtration. Different species of the Rhodospirillaceae have different growth factor requirements, but according to Van Niel (1972), these four factors should support growth of all species. c. Solution C: To 990 mL of distilled water, add 0.5 g of potassium dihydrogen phosphate (KH 2P0 4), 0.2 g of magnesium sulfate heptahydrate (MgS0 4·7H20), 0.015 g of calcium chloride dihydrate (CaCI 2·2H20), and 1.34 g of DL-malic acid (neutralized separately with sodium hydroxide [NaOH]). To solution C, add 1 mL of solution A, and adjust the pH to 6.8. Add 12 g of purified agar (Difeo Noble Agar or Oxoid Ionagar), 1 heat to dissolve, and then autoclave at 121°C for 15 min. Allow to cool to 45°C, add 10 mL of solution B, and mix gently. This medium is then ready for pouring plates.

192

KNOWLES & BARRAQUIO

3. Sterile petri dishes. 4. Anaerobic jar with H 2-C02 GasPak kits and catalyst (Becton, Dickinson and Co., BBL Microbiology Systems, Cockeysville, MD). Alternatively, a jar with gassing ports or a vacuum desiccator. 5. If Gaspak kits are not available, supplies of H 2, CO 2, and N2 or a premixed cylinder containing 10% H 2, 10% CO2, and 80% N2 is required. In these cases, a vacuum pump and gassing manifold with vacuum gauge or gauges are required. 11-7.3 Procedure

Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. From the highest dilution prepared, transfer a I-mL aliquot to each of five petri dishes. Make similar transfers from the successive lower dilutions as necessary. Pour about 20 mL of the 45°C agar medium, into each dish, swirl, and allow to solidify. Place the dishes in the anaerobic jar, and generate the appropriate H 2-C02-N2 atmosphere using the GasPak kit. Alternatively, flush the chamber with the appropriate gas mixture, or evacuate and backfill with N2 and introduce the appropriate partial pressures of H2 (0.1 atm), CO 2 (0.1 atm), and N2 (0.8 atm) using the vacuum pump and gassing manifold (e.g., section 11-4.2.2). Incubate the plates at 25 to 30°C under about 49 !lmol m- 2 S-l (2500 Ix) of fluorescent plus incandescent light for at least 14 d, and then count the purple colonies that have developed. 11-8 METHOD FOR CLOSTRIDIA 11-8.1 Principles

The obligately anaerobic spore-forming clostridia are difficult to enumerate using anaerobic plate count techniques because of gas production, splitting of the agar, and spreading of colonies. Most probable number procedures are therefore used in which a N-deficient liquid medium containing a reducing agent and redox indicator is inoculated with appropriate dilutions of the sample (Brouzes et aI., 1971). The possible criteria for positive tubes are gas production, reduction of the indicator, and detection of C2H 2-reducing activity. Bacilli may grow under the conditions described but, in practice, positive tubes generally contain clostridia. 11-8.2 Prepared Materials

1. Sterile dilution blanks (chapter 8). 2. Medium for N2-fixing clostridia (Brouzes et aI., 1971): To 1000 mL of distilled water, add 20 g of glucose, 2 g of sodium acetate

DINITROGEN·FIXING BACTERIA

193

(NaOAc), 0.5 g of calcium carbonate (CaC03 ), 0.01 g of sodium molybdate dihydrate (Na2Mo04·2H20), 0.25 g of potassium dihydrogen phosphate (KH2P0 4), 0.5 g of sodium thioglycollate, 0.75 g of agar, 8 mL of 0.2% aqueous phenosafranin, and 10 mL of soil extract (see chapter 8 in this book). Dispense the medium in test tubes or Pankhurst tubes (Campbell & Evans, 1969), add a small inverted Durham tube to each, cap, and sterilize at 121°C for 15 min. 3. Sterile rubber stoppers (or serum stoppers) for the above tubes of medium.

11-8.3 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as described in chapter 8. Heat the tubes of medium to almost 100 °C to drive off dissolved O 2, and then cool before use. If the medium is freshly autoclaved, merely cool before use. From the highest serial dilution prepared, transfer a 1.0·mL aliquot to each of five tubes of the liquid degassed medium. Then make similar transfers from the successive lower dilutions as desirable. Insert the sterile stoppers tightly, and incubate the tubes at 30°C for 14 d. The headspace of the tubes may be flushed with sterile N2 before stoppering, but this is not absolutely essential. Record as positive those tubes in which at least a 5-mm column of gas has accumulated in the inverted tube and in which the phenosafranin is more or less decolorized. Confirmatory tests that may be applied and that usually correlate well with the above criteria are as follows: microscopic examination of cells from the bottom of a tube for presence of iodine· stainable cells containing glycogen-like reserve material or typical sporulating Clostridium cells, and C 2 H 2 reduction assay carried out by injecting a small amount of sterile glucose into each tube and then, several hours later, injecting enough C 2 H 2 to give 0.01 to 0.1 atm pC 2 H 2 • One day later a sample of the headspace is analyzed by gas chromatography for the presence of two or three times more C 2H 4 than is present in a similarly treated "blank" tube of uninoculated medium (see section 11-2). Calculate the MPN of N 2 -fixing clostridia by referring to the statistical tables found in section 11-2.

11-8.4 Comments Vegetative cells of clostridia, being extremely O 2 sensitive, are killed during the preparation of the usual aerobic dilution series. The method described above, therefore, represents more likely a spore count rather than a count of total viable cells. The count may be improved, and at least some of the viable vegetative cells included, by careful heating to degas the

KNOWLES & BARRAQUIO

194

dilution blanks and flushing the head space with sterile 02-scrubbed N2 before and after each transfer by pipette. Reducing agents (1.0 g of ascorbic acid and 1.0 g of sodium thioglycollate L -1) may also be included in the dilution fluid. The degassed tubes of liquid medium can also be flushed with N2 before and after inoculation. A suitable N2-flushing device is a Pasteur pipette or stainless steel tube attached to a cotton wool or membrane filter and sterilized before attachment to a cylinder of N2. 11-9 METHOD FOR SULFATE-REDUCING BACTERIA 11-9.1 Principles These bacteria are very strict anaerobes that respire by reducing SOi- to H 2S. Desulfovibrio spp. are highly motile, nonsporulating vibrios that are common in terrestrial and aquatic (freshwater and marine) environments. Desulfotomaculum spp. are spore-forming rods with a more restricted distribution, and several other genera also occur (Postgate, 1984; Widdel & Pfennig, 1984). The SOi- reducers do not proliferate unless the redox potential of the medium is below -100 mV, and therefore the Eh of the growth medium is adjusted to a sufficiently low value (Postgate, 1984). Nitrogen fixation by SOi- -reducing bacteria has been demonstrated (Riederer-Henderson & Wilson, 1970; Postgate et aI., 1985) and a N-deficient medium has been described for enumeration (Patriquin & Knowles, 1972). However, the method described below probably does not provide a true count of N2-fixing SOi--reducing bacteria. 11-9.2 Prepared Materials 1. Sterile dilution blanks prepared generally according to chapter 8. However, the dilution fluid should contain 1.0 g of ascorbic acid and 1.0 g of sodium thioglycollate L -1, and the dilution blanks should be heated to degas, flushed with N2, and then stoppered just before use. 2. Nitrogen-deficient lactate medium (modified from Postgate, 1984; Patriquin & Knowles, 1972; Riederer-Henderson & Wilson, 1970): a. Solution A: To 100 mL of distilled water, add 0.5 g of zinc sulfate heptahydrate (ZnS0 4·7H20), 0.5 g of manganous sulfate monohydrate (MnS0 4·H20), 0.4 g of sodium molybdate dihydrate (Na2Mo04·2H20), 0.4 g of ferrous sulfate heptahydrate (FeS04·7H20 ), 0.005 g of cobaltous sulfate (CoS04), 0.005 g of boric acid (H3 B03), and 0.7 mg of cupric sulfate pentahydrate (CuS04·5H20). b. Solution B: To 900 mL of distilled water, add 0.5 g of potassium dihydrogen phosphate (KH2P04), 1.0 g of calcium sulfate (CaS04), 7.0 g of magnesium sulfate heptahydrate

DINITROGEN-FIXING BACTERIA

195

(MgS04 ·7H20), 3.5 g of sodium lactate, and 0.1 g of yeast extract. c. Solution C: To 100 mL of distilled water, add 1.0 g of ascorbic acid and 1.0 g of sodium thioglycollate. Autoclave at 121°C for 15 min. Add 1 mL of solution A to solution B. Adjust the pH to between 7.0 and 7.5, and autoclave. Add the previously sterilized solution C, and immediately dispense into test tubes, each of which contains a small amount of dry-sterilized steel wool. Close with sterile rubber stoppers.

11-9.3 Procedure Collect and prepare the sample as described in chapter 1, and prepare serial dilutions as in chapter 8, taking care to flush with N2 before and after each transfer. From the highest dilution prepared, transfer a 1-mL aliquot to each of five tubes of the liquid medium. Heat the tubes to degas just before inoculation, flush the head space with N2 , and immediately stopper the tube. Make similar transfers from the successive lower dilutions as seems appropriate. Incubate at 25 to 30°C for 2 to 3 wk, and then record as positive those tubes showing a dense black coloration indicative of FeS formation.

REFERENCES Aleem, M.I.H. 1953. Counting of Azotobacter in soils. Plant Soil 4:248-251. Barraquio, W.L., A. Dumont, and R. Knowles. 1988. Enumeration of free-living aerobic N2-fixing H 2-oxidizing bacteria by using a heterotrophic semisolid medium and mostprobable-number tecltnique. Appl. Environ. Microbiol. 54:1313-1317. Becking, J.H. 1961. Studies on nitrogen-fixing bacteria of the genus Beijerinckia. I. Geographical and ecological distribution in soils. Plant Soil 14:49-81. Billson, S., K. Williams, and J.R. Postgate. 1970. A note on the effect of diluents on the determination of viable numbers of Azotobacteraceae. J. Appl. Bacteriol. 33:270-273. Bowien, B., and H.G. Schlegel. 1981. Physiology and biochemistry of aerobic hydrogenoxidizing bacteria. Ann. Rev. Microbiol. 35:405-452. Brouzes, R., and R. Knowles. 1971. Inhibition of growth of Clostridium pasteurianum by acetylene: Implication for nitrogen fixation assay. Can. J. Microbiol. 17:1483-1489. Brouzes, R., c.1. Mayfield, and R. Knowles. 1971. Effect of oxygen partial pressure on nitrogen fixation and acetylene reduction in a sandy loam soil amended with glucose. p. 481-484. In T.A. Lie and E.G. Mulder (ed.) Biological nitrogen fixation in natural and agricultural habitats. Plant Soil, Spec. Vol., Martinus Nijhoff, the Hague. Brown, M.E., S.K. Burlingham, and R.M. Jackson. 1962. Studies on Azotobacter species in soil. I. Comparison of media and techniques for counting Azotobacter in soil. Plant Soil 17:309-319.

Cameron, R.E., and W.H. Fuller. 1960. Nitrogen fixation by some algae in Arizona soils. Soil Sci. Soc. Am. Proc. 24:353-356. Campbell, N.E.R., and H.J. Evans. 1969. Use of Pankhurst tubes to assay acetylene reduction by facultative and anaerobic nitrogen-fixing bacteria. Can. J. Microbiol. 15:13421343.

Chan, Y.K., L.M. Nelson, and R. Knowles. 1980. Hydrogen metabolism of Azospirillum brasilense in nitrogen-free medium. Can. J. Microbiol. 26:1126-1131. Clark, F.E. 1965. Azotobacter. p. 1493-1497. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI.

196

KNOWLES & BARRAQUIO

David, K.A. V., and P. Fay. 1977. Effects of long-term treatment with acetylene on nitrogenfixing microorganisms. Appl. Environ. Microbiol. 34:640-646. De Bont, J.A.M., and M.W.M. Leijten. 1976. Nitrogen fixation by hydrogen-utilizing bacteria. Arch. Microbiol. 107:235-240. De Bont, J.A.M., and E.G. Mulder. 1974. Nitrogen fixation and co-oxidation of ethylene by a methane-utilizing bacterium. J. Gen. Microbiol. 83:113-121. De Bont, J.A.M., and E.G. Mulder. 1976. Invalidity of the acetylene reduction assay in alkane-utilizing, nitrogen-fixing bacteria. Appl. Environ. Microbiol. 31:640-647. Fedorova, R.I., E.1. Milekhina, and N.1. Il'yukhina. 1973. Evaluation of the method of gas metabolism for detecting extraterrestrial life. Identification of nitrogen-fixing microorganisms. Izv. Akad. Nauk SSSR Ser. BioI. 1973(6):797-806. Finke, L.R., and H.W. Seeley, Jr. 1978. Nitrogen fixation (acetylene reduction) by epiphytes of fresh water macrophytes. Appl. Environ. Microbiol. 36:129-138. Gibson, J. 1975. Uptake of C4 dicarboxylates and pyruvate by Rhodopseudomonas sphaeroides. J. Bacteriol. 123:471-480. Hardy, R.W.F., R.D. Holsten, E.K. Jackson, and R.C. Burns. 1968. The acetylene-ethylene assay for N2 fixation: Laboratory and field evaluation. Plant Physiol. 43:1185-1207. Hill, S. 1973. Method for exposing bacterial cultures on solid media to a defined gas mixture using nylon bags. Lab. Pract. 22:193. Hynes, R.K., and R. Knowles. 1978. Inhibition by acetylene of ammonia oxidation in Nitrosomonas europaea. FEMS Microbiol. Lett. 4:319-321. Imhoff, J.F., and H.G. Triiper. 1989. Purple nonsulfur bacteria. p. 1658-1662. In J.T. Staley et al. (ed.) Bergey's manual of systematic bacteriology. Vol. 3. Williams and Wilkins, Baltimore. Jurgensen, M.F., and C.B. Davey. 1968. Nitrogen-fixing blue-green algae in acid forest and nursery soils. Can. J. Microbiol. 14:1179-1183. Knowles, R. 1978. Free-living bacteria, p. 25-40. In J. D6bereiner et al. (ed.) Limitations and potentials for biological nitrogen fixation in the tropics. Plenum Press, New York. Knowles, R. 1980. Nitrogen fixation in natural plant communities and soils. p. 557-582. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen fixation. John Wiley and Sons, New York. Line, M.A., and M.W. Loutit. 1973. Nitrogen-fixation by mixed cultures of aerobic and anaerobic micro-organisms in an aerobic environment. J. Gen. Microbiol. 74:179-180. Malik, K.A., and H.G. Schlegel. 1981. Chemolithotrophic growth of bacteria able to gtow under N2-fixing conditions. FEMS Microbiol. Lett. 11:63-67. Megraw, S.R., and R. Knowles. 1987. Active methanotrophs suppress nitrification in a humisol. BioI. Fert. Soils 4:205-212. Nohrsted, H.-G. 1983. Natural formation of ethylene in forest soils and methods to correct results given by acetylene-reduction assay. Soil BioI. Biochem. 15:281-286. Oakley, C.J., and J.C. Murrell. 1988. nifH genes in the obligate methane oxidizing bacteria. FEMS Microbiol. Lett. 49:53-57. Okon, Y., S.L. Albrecht, and R.H. Burris. 1977. Methods for growing Spirillum /ipoferum and for counting it in pure culture and in association with plants. Appl. Environ. Microbiol. 33:85-88. Patriquin, D., and R. Knowles. 1972. Nitrogen fixation in the rhizosphere of marine angiosperms. Mar. BioI. (Berlin) 16:49-58. Pedrosa, F.O., J. D6bereiner, and M.G. Yates. 1980. Hydrogen-dependent growth and autotrophic carbon dioxide fixation in Derxia. J. Gen. Microbiol. 119:547-551. Postgate, J.R. 1984. The sulphate-reducing bacteria. Cambridge Univ. Press, Cambridge. Postgate, J.R., H.M. Kent, S. Hill, and T.H. Blackburn. 1985. Nitrogen fixation by Desulfovibrio gigas and other strains of Desulfovibrio. p. 225-234. In P.W. Ludden and J.E. Burris (ed.) Nitrogen fixation and CO 2 metabolism. Elsevier Sci. Publ. Co., New York. Raimbault, M. 1975. Etude de l'infiuence inhibitrice de l'acetylene sur la formation biologique du methane dans un sol de riziere. Ann. Microbiol. (Paris) 126A:247-258. Rennie, R.J. 1980. Dinitrogen-fixing bacteria: computer-assisted identification of soil isolates. Can. J. Microbiol. 26:1275-1283. Rennie, R.J. 1981. A single medium for the isolation of acetylene-reducing (dinitrogen-fixing) bacteria from soils. Can. J. Microbiol. 27:8-14. Ridge, E.H. 1970. Effects of some diluents upon viable counts of Azotobacter chroococcum. J. Gen. Appl. Microbiol. 16:189-192.

DINITROGEN-FIXING BACTERIA

197

Riederer-Henderson, M.-A., and P.w. Wilson. 1970. Nitrogen fixation by sulfate-reducing bacteria. J. Gen. Microbiol. 61:27-31. Salkinoja-Salonen, M.S., E. Vaisanen, and A. Peterson. 1979. Involvement of plasmids in the bacterial degradation of lignin-derived compounds. p. 301-314. In K.N. Timmis and A. Ptihler (ed.) Plasmids of medical, environmental and commercial importance. Elsevier/ North Holland Biomedical Press, Amsterdam. Siefert, E., R.L. Irgens, and N. Pfennig. 1978. Phototrophic purple and green bacteria in a sewage treatment plant. Appl. Environ. Microbiol. 35:38-44. Smith, L.A., S. Hill, and M.G. Yates. 1976. Inhibition by acetylene of conventional hydrogenase in nitrogen-fixing bacteria. Nature (London) 262:209-210. Sprent, J.I., and P. Sprent. 1990. Nitrogen fixing organisms. Pure and applied aspects. Chapman and Hall, London. Stewart, W.D.P. 1973. Nitrogen fixation by photosynthetic microorganisms. Annu. Rev. Microbiol. 27:283-316. Strijdom, B.W. 1966. Counting Beijerinckia spp. by a modification of the dilution tube method. S. Afr. Tydskr. Landbouwet. 9:265-266. Tchan, Y.-T. 1984. Azotobacteraceae. p. 219-234. In N.R. Krieg and J.G. Holt (ed.) Bergey's manual of systematic bacteriology. Vol. 1. Williams and Wilkins, Baltimore. Turner, G.L., and A.H. Gibson. 1980. Measurement of nitrogen fixation by indirect means. p. 111-138. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen-fixation. John Wiley and Sons, New York. Van Neil, C.B. 1972. Techniques for the enrichment, isolation and maintenance of the photosynthetic bacteria. p. 3-28. In A.S. San Pietro (ed.) Methods in enzymology. Vol. 23. Academic Press, New York. Veldkamp, H. 1970. Enrichment cultures of prokaryotic organisms. p. 305-361. In J.R. Norris and D.W. Ribbons (ed.) Methods in microbiology. Vol. 3A. Academic Press, New York. Villemin, G., J. Balandreau, and Y. Dommergues. 1974. Utilisation du test de reduction de l'acetylene pour la numeration des bacteries libres fixatrices d'azote. Ann. Microbiol. 24:87-94. Weaver, P.F., J.D. Wall, and H. Gest. 1975. Characterization of Rhodopseudomonas capsulata. Arch. Microbiol. 105:207-216. Whittenbury, R., K;.C. Phillips, and J.T. Wilkinson. 1970. Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61:205-218. Widdel, F., and N. Pfennig. 1984. Dissimilatory sulfate- or sulfur-reducing bacteria. p. 663679. In N.R. Krieg and J.G. Holt (ed.) Bergey's manual of systematic bacteriology. Vol. 1. Williams and Wilkins, Baltimore. Wiegel, J., and H.G. Schlegel. 1976. Enrichment and isolation of nitrogen-fixing hydrogen bacteria. Arch. Microbiol. 107:139-142. Wieringa, K.T. 1968. A new method for obtaining bacteria-free cultures of blue-green algae. Antonie van Leeuwenhoek 34:54-56. Witty, J.F. 1979. Acetylene reduction assay can overestimate nitrogen fixation in soil. Soil BioI. Biochem. 11:209-210. Wong, T.-Y., and R.J. Maier. 1985. Hz-dependent mixotrophic growth of Nz-fixing Azotobacter vinelandii. J. Bacteriol. 63:528-533.

Published 1994

Chapter 12 Legume Nodule Symbionts R. W. WEAVER, Texas A&M University, College Station, Texas PETER H. GRAHAM, University of Minnesota, St. Paul, Minnesota

We use the word rhizobia in this chapter to define a diverse group of organisms united by a common ability to produce root or stem nodules on leguminous plants. To varying degrees, rhizobia possess the ability to fix N independently or in symbiosis with appropriate host legumes. This ability has influenced agricultural systems since the time of the Romans, and is currently of great importance in developing countries, where the price and limited availability of nitrogenous fertilizers can often preclude their use by subsistence farmers. Interest in the rhizobia is also increasing in developed countries as legumes are used more in minimum tillage and sustainable agricultural systems. Because of their importance in agriculture, the early taxonomy of the rhizobia emphasized host nodulation (Baldwin & Fred, 1929; Fred et al., 1932), dividing these organisms into "cross-inoculation groups", isolates from which were thought to nodulate certain legume species, but not others. Such cross-inoculation groups persist as the basis for recommending inoculant cultures, but the demonstration that nodulation, host specificity, and N 2 -fixation genes can be located on transmissible plasmids, has led to the recent reclassification of the rhizobia (Jordan, 1982, 1984) and to the greater use of other phenotypic and phylogenetic traits in their characterization (Graham et al., 1991). Currently, three genera and 11 species of root and stem-nodule bacteria are distinguished, viz:

• Rhizobium leguminosarum biovars leguminosarum, and trifolii (Jordan, 1984) • R. meliloti (Dangeard, 1926) • R. loti (Jarvis et al., 1982) • R. fredii (Scholla & Elkan, 1984; Chen et al., 1988) • R. galegae (Lindstrom, 1989) • R. tropici (Martinez et al., 1991) Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

199

200

WEAVER & GRAHAM

• R. haukuii (Chen et aI., 1991) • R. etli (Segovia et aI., 1993) • Azorhizobium caulinodans (Dreyfus et aI., 1988) • Bradyrhizobium japonicum (Jordan, 1984) • B. elkanii (Kuykendall et aI., 1992) This is still an interim classification based on isolates from nodules of only a small percentage of the 19 700 species of Leguminosae. Better definition among the slow-growing rhizobia is also needed. Current species are defined principally by genotypic similarities and differences (DNN DNA relatedness, rRNNDNA hybridization, 16S ribosomal RNA analysis and DNA restriction length polymorphisms). Colonial and cultural characteristics and symbiotic performance with selected hosts, however, continue as the first parameters needed for the characterization of particular rhizobia. This chapter presents methods commonly used in the study of root and stem-nodule bacteria. Carefully followed, they should enable a microbiologist, agronomist, or geneticist with little previous experience to isolate and culture rhizobia, and to prepare and use inoculant cultures. It should be emphasized, however, that the slow growth rate of these organisms and the lack of suitable selective media can complicate such studies. Guidance from an experienced individual who has worked with these organisms may be needed. 12-1 NODULE COLLECTION AND THE ISOLATION OF SYMBIONTS 12-1.1 Principles The reasons for isolation of rhizobia from nodules may occur as part of a germplasm collection program, may be undertaken to study their diversity in specific field situations, or may be to recover particular genetic recombinants from a mixed population of cells. Date and Halliday (1987) have prepared excellent guidelines and methods for the collection of rhizobial germplasm. Nodules often contain more than a single strain of Rhizobium or Bradyrhizobium. Double strain occupancy (Lindeman et aI., 1974; Moawad & Schmidt, 1987) and nonrhizobial contaminants in the nodule are surprisingly common (Jansen van Rensberg & Strijdom, 1972; Handelsman & Brill, 1985). The method that follows may be used for the isolation of rhizobia from the nodules of either field or greenhouse-grown plants. 12-1.2 Materials

1. Yeast mannitol agar (YMA) (section 12-2.2). 2. 95% Ethanol.

LEGUME NODULE SYMBIONTS

3. 4. 5. 6. 7.

201

Sterile water. 5.25% Sodium hypochlorite (full-strength commercial bleach). Tweezers and scissors. Petri dishes. Nodules. 12-1.3 Procedure

Cut the nodules from the plant, leaving about 0.5 cm of root attached to the nodule for ease in handling. The nodules should be firm and appear in good physical condition. Avoid older nodules that have begun to senesce, as they may contain large populations of secondary organisms. Thoroughly wash the nodules to remove all traces of soil, then immerse them in 95% ethanol for 30 s. Transfer the nodules to 5.25% sodium hypochlorite solution, and leave them immersed in this solution for 4 min. Prolonged exposure to this substance will result in complete sterilization of the rhizobia within the nodule. Remove all of the nodules from the sodium hypochlorite and place them in a petri dish containing 20 to 25 mL of sterile water. To remove all traces of the sterilant, rinse the nodules in five to six changes of sterile water. A series of petri dishes containing 20 to 25 mL of sterile water allows rapid rinsing. Once the nodule is surface disinfected, crush it between blunt-tipped forceps, then mix the nodule contents with a drop of sterile water in the bottom of a sterile petri dish. Streak a loopful of this suspension onto YMA plates and incubate at 28°C for 3 to 5 d for fast-growing rhizobia and 8 to 10 d for bradyrhizobia before further subculture is attempted. Isolates on agar medium should be white to somewhat translucent, circular and raised, and may produce significant amounts of extracellular polysaccharide. Examine wet mounts of cells from selected colonies using a light- or phase-contrast microscope. Cells should be small rod-shaped organisms 1 to 3 ~m in length and 0.5 to 1.0 ~m in diameter. Colonies that are colored or have a distinctive aroma, and bacteria that are distinctly coccoid, produce endospores, or are grouped in chains are not likely to be rhizobia, though Norris (1958) reported a "red" Rhizobium from Lotononis, and colonial dimorphism (Sylvester-Bradley et aI., 1988b) and "doughnut" -shaped colonies have been reported (Howieson et aI., 1988). Select a representative colony having the characteristics of rhizobia, and subculture it onto fresh plates of YMA. If the culture appears pure, it should then be authenticated using host infection (section 12-4) and preserved. 12-1.4 Comments

Fresh nodules should be used for strain isolations wherever possible. If necessary to delay isolation for some time, nodules should be desiccated

in a vial containing silica gel or anhydrous calcium chloride that is covered with a layer of nonabsorbent cotton or cheesecloth to protect the nodules from direct contact with the desiccant. Nodules may be stored in this way

202

WEAVER & GRAHAM

for several weeks before isolation is attempted, but should then be allowed to imbibe water for 1 h prior to surface disinfection. Alternatively, nodules may be frozen soon after collection. They may be left on roots or removed to conserve freezer space. Alternate disinfectants to sodium hypochlorite include 3 to 5% hydrogen peroxide for 3 to 4 min (Vincent, 1970) or acidified mercuric chloride (0.10 g of mercuric chloride and 0.5 mL of concentrated hydrochloric acid in 100 mL of distilled water) for 3 to 4 min. When hydrogen peroxide is used, there is no need for repeated washing in sterile water. Regulations on disposal of mercuric chloride are stringent. Strains of Rhizobium have generation times of 2 to 4 h on YMA medium and produce colonies that are 2 to 4 mm in diameter after 3 to 5 d incubation. Slightly slower growth rates have been reported for A. caulinodans (Dreyfus et at., 1988). Most bradyrhizobia are even slower growing, having a generation time of 6 to 10 h and producing colonies that do not exceed 1 mm diam. after 5 to 7 d growth. Since the growth rate of the rhizobia is slow relative to that of organisms occurring as contaminants in nodule squashes, the inexperienced worker runs the risk of selecting and propagating contaminants, thinking they are rhizobia. It is advisable to leave all isolates from nodules for at least a week before attempting subculture; even longer may be needed with some strains of Bradyrhizobium. Fungal contaminants will often grow faster than the rhizobia, and overrun plates before colonies of the latter organisms are of size sufficient to identify and subculture. To limit this, it is common to include antifungal compounds in the isolation media. Congo red (10 mL of a 1:400 wt/vol aqueous solution per liter, added to the medium before autoclaving) or 0.02 g per liter of filter sterilized cycloheximide added to the medium after autoclaving may be used (Vincent, 1970). A stock solution of cycloheximide may be made by dissolving 2 g of cycloheximide in 200 mL of ethyl alcohol. Agrobacterium also occurs as a contaminant on YMA plates used for nodule isolations and are easily confused with Rhizobium. The two organisms can, however, be distinguished using the ketolactase test (Bernaerts & De Ley, 1963). The rhizobia from many tropical pasture and tree legumes grow best under slightly acid conditions and may not be recoverable using conventional medium. Date and Halliday (1979) used a defined medium containing arabinose, pH 4.5, as the isolation medium for rhizobia from acid soils, while both Gomez de Souza et at. (1984) and Sylvester-Bradley et al. (1988a) used YMA medium acidified to pH 5.5. While isolates are usually authenticated by their ability to nodulate specific legumes, recent papers have reported the isolation from soil of non-infective organisms with the characteristics of Rhizobium (Jarvis et aI., 1989; Soberon-Chavez & Najera, 1989; Segovia et aI., 1991). A significant percentage of such strains acquire the ability to produce nodules following the introduction of a symbiotic plasmid.

203

LEGUME NODULE SYMBIONTS

12-2 CULTIVATION OF NODULE SYMBIONTS 12-2.1 Principles Rhizobia are aerobic to microaerophilic organisms that under most circumstances behave like typical heterotrophs. They will usually require both a source of energy and combined N for growth, and specific vitamin requirements have been shown for some species (Graham, 1963; Elkan & Kwik, 1968; Chakrabarti et aI., 1981). Less typically, specific rhizobia are capable of nitrate-dependent growth under anaerobic conditions (O'Hara & Daniel, 1985; Van Berkum & Keyser, 1985), or are autotrophic (Hanus et aI., 1979) and may even harvest light energy (Eaglesham et aI., 1990; Ladha et aI., 1990). For most purposes, however, the rhizobia can be grown on relatively simple media. These commonly contain mannitol as the energy source, and yeast or yeast extract to supply combined N, vitamins, and minerals. Glycerol (Balatti, 1982) and gluconate (Kuykendall & Elkan, 1976) have also been used to supply energy, while arabinose and galactose may be preferable energy sources where limited pH change in the medium is desired (Date & Halliday, 1979; Howieson, 1985). The majority of slow-growing rhizobia (bradyrhizobia) cannot use either sucrose or lactose as the sole C source. Organic buffers such as MES (pKa = 6.1 at 25°C, useful pH range 5.5-6.7) and HEPES (pKa = 7.5 at 25°C, useful pH range 6.8-8.2) are increasingly being used to maintain medium pH at a desired level (Cole & Elkan, 1973; Howieson, 1985). 12-2.2 Materials Media used for the cultivation of rhizobia. 1. Yeast Mannitol Agar (YMA). There are several variants of this medium. The medium described by Fred and Waksman (1928) contains (g L -1): mannitol, 10; potassium hydrogen phosphate, 0.5; magnesium sulfate heptahydrate, 0.2; sodium chloride, 0.1; calcium carbonate, 0.01; yeast extract powder, 0.5; agar, 15; distilled water to 1000 mL. For yeast extract mannitol broth (YMB), simply leave out the agar. Combine all of the ingredients except the agar in one-half of the water, then adjust the pH to 7.0 with 1 N hydrochloric acid and warm the solution to 55°C. Separately melt the agar in the remaining water by autoclaving 15 min at 0.10 MPa. Mix the two solutions and dispense in test tubes or media bottles as desired. Sterilize the medium by autoclaving as specified above. 2. Peptone Yeast Extract Agar (PYA). A rich medium that is favored for genetic studies because rhizobia tend to produce less extracellular polysaccharide than in YMA. The medium as described by Noel et al. (1984) contains (g L -1): peptone hydrolysate of casein, 5.0; yeast extract

WEAVER & GRAHAM

204

powder, 3.0; calcium chloride dihydrate, 0.147; agar, 15; and distilled water to 1000 mL. Leave out the agar for peptone yeast extract broth (PYB). The medium is prepared, the pH adjusted, and the medium autoclaved as detailed above. 3. Keyser-Munns Agar (KMA). The KMA is a defined medium used in those studies that require a precise knowledge of medium composition (Keyser & Munns, 1979). The medium contains (mg L -1): mannitol, 10 000; magnesium sulfate heptahydrate, 74; calcium chloride dihydrate, 44; iron EDTA, 37; potassium chloride, 0.75; manganese chloride tetrahydrate, 0.20; zinc sulfate septahydrate, 0.12; cupric chloride dihydrate, 0.017; sodium molybdate dihydrate, 0.0048; cobalt nitrate, 0.00037; sodium glutamate, 1100; potassium dihydrogen phosphate, 68; dipotassium hydrogen phosphate trihydrate, 114; and agar, 15000. For this medium, microelements can be prepared as a 100 x stock solution, with 10 mL added per liter. Vitamins may also be necessary for some strains. Vitamins should be prepared separately (thiamine, 1 mg: calcium pantothenate, 1 mg; biotin, 0.1 mg; distilled water, 1000 mL), and filter-sterilized, using an 0.2 ~m filter, before adding to already autoclaved medium at the rate of 1 mL per liter before the plates are poured. For Keyser-Munns broth (KMB) leave the agar out of KMA. A range of additional media are used in studying specific aspects of rhizobial physiology, including melanin (Cubo et al., 1988) and siderophore (Schwyn & Nielands, 1987; Guerinot, 1991) production, and acid-pH tolerance (Howieson, 1985; Graham et al., 1982).

12-2.3 Procedure Cultures of bradyrhizobia and rhizobia should be maintained in a manner that ensures viability, but minimizes the possibility of mutation or contamination. Frequent subculture provides an opportunity for mutation, and marked variation in the N2-fixing ability of single colony isolates from strains is well documented (Herridge & Roughley, 1975; Weaver & Wright, 1987; Gibson et al., 1990). The ideal is to maintain cultures ofthose strains that are in active use, but to replace these at regular intervals from stocks that are only rarely subcultured (section 12-3). For the former, strains may be maintained for several weeks on agar slants, with tightened screw caps, held at room temperature. Most of the bacteria die during storage but sufficient numbers survive for reisolation and culturing. Lowering the storage temperature to 4 °C after adequate growth has occurred increases the time cells may be stored to several months. For the preparation of broth cultures, the quantity of inoculant should be at least 1% of the final number of cells needed, and 5 to 10% is highly desirable. When low inoculant rates are used, a lag period of several days can occur in the growth of the culture, and culture contamination can be a major problem. One to two loopfuls of a recent (few days old) subculture should be used per 25 mL of broth. Adequate aeration must be provided

LEGUME NODULE SYMBIONTS

205

during the growth of broth cultures. This is most conveniently achieved using a laboratory shaker, but when a shaker is not available, air can be passed through nonabsorbent cotton filters, and sparged through the culture. Optimum temperature for rhizobial growth is 28°C, but room temperature is usually adequate. Extended incubation can result in the production of copious amounts of extracellular polysaccharide, and cause problems in the centrifugation of cultures. The PYB and KMB are less prone to this problem than YMB. Rhizobium or Bradyrhizobium numbers in broth culture will usually exceed 109 cells per milliliter. Counts of total organisms can be made using the Petroff-Hausser counter (Hausser Scientific, Bluebell, PA) and the methodologies detailed by Somas ega ran and Hoben (1985). Viable counts require serial dilution of the culture in sterile tap or distilled water or physiological saline, and the plating of aliquots on the surface of YMA plates. 12-3 MAINTENANCE OF CULTURES 12-3.1 Principles Because of the relatively slow growth of rhizobia and the possibilities for contamination and mutation, many research centers work with a threetier system of culture maintenance. In such a system, cultures in active use (see section 12-2) are backed by stock cultures that are stored at 5 °C and sealed to prevent desiccation, while the long-term survival of strains is ensured by their lyophilization (Vincent, 1970), desiccation on porcelain beads (Norris, 1963; Vincent, 1970), or storage at ultracold temperatures using a cryoprotectant (Dye, 1980; Keyser, 1987). Published data in which these methods are compared is lacking, so the actual method selected will commonly depend on the availability of facilities and personal preference. A description of the procedure for long-term cold storage using glycerol as cryoprotectant is given below. 12-3.2 Materials

1. 2. 3. 4.

Cultures on YMA slants. 20% solution (vol/vol) of glycerol in distilled water. 4-mL screw cap cryogenic storage vials. Low-temperature (-15°C) or ultra-cold (-70 0c) freezer. 12-3.3 Procedures

Prepare the 20% glycerol solution and dispense it in 2-mL amounts in cryogenic vials. Sterilize by autoclaving for 20 min at 0.10 MPa (121°C) on the liquid cycle. Once the glycerol is cool, aseptically transfer it to slants or plates containing the culture to be preserved. Cultures of fast-growing rhizobia should be 3 to 4 d old at the time of preservation; those for

WEAVER & GRAHAM

206

slow-growing strains about 7 d old. Scrape the bacterial growth from the surface of the medium with a sterile loop, or sterile glass beads then use a sterile I-mL pipet to transfer bacteria and glycerol back into the cryogenic vial. Vortex the mixture to ensure even dispersion of the cells. Freeze the culture slowly in a freezer at -15°C, then transfer it to an ultracold freezer at -70°C for long-term storage. Multiple vials should be prepared for each strain, with a new vial used each time a fresh culture of the organism is needed. 12-3.4 Comments The concentration of glycerol used in cryoprotection varies from 10 to 50%. Apparently the concentration of glycerol is not critical, nor is it essential that the glycerol be mixed with buffer or nutrient solution. Scraping the growth from slants or plants rather than using broth cultures ensures a larger initial population of organisms in the cryoprotectant, which allows for die-off during freezing, thawing, and storage. The temperature of storage is largely determined by available facilities. Cultures in glycerol will remain viable for more than a year at -20°C, but with lower temperatures, survival can be extended indefinitely. Best results are obtained with cells stored over liquid N in the gaseous phase. Freezing bacteria somewhat slowly (I-2°C per minute) is generally considered less deleterious than rapid freezing, but thawing should be as rapid as possible. Repeated freezing and thawing will result in lysis of cells. Cultures can, however, be taken from the freezer, some frozen material removed for use as inoculum, and the vial replaced in the freezer. When stored at -20°C, glycerol solutions may not be frozen and can be used directly as an inoculum.

12-4 ENUMERATION OF NODULE SYMBIONTS IN SOIL ANDINOCULANTS 12-4.1 Principles While several selective media have been developed for use with bradyrhizobia or rhizobia (Pattison & Skinner, 1974; Barber, 1979; Habte, 1985), the enumeration of these bacteria from soil by routine dilution count procedures is often impractical. Even in selective media, the growth of bradyrhizobia and rhizobia will commonly be less-than that of other soil organisms, particularly pseudomonads. However, where the rhizobial population is large relative to that of the other organisms present, for example in peat inoculants, rhizobia can be counted by serial dilution and plating. Specific serogroups of Rhizobium or Bradyrhizobium soil populations (e.g., serogroup 123 of B. japonicum) can also be approximated using fluorescent antibody methods (Moawad et aI., 1984; McDermott & Gra-

207

LEGUME NODULE SYMBIONTS Table 12-1. Nutrient solutions for growing plants. t Stock solution Without N 1 2 3 4 5 6* 7

With N 8§

Quantity, g L-l

Chemicals

K2S0 4 Mg S04'7H20 KH2P04 K2 HP0 4 CaCl2 CaS0 4 FeCl 3 Na2H 2EDTA H 3B03 MnS0 4·H 2O ZnS0 4·7H2O CaS0 4·5HzO Na 2 Mo0 4·2HzO CoCI2·6H 2O NiCl 2

93 493 23 145 56

KN0 3 (NH4)zS04

10 133

6.5 13 0.23 0.16 0.22 0.08 0.025 0.034 0.022

Quantity of stock L-l 3mL 1 mL 1 mL 1 mL 1g 1 mL

1 mL

1 mL

t The nutrient solution is a modification of Evans et al. (1972). * Dissolve the two chemicals separately before combining. § For nutrient solution containing N substitute stock solution 8 for 1.

ham, 1989; see chapters 6 and 28 by Bottomley and Wright, respectively, in this book). McDermott and Graham (1989) reported recovery efficiencies with this procedure of from 86 to 114%. The method most commonly used to count rhizobia in the presence of other organisms is a variant of the most probable number (MPN) technique described in chapter 5 by Woomer in this book. The material containing rhizobia to be counted is serially diluted, then aliquots of appropriate dilutions are applied to tubes or pouches containing a suitable, aseptically grown host legume. Obviously, the species of legume used will determine the range of rhizobia counted. 12-4.2 Materials 1. Disposable plastic growth pouches (Vaughn Seed Co., Downers Grove,IL). 2. Pouch holders (record holders are convenient). 3. Seed of an appropriate legume. 4. Nitrogen-free nutrient solution (see Table 12-1). 5. Plastic drinking straws. 6. 95% ethanol solution. 7. 1% sodium hypochlorite solution (20-mL household bleach diluted to 100 mL with distilled water).

WEAVER & GRAHAM

208

Fig. 12-1. Arrangement of divided plastic pouches used in making most probable number (MPN) counts of rhizobia.

8. 9. 10. 11.

Sterile water. Water agar for pregerminating seedlings. Samples with the rhizobia to be enumerated. Greenhouse or growth chamber for plant propagation.

12-4.3 Procedure All materials must be rhizobia free, and extreme care taken to prevent the contamination of plant growth units. Pouches arriving from the manufacturer are usually rhizobia free. They may be moistened and autoclaved prior to use. Place the pouches in a record rack or support stand to hold them while seeding, inoculating, and growing seedlings (Fig. 12-1). Commercial plastic pouches are 16 by 17 cm and are provided with a paper wick (Porter et aI., 1966). To economize on space and supplies, remove the paper wicks, and use a plastic heat sealer to divide the pouches into two to four compartments depending on the size of the legume host being used. Cut the paper wicks so that they again fit into the smaller compartments. The wicks should not extend to the top of the pouches as this facilitates

LEGUME NODULE SYMBIONTS

209

fungal contamination and evaporative water loss. As supplied, the upper section of the wick in which the seedling is placed has small perforations for root penetration. A sterile scalpel can be used to make a somewhat larger hole through which seedling radicals can more easily pass. To kill any bradyrhizobia or rhizobia on the surface of seed, soak the seed in 70 to 95% ethanol for 10 min and rinse with water. Soak the rinsed seed in a solution of 1% sodium hypochlorite for 10 min. Rinse the seed five to six times in sterile water to remove any residual disinfectant. Pregerminate the seeds 2 to 3 d on water agar (agar, 10.0 g L -1) or on blotting paper, then transplant to the growth pouches only those seedlings that are free from contamination and show good root development. In transplanting the seeds, use sterile forceps and make sure that the radical passes through the hole made in the upper section of the wick. Plastic drinking straws, cut so that they extend from halfway down the pouch to 1 to 2 cm above it, greatly facilitate watering the pouches, provide rigidity, and help to limit cross-contamination. Use care during the watering of plants to ensure that rhizobia are not transferred via the tip of the watering device or through splashing. Automatic pipetting machines (e.g., Brewer Model SEPCO 60501-40A-SS, BBL, Baltimore, MD) can also be used to speed watering. Add approximately 50-mL sterile nutrient solution (Table 12-1) to each pouch: use proportionately less where the pouches are compartmentalized. It is convenient to prepare stock solutions for the nutrient solution. When preparing stock solution no. 6, it is important to dissolve the two chemicals separately before combining. The CaS0 4 should be added last to avoid formation of precipitates. If the nutrient solution will be autoclaved, the CaS04 should not be added until after autoclaving or an insoluble precipitate will form. Inoculate plants when 6 to 7 d old and well established. Prepare 10fold serial dilutions of the soil whose rhizobial population is to be counted, then inoculate 1 mL of appropriate soil dilutions onto the root systems of four replicate seedlings. Positive and negative controls should be included with each batch of seedlings. The positive control, inoculated with a pure culture of the appropriate rhizobia, will confirm that growth conditions during the experiment were appropriate for nodule development; the negative control is not inoculated but if seedlings become nodulated this will indicate contamination. Growth pouches should be kept under well-lighted conditions in a growth chamber or greenhouse, and at temperatures appropriate to the plant species being used. Vincent (1970) shows a suitable lighting system with fluorescent and incandescent tubes; high-pressure sodium lamps also provide excellent illumination. Plant husbandry and management techniques for some tropical legumes are discussed by Summerfield et al. (1977). Grow the plants for an additional 1 to 2 wk after nodules first appear on the positive controls. Generally, 2 wk after inoculation is adequate for small-seeded legumes, and 3 wk for larger-seeded species. Plant species can vary significantly, however, in the time needed for nodulation. Record

210

WEAVER & GRAHAM

each replicate as positive (one or more nodules present) or negative (no nodules present). The MPN counts of rhizobia in the sample are computed by statistical procedures. A software program has been developed by Woomer et ai. (1990). Statistical tables for MPN counts have been included in several publications (Brockwell, 1963; Alexander, 1965; Vincent, 1970; Brockwell et aI., 1975; see chapter 5 by Woomer in this book).

12-4.4 Comments The MPN count assumes that a single rhizobial cell can multiply and form at least one nodule on the inoculated root. This assumption is not always valid (Thompson & Vincent, 1967; Boonkerd & Weaver, 1982), and populations are often underestimated. Rhizobia must be uniformly dispersed in the diluent for accurate counting; for peat inoculants, Weaver (1979) has reported that rhizobia desorb readily and become dispersed. It is extremely important to use high-quality seed for planting in plastic growth pouches. Seed of poor quality will not perform well in growth pouches and considerable fungal contamination may occur. It is relatively easy to kill rhizobia on the surface of seed using ethanol or sodium hypochlorite. To actually surface sterilize seed is much more challenging and generally not necessary. If surface sterilization is desired, refer to methods described by Caetano-Anolles et ai. (1990). The host used in the assay can affect the results obtained in MPN counts. Thus, Macroptilium atropurpureum is often used to assay Bradyrhizobium populations in soil, but can overestimate the number of soil rhizobia infective on a particular legume because of its wide host range. Kumar Rao et ai. (1982) reported 190 000 rhizobia per gram in a Kashmir soil when Macroptilium was used as host, but only 3270 when a pigeons pea [Cajanus cajan (L.) Huth] cultivar was used. Glycine ussuriensis is preferable to G. max for counting Bradyrhizobium japonicum because of its small seed size (Brockwell et aI., 1975). Light can affect rooting and nodulation in several legumes, including peanuts, necessitating that the lower portions of pouches or tubes be shielded.

12-5 INOCULANTS FOR FIELD EXPERIMENTATION 12-5.1 Principles Practical investigations with rhizobia frequently culminate with greenhouse or field experiments to determine the need for inoculation, or to observe the effects of inoculation on plant growth or yield. For greenhouse or growth-chamber studies, a broth culture of the inoculant strain will

LEGUME NODULE SYMBIONTS

211

usually be adequate, and may be directly inoculated onto the seed at planting. Even under these conditions, it is important that the rate of inoculation used be a realistic one. Takats (1986) and others have reported reduced nodulation at heavy inoculant dose levels. For field experiments, the large number of seeds used will often require a more sophisticated delivery system, with the inoculant prepared in advance of the planting date and required to maintain high rhizobial numbers for as long as 6 mo. Commercial inoculants are available in most countries, and some inoculant manufacturers will custom prepare peat inoculants using specific strains. With proper storage, purchased peat inoculants will usually contain adequate numbers of rhizobia. Recommended inoculant strains for particular host legumes are shown in Table 12-2. Factors that must be considered in preparing inoculants include suitability of the inoculant carrier, the method of inoculation to be practiced, environmental factors likely to influence rhizobial survival during shipment and application, and additional seed treatments. Peat is the preferred inoculant carrier and can usually be obtained or purchased directly from the inoculant company to ensure its suitability. In some countries, inoculantquality peat may not be available, and a peat source or alternate inoculant carrier may need to be identified and tested. Vegetable oils, charcoal, bagasse, coir, and compos ted organic material have all proved suitable (Corby, 1976; Deschodt & Strijdom, 1976; Philpotts, 1976; Halliday & Graham, 1978; Pazkowski & Berryhill, 1979; Kremer & Peterson, 1983; Graham-Weiss et aI., 1987; Beck, 1991), but each material must be separately tested before use. Methods used in testing such carriers have been detailed in several publications (Roughley & Vincent, 1967; Roughley, 1968) and are beyond the scope of this chapter. Because peat is a proven inoculant carrier and readily available from inoculant manufacturers in the USA, the method described in the next section uses peat as the carrier. 12-5.2 Materials

1. Peat. Peat carriers may be supplied in two grades; a very fine peat with 70 to 95% passing through a 200-mesh sieve for the preparation of seed-applied inoculants, and a more granular preparation (16-40 mesh) for application in the seed furrow (Burton, 1982). 2. Finely ground limestone. 3. Culture of Rhizobium or Bradyrhizobium. 4. Sterile polyethylene bags for packaging the inoculant. 12-5.3 Procedure Most peats are somewhat acid, and finely ground limestone may need to be added to raise the pH toward neutrality. Limestone additions should not exceed 10% by weight of the peat. If only small quantities of peat are needed, the simplest approach may be to buy a commercial peat-based

212

WEAVER & GRAHAM

Table 12-2. Some recommended strains of rhizobium for selected legumes. t Legume

Strain(s)

Aeschynomene falcata Arachis hypogaea A. pintoi Astragalus sinicus Cajanus cajan Calopogonium sp. Centrosema sp. Cicer arietinum Coronilla varia Crotalaria paulina Cyamopsis tetragonolobus Desmodium heterocarpon D. heterophyllum D. intortum Glycine max

CB 2312 CB 756, USDA 3339, USDA 3341 CB 756, CIAT 3101 USDA 3466 CB 756, USDA 3384, USDA 3474 CB756 CB 1923, CIAT 1670, CIAT 3101 CB 2855, USDA 3378, USDA 3379 CB 2012, USDA 3165, USDA 3167 USDA 3384 CB 3035 CIAT 3418 CB 2085, CIAT 3469 SEMIA 656 SEMIA 6003 CB 1809, USDA 110, USDA 142 SEMIA 587, SEMIA 5018 SEMIA 344, SEMIA 360 CB 81, USDA 3404, CIAT 1967 CB 376 CB 2938, SEMIA 806, SEMIA 816 CB 2270 SEMIA 805, SEMIA 806 SEMIA 821 CB 2026 CB 756, SEMIA 656 USDA 1057 SEMIA 138 USDA 1011, USDA 1021a, USDA 1025 SEMIA 115, SEMIA 116 CB 3061 CB2000 CB 756, CIAT 2434 USDA 3259 CB 2899, USDA 2667, USDA 2669, USDA 2674, CIAT 166, CIAT 632, SEMIA 491 CB 1447, USDA 2370, SEMIA 335, SEMIA 374 CB 2898, CIAT 870, CIAT 995, CIAT 2138 CB 756, CIAT 71 CB 756, CB 2126 CB 756 CB 756 CB 2937 CB 2937, SEMIA 208c CB 1990, SEMIA 222 CB 1990, SEMIA 235 CB 787, SEMIA 280 CB 2937, SEMIA 222 SEMIA 354, SEMIA 366 CB 1015 USDA 3447 USDA 3454, SEMIA 634, SEMIA 656 CIAT71

Lens esculenta Leucaena leucocephala Lotononis bainesii Lotus corniculatus L. pedunculatis L. tenuis L. uliginosus Lupinus sp. Macroptilium atropurpureum Medicago lupulina M. polymorphum M. sativa Medicago sp. Onobrychis sativa Peuraria sp. Phaseolus lunatus P. vulgaris Pisum sativum Stylosanthes capitata S. guyanensis S. hamata S. humilis S. scabra Trifolium cherleri T. incarnatum T. pratense T. repens T. semipilosum T. subterraneum Vicia sativa Vigna mungo V. radiata V. unguiculata Zornia sp.

t Recommendations are taken from the CSIRO Division of Tropical Pastures CB Rhizobium Strain Catalogue (1984); the USDA Beltsville Rhizobium Culture Collection Catalog (1987); the CIAT Catalogue of Rhizobium strains for tropical forage legumes (1986); and the Catalogue of Rhizobium strains of the MIRCEN centre, Porto Alegre, Brazil. Addresses for each collection are included in Takishima et al. (1989).

LEGUME NODULE SYMBIONTS

213

inoculant, and autoclave it before adding the strain of interest. Autoclaving can sometimes affect the suitability of peat as an inoculant carrier, but this is not known to be a problem with the peats used by manufacturers in the USA. Grow the inoculant strain in YMB until mid- to late-log phase, as detailed in section 12-2.3, then thoroughly mix approximately equal parts by weight of culture broth and dry peat. Nonsterile peat will generate heat on mixing, so it should be layered to a depth of no more than 2 to 5 cm on trays, and allowed to mature for at least 24 h before packaging. For sterile peat, aseptically transfer 50 to 100 g to a sterile polyethylene bag, then add an equal weight of culture broth. Alternatively, smaller inoculant doses can be applied to already moistened and sealed bags (Somasegaran, 1985), mix the contents by gently squeezing the bag. After mixing and maturation, peats should contain at least 100 million rhizobia per gram, and be moist (0.03 MPa tension) but not saturated. The polyethylene bags used should have a wall thickness of approximately 0.0375 mm and permit some aeration (Roughley, 1976). Store the inoculant under cool conditions until used. 12-5.4 Comments It is important to confirm that the inoculant is of high quality before beginning field experiments. The culture used to inoculate the peat should be checked for contamination, and the viable cell count determined by plate counting. Where some time has elapsed between the preparation and use of the inoculant, plate counts should also be made at planting to verify the level of inoculation.

12-6 INOCULATION OF SEED 12-6.1 Principles

The aim of inoculation is to ensure that the rhizobia applied are sufficient to ensure rapid and abundant nodulation of the host legume. The number required will vary with host and environment, but should not be < 5000 rhizobia per seed. Two basic approaches are used. In one, the inoculant is applied directly to the seed; in the other, it is applied to the soil in the vicinity of the seed. The former method is the more practical in most situations, but has limitations where unfavorable conditions (high soil temperature, acid soil pH, and competition from native soil rhizobia) necessitate the use of higher than normal inoculation rates. In such cases, granular or liquid inoculants are commonly used and applied directly to the soil. For seed-applied rhizobia, it is important to use an adhesive to maintain close contact between rhizobia and the seed, and to reduce the rate of

214

WEAVER & GRAHAM

decline in rhizobial numbers per seed (Vincent, 1970; Materon & Weaver, 1984). A method for seed inoculation is described below. 12-6.2 Materials 1. Seed. 2. Inoculant (see section 12-4.3 for preparation). 3. Adhesive (gum arabic, methyl ethyl cellulose, or commercial product). 4. Finely ground rock phosphate or calcium carbonate. 12-6.3 Procedure Since seed size varies from < 1 mg (Trifolium repens L.) to more than 1000 mg (Stizolobium) per seed, with the surface area per gram of the small-seeded legume significantly greater than for larger-seeded types, the amount of inoculant culture and adhesive applied per kilogram of seed will vary greatly. For small-seeded legumes, use 20 mL of adhesive solution and 109 of peat per kilogram of seed: for larger seeded legumes use 10 mL of adhesive and 5 g of inoculant per kilogram seed. Prepare the adhesive by dissolving 40 g of gum arabic or 5 g of methyl ethyl cellulose per 100 mL of water. Some samples of gum arabic are red and strongly acid and should be avoided. Mix the inoculant culture and adhesive, then apply the inoculant suspension to the seed, mixing until the seed is uniformly coated. Spread the inoculated seed out in a cool location to dry. While the rhizobia should stay viable for several days, it is better to only prepare as much inoculant as needed for the following day. Where seeds are to be planted into acid soil, or face other stresses before germination, it may be advisable to pellet the seed. This protects the rhizobia from temperature and desiccation, and during germination creates a microclimate around the seed that permits nodulation under even quite acid soil conditions (Loneragan et aI., 1955; Graham et aI., 1974; Philpotts, 1977). Finely ground rock phosphate is usually used as the pelleting material for slow-growing rhizobia and calcium carbonate for the fast growers. Gum arabic or methyl ethyl cellulose must be used as the adhesive; most other substances do not adequately bind the pelleting material. Mix the inoculant, adhesive, and seed as before, then add 200 g of the pelleting agent per kilogram seed. Mix rapidly but smoothly for 1 to 2 min, by which time all of the seeds should be evenly coated and separated. Care should be taken in the use of pesticides or molybdenum in association with legume inoculants. While most herbicides and insecticides will show little effect on rhizobia at commercial rates of application, fungicides such as captan are usually lethal (Curley & Burton, 1975; Graham et aI., 1980), and several others will reduce nodulation on prolonged contact with rhizobia. Seed-dressing with Mo can also be toxic to rhizobia

215

LEGUME NODULE SYMBIONTS

(Graham et aI., 1974; Gault & Brockwell, 1980). Where seed-dressing is required for control of root pathogens, granular or liquid inoculant preparations can be used and placed directly in the seed furrow (Weaver & Frederick, 1974; Boonkerd et aI., 1978). Soil-applied inoculants may also be used for crops such as peanuts where the seed coat is fragile, and wetting the seed is likely to damage seed viability. Such inoculants are usually used at the rate of 20 mL of liquid culture or 1 g of granular inoculant per meter of row. Both can be added mechanically via a tube attached behind the furrow opener and connected to a reservoir of the inoculant, or they can be dribbled into the seed row by hand. In each case, the inoculant should lie in the furrow with the seed or 1 to 2 cm below it.

12-7 FIELD EXPERIMENTATION INVOLVING INOCULATION 12-7.1 Principles Field studies with the rhizobia may consider whether inoculation of a particular crop species is necessary, screen promising strains or cultivars for symbiotic response, or consider the effect of environmental, management, or soil factors on some phase of the symbiosis. Because of such differences, it is not possible to provide detailed methodologies in this chapter. Many of the principles involved, however, are similar and should be discussed. Control treatments should be included in studies seeking response to inoculation. An uninoculated treatment will show whether the soil contains indigenous rhizobia and will give an estimate of their effectiveness. It will also provide nodule samples that can be used to characterize indigenous rhizobia by serology or other means. Nitrogen fertilized treatments will show that other constraints to production have been controlled and that response to N addition occurs. This treatment may not be practical for legume-grass mixtures, where use of fertilizer N can change the relative growth of grass and legume. Several major and minor elements are needed for N2 -fixation (Robson, 1978; O'Hara et aI., 1988). Soil at the study site should be sampled, and its chemical analysis undertaken. Where fertility is not part of the study, deficiencies in any of the needed elements should be corrected by appropriate fertilization. Plot size will vary with the plant species being studied and the planned duration and objectives of the experiment, but will commonly provide one or more small areas for subsampling during the growing season, as well as a larger area for yield determinations. For row crops, it is normal to leave at least two rows on either side of the area to be harvested. A further 50 cm to 1 m should be left at the ends of the plot and between areas that will be sampled during the growing season. This is both to limit border effects and reduce the effects of harvest damage on subsequently sampled plants.

216

WEAVER & GRAHAM

The area harvested for yield should be 2 to 4 m 2. A randomized complete block with four to five replications is desirable. This is because several parameters measured in these studies can show high variation. One-meter strips must be left between the plots to ensure that inoculants from one plot do not contaminate others. The parameters measured in inoculation studies will vary with the experiment and may include plant growth and N accumulation measured at several growth stages along with nodule mass, nodule distribution on the root system, and nodule occupancy. Biological yield and N content at physiological maturity is an important parameter in crop legumes where both cultivar and strain can influence N harvest index; yield and N content at each cutting are essential measures in pasture legumes. For nodule determinations, 10 plants per replicate should be sampled to limit plantto-plant variation. Exercise care in digging the samples and cleaning soil from the roots. Nodules detach easily from the root and may be lost. In heavy-textured soils, placing the dug root/soil ball into a bucket of water and letting soak will aid in nodule recovery. Divide the root system into four to five segments (upper and lower tap root, two to three lateral root segments), and if acetylene reduction assays (see chapter 43 by Weaver and Danso in this book) are included do them by root section rather than for the plant as a whole. Following such assays, pick the nodules from each root section, weigh them, and preserve them for strain identification by using serology (see chapter 28 by Wright in this book) or other techniques. Visual ratings of nodulation may be used in breeding nurseries where several samples precludes physical removal and weighing of all plants (Rosas & Bliss, 1986), but is not recommended for smaller studies. Nodule number is more practical and usually of less value than nodule mass. It is an important parameter where environmental conditions (soil acidity, high soil temperature, and fungicide treatment of the seeds) could have affected strain survival in the inoculant and soil. McDermott and Graham (1989) found inoculant strains competitive for nodulation sites in the crown region where the inoculant was placed but of limited mobility and so unable to contribute significantly to lateral root nodulation. Crown nodules produced by the inoculant strain fixed essentially 100% of the N accumulated early in the growth cycle, but most N2 fixation post-flowering was due to nodules on the lateral roots. 12-8 GROWTH-POUCH INFECTION ASSAYS 12-8.1 Principles Bhuvaneswari et al. (1980, 1981) used a modification of the growth pouch procedure for the determination of MPN counts (section 12-5) to study infection. In this procedure, the position of the root tip of growthpouch grown plants is marked at the time of inoculation, and nodulation events are scored relative to the root-tip mark (RTM).

217

LEGUME NODULE SYMBIONTS

Infection in most legumes occurs predominantly in the region in which root hairs are just beginning to differentiate. This is located some 1 to 2 cm behind the root tip, with the developing root hairs only receptive to infection for a period of 4 to 6 h. If conditions for infection are ideal and host and rhizobia are compatible, nodules will be produced in the region above the RTM made at the time of inoculation. If inoculant density is low, if the nutrient solution is too acid, or if the host and strain are limited in compatibility, the uppermost nodules will usually occur below the RTM position. 12-8.2 Materials With slight modifications, the materials used in this procedure are the same as detailed for the MPN procedure (section 12-5.2). 12-8.3 Procedure Moisten the wick material of the growth pouches with distilled water. Lay pouches containing wicks fiat and use a roller to remove air bubbles or bulges. Group the pouches in sets of 25, wrap them in aluminum foil and sterilize normally. Avoid wrinkles by keeping the pouches fiat during autoclaving. Disinfect seeds as described previously (section 12-5.3), then pregermin ate them on water agar until the radicle is a few millimeters long. For large-seeded legumes, the radicle is strong and may be more than l-cm long when planted but for small-seeded legumes the radicle is fragile and should not be more than 2- to 3-mm long when planted. Place the growth pouches in support racks, then aseptically plant two seedlings per pouch, making sure that the root system of each protrudes through the folded section of the paper wick and is in contact with the wick below the fold. Keep the wicks moist by adding 2 to 5 mL of nutrient solution as needed. Prepare the inoculant as previously described, adding 1 mL of a suspension containing 100 000 cells per mL directly to each root in the pouch. Mark the root tip position at the time of inoculation on the outside of the growth pack with a fine tip marker. Grow the plants in a growth chamber or greenhouse until well nodulated, then determine for 20 to 25 replicate pouches per treatment the distance of the uppermost nodule from the RTM made at the time of inoculation. Record also the number of nodules produced above the RTM and the percentage of plants with nodules above the RTM. 12-8.3 Comments Recent studies have shown that speed in nodulation as determined by the RTM methodology correlates closely with the competitiveness of strains determined in pot studies (Stephens & Cooper, 1988; McDermott &

218

WEAVER & GRAHAM

Graham, 1990; Oliveira & Graham, 1990). McDermott et al. (1991) have even found that populations of B. japonicum serogroup 123 from Waukegan (fine-silty over sandy or sandy-skeletal, mixed, mesic Typic Hapludolls) and Webster (fine-loamy, mixed, mesic Typic Haplaquolls) soils showed the same competitive performance in growth pouches as in the field, while Chaverra and Graham (1991) have noted cultivar variation in several traits affecting speed of nodulation. The procedure outlined above has a high coefficient of variation and is the reason for 20 to 25 replications being needed. Further, nodulation may not occur if the developing root system is not in contact with the paper wick. It is for this reason that the root system is kept moist, rather than flooded, and the growth pouches are lain flat until planted.

REFERENCES Alexander, M. 1965. Most-probable number method for microbial populations. p. 14671472. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Baldwin, I.L., and E.B. Fred. 1929. Nomenclature of the root-nodule bacteria of the Leguminosae. J. Bacteriol. 17:141-150. Balatti, A.P. 1982. Culturing Rhizobium in large scale fermentors. p. 127-132. In P.H. Graham and S.C. Hams (ed.) Biological mtrogen fixation technology for tropical agriculture. CIAT, Cali, Colombia. Barber, L.E. 1979. Use of selective agents for recovery of Rhizobium meliloti from soil. Soil Sci. Soc. Am. J. 43:1145-1148. Beck, D.P. 1991. Suitability of charcoal-amended mineral soil as carrier for Rhizobium inoculants. Soil BioI. Biochem. 23:41-44. Bernaerts, M.J., and J. De Ley. 1963. A biochemical test for crown gall bacteria. Nature (London) 197:406-407. Bhuvaneswari, T.Y., A.A. Bhagwat, and W.D. Bauer. 1981. Transient susceptibility of root cells in four common legumes to nodulation by rhizobia. Plant Physiol. 68:1144-1149. Bhuvaneswari, T.Y., B.G. Turgeon, and W.D. Bauer. 1980. Early events in the infection of soybean (Glycine max (L.) Merr) by Rhizobium japonicum. Plant Physiol. 66:10271031. Boonkerd, N., and R. W. Weaver. 1982. Cowpea rhizobia: Comparison of plant infection and plate counts. Soil BioI. Biochem. 14:305-307. Boonkerd, N., D.F. Weber, and D.F. Bezdicek. 1978. Influence of Rhizobium japonicum strains and inoculation methods on soybeans grown in rhizobia-populated soil. Agron. J. 70:547-549. Brockwell, J. 1963. Accuracy of a plant infection technique for counting populations of Rhizobium trifolii. Appl. Microbiol. 11:377-383. Brockwell, J., A. Diatloff, A. Grassia, and A.C. Robinson. 1975. Use of the wild soybean (Glycine ussuriensis Regel and Moack) as a test plant in dilution-nodulation frequency tests for counting Rhizobium japonicum. Soil BioI. Biochem. 7:305-311. Burton, J.C. 1982. Modern concepts in legume inoculation. p. 105-114. In P.H. Graham and S.C. Harris (ed.) Biological nitrogen fixation technology for tropical agriculture. CIAT, Cali, Colombia. Caetano-Anolles, G., G. Favelukes, and W.D. Bauer. 1990. Optimization of surface sterilization for legume seed. Crop Sci. 30:708-712. Chakrabarti, S., M.S. Lee, and A.H. Gibson. 1981. Diversity in the nutritional requirements of strains of various Rhizobium species. Soil BioI. Biochem. 13:349-354. Chaverra, M.H., and P.H. Graham. 1991. Cultivar variation in traits affecting the early nodulation of common bean. Crop Sci. 32:1432-1436. Chen, W.X., G.S. Li, Y.L. Qi, E.T. Wang, H.L. Yuan, and J.L. Li. 1991. Rhizobium huakuii sp. nov. isolated from the roots of Astragalus sinicus. Int. J. Syst. Bacteriol. 41:275-280.

LEGUME NODULE SYMBIONTS

219

Chen, W.X., G.H. Yan, and J.L. Li. 1988. Numerical taxonomic study of fast-growing soybean rhizobia and a proposal that Rhizobium fredii be assigned to Sino rhizobium gen. nov. lnt. J. Syst. Bacteriol. 38:392-397. Cole, M.A., and G.H. Elkan. 1973. Transmissible resistance to penicillin G, neomycin and chloramphenicol in Rhizobium japonicum. Antimicrob. Agents Chemother. 4:248-253. Corby, H.D.L. 1976. A method of making a pure culture, peat-type inoculant, using a substitute for peat. p. 169-173. In P.S. Nutman (ed.) Symbiotic nitrogen fixation in plants. Cambndge Univ. Press, London. Cubo, M.T., A.M. Buendia-Claveria, J.E. Beringer, and J.E. Ruiz-Sainz. 1988. Melanin production by Rhizobium strains. Appl. Environ. Microbiol. 54: 1812-1817. Curley, R.L., and J.e. Burton. 1975. Compatibility of Rhizobium japonicum with chemical seed protectants. Agron. J. 67:807-808. Dangeard, P.A. 1926. Reserches sur les tubercles radicaux des legumineuses. Le Botaniste (Paris) 16:1-270. Date, R.A., and J. Halliday. 1979. Selecting Rhizobium for the acid, infertile soils of the tropics. Nature (London) 277:62-64. Date, R.A., and J. Halliday. 1987. Collection, isolation, cultivation and maintenance of rhizobia. p. 1-27. In G.H. Elkan (ed.) Symbiotic nitrogen fixation technology. Marcel Dekker, New York. Deschodt, C.C., and B.W. Strijdom. 1976. Suitability of a coal bentonite base as carrier of rhizobia in inoculants. Phytophylactica 8:1-6. Dreyfus, B., J.L. Garcia, and M. Gillis. 1988. Characterization of Azorhizobium caulinodans gen. nov., sp. nov., a stem-nodulating nitrogen-fixing bacterium isolated from Sesbania rostrata. lnt. J. Syst. Bacteriol. 38:89-98. Dye, M. 1980. Functions and maintenance of a Rhizobium collection. p. 435-471. In N.S. Subba Rao (ed.) Recent advances in biological nitrogen fixation. Edward Arnold, London. Eaglesham, A.R.J., J.M. Ellis, W.R. Evans, D.E. Fleischman, M. Hungria, and R.W.F. Hardy. 1990. The first photosynthetic N2-fixing Rhizobium. Characteristic. p. 805-811. In P.M. Gresshoff et al. (ed.) Nitrogen fixation: Achievements and objectives. Chapman and Hall, New York. Elkan, G.H., and l. Kwik. 1968. Nitrogen, energy and vitamin nutrition of Rhizobium japonicum. J. Appl. Bacteriol. 31:399-404. Evans, H.J., B. Koch, and R. Klucas. 1972. Preparation of nitrogenase from nodules and separation into components. Methods Enzymol. 24:470-476. Fred, E.B., l.L. Baldwin, and E. McCoy. 1932. Root nodule bacteria and leguminous plants. Univ. of Wisconsin Press, Madison. Fred. E.B., and S.A. Waksman. 1928. Laboratory manual of general microbiology, with special reference to the microorganisms of the soil. McGraw-Hill, New York. Gault, R.R., and J. Brockwell. 1980. Effects of the incorporation of molybdenum compounds in the seed pellet on inoculant survival, seedling nodulation and 81ant growth of lucerne and subterraneum clover. Aust. J. Exp. Agric. Anim. Husb. 2 :63-71. Gibson, A.H., D.H. Demezas, R.R. Gault, T.Y. Bhuvaneswari, and J. Brockwell. 1990. Genetic stability in rhizobia in the field. Plant Soil 129:37-44. Gomez de Souza, L.A., F.M.M. Magalhaes, and L.A. de Oliveira. 1984. Avaliacao do crescimento de Rhizobium de leguminosas florestais tropicais em diferentes meios de cultura. Pesq. Agropec. Bras. 19s/n:165-168. Graham, P.H. 1963. Vitamin requirements of root-nodule bacteria. J. Gen. Microbiol. 30:215-218. Graham, P.H., V.M. Morales, and R. Cavallo. 1974. Materiales excipientes y adhesivos de posible uso en inoculacion de leguminosas en Colombia. Turrialba 24:47-50. Graham, P.H., V.M. Morales, and O. Zambrano. 1974. Seed pelleting of a legume to apply molybdenum. Turrialba 24:335-336. Graham, P.H., G. Ocampo, L.D. Ruiz, and A. Duque. 1980. Survival of Rhizobium phaseoli in contact with chemical seed protectants. Agron. J. 72:625-627. Graham, P.H., M.J. Sadowsky, H.H. Keyser, Y.M. Barnet, R.S. Bradley, J.E. Cooper, J. DeLey, B.D.W. Jarvis, E.B. Roslycky, B.W. Strijdom, and J.P.w. Young. 1991. Proposed minimal standards for the description of new genera and species of root and stem-nodulating bacteria. lnt. J. Syst. Bacteriol. 41:582-587. Graham, P.H., S.E. Viteri, F. Mackie, A.T. Vargas, and A. Palacios. 1982. Variation in acid soil tolerance among strains of Rhizobium phaseoli. Field Crops Res. 5:121-128.

220

WEAVER & GRAHAM

Graham-Weiss, L., M.L. Bennett, and A.S. Paau. 1987. Production of bacterial inoculants by direct fermentation on nutrient supplemented vermiculite. Appl. Environ. Microbiol. 53:2138-2140. Guerinot, M.L. 1991. Iron uptake and metabolism in the rhizobia/legume symbiosis. Plant Soil 130:199-209. Habte, M. 1985. Selective medium for recovering specific populations of rhizobia introduced into tropical soils. Appl. Environ. Microbiol. 50:1553-1555. Halliday, J., and P.H. Graham. 1978. Comparative studies of peat and coal as inoculant carriers for Rhizobium. Turrialba 28:348-349. Handelsman, J., and W.J. Brill. 1985. Erwinia herbicola isolates from alfalfa plants may play a role in nodulation of alfalfa by Rhizobium meliloti. Appl. Environ. Microbiol. 49:818821. Hanus, F.J., R.J. Maier, and H.J. Evans. 1979. Autotrophic growth of H2 uptake positive strains of Rhizobium japonicum in an atmosphere supplied with hydrogen gas. Proc. Natl. Acad. Sci. USA 76:1788-1792. Herridge, D.P., and R.J. Roughley. 1975. Variation in colony characteristics and symbiotic effectiveness of Rhizobium. J. Appl. Bacteriol. 38:19-27. Howieson, J.G. 1985. Use of an organic buffer in the selection of acid tolerant Rhizobium meliloti. Plant Soil 88:367-376. Howieson, J.G., M.A. Ewing, and M.F. D'Antuono. 1988. Selection for acid tolerance in Rhizobium meliloti. Plant Soil 105:179-188. Jansen van Rensburg, H., and B.W. Strijdom. 1972. Information on the mode of entry of a bacterial contaminant into nodules of some leguminous plants. Phytophylactica 4:73-78. Jarvis, B.D.W., C.E. Pankhurst, and J.J. Patel. 1982. Rhizobium loti, a new species of legume root-nodule bacteria. Int. J. Syst. Bacteriol. 32:378-380. Jarvis, B.D. W., L.J .H. Ward, and E.A. Slade. 1989. Expression by soil bacteria of nodulation genes from Rhizobium leguminosarum. Appl. Environ. Microbiol. 55:1426-1434. Jordan, D.C. 1982. Transfer of Rhizobium japonicum Buchanan 1980 to Bradyrhizobium gen. nov. a ~enus of slow growing root-nodule bacteria from leguminous plants. Int. J. Syst. Bactenol. 2:136-139. Jordan, D.C. 1984. Family III. Rhizobiaceae Conn 1938. p. 234-244. In Bergey's manual of systematic bacteriology. Williams and Wilkins, Baltimore. Keyser, H.H. 1987. The role of culture collections in biological nitrogen fixation. p. 413-428. In G.H. Elkan (ed.) Symbiotic nitrogen fixation technology. Marcel Dekker, New York. Keyser, H.H., and D.N. Munns. 1979. Effects of calcium, manganese and aluminum on growth of rhizobia in acid media. Soil Sci. Soc. Am. J. 43:500-503. Kremer, R.J., and H.L. Peterson. 1983. Effects of carrier and temperature on survival of Rhizobium spp. in legume inocula: Development of an improved type of inoculant. Appl. Environ. Microbiol. 45:1790-1794. Kumar Rao, J.V.D.K., P.J. Dart, and M.U. Khan. 1982. Cowpea-group Rhizobium in soils of the semi-arid tropics. p. 291-295. In P.H. Graham and S.C. Harris (ed.) Biological nitrogen fixation technology for tropical agriculture. CIAT, Cali, Colombia. Kuykendall, L.D., and G.H. Elkan. 1976. Rhizobium japonicum derivatives differing in nitrogen-fixing efficiency and carbohydrate utilization. Appl. Environ. Microbiol. 32:511-519. Kuykendall, L.D., B. Saxena, T.E. Devine, and S.E. Udell. 1992. Genetic diversity in "Bradyrhizobium japonicum" Jordan 1982 and a proposal for "Bradyrhizobium elkanii" sp.-nov. Can. J. Microbiol. 38:501-505. Ladha, J.K., R.P. Pareek, R. So, and M. Becker. 1990. Stem nodule symbiosis and its unusual properties. p. 633-640. In P.M. Gresshoff et al. (ed.) Nitrogen fixation: Achievements and objectives. Chapman and Hall, New York. Lindemann, W.C., E.L. Schmidt, and G.E. Ham. 1974. Evidence for double infection within soybean nodules. Soil Sci. 118:274-279. Lindstrom, K. 1989. Rhizobium galegae, a new species of legume root-nodule bacteria. Int. J. Syst. Bacteriol. 39:365-367. Loneragan, J.F., D. Meyer, R.G. Fawcett, and A.J. Anderson. 1955. Lime pelleted clover seeds for nodulation on acid soils. J. Aust. Inst. Agric. Sci. 21:264-265. Martinez, E., L. Segovia, F. Martins, A.A. Franco. P. Graham, and M.A. Pardo. 1991. Rhizobium tropici: A novel species nodulating Phaseolus vulgaris L. beans and Leucaena spp. trees. Int. J. Syst. Bacteriol. 41:417-426.

LEGUME NODULE SYMBIONTS

221

Materon, L.A., and R.W. Weaver. 1984. Survival of Rhizobium trifolii on toxic and non-toxic arrowleaf clover seeds. Soil BioI. Biochem. 16:533-535. McDermott, T.R., and P.H. Graham. 1989. Bradyrhizobium japonicum inoculant mobility, nodule occupancy and acetylene reduction in the soybean root system. Appl. Environ. Microbiol. 55:2493-2498. McDermott, T.R., and P.H. Graham. 1990. Competitive ability and efficiency in nodule formation of strains of Bradyrhizobium japonicum. Appl. Environ. Microbiol. 56:30353039.

McDermott, T.R., P.H. Graham, and M.L. Ferrey. 1991. Competitiveness of indigenous populations of Bradyrhizobium japonicum serocluster 123 as determined using a roottip marking procedure in growth pouches. Plant Soil 135:245-250. Moawad, H.A., W.R. Ellis, and E.L. Schmidt. 1984. Rhizosphere response as a factor in competition among three serogroups of indigenous Rhizobium japonicum for nodulation of field-gro\Vn soybeans. Appl. Environ. Microbiol. 47:607-612. Moawad, H.A., and E.L. Schmidt. 1987. Occurrence and nature of mixed infections in nodules of field-grown soybeans (Glycine max). BioI. Fertil. Soils 5:112-114. Noel, K.D., F. Sanchez, L. Fernandez, J. Leemans, and M.A. Cevallos. 1984. Rhizobium phaseoli symbiotic mutants with transposon Tn5 insertions. J. Bacteriol. 158: 148-155.

Norris, D.O. 1958. A red strain of Rhizobium from Lotononis bainesii Baker. Aust. J. Agric. Res. 9:629-632. Norris, D.O. 1963. A porcelain bead method for storing rhizobium. Emp. J. Exp. Agric. 31:255-259.

O'Hara, G.W., N. Boonkerd, and M.J. Dilworth. 1988. Mineral constraints to nitrogen fixation. Plant Soil 108:93-110. O'Hara, G.W., and R.M. Daniel. 1985. Rhizobial denitrification: A review. Soil BioI. Biochern. 17:1-9. Oliveira, L.A., and P.H. Graham. 1990. Speed of nodulation and competitive ability among strains of Rhizobium leguminosarum by phaseoli. Arch. Microbiol. 153:311-315. Pattison, A.C., and F.A. Skinner. 1974. The effects of antimicrobial substances on Rhizobium spp. and their use in selective media. J. Appl. Bacteriol. 37:239-250. Pazkowski, M.W., and D.L. Berryhill. 1979. Survival of Rhizobium phaseoli on coal based legume inoculants. Appl. Environ. Microbiol. 38:612-615. Philpotts, H. 1976. Filter mud as a carrier of Rhizobium inoculants. J. Appl. Bacteriol. 41:277-281.

Philpotts, H. 1977. Effect of inoculation method on Rhizobium survival and plant nodulation under adverse conditions. Aust. J. Exp. Agric. Anim. Husb. 17:308-315. Porter, F.E., I.S. Nelson, and E.K. Wold. 1966. Plastic pouches. Crops Soils 18:10. Robson, A.D. 1978. Mineral nutrients limiting nitrogen fixation in legumes. p. 277-293. In C.S. Andrew and A.J. Kamprath (ed.) The mineral nutrition of legumes on tropical and subtropical soils. CSIRO, Melbourne. Rosas, J.e., and F.A. Bliss. 1986. Host plant traits associated with estimates of nodulation and nitrogen fixation in common bean. Hortic. Sci. 21:287-289. Roughley, R.J. 1968. Some factors influencing the growth and survival of root nodule bacteria in peat culture. J. Appl. Bacteriol. 31:259-265. Roughley, R.J. 1976. The production of high quality peat inoculants and their contribution to legume yield. p. 125-136. In P.S. Nutman (ed.) Symbiotic nitrogen fixation in plants. Cambridge Univ. Press, London. Roughley, R.J., and J.M. Vincent. 1967. Growth and survival of Rhizobium spp. in peat culture. 1. Appl. Bacteriol. 30:362-376. Scholla, M.H., and G.H. Elkan. 1984. Rhizobiumfredii sp. nov., a fast-growing species that effectively nodulates soybeans. Int. J. Syst. Bacteriol. 34:484-486. Schwyn, B., and J.B. Nielands. 1987. Universal chemical assay for the detection and determination of siderophores. Anal. Biochem. 160:47-56. Segovia, L., D. Pinero, R. Palacios, and E. Martinez-Romero. 1991. Genetic structure of a soil population of nonsymbiotic Rhizobium leguminosarum. Appl. Environ. Microbiol. 57:426-433.

Segovia, L., P.w. Young, and E. Martinez. 1993. Reclassification of American Rhizobium leguminosarum biovar phaseoli type 1 strains as Rhizobium etli sp. nov. Int. J. System. Bacteriol. 43:374-377.

222

WEAVER & GRAHAM

Soberon-Chavez, G., and R. Najera. 1989. Isolation from soil of Rhizobium leguminosarum lacking symbiotic information. Can. J. Microbiol. 35:464-468. Somasegaran, P. 1985. Inoculant production with diluted liquid cultures of Rhizobium spp. and autoclaved peat: Evaluation of diluents, Rhizobium spp., peats, sterility requirements, storage and plant effectiveness. Appl. Environ. Microbiol. 50:398405. Somasegaran, P., and H.J. Hoben. 1985. Methods in legume-Rhizobium technology. Univ. of Hawaii NifTAL, MauL Stephens, P.M., and J.E. Cooper. 1988. Variation in speed of infection of "no root hair zone" of white clover and nodulating competitiveness among strains of Rhizobium trifolii. Soil BioI. Biochem. 20:465-476. Summerfield, R.J., P.A. Huxley, and F.R. Minchin. 1977. Plant husbandry and management techniques for growing grain legumes under simulated tropical conditions in controlled environments. Exp. Agric. 13:81-92. Sylvester-Bradley, R., D. Mosquera, and J.E. Mendez. 1988a. Selection of rhizobia for inoculation of forage legumes in savanna and rainforest soils of tropical America. p. 225-233. In D.P. Beck and L.A. Materon (ed.) Nitrogen fixation by legumes in Mediterranean agriculture. Martinus Nijhoff, Dordrecht, the Netherlands. Sylvester-Bradley, R., P. Thornton, and P. Jones. 1988b. Colony dimorphism in Bradyrhizobium strains. Appl. Environ. Microbiol. 54:1033-1038. Takats, S.T. 1986. Suppression of nodulation in soybeans by superoptimal inoculation with Bradyrhizobium japonicum. Physiol. Plant. 66:669-673. Takishima, Y., J. Shimura, Y. Ugawa, and H. Shugawara. 1989. Guide to world data center on microorganisms with a list of culture collections in the world. WFCC World data center on microorganisms, Riken, Saitama, Japan. Thompson, J .A., and J .M. Vincent. 1967. Methods of detection and estimation of rhizobia in soil. Plant Soil 26:72-84. Van Berkum, P., and H.H. Keyser. 1985. Anaerobic growth and denitrification among different serogroups of soybean rhizobia. Appl. Environ. Microbiol. 49:772-777. Vincent, J.M. 1970. A manual for the practical study of the root-nodule bacteria. IBP Handb. 15. Blackwell Sci. Publ., Oxford, UK. Weaver, R.W. 1979. Adsorption of rhizobia to peat. Soil BioI. Biochem. 11:545-546. Weaver, R.W., and L.R. Frederick. 1974. Effect of inoculum rate on competitive nodulation of Glycine max. L. Merrill. ii. Field studies. Agron. J. 66:233-235. Weaver, R.W., and S.F. Wright. 1987. Variability in effectiveness of rhizobia during culture and in nodules. Appl. Environ. Microbiol. 53:2972-2974. Woomer, P., J.B. Bennett, and R. Yost. 1990. Overcoming the inflexibility of most-probablenumber procedures. Agron. J. 82:349-353.

Published 1994

Chapter 13 Anaerobic Bacteria and Processes HEINRICH F. KASPAR, Cawthron Institute, Nelson, New Zealand JAMES M. TIEDJE, Michigan State University, East Lansing, Michigan

With the exception of rice paddies, most agricultural soils can generally be termed aerobic. But this does not mean that aerobic soils never experience anaerobiosis. Even very dry and well-structured soils may contain anaerobic microsites, and most agricultural land experiences seasonal variation of aeration state that may range from true aerobiosis to complete anaerobiosis. At any site in a soil, anaerobic conditions are created as soon as the O 2 demand exceeds the O 2 diffusion to this site. Thus, there are two main causes of soil anaerobiosis: (i) a high rate of O2 consumption caused by a high respiration rate that generally correlates with a relatively high organic matter content, and (ii) a low rate of gas diffusion. Anaerobiosis occurs characteristically in poorly drained or water-logged soils. It also is to be expected in soils with large aggregates or poor, massive structure, which may be enhanced through compaction caused by frequent use of heavy equipment or by grazing cattle on wet soils containing large amounts of clay. Some benefits of microbial activities due to soil anaerobiosis have been demonstrated. Microbial degradation of DDT and dechlorination of many organochlorine compounds occur more rapidly under anaerobic conditions. However, soil anaerobiosis is generally undesirable; it retards plant growth due to production of phytotoxic compounds such as fatty acids and H 2S, lowers the pH, reduces available N through denitrification, and restricts root respiration. Bacteria that grow anaerobically can be divided into two groups: facultative and obligate anaerobes. Facultative anaerobes can grow under both aerobic and anaerobic conditions. This chapter is not directed at this large group of microorganisms, but the methods given here can be used for the study of anaerobic metabolism of these facultative organisms. Among the obligate anaerobes of special interest to the agricultural microbiologist Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

223

224

KASPAR & TIEDJE

are those that fix N2 , decompose cellulose, reduce SO~-, and produce CH4 . But very little is known about soil anaerobes, though they may play an important role in nutrient transformations as well as in the degradation of organic matter and xenobiotics. A review on soil anaerobes and their activities has been written by Skinner (1975). Two more recent and prominent books on anaerobes in nature are by Zehnder (1988) and the second edition of The Prokaryotes (Balows et aI., 1992). This chapter is an updated version of chapter 46 published in the 1982 edition of Methods of Soil Analysis.

13-1 PRINCIPLES The basic limitation of anaerobic metabolism is the disposal of reducing equivalents created by the oxidation of the energy-yielding substrate. The four chemical classes of electron acceptors used give rise to four major groups of obligate anaerobes: fermenters, SOa- reducers, proton reducers and CO2 reducers (most N0 3- reducers are not obligate anaerobes [see chapter 14 by Tiedje in this book]). 1. Obligate anaerobic fermenters are heterotrophs that dispose of the excess electrons by producing reduced fermentation products, such as H 2, alcohols and fatty acids. Thus, they behave in the same way as facultative anaerobic fermenters. A test for aerobic growth allows the separation of these two groups. Most of the obligate anaerobic fermenters in soil appear to be clostridia. Most species of this genus are strict anaerobes, though some are aero tolerant. They are spore formers and have a chemoorganotrophic metabolism. Based on their hydrolytic capacities, they can be divided into saccharolytic, proteolytic, pectinolytic, and chitinolytic clostridia. Some species commonly found in soils are disease agents, namely, Clostridium tetani (tetanus), C. perfringens (gangrene), and C. botulinum (botulism, intoxication rather than infectious agent). Some of the saccharolytic clostridia are able to fix N2 (e.g., c. pasteurianum), and the dissimilatory reduction of N0 3- to NH/ can be carried out by many soil clostridia. 2. Sulfate reducers dispose of the electrons generated by the oxidation of the energy-yielding substrate by reducing S compounds, mainly SO~- , SOj-, S20j-, and So. In recent years, much more diversity has been discovered among the sulfate-reducing bacteria (Widdel & Hansen, 1992). There are now at least 14 genera known including gram-positive and gramnegative classes. Since SOa- reducers are responsible for the anaerobic corrosion of Fe, they are locally important in agriculture (corrosion of submerged structures containing Fe, such as drainage tiles, irrigation devices, and waste pits). Moreover, the reduction product, H 2S, is toxic to plant roots. Some bacteria related to sulfur and sulfate-reducing bacteria also can use metals such as Fe (III) oxides and Mn(IV) oxides as terminal electron

ANAEROBIC BACTERIA AND PROCESSES

225

acceptors. These organisms appear to be important particularly in terrestrial iron geochemistry (Lovley, 1991). 3. Proton reducers oxidize fatty acids and alcohols to acetate and CO2. The reducing equivalents released are transferred to protons, with molecular H2 formed as the reduced product. They are chemoorganotrophs with a limited substrate range (fatty acids and alcohols) and require a low oxidation-reduction potential (Eh). Since they are very difficult to grow in media, little information is available on this group. However, proton reducers appear to be the only organisms capable of anaerobic fatty acid and alcohol decomposition in absence of inorganic electron acceptors and thus play an essential role in the complete degradation of organic matter to CO2 and CH4 . 4. Carbon dioxide reducers use CO2 as a terminal electron acceptor. The products of the CO2 reduction are CH 4 (methanogens) or acetate (acetogens). Methane can also be produced by reductive decarboxylation of acetate. All CO2 reducers require a low Eh for growth. They are either chemolithotrophs or chemoorganotrophs. Some strains of acetogens form spores. Acetogens and methanogens play an important role in the regulation of the pH of anaerobic environments by their production and consumption of acetate. Methanogens mediate the last and often rate-limiting step of anaerobic mineralization of organic matter. The requirement for anaerobic conditions is the only common characteristic of all anaerobes, and yet even with this requirement, anaerobes vary in their sensitivity to oxygen. Generally, members of the group of 02-sensitive anaerobes lack the enzymes superoxide dismutase and catalase (strict anaerobes) or only superoxide dismutase (aerotolerant anaerobes). The absence of these enzymes leads to the accumulation of the toxic compounds O2- and H 20 2, respectively. A second group of anaerobes is sensitive to the high redox potential created by O 2 rather than to O 2 itself. These organisms contain essential enzymes that require a low Eh (strict anaerobes, but are not necessarily killed under aerobic conditions). Since environments-including defined growth media-are very rarely in chemical equilibrium, it is usually not possible to conclude from a measured O 2 concentration the Eh of this particular environment, and vice versa. In addition, the tolerance of many anaerobes to O 2 , high Eh, or both is not known; therefore, to ensure "sufficiently anaerobic" conditions, one should remove as much O 2 as possible and lower the Eh by addition of reducing agents. Generally, soils are aerobic, and anaerobic microsites are the result of a delicate equilibrium between O 2 supply and consumption. A change of the aeration conditions during sampling is likely and may lead to changes of microbial activities and population sizes. To keep these changes at a minimum, samples should be analyzed immediately after collection. If this is impossible, they are best collected and stored as intact cores in plastic bags at 4°C. As a rule, aeration conditions should be changed as little as possible between sampling and analysis. Thus, anaerobic techniques are generally unnecessary for collection and storage of soil samples.

226

KASPAR & TIEDJE

Fig. 13-1. Equipment for working with anaerobes. Opposite page -> A-(l) Sterilizable gas filter used for filing and evacuating septum-stoppered incubation vessels; consists of Cu tubing packed with cotton, Swagelok fittings, syringe needle connector (Teflon), and disposable needle. (2) Anaerobic culture tube with long butyl rubber stopper and Al crimp seal (Balch & Wolfe, 1976). (3) Hungate culture tube that has butyl rubber septum and screw cap with hole in center to provide access to the septum. B-Serum bottle with long butyl rubber stopper and crimp seal. Some manufacturers produce serum bottles with a longer neck that allows a more extensive seal between stopper and bottle (shown here). C-Hungate gas probe apparatus for providing 02-free gas. (4) Furnaces heat Cu filings to 350 °C to remove O 2 from the flowing gas. (5) In raised tube, the zone of reduced Cu (CUO) is noted by its light (orange) color. (6) An oxidized zone (CuO) is apparent from its dark (black) color. (7) The 02-free gas is passed to a manifold that is used to distribute it to gas probes (8). (8) The gas probes consist of glass syringes filled with cotton to sterilize the gas and hypodermic needles (15.24 cm [6 inch], 8 gauge) bent to hang in tubes or flasks. (9) Conventional pressure cooker adapted for use as anaerobic incubator. (10) Inlet and outlet tapped into cover allow evacuation and refilling with 02-free gas.

13-2 METHODS FOR REMOVAL OF OXYGEN 13-2.1 Removal of Oxygen from Gas Lines and Filter-Sterilization of Gases (Martin, 1971; Zehnder, 1976) 13-2.1.1 Materials 1. Gastight tubing, quartz glass, neoprene, Teflon, steel, Cu (refrigerator tubing) with Swagelok or Gyrolok fittings (Crawford Fittings, Niagara Falls, Canada; Hoke Inc., Cresskill, NJ). 2. Oven, thermostatically controlled, able to keep about 500 g of Cu filings at 300 to 350°C (e.g., Fig. 13-1, C; Sargent-Welch, Skokie, IL). 3. Copper filings, obtainable as CuO wire (has to be reduced before use, e.g., VWR Scientific Inc., Columbus, Ohio). 4. Syringe needle connectors (Hamilton Co., Reno, Nev.).

13-2.1.2 Procedure Slowly pass commercially available high-purity gas over the hot, reduced Cu. Sterilize the gas by passing it through disposable 0.22 ~m filters. An alternative is to make a filter using a 10- to 20-cm long heat-resistant tube (e.g., Cu, 6.35 mm [V4 in.] o.d.) filled with cotton and provided with a Teflon syringe needle connector that allows an easy mount of sterile needles (Fig. 13-1A). The homemade filter needs to be sterilized by dry heat before use.

228

KASPAR & T1EDJE

a mixture of H2 with an inert gas over the filings until they are again bright Cu colored. The reaction of H2 with CuO is exergonic. During generation of the reduced Cu, care must be taken that the temperature does not melt the glass (for this reason quartz is recommended, though Vycor will work if the temperature is kept below 600 to 650°C). Mixtures of H2 and air are explosive. Before regeneration with H 2, the Cu filings must be freed of air by sparging with an inert gas. If H2 is allowed to escape into air, it has to be diluted immediately, and sparks must be prevented by all means. In the last decade, the expansion in work with anaerobes has led to the general commercial availability of anaerobe quality gases. Most specialists working with anaerobes prefer to pass these gases through hot Cu before use. 13-2.2 Cold Catalytic Oxygen Removal with Hydrogen 13-2.2.1 Materials

1. Container with gastight closure, able to be flushed with gas from a tank (e.g., modified pressure cooker with sealable inlet and outlet, Fig. 13-1, C). 2. Hydrogenation catalyst, usually Pd-coated asbestos (e.g., Becton, Dickinson and Co., BBL Microbiology Systems, Cockeysville, MD). 3. Nitrogen gas. 4. Gas mixture containing 7% H 2, 13% CO2 and 80% 02-free N2 (section 13-3; Aranki & Freter, 1972). 13-2.2.2 Procedure Place the cultures to be incubated anaerobically and the catalystiJ;l the containers. After closing the jar tightly, flush with N2 (about five times its volume). Repeat this procedure with the H 2-N2 mixture. After the inlet and outlet are closed, the remaining O 2 is consumed by catalytic hydrogenation at normal incubation temperatures (2H2 + O 2 ~2H20). 13-2.2.3 Comments This method provides good anoxic conditions in a relatively short time. The initial flushing with N2 is done to prevent explosive mixtures of air and H 2. The system can be improved by alternate evacuation by pump or aspirator and refilling with the 02-free gas. In this case, plates should be placed upright in the container (agar may fall off under vacuum if inverted). GasPak and Bio-Bag are convenient modifications of the cold catalyst method and are commercially available (Becton, Dickinson and Co., BBL Microbiology Systems, Cockeysville, MD; Marion Scientific Co., Kansas City, MO). They have the advantage that no flushing is necessary since the H2 needed for O 2 consumption is chemically produced in

ANAEROBIC BACTERIA AND PROCESSES

229

the container. However, anaerobic conditions are obtained more slowly than in the flushed jar. 13-2.3 Other Methods Specialists in anaerobic microbiology use the above methods, especially that in section 13-2.1, but the following classical methods are briefly mentioned because they may be the most reasonable for the occasional, simple anaerobic experiment. Cultures of fast-growing aerobes (e.g., Escherichia coli) can be used to remove O 2 from closed containers (Soc. Am. Bacteriol., 1957). This method requires the least special equipment, but anaerobic conditions are established slowly. Thus, fast-growing aerobes may be able to grow in the test cultures. The method can be improved by flushing the jar at the beginning with 02-free gas. It has been modified for use with single plates (Snieszko, 1930). Parker (1955) used activated steel wool to remove O 2 from anaerobic culture chambers. The untreated steel wool is dipped into a detergent solution and then placed in a sealed chamber; the rusting consumes O 2, This method appears to have some advantages over the pyrogallol method in that CO2 is not absorbed and no CO is produced. The candle jar method also creates anaerobic conditions economically. A desiccator or other tightly closing jar is used. Add inoculated plates or open culture tubes, then add candle, light, and seal. Candle will extinguish when O 2 tension is low. This method is good for microaerophiles and those organisms that prefer high CO2, This method is not adequate for strict anaerobes. Some CO will be produced which may inhibit some strains. Pyrogallol has also been used to remove O 2 (Soc. Am. Bacteriol., 1957). O 2 is consumed during the alkali-catalyzed polymerization of pyrogallol (1, 2, 3-trihydroxybenzene). The alkali also removes CO2 that can limit growth of many anaerobes. Some CO is also produced. The method is simple, effective, and appropriate for the occasional need for anaerobic conditions. Pyrogallol is toxic and contact with workers as well as cultures must be avoided.

13-3 METHODS FOR REDUCTION OF MEDIA 13-3.1 Materials

1. Reducing agents: Table 13-1 summarizes standard redox potentials at pH 7 (Eo') and stock solution preparation of reducing agents commonly used in microbiology. 2. Gassing probe (section 13-2.1, Fig. 13-1, C).

230

KASPAR & TIEDJE

Table 13-1. Standard redox potentials at pH 7 (Eo') and stock solution preparation of reducing agents.

Reducing agent Sodium thioglycollate

Eo' [mY]

< -100

Preparation of stock solution

Final concentration in medium

1% (wt/vol) in distilled water, autoc1aved and stored under 02-free gas

0.05%

Sodium sulfide nanohydrate

-225

1.2% (wt/vol) Na2S·9H20 in distilled water, autoclaved and stored under O 2free gas

0.025%

Cysteine

- 340

1% (wt/vol) in distilled water, autoc1aved and stored under 02-free gas

0.025%

Titanium(III) citrate

-480

5 mL (wt/vol) aqueous solution of TiCl3 added to 50 mL of 2.36% (wtlvol) aqueous solution of sodium citrate dihydrate and neutra1ized by addition of saturated Na2C03 solution, then filter-sterilized; must be used within 1 d at concentration of 30 mUL medium

0.5-2 mM

3. Thick-walled containers that are able to be tightly stoppered (Fig. 13-1). 4. Microbiological medium. 13-3.2 Procedure For prereduced media, combine the heat-stable ingredients of the medium in a flask, and gently boil while sparging with Oz-free gas (e.g., Nz + 3% Hz) through the gassing probe (section 13-2.1). Continue gassing while dispensing the medium into thick-walled containers; then tightly stopper and autoclave. After slow cooling, add by syringe the heat-labile ingredients and reducing agents that have been previously filter-sterilized. 13-3.3 Comments It is important to remove as much O 2 from the medium as possible before adding the reducing agents to minimize the toxicity that can come from use of larger quantities of reducing agent and from products produced by the reaction of reductant with Oz. To prevent precipitation, the media must be cooled before addition of the reducing agents. Cold sterilization of the reducing agents is most conveniently done by using syringes and disposable membrane filters (0.22 !lm, e.g., Millipore Corp., Bedford, MA). Besides the above-mentioned S compounds, HzS, dithionite, and amorphous FeS have been used as reducing agents. Amorphous FeS has

ANAEROBIC BACTERIA AND PROCESSES

231

been shown to remove O 2 faster than cysteine or HS- (Brock & O'Dea, 1977). The use of S compounds may lead to false positive tubes in the enumeration of SO~- reducers (Richard Smith, 1978, personal communication). Titanium(III) citrate provides the lowest Eh (Zehnder & Wuhrmann, 1976). If the citrate is utilized by bacteria, it may be replaced by other ligands, such as tartrate, rhodanide, or oxalate. The Ti(III) citrate complex has a blue-violet color that is lost upon oxidation. Thus, it may also serve as a redox indicator. It is possible to use reducing agents in combinations, e.g., Na2S, cysteine, and Ti(III) citrate (Zehnder & Wuhrmann, 1977). Convenient thick-walled containers with secured rubber stoppers are the Hungate tube, the anaerobe culture tube, (both shown in Fig. 13-1A and obtainable from Belleo Glass, Inc., Vineland, NJ) and the serum bottle shown in Fig. 13-lB. Long, black butyl rubber stoppers provide the best seal. The common red rubber, thin-walled, sleeve-type serum bottle stoppers and silicone septa are inadequate for anaerobic work. 13-4 REDOX INDICATORS 13-4.1 Methylene Blue (Skinner, 1971) 13-4.1.1 Materials 1. Solution A: Combine 3 mL of 0.5% aqueous methylene blue solution with 100 mL of distilled water (stored in dark). 2. Solution B: Combine 0.5 g of glucose, one small crystal of thymol, and 100 mL of distilled water. 3. Solution C: Combine 1.06 g of anhydrous sodium carbonate (Na2C0 3), 0.84 g of sodium bicarbonate (NaHC0 3), and 100 mL of distilled water.

13-4.1.2 Procedure Add solution C to solution B until pH 10 is reached. (This mixture is stored cold and dark.) Mix the product with solution A to give a light blue color. Boil portions of this methylene blue indicator fluid in small beakers or test tubes until it is colorless, and then place in the containers to be monitored for anaerobiosis. Close the containers immediately, and make the atmosphere anaerobic (section 13-2). 13-4.2 Resazurin Filter-sterilize a 0.1 % aqueous stock solution of resazurin, and add to the growth medium to give a final concentration of 0.0001 to 0.0003%. Resazurin can also be autoclaved directly in the medium.

KASPAR & TIEDJE

232

13-4.3 Pbeliosafranine (Bryant, 1963) Filter sterilize a 0.1 % aqueous stock solution of phenosafranine, and add to the growth medium to give a final concentration of 0.0001 to 0.0002%. 13-4.4 Comments As mentioned in section 13-3.3, Ti(I1I) citrate loses its blue color upon oxidation. Its standard redox potential at pH 7 and 25 °C (Eo') is -480 mV (Zehnder & Wuhrmann, 1976). Thus, if Ti(III) citrate is used as a reducing agent and the medium stays blue-violent, highly reducing conditions are guaranteed. Phenosafranine has a Eo' of -252 mV (Jacob, 1970) and therefore changes color under less reduced conditions. Resazurin has a Eo' of -51 mY, and its colorless state may not mean sufficiently reduced conditions for growth of all obligate anaerobes. This is even more the case with methylene blue, which has a Eo' of + 11 mV (equivalent to 0.05 atm 02)' Methylene blue, resazurin, and phenosafranine gradually change color within a range of about 120 mV around their Eo' and become completely colorless 60 mV below their Eo'. The Eo of these dyes is strongly dependent on the pH of the medium. Resazurin is the most commonly used redox indicator and is the one we recommend for most soil microbiology studies. Its properties are summarized below: Resazurin (Blue)

pH 70% homology) within the Alnus isolates, five genomic species within the Elaeagnus isolates, and one genomic species for the Casuarina isolates. Subsequent work with 27 Frankia strains from the Alnus and Elaeagnus host specificity groups defined five genomic species within the Alnus isolates and four genomic species within the Elaeagnus isolates (Akimov & Dobritsa, 1992). Large variation in DNA:DNA homology (12 to 99%) has been found for Frankia isolated from Myrica pensylvanica (Bloom et aI., 1989), perhaps because it is a promiscuous host. Nevertheless, this large diversity within Frankia is undoubtedly an underestimate of the true diversity, because only isolated strains from just a few of the actinorhizal hosts have been used for DNA:DNA hybridization. 16-3.2.7 Restriction Fragment Length Polymorphism Patterns A more discriminating test of diversity that uses DNA is the use of RFLP patterns (see chapter 31 by Sadowsky in this book). The earliest work with Frankia compared ethidium bromide-stained gels of total chromosomal DNA digested with restriction enzymes (An et aI., 1985b). Bloom et ai. (1989) applied this method to 16 Frankia isolates from Myrica pensylvanica and found nine distinct RFLP groups, which illustrates the sensitivity of this method. Application of restriction enzymes that cut at low frequency allowed more than 100 Frankia isolates to be grouped into 15 clusters (Beyazova & Lechevalier, 1992). Researchers also have used specific gene probes (e.g., nifHDK or 16S rDNA) to hybridize Southern transfers of restricted DNA (Normand et aI., 1988; Nazaret et aI., 1989; Nittayajarn et aI., 1990; Jamann et aI., 1992). This approach is very selective and has been found to give results consistent with the proposed speciation of Frankia (Lalonde et aI., 1988; Nazaret et aI., 1989) or previously described host specificity groups (Nittayajarn et aI., 1990). A new modification of this approach combines the polymerase chain reaction (peR) with subsequent restriction enzyme digestion to generate RFLP patterns (Maggia et aI., 1992; J amann et aI., 1993). 16-3.2.8 DNA Sequencing and Strain Specific Probes The most recent advance in differentiating Frankia strains has involved sequencing specific genes and synthesizing strain-specific gene probes (see chapter 32 by Ogram in this book). To date, two genes have

FRANKIA & THE ACTINORIDZAL SYMBIOSIS

309

been used. Based on the DNA sequence of the nifH gene, Simonet et ai. (1990b) were able to construct 15-bp oligonucleotides that were capable of distinguishing between two Frankia strains isolated from Alnus. Hahn et ai. (1989b) sequenced the portion of the rrn gene, which codes for rRNA's, and used this information to construct oligonucleotide probes that could differentiate between an effective and an ineffective strain (Hahn et aI., 1989a). Recently, additional rrn sequences have been determined that allow other strains of Frankia to be differentiated with oligonucleotide probes (Harry et aI., 1991; Nazaret et aI., 1991; Bosco et aI., 1992; Normand et aI., 1992). Greater use will undoubtedly be made of strain-specific probes designed on the basis of sequence information. It should be noted, however, that other approaches, such as using randomly cloned pieces of Frankia DNA as probes, may be equally useful in differentiating Frankia strains. 16-4 CHARACTERIZATION OF FRANKIA IN SYMBIOSIS

Frankia strains can be characterized using actinorhizal host plants. Plants can be used to assess host specificity and Nrfixing potential, or to measure differential characteristics of Frankia strains that are peculiar to actinorhizae, e.g., sp + vs. sp- nodules. In addition, actinorhizal hosts can be used as Frankia 'traps' and their nodules as enrichment sites for testing Frankia DNA and RNA with gene probes. 16-4.1 Host Specificity Torrey (1990) provides an excellent guide to testing for cross-inoculation groups. Although the basic procedure is straightforward, skill is needed in raising test plants and taking necessary precautions to prevent contamination. Testing for host specificity involves inoculation of one or more actinorhizal host plants with one or more sources of Frankia inoculum. It is important that both positive (inoculating with a Frankia strain known to cause nodulation) and negative (not inoculated) controls be done. Plants are checked periodically for nodulation and sometimes the nodules are assessed for sporulation, effectiveness (nitrogenase activity), or other physiological characteristics. Patience is often required before consistent results are obtained. 16-4.1.1 Inoculum Preparation The source of inoculum may be pure culture isolates of Frankia, crushed nodule suspensions, or soil. Pure cultures are probably the easiest to work with, however, standardization is needed because of the different morphologic forms of Frankia. Torrey (1990) suggests growing Frankia cultures to stationary phase in a medium that promotes spore formation. The Frankia should be harvested, washed, homogenized, and centrifuged

310

MYROLD

to determine pcv (see section 16-3.1.1.2). About 1 to 10 !J.L pcv per seedling is adequate for nodulation. Experience has shown that spore suspensions are much more infective than hyphal suspensions on a pcv basis and retain a degree of viability following desiccation (Burleigh & Torrey, 1990). Preparation of crushed nodule suspensions is similar to that described for isolation of Frankia (see section 16-2.1). Young, healthy nodules should be collected, washed, surface sterilized (usually H 20 2 or NaOCI), homogenized, and the homogenate diluted in water or 1% (wt/vol) NaCl. Torrey (1990) suggests 1 g of nodule per 20 mL of diluent. Roots of the test plants may be dipped in the crushed nodule suspension or the suspension can be applied to the roots. About 0.25 to 2.5 mg crushed nodule is generally sufficient for nodulation. Soil can be applied directly or as a suspension at a rate of 1 to 5 g soil per seedling (Benoit & Berry, 1990). 16-4.1.2 Plant Propagation and Assay Systems The details of propagating and growing all of the different types of actinorhizal plants is beyond the scope of this chapter; each has different requirements. An overview is given in Table 16-4, but the review by Benoit and Berry (1990), and the references therein, should be consulted for more complete information on growing actinorhizal plants from seed or through vegetative propagation. Techniques for the production of clonal plant material using tissue culture methods are described by Seguin and Lalonde (1990). Commercial sources of seed are available for some actinorhizal plants, but often collection is required. Many actinorhizal hosts grow well in water culture, although some may require a solid matrix, such as sand. What follows is a general description of a water culture system. Torrey (1990) should be referred to for additional details. Healthy seedlings with their first true leaves are inoculated and transferred to nutrient solution containers. Containers should not allow light to enter to prevent algal growth. Inoculated seedlings are started in low-N ( < 10 mg-N L -1) nutrient solution and transferred in a few weeks to N-free nutrient solution. Torrey (1990) points out that use of a proper nutrient solution is critical and recommends using a modified Hoagland's solution (Table 16-5). Nutrient solution is replenished as needed and renewed periodically (e.g., biweekly). Aeration of the solution is often desirable. Conditions suitable for growth of plants (adequate temperature and light) should be maintained (i.e., a growth chamber or greenhouse). Plants are checked periodically for nodulation. Usually at least 14 d is required, and sometimes several months, before nodulation occurs (Torrey, 1990). 16-4.2 Nodule Metabolism (Nitrogen fixation) Nodule respiration, nitrogenase activity, and hydrogenase activity have been assayed to determine the efficiency of N2 fixation (Huss-Danell,

FRANKlA & THE ACTINORHIZAL SYMBIOSIS

311

Table 16-4. Guidelines for propagation of actinorhizal plants. (Adapted from Benoit and Berry, 1990.) Genus

Seed treatment

Germination conditionst

Alnus

Overnight soak in aerated water

20-30 °C, 16-h daylight, 7-12 d

Casuarina

None

20-30 °C, 16-h daylight, 11-40 d

Ceanothus

Hot water soak

25 °C, 12-30 d

Cercocarpus

Aerated water soak for 24 h, stratify 5 wk

15-20 °C, 1-2 wk

Comptonia

Scarify plus 500 ppm GA3

15-20 °C, 8 wk

Cowania

Aerated water soak for 24 h, stratify 5 wk

15-20 °C, 1-2 wk

Discaria Dryas Elaeagnus Hippophae

Stratify 90 d

Myrica

Stratify 14-90 d, 500 ppm GA3 24 h Stratify 5 wk

Purshia

Shepherdia

Stratify 60-90 d, 0.5-1 h H 2 S0 4 treatment for hard seeds

30 °C d, 20 °C night 30 °C d, 20 °C night, 40 d 23-25 °C, 16 h daylight, 11-150 d 15-20 °C, 14 d

20-30 °C, 21-60 d

Vegetative

propagation~

Stem cuttings, 5000 ppm IBA or powdered rooting aid, 3-6 wk in mist, sand, or water culture Softwood cuttings, 50 ppm IBA, 5 wk in mist or water culture Softwood cuttings, 5000 ppm IBA or powdered rooting aid, 4-10 wk Semihardwood cuttings, 5000 ppm IBA, 6 wk in mist with bottom heat Softwood cuttings, stem layering, root pieces covered by rooting medium

Semihardwood cuttings Semihardwood cuttings, layering horizontal stems Hardwood cuttings, layering, root cuttings Hardwood cuttings, layering, root cuttings Semihardwood cuttings with 5000 ppm IBA, layering Semi hardwood cuttings, 5000 ppm IBA or powdered rooting aid, 8-10 wk on shade bench Semihardwood cuttings, 8000 ppm IBA dip

t GA 3 , gibberellic acid. ~ IBA, indole-3-butyric acid.

1990). With the one exception of the "local source" of Frankia from Sweden (Sellstedt & Huss-Danell, 1984; Sellstedt, 1989), hydrogenase activity is universally present in Frankia and actinorhizal nodules. Consequently, the focus of this section will be measurement of nitrogenase activity. Ecological measurements of N2 fixation have been made to: discriminate between effective and ineffective nodules, test strains for their N2 fixing ability, compare the effectiveness of different Frankia strain-plant combinations, and determine the amount of N2 fixed in the field. In actinorhizal symbioses, N2 fixation has been assessed by mass balance, 15N

MYROLD

312

Table 16-5. Recipes for nutrient solutions commonly used for growing actinorhizal plant seedlings. Solution 114 Hoagland'st

Ingredient Macronutrient solutions Ca(H2 P04 )z· H 2 O

CaS0 4 ·2H2 O K2 S0 4 MgS0 4 ·7H 2 O

Chelated iron

FeS0 4 ·7H2 O NazEDTA

Micronutrients CoCl z·6H2 O CuS0 4 ·5H zO

H 3 B03 MnCl z·4HzO Na2 Mo0 4 ·2H2 O ZnS0 4 ·7HzO

Nitrogen

NH4 N0 3

1110 Evan's*

Stock solution, g L-l 12.6 1.72 87 246 5.56 7.45

0.D25 0.080 2.86 1.81 0.D25 0.220 80.05

Solution 1

K2S0 4 MgS0 4 ·7H 2 O

Solution 2

KH2P0 4 K2 HP0 4

Solution 3

CaClz·2HzO CaS0 4 ·2HzO Solution 4

FeCL3 ·6H2 O Na 2 EDTA

Solution 5

CoS0 4 ·7H2 O

CuS0 4 ·5HzO

H 3 B03 MnS0 4 ·H2 O NazMo0 4 ·2HzO ZnS04 ·7HzO Solution 6

NH4 N0 3

27.88 49.28

Final solution, mLL-l 1.25 50 1.25 0.50 1.25

0.25

0.179 1

2.50 14.5 7.35

10.33

0.483 0.628 0.0141 0.0079 0.143 0.0773 0.0050 0.0220 2.86

10

t Modified from Hoagland and Arnon (1950). Because of its low solubility, it is most convenient to add CaS04 ·2HzO directly to the final solution volume in the desired amount, e.g., 0.86 g for 1 L of 114 Hoagland's solution. Omit NH4 N0 3 for N-free solution. When adjusted to pH 6.0 (two to three drops of 0.1 N NaOH for 1 L of 114 Hoagland's solution), I have found this solution to work well in the MPN bioassay. Others have successfully used similar Hoagland's resolutions at one-fourth to full strength and pH values from 5.4 to 6.8 (Quispel, 1958; van Dijk, 1984; van Dijk & Sluimer-Stolk, 1990). :j: This is Huss-Danell's (1978) modification of Evan's et al. (1972) solution. Solution 3 does not completely dissolve, so it must be thoroughly mixed before use. Omit NH4 N0 3 for N-free solution. The pH of 1110 Evan's solution is about 7.0, however, it is weakly buffered. I have found 1110 Evan's solution to be comparable to 114 Hoagland's solution in the MPN bioassay, however, it has been successfully used by others at one-half strength (Smolander & Sundman, 1987).

FRANKIA & THE ACTINORHIZAL SYMBIOSIS

313

methods, and the acetylene reduction assay. Details about these methods can be found in chapter 43 or, with special reference to actinorhizal plants, in Winship and Tjepkema (1990). In addition, there have been several recent papers devoted to measuring N2 fixation of actinorhizal plants in the field (e.g., Beaupied et aI., 1990; Sougoufara et aI., 1990). One important point that should not be forgotten, is that the activity of root nodules as measured on intact plants is likely to be different from that measured in excised root nodules. Some Frankia strains are infective but ineffective, i.e., they are capable of forming nodules but do not fix N 2. Ineffective nodules are typically small in size, contain relatively few Frankia, and do not normally contain vesicles (Mian et aI., 1976; Hahn et aI., 1988; Berry & Sunell, 1990). Ineffective nodules often occur because of intergeneric or interspecific incompatibilities (Weber et aI., 1987; van Dijk et aI., 1988), however, some Frankia isolates form ineffective nodules even when inoculated on the same host species from which they were isolated (Hahn et aI., 1988). Van Dijk and Sluimer-Stolk (1990) report that some ineffective Frankia have no nitrogenase activity in pure culture and their DNA does not hybridize with a nifHDK probe. Hahn et ai. (1989a) found significant differences in the 16S rDNA sequence of an ineffective compared to an effective Frankia strain. In some cases, effectiveness can be lost by mutation (FaureRaynaud et aI., 1990b). The N 2-fixing effectiveness of host-Frankia combinations has been studied primarily with Casuarina and Alnus spp. Over threefold differences in growth and N2 fixation have often been found for various CasuarinaFrankia combinations (Reddell & Bowen, 1985; Fleming et ai. 1988; Reddell et aI., 1988; Sanginga et aI., 1990), although Rosbrook and Bowen (1987) found no significant Frankia strain effect. Differences in host provenance may be greater determinants of growth and N2-fixing effectiveness than variation among Frankia strains (Sougoufara et aI., 1992). Similar differences in growth and N2 fixation have been found for Alnus-Frankia combinations (Normand & Lalonde, 1982; Wheeler et aI., 1986; Hooker & Wheeler, 1987; Weber et aI., 1989). In the Alnus-Frankia symbiosis, greater N2-fixing effectiveness has been reported for sp- compared to sp+ symbioses (Normand & Lalonde, 1982; Wheeler et aI., 1986), although this relationship is not always observed (Kurdali et aI., 1990). Despite the large variation in N2-fixing capacity of various host-Frankia combinations, no clearly superior host genotypes or Frankia strains have been recognized and any positive effect of inoculation with a superior Frankia strain may soon be overshadowed in the field by nodulation by indigenous Frankia (Hooker & Wheeler, 1987). 16-4.3 Nodule Morphology (Sporulation) Actinorhizal nodules come in a variety of shapes and sizes: some have discrete branched lobes, some are compact, and nodules of Casuarina and Myrica can form nodule roots (Berry & Sunell, 1990). Recently, aerial

314

MYROLD

nodules have been discovered on Casuarina cunninghamiana (Prin et aI., 1991a). Nodule morphology is probably under host plant control but may reflect environmental conditions. The major feature of nodule morphology of interest to microbial ecologists, however, is sporulation. Although virtually all Frankia can be induced to form sporangia in pure culture, this is not necessarily the case in planta. Van Dijk and Merkus (1976) observed that Frankia within actinorhizal root nodules either form many sporangia (sp+) or no (or at least very few) sporangia (sp-). Smolander and Sundman (1987) proposed an "intermediate" class for nodules containing few sporangia and reserving sp- only for nodules devoid of sporangia, however, Schwintzer (1990) suggests that "intermediate" nodules are probably better classified as sp-. The spore type of actinorhizal nodules is of interest for at least three reasons: (i) It is doubtful if any Frankia have been isolated from sp + nodules (see section 16-1.2), (ii) Interesting and complex ecological relationships have been observed between sp - and sp + strains of Alnus sp. and Myrica gale (Weber, 1986; Holman & Schwintzer, 1987; Kashanski & Schwintzer, 1987; Smolander & Sundman, 1987), and (iii) There is some evidence that sp- and sp+ nodules differ in both absolute and relative nitrogenase efficiency (Wheeler et aI., 1986). Nodule spore type is determined microscopically. Either hand or microtome sections of nodules are prepared. Hand sections are easily prepared with a razor blade. A description of microtome sectioning is given by Smolander and Sundman (1987). Sections should be prepared from the base of peripheral nodule lobes, because older tissues are more likely to contain sporangia (Schwintzer, 1990). Several sections from each lobe should be examined. Once cut, sections are mounted in dilute Fabil reagent (van Dijk, 1978) or lactophenol-cotton blue (Torrey, 1987) and examined at 400 x magnification. Schwintzer (1990) states that free-hand sections from sp + nodules often have a cloud of sporangia and free-spores around the edge of the nodule tissue (Fig. 16-4). When sporangia are not abundant, enough sections must be examined so that at least 50 infected cells are examined (Schwintzer, 1990). 16-4.4 Nodule DNAIRNA Assays

Immunological methods, such as the fluorescent antibody technique that is used widely with rhizobia, are not specific enough for differentiating Frankia strains in nodules. But with the development of molecular techniques, it is now possible to determine nodule occupancy. To date, three reports in the literature have used gene probes to detect specific Frankia strains in nodules. Simonet et al. (1988) used a radiolabeled Frankia plasmid cloned into pBR322 to probe blots of nodule DNA obtained from Alnus stands. This allowed them to distinguish Alnus stands that had the plasmid-containing Frankia strain from those that did not. But there are problems associated with using plasmids as strain markers. Plasmids may be lost or transferred

316

MYROLD

and it is possible for the same plasmid to be present in strains with a different chromosomal background or vice versa (Mullin & An, 1990). Hahn et al. (1990b) used oligonucleotide probes based on sequence differences in the variable region of 16S rRNA to detect a particular Frankia strain in Alnus glutinosa nodules. RNA was extracted from nodules and Frankia was detected in nodules as small as 1 mg. Simonet et al. (1990b) used oligonucleotide probes based on sequence differences in the nifH gene to detect and differentiate between two Frankia strains in Alnus glutinosa and A. incana nodules. DNA was extracted from individual nodule lobes and the nifH gene was specifically amplified with the PCR by using primers to the highly conserved region of the nifH gene. The amplified DNA was hybridized with the oligonucleotide probes to differentiate between the strains. Simonet et al. (1990b) reported that the PCR amplification step was necessary, because insufficient target DNA was obtained by extraction alone, however, D.D. Baker (1992, unpublished data) has been able to differentiate among Frankia strains using RFLP analysis of DNA extracted from nodules using the CTAB protocol (Murray & Thomson, 1980). These methods all require the extraction of nucleic acids and filter hybridization. An alternative approach, which allows visualization of Frankia in planta by using ftuorescently tagged oligonucleotide probes (e.g., Amann et aI., 1990), was recently reported (Prin et aI., 1993). The diversity of Frankia in nodules can also be studied using PCR-RFLP (N.J. Ritchie and D.D. Myrold, 1993, unpublished data) or DNA sequencing of PCR products (Nick et aI., 1993). 16-5 QUANTIFICATION IN SOIL Many of the methods used to measure Frankia in soil are extensions of those used in pure culture or in planta. But following a small population of Frankia in soil against a large background of other microorganisms presents some special problems. Methods for detecting Frankia in soil must be selective or be able to specifically enrich for Frankia. 16-5.1 Plants as a Bioassay System Plants meet the criteria given above. Within host specificity limits, actinorhizal plants select for Frankia when they become nodulated and the nodules are an enrichment of Frankia. 16-5.1.1 Use of Trap Plants The simplest use of plants is to serve as trap plants. Actinorhizal plants can be planted in soil and then checked for nodulation. Such an approach is only qualitative, although it may be possible to make it quantitative if a dilution series of the soil with a sterile potting medium is made. In that case, an MPN estimate of Frankia numbers might be made.

FRANKIA & THE ACTINORHIZAL SYMBIOSIS

317

16-5.1.2 Plant Bioassay and Frankia Infective Units Virtually all of the ecological information about Frankia populations in soil has been collected by using some variation of a plant bioassay system. The statistical basis for most probable number (MPN) counts is given in chapter 3. After a brief summary of the autecology of Frankia in soil, the MPN methodology as it has been used with Frankia will be described. Quantitative estimates of Frankia populations in soil are based on IUs as measured with a plant bioassay and to date have been devoted to Alnuscompatible Frankia. Surveys of soils from a variety of forest sites in Finland (Smolander & Sundman, 1987; van Dijk et aI., 1988; Smolander, 1990), Sweden (Myrold & Huss-Danell, 1994), and the Pacific Northwest of the USA (A.B. Hilger & D.D. Myrold, 1992, unpublished data) have shown Frankia populations to range from 0 to 4600 IU g-l soil. Frankia populations vary according to plant species present and soil conditions. It is particularly interesting that higher numbers have been found associated with non-actinorhizal plants (e.g., birch) than with actinorhizal plants (Smolander & Sundman, 1987; van Dijk et aI., 1988; Smolander, 1990). Soil pH is positively correlated with Frankia populations (Smolander & Sundman, 1987). Laboratory experiments that have investigated soil and rhizosphere effects on Frankia populations have largely confirmed field observations. When Frankia were added to limed (pH 6.0) and unlimed soil (pH 4.2), survival was greater in limed soil (Smolander et aI., 1988; Smolander & Sarsa, 1990; A.B. Hilger & D.D. Myrold, 1992, unpublished data). Rhizosphere effects, however, have given mixed results with some studies showing Frankia populations enhanced in the rhizospheres of some plants, such as birch (Smolander & Sarsa, 1990) and others showing no effect of several plant species, including birch (A.B. Hilger and D.D. Myrold, 1992, unpublished data). A survey of sand dunes in the Netherlands, which took advantage of being able to distinguish effective from ineffective Frankia strains, found populations of ineffective Frankia to be present at much higher populations than effective Frankia (van Dijk & Sluimer-Stolk, 1990). 16-5.1.2.1 Nodulation Capacity. The most widely used plant bioassay was developed by van Dijk (1984). The procedure involves germinating alder seedlings and growing them for 6 wk (until they have three to four leaves) in one-fourth-strength complete Hoagland's solution (Table 16-5). Nutrient solution is changed weekly. Seedlings are transferred to 250-mL culture jars containing one-half-strength N-free Hoagland's solution (eight seedlings per jar) and inoculated 2 d later. Serial dilutions of inoculum are made up in one-half-strength N-free Hoagland's solution and mixed into the culture jars. Plants are grown in a growth chamber. After 5 d, nutrient solution is replaced and thereafter replaced weekly. Nodulation is assessed 3 wk following inoculation (longer periods did not result in any more nodules forming).

318

MYROLD

Nodulation capacity (analogous to IU) was defined by van Dijk (1984) to be the number of nodules formed per unit of inoculum. It is equivalent to the slope of the line when number of nodules per jar is plotted against amount of inoculum per jar. This empirical relationship is linear up to the point where root biomass begins to limit nodulation. van Dijk (1984) chose to calculate nodulation capacity by summing the number of nodules in all jars of all dilutions in the linear portion of the curve (r) and dividing r by the fraction of the total amount of inoculum that was added to these jars (P). The resulting number is divided by the original inoculum quantity to get the nodulation capacity in terms of nodules per amount of inoculum. An estimate of the variance of the nodulation capacity is given by multiplying (l-p) by (r+ 1) and dividing by p2. No derivation of these formulae is provided by van Dijk (1984) and it seems that doing a linear regression should provide similar estimates of the mean and variance of nodulation capacity using more widely recognized statistical techniques. 16-5.1.2.2 Most Probable Number Bioassay. Another approach for estimating infective units is based on MPN statistics. Instead of using several seedlings per container and counting total numbers of nodules, a single seedling per container is used and nodulation is scored in a plus/minus fashion. These data are analyzed using MPN tables or computer programs that calculate the appropriate MPN statistics (see chapter 5 by Woomer in this book). A convenient experimental system for Frankia MPN tests was described by Hilger et al. (1991) and is shown in Fig. 16-5. Germinated seedlings of the one-leaf stage are placed in modified centrifuge tubes containing one-fourth strength N-free Hoagland's solution plus 5 mg NH4 N0 3-N L -1, and inoculated with a serial dilution of soil. After 1 wk, and at 2 wk intervals thereafter, the solution is replaced with one-fourth strength N-free Hoagland's solution. Six to 8 wk following inoculation, seedlings are checked for nodule formation. Negative (not inoculated) and positive (inoculated with an effective Frankia strain) controls should always be included. This system works well with Alnus rubra and A. incana. 16-5.1.2.3 Comparison of Methods. The nodulation capacity assay is more widely used for measuring Frankia IUs, but the MPN assay may be based on a better established statistical foundation and is quite space efficient. A five-dilution, five tubes per dilution MPN assay occupies only about 0.08 m- 2 , which may enhance the number oftrue replicates that can be done in a given bioassay. Both of the plant bioassay methods work well, however, and give comparable estimates of Frankia IUs (A.B. Hilger and D.D. Myrold, 1992, unpublished data; C. van Dijk, 1992, personal communication; Huss-Danell & Myrold, 1994).

319

FRANKIA & THE ACTINORHIZAL SYMBIOSIS

- - - - Screw cap _ _ _ --'-_ 50-ml Polypropylene tube

'/-+--- --

-

Nutrient solution

- - - Styrofoam block

Fig. 16-5. Schematic diagram of an MPN seedling bioassay system for determining Frankia IV.

16-5.2 Fluorescent Antibodies Several Frankia research groups have tried to use serological methods for detecting Frankia (see section 16-3.2.1), but no one has yet been able to produce antibodies of sufficient specificity for identifying a particular Frankia strain. These less specific probes might be useful for studies of Frankia in soil at the genus level, however, difficulties will be faced because of low soil populations. 16-5.3 DNAIRNA Probes Perhaps the most promising approach for detecting and quantifying Frankia in soil will involve the use of molecular probes to detect Frankia

DNA or RNA extracted from soil, possibly following DNA amplification by the peR (see chapters 32, 34, and 35). 16-5.3.1 Frankia-Specific Probes and Polymerase Chain Reaction Primers Frankia-specific probes and primers are based on the 16S rRNA se quences (Hahn et aI., 1989a; Nazaret et aI., 1991). Additional sequences are becoming available (Harry et aI., 1991) and checks of the GenBank, or other sequence databases will likely turn up others. Using such information, several primers have been developed that are specific to gram-positive

MYROLD

320

organisms (Table 16-6) and can be used as probes or primers in conjunction with the Frankia-specific primers. 16-5.3.2 DNAIRNA Extraction There are numerous protocols for extracting DNA and RNA from soil (see chapter 35 in this book), however, only the direct extraction methods are likely to be useful for a filamentous bacterium like Frankia. Another important consideration when working with Frankia is their gram-negative cell wall structure, which can be more difficult to lyse than that of the gram-negative bacteria often used as test organisms in DNA extraction protocols. Hilger and Myrold (1991) have developed such a method to extract Frankia DNA from soil. The critical steps of this method remove humic materials that could interfere with subsequent probing or amplification techniques. A protocol for extracting Frankia RNA from soil has been published (Hahn et al., 1990a) which allows the detection of about 1()4 cells g-l soil. 16-5.3.3 Polymerase Chain Reaction Applications of PCR to Frankia have primarily used DNA extracted from Frankia isolates (Simonet et aI., 1991; Bosco et aI., 1992; Normand et aI., 1992) or actinorhizal nodules (Simonet et aI., 1990b; Nazaret et aI., 1991). The PCR can be used to amplify DNA extracted from soil (Myrold et aI., 1990). By using the MPN approach of dilution to extinction, the number of Frankia genomes can be quantified (Hilger & Myrold, 1992; Picard et aI., 1992). The PCR-MPN has been used with the plant bioassay MPN to study Frankia populations in soil (Myrold & Huss-Danell, 1994). 16-6 CONCLUSION Research on Frankia, especially its ecology in soil, is coming of age. Although there will still be methodological hurdles to overcome, many techniques are now available to allow long-standing questions to be answered. The use of molecular tools will greatly enhance these efforts, but proven methods using host plants will still playa vital role. One of the interesting areas that is just beginning to be explored is the diversity of Frankia in natural populations.

ACKNOWLEDGMENTS I express appreciation to C-Y. Li, who helped me begin my research with Frankia, and Dwight Baker who has often given me valuable advice. Arlene Hilger and Kendall Martin have been stalwarts in my lab since we began studying Frankia and have been the source of much inspiration. I

Eubacteria Frankia Frankia Frankia Eubacteria Eubacteria Actinomycetes Frankia. Alnus and Casuarina groups Frankia, Elaeagnus group Frankia Frankia Actinomycetes Eubacteria Eubacteria Gram positive Frankia (putative) Frankia§§ Frankia (putative) Eubacteria Frankia§§

FGPS6 FGPS59 FR183# FGPS305' FR485 FGPS849tt FGPS958 FGPS989ac

990-1009 1005-1025 1074-1093 1116-1176 1356-1373 1377-1401 1479-1497 1493-1512 1911-1933 1948-1973 2038-2055

989-1004

6-25 59-76 183-204 281-305 485-502 849-868 958-976 989-1004

Numbering§

TGC AGG ACC CTT ACG GA(Clt) CC AGC CAT GCA CCA CCT GTG CAG GCA ACA TAG GAC GAG GGT TG GGG GCA TGA TGA CTT GAC GTC ACG GGC GGT GTG TAC AAG TIC GGG TGT TAC CGA CTT TCG TGA C GTA CCG GAA GGT GCG GCT G GGC TGG ATC ACC TCC TTT CT TAA CTT GGC CAC AAA GAT GCT CG ATC GGC TCg aGG TGC CAA GGg ATC CA CCG GGT TTC CCC ATT CGG

GGG GTC CtT AgG GGc t

GGA GAG TTa GAT CtT GGC TC AAG TCG AGC GGG GAG CTT CTG GTG GTG TGG AAA GAT TTA T CCA GTG TGG CCG GTC GCC CTC TCA G CAG CAG CCG CGG TAA TAC GCC TGG GGA GTA CGG CCG CA CTI GAC ATG CAG GGA AAT C GGG GTC CGT AAG GGT C

Sequence~

Modified from Hahn et aI., 1989b L. Simon, 1992, personal communication Simonet et aI., 1991 Simonet et aI., 1991 Hilger and Myrold, 1992, unpublished data Hilger and Myrold, 1992, unpublished data Martin and Myrold, 1992, unpublished data Simonet et aI., 1991 Martin and Myrold, 1992, unpublished data Bosco et aI., 1992 Simonet et aI., 1991

Bosco et aI., 1992

Bosco et aI., 1992 Picard et aI., 1992 Modified from Hahn et aI., 1989b Picard et aI., 1992 Hilger and Myrold, 1992, unpublished data Simonet et aI., 1991 Simonet et aI., 1991 Bosco et aI., 1992

Reference

:j:

t No suffix dcsignates a forward primer, homologous to the coding strand; a ' designates a reverse primer, complementary to the coding strand. Primer specificity determined either from empirical tests with other bacterial species and strains, or by comparison with rrn sequence information. § Numbering based on sequence for DNA of Frankia CeD isolated from Casuarina equisetifolia (Normand et ai., 1992). This sequence is accessible from the GenBank databank, accession number M55343. ~ Lower case bases designate disagreement with the rrn sequence of Frankia CeD (Normand et ai., 1992). Bases in parentheses designate an equal mixture of the two bases at this position. # Also known as SSU200 (L. Simon, 1992, personal communication). tt Also known as SSU901 (Normand et ai., 1992). :j::j: Also known as FGPS1146' (Nazaret et ai., 1991) and SSUI190' (Normand et ai., 1992). §§ When FGPS1493 and FGPL2054' are used together, Geodermatophilus DNA is amplified, however, the resulting product is different than the 561-bp product obtained with Frankia DNA (Simonet et ai., 1991).

FR1009' SSU1000' FGPSI093' FGPS1176':j::j: FR1373 , FR1401' FR1479 FGPS1493 FR1933' FGPL1948, FGPL2054,

FGPS98ge

Specificity:j:

Labelt

Table 6-6. PCR primers that can be used to specifically amplify Frankia DNA. Primer pairs FR1831FR1401, , FR485IFR1009', FR183IFR1009', and FR14791FR1933' have been used successfully in my laboratory. The references cited should be consulted for other compatible primer pairs. Other possible primer pairs should be assessed for similar melting temperatures and potential to form primer dimers prior to use.

...~

rJ> rJ>

0

::::

--=

-


t""

N ;I>-

==

~

-

Z 0

>-3

("l

-

;I>-

trl ==

>-3

f(o

s:~

~

MYROLD

322

thank Kerstin Huss-Danell for providing an excellent sabbatical environment while I wrote this paper. While on sabbatical, I was supported by National Science Foundation grant INT-9025112 and a grant from the Swedish Council for Forestry and Agricultural Research to K. HussDanell. Lastly, thanks must go to the collective group of Frankia researchers throughout the world, who are always ready to help and advise. This article is Technical Paper No. 10,266 of the Oregon Agricultural Experiment Station.

REFERENCES Akimov, V.N., and S.V. Dobritsa. 1992. Grouping of Frankia strains on the basis of DNA relatedness. System. Appl. Microbiol. 15:372-379. Amann, R.I., L. Krumholz, and D.A. Stahl. 1990. Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J. Bacteriol. 172:762-770. An, C.S., J.W. Wills, W.S. Rig~sby, and B.C. Mullin. 1983. Deoxyribonucleic acid base composition of 12 Frankia Isolates. Can. J. Bot. 61:2859-2862. An, C.S., W.S. Riggsby, and B.C. Mullin. 1985a. Relationships of Frankia isolates based on deoxyribonucleic acid homology studies. Int. J. Syst. Bacteriol. 35:140-146. An, C.S., W.S. Riggsby, and B.C. Mullin. 1985b. Restriction pattern analysis of genomic DNA of Frankia isolates. Plant Soil 87:43-48. Baker, D. 1987a. Modification of catalog numbering system for Frankia strains and revision of strain numbers for DDB Frankia collection. Actinomycetes 20:85-88. Baker, D. 1987b. Relationships among pure cultured strains of Frankia based on host specificity. Physiol. Plant. 70:245-248. Baker, D. 1988. Opportunities for autecological studies of Frankia, a symbiotic actinomycete. p. 271-276. In Y. Okami et aI. (ed.) Biology of actinomycetes. Japan Sci. Soc. Press, Tokyo, Japan. Baker, D. 1989. Methods for the isolation, culture, and characterization of the Frankiaceae: Soil actinomycetes and symbionts of actinorhizal plants. p. 213-236. In D.P. Labeda (ed.) Isolation of biotechnological organisms from nature. McGraw-Hill, New York. Baker, D.D. 1993. A bibliography on Frankia and actinorhizal plants, March 1993 edition. Panlabs, Inc., South Bothell, WA. Baker, D., and D. O'Keefe. 1984. A modified sucrose fractionation procedure for isolation of frankiae from actinorhizal root nodules and soil samples. Plant Soil 78:23-28. Baker, D., W.L. Pengelly, and J.G. Torrey. 1981. Immunochemical analysis of relationships among isolated frankiae (Actinomycetales). Int. J. Syst. Bacteriol. 31:148-151. Baker, D.D., and C.R Schwintzer. 1990. Introduction. p. 1-13. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Baker, D., and E. Seling. 1984. Frankia: New light on an actinomycete symbiont. p. 563-574. In L. Ortiz-Ortiz et al. (ed.) Biological, biochemical and biomedical aspects of actinomycetes. Academic Press, New York. Baker, D., J.G. Torrey, and G.H. Kidd. 1979. Isolation by sucrose-density fractionation and cultivation in vitro of actinomycetes from nitrogen-fixing root nodules. Nature (London) 281:76-78. Beaupied, H., A. Moiroud, A.M. Domenach, F. Kurdali, and R. Lensi. 1990. Ratio of fixed and assimilated nitrogen in a black alder (Alnus glutinosa) stand. Can. J. For. Res. 20:1116-1119. Becking, J.H. 1970. Frankiaceae fam. nov. (Actinomycetales) with one new combination and six new species of the genus Frankia Brunchorst 1886. Int. J. Syst. Bacteriol. 20:201220. Benoit, L.F., and A.M. Berry. 1990. Methods for production and use of actinorhizal plants in forestry, low-maintenance landscapes, and revegetation. p. 281-297. In C.R. Scliwintzer and J.D. Tjepkema (ed.) The biology of Frankia and Actinorhizal plants. Academic Press, San Diego.

FRANKlA & THE ACTINORHIZAL SYMBIOSIS

323

Benson, D.R. 1982. Isolation of Frankia strains from alder actinorhizal nodules. Appl. Environ. Microbiol. 44:461-465. Benson, D.R., and D.G. Hanna. 1983. Frankia diversity in an alder stand as estimated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis of whole-cell proteins. Can. J. Bot. 61:2919-2923. Benson, D.R., and N.A. Schultz. 1990. Physiology and biochemistry of Frankia in culture. p. 107-127. In C.R Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San DIego. Benson, D.R., and W.B. Silvester. 1993. Biology of Frankia strains, actinomycete symbionts of actinorhizal plants. Microbiol. Rev. 57:293-319. Berry, A.M., R.A. Moreau, and A.D. Jones. 1991. Bacteriohopanetetrol: Abundant lipid in Frankia cells and in nitrogen-fixing nodule tissue. Plant Physiol. 95:111-115. Berry, A.M., and L.A. Sunell. 1990. The infection process and nodule development. p. 61-8l. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Beyazova, M., and M.P. Lechevalier. 1992. Low-frequency restriction fragment analysis of Frankia strains (Actinomycetales). Int. J. System. Bacteriol. 42:422-433. Bloom, R.A., B.C. Mullin, and R.L. Tate, III. 1989. DNA restriction patterns and solution hybridization studies of Frankia isolates from Myrica pensylvanica. Appl. Environ. Microbiol. 55:2155-2160. Bosco, M., M.P. Fernandez, P. Simonet, R. Materassi, and P. Normand. 1992. Evidence that some Frankia sp. strains are able to cross boundaries between Alnus and Elaeagnus host specificity groups. Appl. Environ. Microbiol. 58:1569-1576. Burggraaf, A.J.P. 1984. Isolation, cultivation and characterization of Frankia strains from actinorhizal root nodules. Ph.D. thesis. State University, Leiden, the Netherlands. Burggraaf, A.J.P., and W.A. Shipton. 1982. Estimates of Frankia growth under various pH and temperature regimes. Plant Soil 69:135-147. Burggraaf, A.J.P., and W.A. Shipton. 1983. Studies on the growth of Frankia isolates in relation to infectivity and nitrogen fixation (acetylene reduction). Can. J. Bot. 61:27742782. Burleigh, S., and J.G. Torrey. 1990. Effectiveness of different Frankia cell types as inocula for the actinorhizal plant Casuarina. Appl. Environ. Microbiol. 56:2565-2567. Callaham, D., P. Del Tredici, and J.G. Torrey. 1978. Isolation and cultivation in vitro of the actinomycete causing root nodulation in Comptonia. Science 199:899-902. Cruz-Cisneros, R., and M. Valdes. 1991. Actinorhizal nodules on Aldolphis infesta (H.B.K.) Meissner (Rhamnaceae). Nitrogen Fix. Tree Res. Rep. 9:87-89. Dawson, J.O. 1990. Interactions among actinorhizal and associated plant species. p. 299-316. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Diem, H.G., and Y.R Dommergues. 1983. The isolation of Frankia from nodules of Casuarina. Can. J. Bot. 61:2822-2825. Diem, H.G., and Y.R. Dommergues. 1988. Isolation, characterization and cultivation of Frankia. p. 227-254. In N.S. Subba Rao (ed.) Biological nitrogen fixation, recent developments. Gordon and Breach Sci. Publ., New York. Diem, H.G., and Y.R. Dommergues. 1990. Current and potential uses and management of Casuarinaceae in the tropics and subtropics. p. 317-342. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Diem, H.G., Gauthier, D., and Y.R. Dommergues. 1982. Isolation of Frankia from nodules of Casuarina equisetifolia. Can. J. Microbiol. 28:526-530. Dobritsa, S.V. 1982. Extrachromosomal circular DNAs in endosymbiont vesicles from Alnus glutinosa nodules. FEMS Microbiol. Lett. 15:87-9l. Dobritsa, S.v., S.N. Novik, and o.s. Stupar. 1990. Infectivity and host specificity of strains of Frankia. Mikrobiologiya 59:314-320. Evans, H.J., B. Koch, and R. Klucas. 1972. Preparation of nitrogenase from nodules and separation into components. Meth. Enzymol. 24:470-476. Faure-Raynaud, M., M.A. Bonnefoy-Poirier, and A. Moiroud. 1990a. Diversity of Frankia stains [sic] isolated from actinorhizae of a single Alnus rubra cultivated in nursery. Symbiosis 8:147-160.

324

MYROLD

Faure-Raynaud, M., C. Daniere, A. Moiroud, and A. Capellano. 1990b. Preliminary characterization of an ineffective Frankia derived from a spontaneously neomycin-resistant strain. Plant Soil 129:165-172. Faure-Raynaud, M., C. Daniere, A. Moiroud, and A. Capellano. 1991. Diversity of Frankia stratns isolated from single nodules of Alnus glutinosa. Plant Soil 132:207-211. Fernandez, M.P., H. Meugnier, P.A.D. Grimont, and R. Bardin. 1989. Deoxyribonucleic acid relatedness among members of the genus Frankia. Int. J. Syst. Bacteriol. 39:424429. Fleming, P.I., E.R. Williams, and J.W. Trunbull. 1988. Growth and nodulation and nitrogen fixation of provenances of Casuarina cunninghamiana inoculated with a range of Frankia sources. Austral. J. Bot. 36:171-181. Fontaine, M.S., P.H. Young, and J.G. Torrey. 1986. Effects of long term preservation of Frankia strains on infectivity, effectivity and in vitro nitrogenase activity. Appl. Environ. Microbiol. 51:694-698. Gardes, M., J. Bousquet, and M. Lalonde. 1987. Isozyme variation among 40 Frankia strains. Appl. Environ. Microbiol. 53:1596-1603. Gardes, M., and M. Lalonde. 1987. Identification and subgrouping of Frankia strains using sodium dodecyl sulfate polyacrylamide gel electrophoresis. Physiol. Plant. 70:22372244. Hahn, D., M. Dorsch, E. Stackebrandt, and A.D.L. Akkermans. 1989a. Synthetic nucleotide probes for identification of Frankia strains. Plant Soil 118:211-219. Hahn, D., R. Kester, M.J.C. Starrenburg, and A.D.L. Akkermans. 199Oa. Extraction of ribosomal RNA from soil for detection of Frankia with oligonucleotide probes. Arch. Microbiol. 154:329-335. Hahn, D., M.P. Lechevalier, A. Fischer, and E. Stackebrandt. 1989b. Evidence for a close phylogenetic relationship between members of the genera Frankia, Geodermatophilus, and "Blastococcus" and emendation of the family Frankiaceae. Syst. Appl. Microbiol. 11:236-242. Hahn, D., M.J.C. Starrenburg, and A.D.L. Akkermans. 1988. Variable compatibility of cloned Alnus glutinosa ecotypes against ineffective Frankia strains. Plant Soil 107:233243. Hahn, D., M.J.C. Starrenburg, and A.D.L. Akkermans. 1990b. Oligonucleotide probes against rRNA as a tool to study Frankia strains in root nodules. Appl. Environ. Microbiol. 56:1324-1346. Harry, D.E., D.C. Yang, and J.O. Dawson. 1991. Nucleotide sequence and diversity in 16S ribosomal RNA from Frankia. Plant Soil 131:143-146. Hibbs, D.E., and K. Cromack, Jr. 1990. Actinorhizal plants in Pacific Northwest forests. p. 343-363. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Higgins, M.L., and M.P. Lechevalier. 1969. Poorly lytic bacteriophage from Dactylosporangium thailandensis (Actinomycetales). J. Virol. 3:210-216. Higgins, M.L., M.P. Lechevalier, and H.A. Lechevalier. 1967. Flagellated actinomycetes. J. Bacteriol. 94:1446-1451. Hilger, A.B., and D.D. Myrold. 1991. Method for extraction of Frankia DNA from soil. Agric. Ecosys. Environ. 34:107-113. Hilger, A.B., and D.D. Myrold. 1992. Quantitation oLfoil Frankia by bioassay and gene probe methods: response to host and non-host rhizospheres and liming. Acta CEcol. 13:505-506 Hilger, A.B., Y. Tanaka, and D.D. Myrold. 1991. Inoculation of fumigated nursery soil increases nodulation and yield of bare-root red alder (Alnus rubra Bong.). New Forests 5:35-42. Hoagland, R.D., and DJ. Arnon. 1950. The water culture for growing plants without soil. California Agric. Exp. Stn. Circ. 347. Holman, R.M., and C.R. Schwintzer. 1987. Distribution of spore-positive and spore-negative nodules of Alnus incana spp. rugosa in Maine, USA. Plant Soil 104:103-111. Hooker, J.E., and C. T. Wheeler. 1987. The effectivity of Frankia for nodulation and nitrogen fixation in Alnus rubra and A. glutinosa. Physiol. Plant. 70:333-341. Huss-Danell, K. 1978. Nitrogenase activity measurements in intact Alnus incana. Physiol. Plant. 43:372-376. Huss-Danell, K. 1990. The physiology of actinorhizal nodules. p. 129-156. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego.

FRANKIA & THE ACTINORHIZAL SYMBIOSIS

325

Huss-Danell, D., and D.D. Myrold. 1994. Intrageneric variation in nodulation of Alnus: Consequences for quantifying Frankia infective units in soil. Soil BioI. Biochem. 26:525-531. Jamann, S., M.P. Fernandez, and A. Moiroud. 1992. Genetic diversity of Elaeagnaceaeinfective Frankia strains isolated from various soils. Acta CEcol. 13:395-406. Jamann, S., M.P. Fernandez, and P. Normand. 1993. Typing method for N2-fixing bacteria based on PCR-RFLP-application to the characterization of Frankia strains. Mol. Ecol. 2:17-26. Kashanski, C.R., and C.R. Schwintzer. 1987. Distribution of spore-positive and spore-negative nodules of Myrica gale in Maine, USA. Plant Soil 104:113-120. Kurdali, F., F. Rinaudo, A. Moiroud, and A.M. Domenach. 1990. Competition for nodulation and 15N2-fixation between a Sp+ and a Sp- Frankia strain in Alnus incana. Soil BioI. Biochem. 22:57-64. Lalonde, M., and H.E. Calvert. 1979. Production of Frankia hyphae and spores as an infective inoculant for Alnus species. p. 95-110. In J.C. Gordon et al. (ed.) Symbiotic nitrogen fixation in the management oftemperate forests. For. Res. Lab., Oregon State Univ., Corvallis. Lalonde, M., L. Simon, J. Bousquet, and A. Seguin. 1988. Advances in the taxonomy of Frankia: Recognition of species alni and elaeagni and novel subspecies pommerii and vandijkii. p. 671-680. In H. Bothe et al. (ed.) Nitrogen fixation: Hundred years after. Fischer, Stuttgart, Germany. Lechevalier, M.P. 1983. Cataloging Frankia strains. Can. J. Bot. 61:2964-2967. Lechevalier, M.P. 1984. The taxonomy of the genus Frankia. Plant Soil 78:1-6. Lechevalier, M.P. 1986. Catalog of Frankia strains. Actinomycetes 19:131-162. Lechevalier, M.P., D. Baker, and F. Horriere. 1983. Physiology, chemistry, serology, and infectivity of two Frankia isolates from Alnus incana subsp. rugosa. Can. J. Bot. 61:2826-2833. Lechevalier, M.P., F. Horriere, and H.A. Lechevalier. 1982. The biology of Frankia and related organisms. Dev. Ind. Microbiol. 23:51-60. Lechevalier, M.P., and H.A. Lechevalier. 1990. Systematics, isolation, and culture of Frankia. p. 35-60. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Maggia, L., S. Nazaret, and P. Simonet. 1992. Molecular characterization of Frankia isolates from Casuarina equisetifolia root nodules harvested in West Africa (Senegal and Gambia). Acta CEcol. 13:453-462. Mansour, S.R., A. Dewedar, and J.G. Torrey. 1990. Isolation, culture, and behavior of Frankia strain HFPCgI4 from root nodules of Casuarina glauca. Bot. Gaz. 151:490-496. Mian, S., G. Bong, and C. Rodriquez-Barrueco. 1976. Effective and ineffective root-nodules in Myrica faya. proc. R. Soc. London B194:285-293. Mort, A., P. Normand, and M. Lalonde. 1983. 2-O-methyl-D-mannose, a key sugar in the taxonomy of Frankia. Can. J. Microbiol. 29:993-1002. Mullin, B.C., and C.S. An. 1990. The molecular genetics of Frankia. p. 195-214. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Murray, M.G., and W.F. Thomson. 1980. Rapid isolation of high molecular weight plant DNA. Nucleic Acids Res. 8:4321-4324. Myrold, D.D., and K. Huss-Danell. 1994. Population dynamics of Alnus-infective Frankia in a forest soil with and without host trees. Soil BioI. Biochem. 26:533-540. Myrold, D.D., A.B. Hilger, and S.H. Strauss. 1990. Detecting Frankia in soil using PCR. p. 429. In P.M. Gresshoff et al. (ed.) Nitrogen fixation: Achievements and objectives. Chapman and Hall, New York. Nazaret, S., B. Cournoyer, P. Normand, and P. Simonet. 1991. Phylogenetic relationships among Frankia genomic species determined by use of amplified 16S rDNA sequences. J. Bacteriol. 173:4072-4078. Nazaret, S., P. Simonet, P. Normand, and R. Bardin. 1989. Genetic diversity among Frankia isolated from Casuarina nodules. Plant Soil 118:241-247. Nick, G., E. Paget, P. Simonet, A. Moiroud, andP. Normand. 1992. The nodular endophytes of Coriaria spp. form a distinct lineage within the genus Frankia. Mol. Ecol. 1:175-182. Nittayajarn, A., and D.D. Baker. 1989. Methods for the quantification of Frankia cell biomass. Plant Soil 118:199-204.

326

MYROLD

Nittayajarn, A., B.C. Mullin, and D.D. Baker. 1990. Screening of symbiotic frankiae for host specificity by restriction fragment length polymorphism analysis. Appl. Environ. Microbiol. 56:1172-1174. Normand, P., B. Cournoyer, P. Simonet, and S. Nazaret. 1992. Analysis of a ribosomal RNA operon in the actinomycete Frankia. Gene 111:119-124. Normand, P., and M. Lalonde. 1982. Evaluation of Frankia strains isolated from provenances of two Alnus species. Can. J. Microbiol. 28:1133-1142. Normand, P., and M. Lalonde. 1986. The genetics of actinorhizal Frankia: A review. Plant Soil 90:429-453. Normand, P., P. Simonet, and R. Bardin. 1988. Conservation of nif sequences in Frankia. Mol. Gen. Genet. 213:238-246. Normand, P., P. Simonet, J.L. Butour, C. Rosenberg, A. Moiroud, and M. Lalonde. 1983. Plasmids in Frankia sp. J. Bacteriol. 155:32-35. Parson, W.L., L.R. Robertson, and C.V. Carpenter. 1985. Characterization and infectivity of a spontaneous variant isolated from Frankia sp. WEY 0131391. Plant Soil 87:31-42. Picard, C., C. Ponsonnet, E. Paget, X. Nesme, and P. Simonet. 1992. Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction. Appl. Environ. Microbiol. 58:2717-2722. Prin, Y., E. Duhoux, H.G. Diem, Y. Roederer, and Y.R. Dommergues. 1991a. Aerial nodules in Casuarina cunninghamiana. Appl. Environ. Microbiol. 57:871-874. Prin, Y., L. Maggia, B. Picard, H.G. Diem, and Ph. Goullet. 1991b. Electrophoretic comparison of enzymes from 22 single-spore cultures obtained from Frankia strain DRS 140102. FEMS Microbiol. Lett. 77:223-228. Prin, Y., F. Mallein-Gerin, and P. Simonet. 1993. Identification and localization of Frankia strains in Alnus nodules by in situ hybridization of nifH mRNA with strain-specific oligonucleotide probes. J. Exp. Bot. 44:815-820. Prin, Y., M. Neyra, and H.G. Diem. 1990. Estimation of Frankia growth using Bradford protein and INT reduction activity estimates: Application to inoculum standardization. FEMS Microbiol. Lett. 69:91-96. Puppo, A., L. Dimitrijevic, H.G. Diem, and Y.R. Dommergues. 1985. Homogeneity of superoxide dismutase patterns in Frankia strains from Casuarinaceae. FEMS Microbiol. Lett. 30:43-46. Quispel, A. 1958. Symbiotic nitrogen fixation in non-leguminous plants. IV. The influence of some environmental conditions on different phases of the nodulation process in Alnus glutinosa. Acta Bot. Nerd. 7:191-204. Quispel, A., and A.J.P. Burggraaf. 1981. Frankia, the diazotrophic endophyte from actinorhizas. p. 229. In A.H. Gibson and W.E. Newton (ed.) Current perspectives in nitrogen fixation. Austral. Acad. Sci., Canberra, Australia. Quispel, A., A.B. Svedsen, J. Schripsema, W.J. Bass, C. Erkelens, and J. Lugtenburg. 1989. Identification of dipterocarpol as isolation factor for the induction of primary isolation of Frankia from root nodules of Alnus glutinosa (L.) Gaertner. Mol. Plant-Microbe Interact. 2: 107 -112. Racette, S., and J.G. Torrey. 1989. The isolation and infectivity of a Frankia strain from Gymnostoma papuanum (Casuarinaceae). Plant Soil 118:165-170. Reddell, P., and G.D. Bowen. 1985. Frankia source affects growth, nodulation and nitrogen fixation in Casuarina species. New Phytol. 100:115-122. Reddell, P., P.A. Rosbrook, G.D. Bowen, and D. Gwaze. 1988. Growth responses in Casuarina cunninghamiana plantings to inoculation with Frankia. Plant Soil 108:79-86. Rosbrook, P.A., and G.D. Bowen. 1987. The ability of three Frankia isolates to nodulate and fix nitrogen with four species of Casuarina. Physiol. Plant. 70:373-377. Rosbrook, P.A., A.J.P. Burggraaf, and P. Reddell. 1989. A comparison of two methods and different media for isolating Frankia from Casuarina root nodules. Plant Soil 120:187193. Sanginga, N., G.D. Bowen, and S.K.A. Danso. 1990. Genetic variability in symbiotic nitrogen fixation within and between provenances of two Casuarina species using the 15N_ labeling methods. Soil BioI. Biochem. 22:539-547. Schultz, N.A., and D.R. Benson. 1989. Developmental potential of Frankia vesicles. J. Bacteriol. 171:6873-6877. Schwintzer, C.R. 1990. Spore-positive and spore-negative nodules. p. 177-193. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego.

FRANKIA & THE ACTINORlllZAL SYMBIOSIS

327

Schwintzer, c.R., and J.D. Tjepkema. 1990. The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Seguin, A., and M. Lalonde. 1990. Micropropagation, tissue culture, and genetic transformation of actinorhizai plants and Betu{a. p. 215-238. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Sellstedt, A. 1989. Occurrence and activity of hydrogenase in symbiotic Frankia from fieldcollected Alnus incana. Physiol. Plant. 75:304-308. Sellstedt, A., and K. Huss-Danell. 1984. Growth, nitrogen fixation and relative efficiency of nitrogenase in Alnus incana grown in different cultivation systems. Plant Soil 78:147158. Silvester, W.B., S.L. Harris, and J.D. Tjepkema. 1990. Oxygen regulation and hemoglobin. p. 157-176. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Simon, L., S. Jabaji-Hare, J. Bousquet, and M. Lalonde. 1989. Confirmation of Frankia species using cellular fatty acids analysis. System. Appl. Microbiol. 11:229-235. Simonet, P., A. Capellano, E. Navarro, R. Bardin, and A. Moiroud. 1984. An improved method for lysis of Frankia with achromopeptidase allows detection of new plasmids. Can. J. Microbiol. 30:1292-1295. Simonet, P., M.-C. Grosjean, A.K. Misra, S. Nazaret, B. Cournoyer, and P. Normand. 1991. Frankia genus-specific characterization by polymerase chain reaction. Appl. Environ. Microbiol. 57:3278-3286. Simonet, P., J. Haurat, P. Normand, R. Bardin, and A. Moiroud. 1986. Localization of nif genes on a large plasmid in Frankia sp. strain ULQ 0132105009. Mol. Gen. Genet. 204:492-495. Simonet, P., N.T. Le, E.T. du Cros, and R. Bardin. 1988. Identification of Frankia strains by direct DNA hybridization of crushed nodules. Appl. Environ. Microbiol. 54:25002503. Simonet, P., P. Normand, A.M. Hirsch, and A.D.L. Akkermans. 1990a. The genetics of the Frankia-actinorhizal symbiosis. p. 77-109. In P.M. Gresshoff (ed.) The molecular biology of symbiotic nitrogen fixation. CRC Press, Boca Raton, FL. Simonet, P., P. Normand, A. Moiroud, and R. Bardin. 1990b. Identification of Frankia strains in nodules by hybridization of polymerase chain reaction products with strainspecific oligonucleotide probes. Arch. Microbiol. 153:235-240. Simonet, P., P. Normand, A. Moiroud, and M. Lalonde. 1984. Restriction enzyme digestion patterns of Frankia plasmids. Plant Soil 87:49-60. Smolander, A. 1990. Frankia populations in soils under different tree species-with special emphasis on soils under Betula pendula. Plant Soil 121:1-10. Smolander, A., and M-L. Sarsa. 1990. Frankia strains of soil under Betula pendula: Behaviour in soil and in pure culture. Plant Soil 122:129-136. Smolander, A., and V. Sundman. 1987. Frankia in acid soils of forests devoid of actinorhizal plants. Physiol. Plant. 70:297-303. Smolander, A., C. van Dijk, and V. Sundman. 1988. Survival of Frankia strains introduced into soil. Plant Soil 106:65-72. Sougoufara, B., S.K.A. Danso, H.G. Diem, and Y.R. Dommergues. 1990. Estimatin~ Nz fixation and N derived from soil by Casuarina equisetifolia using labelled 15N fertilizer: Some problems and solutions. Soil BioI. Biochem. 22:695-701. Sougoufara, B., L. Maggia, E. Duhoux, and Y.R. Dommergues. 1992. Nodulation and Nz fixation in nine Casuarina clone-Frankia strain combinations. Acta fficol. 13:497-503. Stowers, M.D. 1987. Collection, isolation, cultivation and maintenance of Frankia. p. 29-53. In G.H. Elkan (ed.) Symbiotic nitrogen fixation technology. Marcel Dekker, New York. Tjepkema, J.D., and J.G. Torrey. 1979. Symbiotic nitrogen fixation in actinomycete-nodulated plants. Preface. Bot. Gaz. (Chicago) Suppl. 140:i-ii. Tjepkema, J.D., C.R. Schwintzer, and D.R. Benson. 1986. Physiology of actinorhizal nodules. Annu. Rev. Plant Physiol. 37:209-232. Torrey, J.G. 1987. Endophyte sporulation in root nodules of actinorhizal plants. Physiol. Plant. 86:581-583. Torrey, J.G. 1990. Cross-inoculation groups within Frankia and host-endosymbiont associations. p. 83-106. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego.

328

MYROLD

Tortosa, R.D., and M. Cusata. 1991. Effective nodulation of rhamnaceous actinorhizal plants induced by air dry soils. Plant Soil 131:229-233. Tunlid, A., N.A. Schultz, D.R. Benson, D.B. Steele, and D.C. White. 1989. Differences in fatty acid composition between vegetative cells and N2-fixing vesicles of Frankia sp. strain CpIl. Proc. Natl. Acad. Sci. USA 86:3399-3403. Tzean, S.S., and J.G. Torrey. 1989. Spore germination and the life cycle of Frankia in vitro. Can. J. Microbiol. 35:801-806. van Dijk, C. 1978. Spore formation and endophyte diversity in root nodules of Alnus glutinosa (L.) Vill. New Phytol. 81:601-615. van Dijk, C. 1984. Ecological aspects of spore formation in the Frankia-Alnus symbiosis. Ph.D. thesis. State University, Leiden, the Netherlands. van Dijk, C., and E. Merkus. 1976. A microscopical study of the development of a spore-like stage in the life cycle of the root-nodule endophyte of Alnus glutinosa (L.) Gaertn. New Phytol. 77:73-91. van Dijk, C., A. Sluimer, and A. Weber. 1988. Host range differentiation of spore-positive and spore-negative strain types of Frankia in stands of Alnus glutinosa and Alnus incana in Finland. Physiol. Plant. 72:349-358. van Dijk, C., and A. Sluimer-Stolk. 1990. An ineffective strain type of Frankia in the soil of natural stands of Alnus glutinosa (L.) Gaertner. Plant Soil 127:107-121. Weber, A. 1986. Distribution of sl?ore-positive and spore-negative nodules in stands of Alnus glutinosa and Alnus incana In Finland. Plant Soil 96:205-213. Weber, A., E.L. Nurmiaho-Lassila, and V. Sundman. 1987. Features of intrageneric AlnusFrankia specificity. Physiol. Plant. 70:289-296. Weber, A., M-L. Sarsa, and V. Sundman. 1989. Frankia-Alnus incana symbiosis: Effect of endophyte on nitrogen fixation and biomass production. Plant Soil 120:291-297. Wheeler,C.T., J.E. Hooker, A. Crowe, and A.M.M. Berrie. 1986. The improvement and utilization in forestry of nitrogen fixation by actinorhizal plants with special reference to Alnus in Scotland. Plant Soil 90:393-406. Wheeler, C.T., and I.M. Miller. 1990. Current and potential uses of actinorhizal plants in Europe. p. 365-389. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego. Winship, L.J., and J.D. Tjepkema. 1990. Techniques for measuring nitrogenase activity in Frankia and actinorhizal plants. p. 263-280. In C.R. Schwintzer and J.D. Tjepkema (ed.) The biology of Frankia and actinorhizal plants. Academic Press, San Diego.

Published 1994

Chapter 17 Filamentous Fungi DENNIS PARKINSON, The University of Calgary, Calgary, Alberta, Canada

Fungi play important roles in several soil processes. These range from organic matter decomposition and nutrient cycling to interactions with plant roots (symbiotic or parasitic). A wide diversity of fungi have been shown to be present in soils and they represent a substantial portion of the total soil microbial biomass. Investigators of soil fungi encounter considerable problems in qualitative and quantitative studies because the fungi exist in soil in a variety of morphological and physiological states (e.g., active, dormant and even dead hyphae, various types of spores, and resting structures such as sclerotia). In many studies, it is important to assess the active hyphal fungi in soil or organic matter samples; but, despite some progress, this is still a difficult task. In the past decade, several reviews on methods for studying soil fungi have appeared (e.g., Parkinson, 1982; Kendrick & Parkinson, 1990; Frankland et aI., 1990; Seifert, 1990; Parkinson & Coleman, 1991). These emphasize the problems stated above. The present chapter deals particularly with methods for studying communities of filamentous fungi in soil and organic matter samples. Topics such as sampling (chapter 1 in this book), statistical analyses (chapter 2 in this book), direct microscopic methods (chapter 6 in this book) and mycorrhizal fungi (chapter 18 in this book) are dealt with in other chapters of the present volume as are immunological methods (chapter 28 in this book) which are so valuable for studies on individual species. It must be emphasized here that, prior to embarking on a detailed study of soil fungi (as for any group of soil organisms), there should be a clear definition of the aims of the study. Subsequently, methods can be chosen that allow fulfillment of these aims. The details of the various methods provided in this chapter should be regarded as examples and not as hard-and-fast protocols. All methods will probably need modification for use with different soils. Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 329

330

PARKINSON

17-1 QUALITATIVE STUDIES: ISOLATION METHODS 17-1.1 Choice of Appropriate Isolating Media For studies on the community structure of soil fungi in the vast majority of cases, it is necessary to isolate these organisms onto nutrient media. Therefore, the choice of appropriate media for these studies is of prime importance. At one time, the aim in synecological studies of soil fungi was to develop a nonselective medium, i.e., one which would allow the isolation of any fungus present in the soil. Therefore, media such as plain water agar were used as the primary isolation medium, from which isolates were transferred to nutrient agar media for subsequent identification. This hope for the development of nonselective media has given way to the more realistic approach of using a medium (or small group of media) which allows the isolation of the maximum number of fungal taxa from the soil under study. Probably the use of a wide range of selective media would be the most efficient approach to a complete study of soil fungal community structure, but this is rarely (if ever) feasible because of the required personnel and time demands. The most appropriate isolation medium may vary depending on the soil type being studied (and the prior experiences of the individual investigator). Therefore, it is dangerous to recommend dogmatically any particular medium. Before carrying out a large study on the mycota of a particular soil, a proper comparative study of a range of media should be carried out to determine the medium that allows isolation of the maximum number of fungal taxa. Numerous nutrient media have been used in studies of filamentous fungi in soil. Among the most frequently used media are Czapek-Dox agar with or without small amounts of yeast extract; malt extract agar with or without materials such as peptone, dextrose, and yeast extract; and, dextrose-peptone agar with or without growth retardants (see later). Media incorporating soil extract have not been used as frequently for isolating soil fungi as for isolations of soil bacteria. Recipes for a wide range of media have been well documented elsewhere (Parkinson et aI., 1971; Johnson & Curl, 1972; Smith & Onions, 1983; Gams et aI., 1987; Seifert, 1990). Recipes for some commonly used types of media are provided here: 1. Czapek-Dox medium-30.0 g of sucrose, 3.0 g of sodium nitrate (NaN03), 1.0 g of potassium monohydrogen phosphate (K2HP04), 0.5 g of magnesium sulfate heptahydrate (MgS0 4·7H20), a trace of ferrous sulfate (FeS04)' 15.0 g of agar, 1000 mL of distilled water, and 0.5 to 1.0 g of yeast extract (if desired): Dissolve inorganic constituents separately; add FeS0 4 last. Add sucrose just before sterilization.

FILAMENTOUS FUNGI

331

2. Malt extract agar: Variants of this medium range from 20.0 g of malt extract, 15.0 g of agar, and 1000 mL of distilled water to 20.0 g of malt extract, 20.0 g of dextrose, 1.0 g of peptone, 15.0 g of agar, and 1000 mL of distilled water. The amount of malt extract can be decreased if desired, and small amounts of yeast extract can be added (0.5-1.0 g.L -1). 3. Dextrose-peptone agar: a. Dissolve 10.0 g of dextrose, 5.0 g of peptone, 1.0 g of potassium dihydrogen phosphate (KH2P0 4), 0.5 g of magnesium sulfate heptahydrate (MgS0 4·7H20) and 20.0 g of agar in 1000 mL of distilled water, and heat. Add 3.3 mL of 1% rose bengal. Autoclave the medium, cool to about 50°C, and add 30.0 mg of streptomycin (Martin, 1950). b. Dissolve 5.0 g of dextrose, 1.0 g of peptone, 2.0 g of yeast extract, 1.0 g of ammonium nitrate (NH4N0 3), 1.0 g of potassium monohydrogen phosphate (K2HP0 4), 0.5 g of magnesium sulfate heptahydrate (MgSO 4· 7H20) , a trace of ferric chloride hexahydrate (FeCI3 ·6H2 0), 5.0 g of oxgall (dehydrated fresh bile), 1.0 g of sodium propionate (NaC3H s0 2), and 20.0 g of agar in 1000 mL of distilled water, and heat. Autoclave the medium, and after cooling to about 50°C, add 30.0 mg of aureomycin and 30.0 mg of streptomycin (Papavizas & Davey, 1959). If soil extracts are required for attempts to make media of low selectivity, the usual procedure in their preparation is to autoclave 1 kg of soil with 1 L water (20 min at either 6.8 or 9.1 kg of pressure). The material is then filtered, and the filtrate is made up to 1 L. If the resultant liquid is still cloudy, a little CaS04 is added, the liquid is allowed to stand, and then filtration is repeated. The soil extract can be solidified with agar (1.5%) and used with or without the addition of other nutrients, such as 0.02% K2 HP0 4 or 0.1 % glucose. In soils containing large numbers of bacteria, it is frequently necessary to amend the chosen isolation medium with antibacterial substances in an attempt to reduce competition from the soil bacteria and thus increase the likelihood of isolating slow growing fungi of low competitive ability. Acidification of the medium (pH 3.5-5.0) was frequently used, and is still used in some investigations. Acids should be added to the medium after sterilization and include 0.5% malic acid, 0.1 % lactic acid, or 0.1 to 1.0 MHCl or H 2S0 4. If acidification of media is applied the investigator should ensure that this practice is not having a selective effect on the isolation of fungi. Bacterial growth retardants such as crystal violet (10 mg L -1), potassium tellurite (100 mg L -1), oxgall (0.5-1.5%), rose bengal (30-700 mg L -1), sodium deoxycholate and sodium propionate have been used on

332

PARKINSON

isolation media (Littman, 1947; Martin, 1950; Bakerspigel & Miller, 1953; Papavizas & Davey, 1959; Watling, 1971; see review by Seifert, 1990). Fungal growth retardants have been used to restrict rapidly growing isolates and prevent slower growing fungi from being overgrown. Orthophenyl phenol, rose bengal, oxgall, dichloran (2-6-dichloro-4-nitroaniline), and benomyl (1-butycarbonyl-2-benzimadazole carbonic acid methyl ester) are examples of such retardants and these have been discussed in detail by Seifert (1990). Suggested concentrations of these compounds in nutrient media are given below. The use of antibiotics for the suppression of bacterial development during isolations of soil fungi is now common and has tended to supplant the use of the agents mentioned above. Examples of antibiotic additions to some standard media have been given in the recipes outlined above. Streptomycin and aureomycin are commonly used antibiotics at concentration ranges of 100 to 200 and 25 to 100 mg L -1, respectively. These antibiotics should be added to the nutrient agar after it has been autoclaved and cooled (prior to pouring). A wide range of media have been developed for the selective isolation of either specific physiological groups of soil fungi (e.g., cellulolytic spp., chitinolytic spp. etc.) or specific taxa (e.g., basidiomycetes, Fusarium spp., Phytophthora spp. etc.). New media for specific studies are being developed regularly, and examples of a range of selective media were given by Johnson and Curl (1972). The most recent, detailed survey has been given by Seifert (1990). Basidiomycetes are usually isolated from soil with relative infrequency. This is disturbing, particularly when studies are being made on forest litter soil systems where production of fruit bodies of a range of species of basidiomycetes is observed. This problem may center on the nutritional requirements of these fungi or their slow growth rates and inability to compete with faster growing more aggressive species or both. Attempts to isolate basidiomycetes from soil usually involve the addition of chemical retardants to the growth of faster growing fungi (see earlier). Orthophenyl phenol, pentachloronitrobenzene, dichloran, phenol and benomyl (to maximum of 5 mg L -1) are examples of such retardants. Worrall (1991), in a comparison of several media for the selective isolation of wood-decay hymenomycetes, found that malt extract agar with benomyl (2 mg L -1) and dichloran (2 mg L -1) and with 100 mg L -1 streptomycin added after autoclaving was the most suitable medium. Special media have been developed for isolating and culturing mycorrhizal fungi (Molina & Palmer, 1982). Three media commonly used for these purposes are:

1. Hagem's agar (Modess, 1941): 5.0 g glucose, 5.0 g malt extract, 0.5 g potassium dihydrogen phosphate (KH2P0 4 ), O.S g magnesium sulfate heptahydrate (MgS0 4 ·7H2 0), 0.5 g ammonium chloride (NH4CI), 0.5 mL of a 1% ferric citrate (FeC6 H s0 7 ·SH2 0) solution, 15.0 g agar, 1000 mL distilled water.

FILAMENTOUS FUNGI

333

2. Melin-Norkrans medium (Norkrans, 1949): 2.5 g sucrose, 25 /lg thiamine hydrochloride, 0.5 g potassium dihydrogen phosphate (KH 2P04 ), 0.15 g magnesium sulfate heptahydrate (MgS0 4 ·7H2 0), 0.25 g ammonium monohydrogen phosphate [(NH4 hHP0 4 ], 1.2 mL of a 1% solution of ferric chloride (FeCI3 ), 15.0 g agar, 1000 mL distilled water. 3. Modified Melin-Norkrans medium (Marx, 1969): 10 g sucrose, 3.0 g malt extract, 100 /lg thiamine hydrochloride, 0.5 g magnesium sulfate heptahydrate (MgS0 4 ·7H20), 0.25 g ammonium monohydrogen phosphate [(NH4 hHP0 4 ], 0.05 g calcium chloride (CaCI 2 ), 0.025 g sodium chloride (NaCl), 1.2 mL of a 1% solution of ferric chloride (FeCI 3 ), 15.0 g agar, 1000 mL distilled water. Modified Melin-Norkrans medium has been further modified by various workers, e.g., Mexal and Reid (1973) substituted glucose for sucrose and 48 ppm Sequestrene 330 (Ciba-Geigy Corp., Greensboro, NC) for FeCl3 solution. In an attempt to isolate only basidiomycete mycorrhizal fungi, Danielson (1982) added benomyl (10 mg L -1) to Modified Melin-Norkrans medium. Following primary isolation of fungi from soil, individual colonies are subcultured to provide pure cultures for identification. A wide range of media is available for the maintenance of pure cultures, many of which are "natural" media, e.g., potato extract, potato dextrose, potato-carrot, V8 juice (Campbell Soup Co., Camden, NJ) cherry extract and malt extract. For identification of specific taxa, it is important to use the media recommended by the monographs being used for identification. Details on culture storage, maintenance, and observations have been given by Gams et aI., 1987; Smith, 1988; Kendrick & Parkinson, 1990, and Seifert, 1990. 17-1.2 General Soil Studies 17-1.2.1 Soil Dilution Plate Method

Until the 1950s, this method was used almost exclusively for studies on soil fungal communities. For reasons which are given later, the use of other methods has increased in the past three decades, however the soil dilution plate method continues to be used in some studies. Ever since the work of Brierley et al. (1928), every facet of this method has been subjected to close scrutiny, e.g., initial weight of the soil sample, volume of initial suspension, type of suspending fluid, type and time of agitation to produce a homogenous soil suspension, type of diluent, type of nutrient medium, and method of plating (i.e., incorporation into the agar medium or surface plating). Wollum (1982) provided clear details on the use of this method for isolating various groups of soil microorganisms from non-rhizosphere and rhizosphere soil. It has been suggested (Parkinson et aI., 1970) that if this method is used, individual workers should vary the details of the method to suit the soil type and so forth with

334

PARKINSON

which they are working or the types of organisms being studied. Therefore, the postulation of a standard procedure for this method would provide one of only dubious value. One example of a methodological procedure used for a forest soil is as follows: 1. 10 g soil are dispensed in 90 mL diluent (0.2% solution of dextrin in soil extract) using mechanical shaking (250 strokes min- 1 for 15 min). 2. From the initial suspension a dilution series is made in the conventional way using I-mL blowout pipettes of known accuracy and 0.2% dextrin soil extract as the diluent. 3. Aliquots (0.1 mL) of chosen dilutions (e.g., 10- 5 and 10- 6) are dispensed onto a chosen nutrient agar (section 17-1.1) solidified in petri dishes and spread plates are prepared. 4. The plates are incubated at the temperature required by the investigator (for studies of mesophilic fungi this is normally 22-25 0c) for at least 14 d. 5. Colonies developing on the plates are subcultured for subsequent identification. Before embarking on a large-scale use of this method, it is important to determine the exact details of this method that are optimal for use with the soil under study. If this method is used for soil fungal community studies, it should be remembered that: 1. It has been shown (Warcup, 1957) that the large majority of fungal colonies developing on soil dilution plates originate from spores or other propagules and not from hyphal fragments. 2. In view of the foregoing comment the method can be used for assessment of fungal spore content of soil samples but gives little information on fungi present as hyphae in those samples. 3. When this method is used, it is usually impossible to ascertain from which microhabitats in the soil the fungi originated. 17-1.2.2 Soil Plate Methods This method, developed by Warcup (1950) is essentially a simple variation of the soil dilution plate method in which small amounts of soil are dispersed in known volumes of an appropriate nutrient agar medium. 1. A small amount of soil (0.005-0.015 g) is taken from the large soil sample, using a sterile needle with a flattened tip, and is placed in the base of a petri dish. 2. The small soil sample is thoroughly broken up (using sterile needles) and dispersed over the base of the dish. Addition of a drop of sterile water can enhance this dispersion.

FILAMENTOUS FUNGI

335

3. Molten but cooled (45°C) agar medium (8-10 mL) is added to the dish, which is then rotated gently to disperse the soil particles in the agar medium. 4. The plates prepared in this way are then incubated (usually 2225°C) and fungi developing from the soil particles are isolated into pure culture for identification. Like the soil dilution plate method, this method is selective for fungi present as spores in soil samples. However, because of the simplicity and speed of execution of this method, replicate soil plates from each of several soil samples can be prepared quickly, it is valuable for preliminary examination of soil mycofloras prior to using other isolation methods. In attempts to isolate fungi present in soil as actively growing or dormant hyphae three approaches have been developed: Immersion methods, direct hyphal isolation, and washing methods. 17-1.2.3 Immersion Methods

In an attempt to achieve direct isolation of fungi present in soil as actively growing hyphae, Chesters (1940) developed the immersion tube method. Subsequently, this method has been modified in a variety of ways (e.g., Thornton, 1952; Mueller & Durrell, 1957; Parkinson, 1957; Wood & Wilcoxson, 1960; Anderson & Huber, 1965; Luttrell, 1967). In these methods, the isolating agar medium (frequently water agar or a chosen nutrient agar) is placed at the desired position(s) in the soil profile under study and is left in situ for a suitable period (which is determined by preliminary tests but is usually 5-7 d). The isolating medium is usually separated from the soil by an air gap, so that any fungi entering that medium must have grown actively across the air gap. After the appropriate time in the soil, the pieces of immersion apparatus are collected from the field, and in the laboratory, small pieces of the isolation medium are plated onto an appropriate nutrient agar medium. Fungi developing on these plates are assumed to have been present in the soil in an active hyphal state. Probably the simplest form of this method and the one requiring least special equipment is that described by Mueller and Durrell (1957). 17-1.2.3.1 Procedure 1. Immersion tubes are made from autoclavable plastic centrifuge tubes in the walls of which holes (0.25 cm diam.) have been bored at desired positions. The tube is wrapped with plastic tape. 2. The tubes are filled, to within 4 cm of the top, with the chosen nutrient medium and then plugged and autoclaved. 3. The tubes are taken to the field study site, the plastic tape round each tube is pierced with a large sterile needle at positions corresponding to the holes bored in the centrifuge tube.

336

PARKINSON

4. The tubes are embedded in the soil and left in situ for a predetermined number of days. 5. Following removal from the soil, the tubes are taken to the laboratory where a core of agar is taken from each tube. Each agar core is cut into small pieces each of which is plated onto the chosen nutrient medium. 6. The isolation plates are incubated and fungi developing on the plates are subcultured for subsequent identification. The defects of this type of method appear to be: 1. Placing tubes, plates etc. into soil can cause germination of spores in the soil around the immersed materials, so that fungi isolated may be derived from these spores and not from hyphae previously active in the soil. 2. There may be interspecific competition for entry into the immersion apparatus and for subsequent colonization of the medium in the apparatus. In such cases, only fungi with high competitive abilities may be isolated. 3. Small soil invertebrates may carry fungal spores into the nutrient medium in the immersion apparatus. Notwithstanding a small number of excellent studies (e.g., Thornton, 1956; Sewell, 1959), these immersion methods have been used too infrequently to allow full evaluation of their value in comparison with other, more frequently used, methods. 17-1.2.4 Direct Hyphal Isolation

Warcup (1955) described a method for picking hyphae from small soil samples and plating these hyphae onto nutrient medium: 1. Small soil samples or soil crumbs are saturated with sterile water and then broken with a fine jet of sterile water. 2. Heavier soil particles are allowed to sediment, and the fine particles are decanted off. 3. This procedure is repeated several times until only the heavier particles remain. 4. The remaining particles are spread in a film of sterile water and examined under a dissecting microscope. 5. Any fungal hyphae observed are picked out using sterile needles or very fine forceps, and plated onto nutrient agar. 6. It is frequently necessary to attempt to remove adherent bacteria, other propagules and organic debris from the hyphae prior to plating. 7. When plating, the position of each hyphal inoculum is marked on the base of the plate. The plates are incubated and the plated fragments are observed at least daily under the microscope to ensure that any fungal growth recorded has originated from the plated hyphae and not from adherent propagules etc.

FILAMENTOUS FUNGI

337

This method is useful for restricted, specific studies; however, because of the time required to deal with each small soil sample, it would appear to be impractical for investigations that entail regular, replicate isolations from large numbers of soil samples. When it is applied, unless great care is taken, the method tends to be selective for hyphae of large diameter, for hyphae that do not fragment easily, and for dark-pigmented hyphae. It tends to be selective against fine hyaline hyphae and for hyphae that are closely associated with organic fragments. Obviously, considerable skill is required for the efficient application of this method. 17-1.2.5 Soil Washing

In an attempt to achieve a simpler method that requires less skill and experience for isolation of fungi present as hyphae in soil samples, various methods for soil washing have been developed. These methods range from the simple washing-decanting method (Watson, 1960), where soil samples are held in 500-mL Erlenmeyer flasks and are washed with numerous changes of sterile water, to the use of automated, multisample washing machines (Hering, 1966; Bissett & Widden, 1972). All these methods attempt to achieve the removal of as many fungal spores as possible from a wide range of soil microhabitats (i.e., soil particles of different sizes and organic fragments) that can then be plated separately. In most soil washing methods the following features are evident: 1. Soil samples are placed in separate sterile boxes, each of which contains a number of sieves of graded size (the sieve sizes being determined by the soil type being studied and by the aims of the investigation) . 2. The soil samples are vigorously washed (about 2 min. wash-I) with numerous changes of sterile water. The washing action is achieved either by passing sterile air through the system or by physical shaking of the apparatus. The number of washings required to achieve efficient removal of fungal spores from the soil samples being studied must be determined by preliminary tests, i.e., by plating aliquots of washing water after each washing onto replicate nutrient agar plates and counting fungal colonies developing on these plates (see Harley and Waid, 1955 for details). 3. Following thorough washing, soil from the sieves is removed onto sterile filter paper to remove excess water. 4. Individual particles of known size and nature (inorganic or organic) are then plated onto nutrient agar medium. The number of particles per plate depends on the soil type and degree of fungal development (usually one to four particles per plate). 5. Fungi developing on the plates, following incubation, are subcultured for subsequent identification. The efficiency of soil washing methods will be related to the physical nature of the soil under study. Williams et al. (1965) studied this matter and

338

PARKINSON

demonstrated that sandy soil could be washed more efficiently than soils with high clay (or humus colloid) content. In all cases, it is unlikely that all spores are removed from soil samples by washing; however, a high proportion are removed. For extensive studies on the mycofiora of soil where replicate soil samples taken at each of numerous sample times are to be studied, the construction of an automatic, multibox, washing apparatus is thoroughly worthwhile. However, in small (or very restricted) studies, the use of the simple washing-decantation method is probably appropriate. Whatever method is used, information can be gained on fungi present as hyphae (by plating washed particles) and fungi present as spores (by plating washing water). 17-1.3 Special Substrates 17-1.3.1 Roots 17-1.3.1.1 Root Washing. While various washing methods have been used since the 1930s the Harley and Waid (1955) method and its variations are the most frequently used for general studies of root surface fungi: 1. Roots taken from soil are thoroughly washed in tap water to remove macroscopic debris. 2. The root system is cut into 1 to 2 cm. pieces, the choice of the region(s) of the root system cut in this way depends on the aims of the study. 3. The 1 to 2 cm pieces are placed in 25-mL screw-capped vials and thoroughly washed in several changes of sterile water. 4. The exact number of washings required is determined by preliminary tests as for soil washing but is usually between 15 and 40 washings. 5. Each washing is usually of 2 min duration and is vigorous-being effected manually, mechanically, or in an automated apparatus. 6. After the appropriate number of washings the root pieces are transferred to petri dishes containing sterile filter paper to remove excess water from the roots. 7. The washed, surface-dried roots are cut into small segments. Twomillimeter segments have been used frequently, but smaller segments < 1 mm allow isolation of larger number of species. 8. The small segments are plated onto nutrient medium (one to four segments per plate, depending on degree of fungal colonization of the roots). 9. The plates are incubated and fungi developing from the root segments are isolated into pure culture. 17-1.3.1.2 Surface Disinfestation. This has been used commonly by plant pathologists for isolating microorganisms from internal plant tissues. When isolating fungi from roots the following general procedure can be followed:

FILAMENTOUS FUNGI

339

1. Pieces of root are placed in a suitable surface disinfestant of standard concentration for standard time. 2. Then the roots must be thoroughly washed with sterile water to remove all traces of the disinfestant. 3. Excess water is removed from the roots using sterile filter paper, the roots are cut into small pieces (1-2 mm) which are plated onto nutrient agar. Before using this method, each step must be tested to ensure that appropriate concentrations of disinfestant, time of application, and amount of washing necessary for removal of disinfectant from the roots are determined. Commonly used sterilants include NaOCI solution, 0.1% (wt/vol) HgCl 2 and Nance's solution (0.1% (wt/vol) HgCl 2 with 0.5% TeepoP in distilled water). Times of surface sterilization vary greatly depending on the type of tissue being studied but are usually between 10 sand 2 min. Zak and Bryan (1963) described a surface disinfestation method for isolating ectomycorrhizae fungi from plant roots: 1. Root samples from the field are transported to the laboratory in such a way as to prevent root drying and the temperature of the roots rising above that of the field soil. 2. In the laboratory, the roots are washed under tap water to remove as much adhering soil as possible. 3. 1.5 to 3.0 cm lengths of roots bearing mycorrhizae are cut off and thoroughly washed in detergent (sometimes ultrasonic cleansing may be needed to remove fine debris). For detergent washing 15 to 20 pieces of root are placed in a screw-capped plastic vial (4 by 2 cm) that has been perforated (top, bottom, and sides). The vial is placed in a jar of detergent solution and thoroughly shaken. 4. The vial containing root pieces is removed from the detergent and the roots are washed in tap water to remove all detergent. 5. The washed roots are then placed in 1% NaOCI solution and shaken for 10 min. (in the same manner as the detergent washing). 6. The disinfectant is removed by thorough washing with sterile distilled water. 7. Root pieces are removed aseptically from the perforated vial and placed in sterile dishes (excess water can be removed from the roots using sterile filter paper). Mycorrhizae are cut from the roots and plated onto an appropriate agar medium (see section 17-1.1). Danielson (1991) surface sterilized root segments by dipping them in 95% ethanol and soaking in 30% H 20 2 before rinsing in two changes of ice-cold sterile water for 1 h. Ice-cold water was used because it stops the action of H 20 2 sooner than would water at room temperature.

1 British

Drug Houses, Ville St. Laurent, Montreal; or substitute sodium dedocyl sulfate.

PARKINSON

340

17-1.3.1.3 Root Fragmentation. Various types of root fragmentation with or without preliminary surface sterilization have been used to attempt to isolate fungi from the inner tissues of roots. The simplest method (Warcup, 1960) involves placing small pieces of thoroughly washed root (washed as described in section 17-1.3.1.1) in drops of sterile water in petri dishes (1 drop root piece- I dish-I) and then dissecting the roots into as many small fragments as possible using sterile needles. The chosen nutrient agar medium is poured into each dish, after which the dishes are carefully rotated to ensure dispersal of the root fragments in the nutrient medium. More violent disruption of washed roots can be achieved either by using a mortar and pestle or by using some type of blender (Stover & Waite, 1953; Clarke & Parkinson, 1960). Following disruption, small amounts of macerate are plated as described above. In such fragmentation methods, it is frequently difficult to discern from which root tissues the developing fungal colonies have arisen. The possibility exists that in the maceration process, substances inhibitory to the growth of fungi may be released from the plant tissues. If more exact information is required on the location of fungi within roots, root dissection is necessary. Waid (1957) described a method in which roots previously washed (described above) are placed in sterile water and, using a sterile scalpel etc., are dissected (working under a binocular dissecting microscope). Vascular tissue may be separated from the cortex, and these fractions can be plated separately onto the chosen nutrient agar medium. 17-1.3.2 Other Organic Matter 17-1.3.2.1 Direct Observation Followed by Isolation. Frankland et al. (1990) pointed out the value of maintaining samples (macroscopic) of different types of organic material (e.g., leaves, stems, pieces, or sections of wood) in damp chambers and recording the various fungi that sporulate or the material under these conditions. Careful use of this simple method can frequently allow isolation of species that are not obtained by use of the methods described in the previous sections. Seifert (1990) described a method that includes the following steps: 1. The base of a petri dish is lined with filter paper and with paper towelling or pulp around the inner rim. All this material is moistened with sterile water. 2. The organic material is placed in the dish which is then covered with a loosely fitting lid. 3. Dishes prepared in this way are incubated (room temperature) and examined regularly, using a dissecting microscope, for fungal sporulation that may begin as early as 2 d after initiation of damp chambers. 4. Spores of the fungi on the organic matter can be removed, using an agar-tipped needle, to nutrient agar plates.

341

FILAMENTOUS FUNGI

5. If insect development on the organic matter is likely to become a problem, a solid insecticide (e .g., Vapona strips) can be placed in the damp chamber. 17-1.3.2.2 Direct Observation of Ectomycorrhizal Roots. The importance of direct observation of ectomycorrhizal roots in attempting identification of the fungal symbionts was emphasized by Zak (1973). Recent publications (e.g., Agerer, 1986; Ingleby et aI., 1990; Danielson, 1991) have given details on methods for observing roots and the characters of the mycorrhizas that allow symbiont identification. Both macroscopic and microscopic characters of mycorrhizas are important in identification of symbionts. Macroscopic characters include: color, size, form associated hyphae, hyphal strands, rhizomorphs, and sclerotia. Microscopic characters, seen in squash preparations or transverse sections include: hyphal size, presence of cystidia, hyphal ornamentation, crystals or other hyphal exudates, and hyphal wall pigmentation. 17-1.3.2.3 Serial Washing, Surface Sterilization, Fragmentation, and Dissection. All these methods have been described, above, for the isolation of fungi from roots. They can be used equally well for studies of fungi associated with other plant parts and organic materials in various stages of decomposition. For large pieces of organic matter a procedure that involves coarse blending followed by serial washing (predetermined number of washings), air drying and plating fragments (one to two per plate) onto the chosen agar medium has been used frequently. 17-1.4 Selective Methods All the methods described above are used for synecological studies of soil fungi, i.e., where an attempt is made to isolate as many species as possible from a soil sample. However, it is desirable frequently to study the distribution of specific taxa or of specific physiologicaVsubstrate groups of fungi in soil. Therefore, methods selective for different taxa etc. have been developed. To itemize a list of such methods is impossible in the present restricted chapter on the filamentous fungi. Scrutiny of journals such as Phytopathology over the past two decades would show that a considerable number of media have been developed for isolating specific soil-borne pathogens. Johnson and Curl (1972) provided a general account of methods for isolating these organisms. Parkinson et ai. (1971) provided data on selective media useful for studying different physiologicaVsubstrate groups of soil microorganisms. More recent information on these groups of soil fungi has been provided by Frankland et ai. (1990) and Seifert (1990).

342

PARKINSON

17-2 QUANTITATIVE METHODS 17-2.1 Introduction For several decades, the soil dilution plate method was used exclusively in attempts to quantify soil fungal communities. However, the data obtained using this method, expressed as numbers of fungi per unit weight of soil (an unreasonable expression, given the hyphal growth form of the vast majority of the soil fungi), in reality give an indication of the number of spores present in the soil sample (see comments in section 17-1). Other isolation methods may indicate the degree of fungal colonization of different substrates in soil and may allow comparisons of different soils using calculations of frequency of occurrence of fungi or other mathematical analyses (see Frankland et aI., 1990). However, these approaches give no indications of fungal biomass. It appears that three general types of methods are available for measuring fungal biomass: (i) direct observation methods, (ii) chemical methods, and (iii) physiological methods. Other chapters in this volume deal in detail with these three approaches (see chapters 6 and 35 in this book). Therefore, only general comments on these methods are given here.

17-2.2 Direct Observation Methods In the early studies of soil fungi, when there was some debate on whether or not fungi were active in soil, direct observation of soil smears and of slides which had been buried in soil for appropriate time periods demonstrated fungal growth in soil. However, attempts to convert such observations into mass of fungal hyphae in soil samples were rare. Nicholas and Parkinson (1967) discussed the relative merits of several direct observation methods. The methods discussed were modified impression slides, soil sections and soil-agar films and the conclusion was that, while not ideal, the soil-agar film method was the least prone to experimental and observational errors. With the development of superior methods for preparing soil sections, application of fluorochromes and the use of fluorescence microscopy (Altemiiller & van Vliet-Lanoe, 1990), the use of soil sectioning has been shown to be valuable for studies on the spatial distribution of soil microorganisms (Postma & Altemiiller, 1990). It may well prove valuable for obtaining data on microbial biomass associated with specific soil microhabitats.

17-2.2.1 Soil-Agar Film Method The soil-agar film method (Jones & Mollison, 1948 as modified by Thomas et aI., 1965) has been described for use with soil samples (Parkinson, 1982) and for litter (Frankland, 1978; Parkinson, 1982; Frankland et aI., 1990). Lodge and Ingham (1991) have compared various modifications

FILAMENTOUS FUNGI

343

of the soil-agar film method for estimating fungal biovolumes in litter and soil, and report a coverslip well technique to be least variable and most preferable for making agar films. The recommendations and comments made by Frankland et aI. (1990) and by Lodge and Ingham (1991) for preparation and observation of agar films can be summarized as follows: 1. The duration and intensity of methods used for dispersal of hyphae in diluent are critical factors. These must be determined in preliminary experiments. 2. For soil samples, shaking or sonication are much preferable to maceration (which breaks hyphae into small fragments requiring high magnification for detection). Litter samples require maceration, the times for which must be predetermined. 3. It is necessary to determine the optimal dilution of the sailor litter samples, i.e., the dilution giving the maximum estimate of hyphal length. 4. Different investigators recommended different ranges of magnification for measuring hyphallengths. Baath and SOderstrom (1980) found that increasing the magnification from 800 x to 1250 x doubled the hyphallengths measured. Lodge and Ingham (1991) recommended using the lowest magnification under which hyphae can be readily identified, which will depend on the predominant size of hyphae (particularly diameter) and the substrate relationships of the hyphae. 5. The experimental design is important. Frankland et aI. (1978) and Lodge and Ingham (1991) have discussed the importance of properly replicated observations of several replicate films (these numbers of replicates being predetermined.) Frankland et aI. (1990) observed that optimum replication is usually achieved by increasing the number of field samples of soil or litter relative to the number of soil-agar films made and microscopic fields observed. 6. Conversions of hyphal length measurements to biomass require data on the proportions of hyphae falling into various diameter classes, their density, and water content. It is unsatisfactory to use values for these parameters that are taken from the older literature (e.g., Parkinson et aI., 1971). Of the factors mentioned above hyphal diameter is most important (see Bfulth & SOderstrom, 1979). Measurements should be made with care, and should be made on fresh agar films to avoid errors due to shrinkage (Jenkinson & Ladd, 1981). Moisture content and density of hyphae vary with species, age of hyphae and growth conditions, and measurements of these values should be made using mycelium collected from the field (Lodge, 1987) or using hyphae grown under carefully designed, low nutrient conditions (Frankland et aI., 1990). 7. Estimates of biomass in living hyphae can be made by assuming that the presence of cell contents (observed using phase contrast microscopy) indicates live hyphae (Frankland, 1975; Frankland et

PARKINSON

344

al., 1990). However, various staining methods (fluorescent stains) are used extensively (see below). 17-2.2.2 Membrane Filter Method Hansen et al. (1974) developed a membrane filter method which is more rapid than the soil-agar film method for measuring hyphallengths in soil samples (which can be converted to biomass as for the soil-agar films). Variations on this method have been described by Paul and Johnson (1977) and Sundman and SiveHi (1978). More recently, Elmholt and Kj~ller (1987) described a general procedure for studying hyphal biomass in cultivated field soils. Details on membrane filter methods for studying general soil fungi and mycorrhizal fungi are given elsewhere (chapters 6 and 18 in this book). Baath and Soderstrom (1980) compared the soil-agar film and membrane filter methods for making measurements of hyphallength in several soils. They found that the former method gave higher hyphallength values, but in studies comparing different soils the membrane filter method was preferred because of the ease and rapidity of its execution. West (1988) found significantly greater hyphallengths on membrane filters of grassland soil suspensions treated with fluorescent brightener than on soil-agar films stained with phenolic aniline blue and viewed with phase contrast microscopy. Bardgett (1991) found the membrane filter method particularly useful in a comparative study of different grassland soils. Vital staining, using fluorescein diacetate (FDA), of membrane filtered soil suspensions, has been used to assess the lengths of live hyphae in soil samples (SOderstrom, 1977). Using this method, FDA-stained hyphae are frequently found to comprise only a small percentage of the total hyphae. Soderstrom (1979) suggested that the method of preparing soil suspensions (vigorous homogenization) could cause cytoplasm leakage from some hyphal fragments and thereby underestimate the length of viable hyphae. However, Ingham and Klein (1984) used a gentle extraction procedure and obtained significant correlations between lengths of FDAstained hyphae and metabolic activity. Gray (1990), in a detailed account of the FDA-staining method, reiterates the problem of the effect of drastic homogenization destroying the integrity of some hyphae. In the preparation of soil suspensions for membrane filtration and calculation of fungal biomass from hyphallength measurements, many of the comments made regarding soil-agar film preparation are equally relevant. Morgan et al. (1991) described an automated image analysis method for use in the membrane filter method, in which Calcaftuor M2R and FDA were used to stain total and viable hyphae, respectively. The results showed excellent correlation with those obtained using manual microscopic hyphallength measurements. Using this method "operator differences" and "fatigue-induced bias" are eliminated.

FILAMENTOUS FUNGI

345

17-2.3 Chemical Methods The quantitative extraction from soil of chemical constituents of fungal cells, and the subsequent use of such data to estimate total fungal biomass is an attractive possibility. In the choice of such a cell constituent several criteria must be taken into account (Jenkinson & Ladd, 1981): 1. It should be present in live cells at concentrations that fall within a known range. 2. It should be absent from dead cells and other non-living materials in soil. 3. An accurate method must be available for assaying the compound. With respect to studies on filamentous fungi, estimations of chitin and ergosterol have received most attention (see Frankland et aI., 1990). Chitin determinations involve measuring the quantity of glucosamine released by hydrolysis of chitin (Ride & Drysdale, 1972). Unfortunately, chitin does not fulfill the first two criteria listed above. For general studies of mixed soil fungal communities where substantial arthropod communities are also present, it is difficult to obtain valid conversion factor (glucosamine content to fungal biomass) and, at present, such conversion is only appropriate in studies of single species of fungi (Herbert, 1990). In some soils, most of the glucosamine measured is present in dead organic matter (West et aI., 1987). On the other hand, the use of high pressure liquid chromatography (HPLC) has enhanced the speed and sensitivity of chitin analyses (Zelles, 1988). Ergosterol determinations are considered to be more specific indicators of fungal biomass than are glucosamine measurements (Newell et aI., 1988). Various analytical methods are available for determining ergosterol; however, it is difficult to convert estimates of ergosterol to fungal biomass because of considerable interspecific and age variation in ergosterol content of hyphae (Newell et aI., 1987). Nevertheless it has been used to monitor changes in fungal biomass in soil (West et aI., 1987) and in litter (J. Lodge, personal communication). Despite the attraction of chemical methods, it appears that, for studies of soil fungal or plant litter communities, direct observation methods are superior for fungal biomass determinations. 17-2.4 Physiological Methods Anderson and Domsch (1978) used glucose stimulated respiration to determine total microbial biomass in soil samples. This method, now called substrate-induced respiration (SIR) is discussed elsewhere (chapter 36 in this book) as is the method for estimating active soil microbial biomass by mathematical analysis of respiration curves (Van de Werf & Verstraete, 1987).

346

PARKINSON

The SIR method becomes important for determining fungal biomass in soil when it is coupled with the addition of selective inhibitors (e.g., actidione [cycloheximide] and streptomycin, protein synthesis inhibitors for fungi and bacteria, respectively) applied at predetermined optimal concentrations (Anderson & Domsch, 1973, 1975). Differences in respiration rates of antibiotic treated and untreated glucose-amended soil samples allow calculations of the relative proportions of fungi and bacteria in the total microbial biomass (Anderson & Domsch, 1975). Data obtained using this method generally show fungi to be the dominant component of the total microbial biomass, with bacterial to fungal biomass ratios ranging from 40:60 to 10:90. For each soil type being studied it is essential to determine carefully the required concentrations of each inhibitor. The criterion for these determinations is that the sum of the inhibition effects of actidone and streptomycin when applied individually should equal the effect when these inhibitors are applied as a mixture (Anderson & Domsch, 1973). Following determination of these inhibitor concentrations, experiments with short incubation periods (6-8 h) should be used to eliminate problems with population shifts and inhibitor degradation (Anderson & Domsch, 1975). Anderson and Domsch (1973, 1975) applied actidone and streptomycin in powder form to glucose (powder) amended soil samples. West (1986) recommended the addition of glucose and inhibitors in solution, to make them more readily available to the whole soil microbial community, particularly in soil where moisture is limiting. West (1986) also recommended that the soil + glucose and inhibitors be incubated for 3.5 h prior to the assays of inhibition. Beare et al. (1991) preincubated plant litter samples in solutions of each of the inhibitors individually or the mixture of inhibitors for 12 h at 4 °C prior to the addition of the optimal glucose concentration after the samples had been brought to room temperature (about 22°C). It is unclear, as yet, what proportion of the total soil microbial biomass responds quickly to glucose. But the use of the selective inhibition method can be used only with reference to the glucose-responsive component of the microbial biomass, i.e., that component which is undergoing protein synthesis immediately following glucose addition (Van de Werf & Verstraete, 1987; Wardle & Parkinson, 1990). From the data provided by Anderson and Domsch (1975), West (1986) and Wardle and Parkinson (1990) it can be seen that the method applies only to a fraction of the microbial biomass-the inhibition caused by the mixture of actidione and streptomycin is typically < 60% of glucose stimulated respiration. Therefore, two assumptions are necessary when this method is used: (i) that the bacterial to fungal biomass ratio is the same in both inhibitor sensitive and insensitive components of the total microbial biomass (West, 1986); and, (ii) that bacterial and fungal components of the total biomass respire at the same rate, per unit biomass, following addition of glucose. Domsch et al. (1979) discussed the applicability of this method for use with a range of soils. They considered it to be objective, but commented on the need for more work to elucidate its sensitivity. This comment still holds

347

FILAMENTOUS FUNGI

good, particularly for studies on highly organic soils and decomposing plant litter. West (1986) found that, while the modified method was not applicable to all soils, it could be used as complementary to studies using direct observation methods.

REFERENCES Agerer, R. 1986. Studies on ectomycorrhizae. II. Introducing remarks on characterization and identification. Mycotaxon 26:473-492. Altemiiller, H -J ., and B. van Vliet-Lanoe. 1990. Soil thin section fluorescence microscopy. p. 565-578. In L.A. Douglas (ed.) Soil micromorphology. Elsevier Sci. Pub I. B.Y., Amsterdam. Anderson, J.P.E., and K.H. Domsch. 1973. Quantification of bacterial and fungal contributions to soil respiration. Arch. Mikrobiol. 93:113-127. Anderson, J.P.E., and K.H. Domsch. 1975. Measurements of bacterial and fungal contributions to respiration of selected agricultural and forest soils. Can. J. Microbiol. 21:314322. Anderson, J.P.E., and K.H. Domsch. 1978. A physiological method for the quantitative measurement of microbial biomass in soils. Soil BioI. Biochem. 10:215-221. Baath, E., and B. Soderstrom. 1979. The significance of hyphal diameter in calculations of fungal biovolume. Oikos 33:11-14. Baath, E., and B. Soderstrom. 1980. Comparison of the agar-film and membrane-filter methods for estimation of hyphal lengths in soil, with particular reference to the effect of magnification. Soil BioI. Biochem. 12:385-387. Bakerspigel, A., and J.J. Miller. 1953. Comparison of oxgall, crystal violet, streptomycin, and penicillin as bacterial growth inhibitors in platings of soil fungi. Soil Sci. 76: 123-126. Bardgett, R.D. 1991. The use of the membrane filter technique for comparative measurements of hyphallengths in different grassland sites. Agric. Ecosystems Environ. 34:115119. Beare, M.H., C.L. Neely, D.C. Coleman, and W.L. Hargrove. 1991. Characterization of a substrate-induced respiration method of measuring fungal, bacterial and total microbial biomass on plant residues. Agric. Ecosystems Environ. 34:65-73. Bissett, J., and P. Widden. 1972. An automatic, multi-chamber soil-washing apparatus for removing fungal spores from soil. Can. J. Microbiol. 18:1399-1409. Brierley, W.B., S.T. Jewson, and M. Brierley. 1928. The quantitative study of soil fungi. Int. Congr. Soil Sci. Trans. 1st (Washington, DC) 3:48-71. Chesters, C.G.C. 1940. A method for isolating soil fungi. Trans. Br. Mycol. Soc. 24:352-355. Clarke, J.H., and D. Parkinson. 1960. A comparison of three methods for the assessment of fungal colonization of seedling roots of leek and broad bean. Nature (London) 188: 166167. Danielson, R.M. 1982. Taxonomic affinities and criteria for identification of the common ectendomycorrhizal symbiont of pines. Can. J. Bot. 60:7-18. Danielson, R.M. 1991. Temporal changes and effects of amendments on the occurrence of sheathing (ecto-) mycorrhizas of conifers in oil sands tailings and coal spoil. Agric. Ecosystems Environ. 35:261-281. Domsch, K.H., Th. Beck, J.P.E. Anderson, B. Soderstrom, D. Parkinson, and G. Trolldenier. 1979. A comparison of methods for soil microbial population and biomass studies. Z. Pflanzenernaehr. Bodenkd. 142:520-533. Elmholt, S., and A. Kj¢ller. 1987. Measurement of the length of fungal hyphae by the membrane filter technique as a method for comparing fungal occurrence in cultivated soils. Soil BioI. Biochem. 19:679-682. Frankland, J.e. 1975. Estimation of live fungal biomass. Soil BioI. Biochem. 7:339-340. Frankland, J.C., J. Dighton, and L. Boddy. 1990. Methods for studying fungi in soil and forest litter. p. 343-404. In R. Grigorova and J.R. Norris (ed.) Methods in microbiology. Vol. 22. Techniques in microbial ecology. Academic Press, New York. Frankland, J.C., D.K. Lindley, and M.J. Swift. 1978. A comparison of two methods for the estimation of mycelial biomass in leaf litter. Soil BioI. Biochem. 10:323-333.

348

PARKINSON

Gams, W., H.A. van der Aa, A.J. van der Plaats-Niternik, R.A. Samson, and J.A. Stalpers. 1987. CBS course of mycology. 3rd ed. Centraalbureau voor Schimmelcultures. Baarn, the Netherlands. Gray, T.R.G. 1990. Methods for studying the microbial ecology of soil. p. 309-342. In R. Grigorova and J.R. Norris (ed.) Methods in microbiology. Vol. 22. Techniques in microbial ecology. Academic Press, New York. Hansen, J .F., T.F. Thingstad, and J. Goks\:1yr. 1974. Evaluations of hyphallengths and fungal biomass in soil by a membrane filter technique. Oikos 25:102-107. Harley, J.L., and J.w. Waid. 1955. A method for studying active mycelia on living roots and other surfaces in the soil. Trans. Br. Mycol. Soc. 38:104-118. Herbert, R.A. 1990. Methods for enumerating microorganisms and determining biomass in natural environments. p. 1-39. In R. Grigorova and J.R. Norris (ed.) Methods in microbiology. Vol. 22. Techniques in microbial ecology. Academic Press, New York. Hering, T.F. 1966. An automatic soil-washing apparatus for fungal isolation. Plant Soil 25:195-200. Ingham, E.R., and D.A. Klein. 1984. Soil fungi: relationships between hypha I activity and staining with fluorescein diacetate. Soil BioI. Biochem. 16:273-278. Ingleby, K., P.A. Mason, F.T. Last, and L.V. Fleming. 1990. Identification of ectomycorrhizas. ITE Res. Publ. No.5. HMSO, London. Jenkinson, D.S., and J.N. Ladd. 1981. Microbial biomass in soil. p. 415-471. In E.A. Paul and J.N. Ladd (ed.) Soil biochemistry. Marcel Dekker, New York. Johnson, L.F., and E.A. Curl. 1972. Methods for research on the ecology of soil-borne plant pathogens. Burgress Publ. Co., Minneapolis. Jones, P.C.T., and J.E. Mollison. 1948. A technique for the quantitative estimation of soil micro-organisms. J. Gen. Microbiol. 2:54-69. Kendrick, W.B., and D. Parkinson. 1990. Soil fungi. p. 49-68. In D.L. Dindal (ed.) Soil biology guide. John Wiley and Sons, New York. Littman, M.L. 1947. A culture medium for the primary isolation of fungi. Science 106:109111. Lodge, D.J. 1987. Nutrient concentrations, percentage moisture and density of field collected fungal mycelia. Soil BioI. Biochem. 19:727-734. Lodge, D.J., and E.R. Ingham. 1991. A comparison of agar film techniques for estimating fungal biovolumes in litter and soil. Agric. Ecosystems Environ. 34:131-144. Luttrell, E.S. 1967. A strip bait for studying the growth of fungi in soil and aerial habitats. Phytopathology 57: 1266-1267. Martin, J.P. 1950. Use of acid, rose bengal and streptomycin in the plate method for estimating soil fungi. Soil Sci. 69:215-232. Marx, D.H. 1969. The influence of ectotrophic mycorrhizal fungi on resistance of pine roots to pathogenic infections. I. Antagonism of mycorrhizal fungi to root pathogenic fungi and bacteria. Phytopathology 59:153-163. Mexal, J., and C.P.P. Reid. 1973. The growth of selected mycorrhizal fungi in response to induced water stress. Can. J. Bot. 51:1579-1588. Modess, O. 1941. Zur Kenntnis der Mykorrhizabildner von Kiefer und Fichte. Symb. Bot. Usal. 5:1-146. Molina, R., and J.G. Palmer. 1982. Isolation, maintenance, and pure culture manipulation of ectomycorrhizal fungi. p. 115-112. In N.C. Shenck (ed.) Methods and principles of mycorrhizal research. The American Phytopathol. Soc., St. Paul. Morgan, P., C.J. Cooper, N.S. Battersby, S.A. Lee, S.T. Lewis, T.M. Machin, S.C. Graham, and R.J. Watkinson. 1991. Automated image analysis method to determine fungal biomass in soils and on solid matrices. Soil BioI. Biochem. 23:609-616. Mueller, K.E., and L.W. Durrell. 1957. Sampling tubes for soil fungi. Phytopathology 47:243. Newell, S.Y., T.L. Arsuffi, and R.D. Fallon. 1988. Fundamental procedures for determining ergosterol content of decaying plant material by liquid chromatography. Appl. Environ. Microbiol. 54: 1876-1879. Newell, S.Y., J.D. Miller, and R.D. Fallon. 1987. Ergosterol content of salt marsh fungi: effect of growth conditions and mycelial age. Mycologia 79:688-695. Nicholas, D.J., and D. Parkinson. 1967. A comparison of methods for assessing the amount of fungal mycelium in soil samples. Pedobiologia 7:23-41. Norkrans, B. 1949. Some mycorrhiza-forming Tricholoma species. Sven. Bot. Tidskr. 43:485490.

FILAMENTOUS FUNGI

349

Papavizas, G.c., and C.B. Davey. 1959. Evaluation of various media and antimicrobial agents for isolation of soil fungi. Soil Sci. 88:112-117. Parkinson, D. 1957. New methods for the qualitative and quantitative study of fungi in the rhizosphere. Pedologie 7 (no. Spec.):146-154. Parkinson, D., T.R.G. Gray, J. Holding, and H.M. Nagel-de-Boois. 1970. Heterotrophic microflora. p. 34-50. In J. Phillipson (ed.) IBP Handbook 18. B1ackwells Sci. Publ. Ltd., Oxford. Parkinson, D., T.R.G. Gray, and S.T. Williams. 1971. Methods for studying the ecology of soil micro-organisms. IBP Handbook 19. Blackwells Sci. Publ. Ltd., Oxford. Parkinson, D. 1982. Filamentous fungi. p. 949-968. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. Chemical and microbiological properties. 2nd ed. ASA, and SSSA Madison, WI. Parkinson, D., and D.C. Coleman. 1991. Microbial communities, activity and biomass. Agric. Ecosystems Environ. 34:3-33. Paul, E.A., and R.L. Johnson. 1977. Microscopic counting and adenosine 5'-triphosphate measurement in determining microbial growth in soils. Appl. Environ. Microbiol. 34:263-269. Postma, J., and H.-J. Altemiiller. 1990. Bacteria in thin soil sections stained with the fluorescent brightener calcafluor white M2R. Soil BioI. Biochem. 22:89-96. Ride, J.P., and R.B. Drysdale. 1972. A rapid method for the chemical estimation of filamentous fungi in plant tissue. Physiol. Pathol. 2:7-15. Seifert, K.A. 1990. Isolation of filamentous fungi. p. 21-51. In D.P. Labeda (ed.) Isolation of biotechnological organisms from nature. McGraw-Hili Publ. Co., New York. Sewell, G.w.F. 1959. Studies of fungi in a Calluna-heathland soil. II. By the complementary use of several isolation methods. Trans. Br. Mycol. Soc. 42:354-369. Smith, D., and A.H.S. Onions. 1983. The preservation and maintenance of living fungi. Commonwealth Mycol. Inst., Kew, England. Smith, D. 1988. Culture and preservation. p. 75-99. In D.L. Hawksworth and B.E. Kirsop (ed.) Living resources for biotechnology. Cambridge TJniv. Press, Cambridge, England. Soderstrom, B.E. 1977. Vital staining of fungi in pure cultures and in soil with fluorescein diacetate. Soil BioI. Biochem. 9:59-63. Soderstrom, B. 1979. Some problems in assessing the fluorescein diacetate-active fungal biomass in the soil. Soil BioI. Biochem. 11:147-148. Stover, R.H., and B.H. Waite. 1953. An improved method of isolating Fusarium spp. from plant tissue. Phytopathology 43:700-701. Sundman, V., and S. SiveUi. 1978. A comment on the membrane filter technique for estimation of length of fungal hyphae in soil. Soil BioI. Biochem. 10:399-401. Thomas, A., D.P. Nicholas, and D. Parkinson. 1965. Modifications of the agar film technique for assaying lengths of mycelium in soil. Nature (London) 205:105. Thornton, R.H. 1952. The screened immersion plate. A method for isolating soil microorganisms. Research (London) 5:190-191. Thornton, R.H. 1956. Fungi occurring in mixed oakwood and heath soil profiles. Trans. Br. Mycol. Soc. 39:485-494. Van de Werf, H., and W. Verstraete. 1987. Estimation of active soil microbial biomass by mathematical analysis of respiration curves: relation to conventional estimation of total biomass. Soil BioI. Biochem. 19:267-271. Waid, J.S. 1957. Distribution of fungi within the decomposing tissues ofryegrass roots. Trans. Br. Mycol. Soc. 40:391-406. Warcup, J.H. 1950. The soil plate method for isolation of fungi from soil. Nature (London) 166:117-118. Warcup, J.H. 1955. Isolation offungi from hyphae present in soil. Nature (London) 175:953954. Warcup, J.H. 1957. Studies on the occurrence and activity offungi in a wheatfield soil. Trans. Br. Mycol. Soc. 40:237-262. Warcup, J.H. 1960. Methods for isolation and estimation of activity of fungi in soil. p. 3-31. In D. Parkinson and J .S. Waid (ed.) The ecology of soil fungi. Univ. of Liverpool Press, Liverpool. Wardle, D.A., and D. Parkinson. 1990. Response of the soil microbial biomass to glucose, and selective inhibitors, across a soil moisture gradient. Soil. BioI. Biochem. 22:825-834.

350

PARKINSON

Watling, R. 1971. Basidiomycetes, Homobasidiomycetes. p. 216-236. In C. Booth (ed.) Methods in microbiology. Vol. 4. Academic Press, New York. Watson, R.D. 1960. Soil washing improves the value of the soil dilution and plate count method of estimating populations of soil fungi. Phytopathology 50:792-794. West, A.W. 1986. Improvement of the selective respiratory inhibition technique to measure eukaryote: prokaryote ratios in soils. J. Microbiol. Methods. 5:125-138. West, AW., W.D. Grant, and G.P. Sparling. 1987. Use of ergosterol, diaminopimelic acid and glucosamine content of soils to monitor changes in microbial populations. Soil BioI. Biochem. 19:607-612. West, A.W. 1988. Specimen preparation, stain type, and extraction and observation procedures as factors in the estimation of soil mycelial lengths and volumes by light microscopy. BioI. Fert. Soils 7:88-94. Williams, S.T., D. Parkinson, and N.A. Burges. 1965. An examination of the soil washing technique by its application to several soils. Plant Soil 22:167-186. Wollum, AG., II. 1982. Cultural methods for soil microorganisms. p. 781-802. In AL. Page et al. (ed.) Methods of soil analysis. Part 2. Chemical and ffilcrobiological properties. 2nd ed. ASA, Madison, WI. Wood, F.A., and R.D. Wilcoxson. 1960. Another screened immersion plate for isolating fungi from soil. Plant Dis. Rep. 44:594. Worrall, J.J. 1991. Media selective for the isolation of Hymenomycetes. Mycologia 83:296302. Zak, B., and W.e. Bryan. 1963. Isolation of fungal symbionts from pine mycorrhizae. For. Sci. 9:270-278. Zak, B. 1973. Classification of ectomycorrhizae. p. 43-78. In G.C. Marks and T.T. Kozlowski (ed.) Ectomycorrhizae, their ecology and physiology. Academic Press, New York. Zelles, L. 1988. The simultaneous determination of muramic acid and glucosamine in soil by high-performance liquid chromatography with precolumn fluorescence derivatization. BioI. Fert. Soils 6:125-130.

Published 1994

Chapter 18 Vesicular-Arbuscular Mycorrhizal Fungi DAVID M. SYLVIA, University of Florida, Gainesville, Florida

Mycorrhizae are symbiotic associations between beneficial soil fungi and plant roots. They have an important role in increasing plant uptake of P and other poorly mobile nutrients (O'Keefe & Sylvia, 1991). The vesiculararbuscular mycorrhizal (VAM) fungi have a very broad host range-members of more than 90% of all vascular plant families are colonized by them. Due to the obligate nature of these organisms, however, much is still unknown about their biology in natural and managed ecosystems. An early impetus for mycorrhizal research was the prospect for practical utilization of these fungi in forestry and agriculture. This research served mostly to demonstrate the complex biology of the symbiosis, and many efforts at utilization have been frustrated (Jeffries & Dodd, 1991). Today, there is increased research interest in understanding the basic physiology and ecology of mycorrhizae in vivo, in vitro, and in situ. Only through such efforts can we hope to reliably manage mycorrhiza in forestry and agriculture. The purpose of this chapter is to summarize basic methods to: (i) quantify propagules of VAM fungi, (ii) select effective VAM fungi from soil, and (iii) propagate VAM inoculum and apply it in the nursery and field. A wide range of methods are currently used successfully by various research groups-each method needs to be assessed for the particular application required. For further detail and explanation, the reader is referred to several excellent reviews of the methods presented here (Schenck, 1982; Hayman, 1984; Powell & Bagyaraj, 1984; Kendrick & Berch, 1985; Jeffries, 1987; Jeffries & Dodd, 1991; Schenck & Perez, 1990a; Jarstfer & Sylvia, 1992a; Varma et ai., 1992). Methods for manipulating ectomycorrhizal fungi are not discussed here.

Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

351

352

SYLVIA

18-1 QUANTIFICATION OF VESICULAR·ARBUSCULAR MYCORRIDZAL PROPAGULES IN SOIL 18-1.1 Introduction Even though VAM fungi are among the most common fungi in soil, they are often overlooked because they do not grow on standard soildilution plating media. The spores of VAM fungi are larger than those of most other fungi (ranging from 10-1000 I-tm in diam.) and can easily be observed with a dissecting microscope. However, spore counts often underestimate numbers of VAM fungi since colonized roots and hyphae can also serve as propagules. For this reason, various assays have been used to obtain an estimate of total propagule number. 18-1.2 Most Probable Number Assay The most probable number (MPN) assay was developed as a method for estimating the density of organisms in a liquid culture (Cochran, 1950). It was first used to estimate the propagule density of VAM fungi in soil by Porter (1979). Values for a MPN assay can be obtained from published tables (Halvorson & Ziegler, 1933; Fisher & Yates, 1963; de Man, 1975; Alexander,. 1982); however, these tables restrict experimental design, thereby reducing the accuracy that can be obtained. A better approach is to program the equation into a computer and directly solve for the MPN value based on optimal experimental design-increased replication and decreased dilution rates will greatly enhance the accuracy of the result (chapter 5 by Woomer in this book). The general equation for calculating MPN values is presented in chapter 5 of this volume. Numerous factors affect the outcome of an MPN assay (Wilson & Trinick, 1982; Morton, 1985; Adelman & Morton, 1986; Graham & Fardelmann, 1986; An et aI., 1990; O'Donnell et aI., 1992); therefore, the values obtained should be considered relative rather than absolute. Nonetheless, this assay has been a useful tool for estimating propagule numbers in field soil, pot cultures, and various forms of inocula. 18-1.2.1 Important Considerations for Implementing a Most Probable Number Assay 1. Dilution factor - Preliminary studies should be conducted so that

the lowest possible dilutions are used to bracket actual numbers found in the soil. 2. Sample processing-Samples should be kept cool and processed as soon as possible after collection. The sample soil needs to be relatively dry and root pieces > 2 mm in diam. removed from the sample to allow thorough mixing with the diluent soil. Obviously, these treatments will affect propagule numbers and viability, so all samples must be treated in a similar manner.

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

353

3. Diluent soil- The soil preferably should be the same as the original sample and should be pasteurized rather than sterilized. Controls with no sample added should be set up with the pasteurized soil to ensure that all VAM propagules have been eliminated. 4. Host plant-The host must be highly susceptible to VAM colonization, produce a rapidly growing, fibrous root system, and be readily cleared for observation of colonization. Zea mays L. is a good choice. S. Length of assay - Plants need to be grown long enough so that roots fully exploit the soil in each container. It is better to err on the conservative side and grow plants until they are pot bound. Roots with well-developed mycorrhizae are also more easily evaluated. A typical assay may run for 6 to 8 wk. 6. Confirming negative colonization - The entire root system must be examined to confirm a negative reading. 18-1.3 Infectivity Assays A more straight-forward approach for comparing VAM populations among soils is an infectivity assay. The draw back is that actual propagule numbers are not estimated. Plants are grown under standard conditions in soil collected from pot cultures or a field, and root colonization is estimated after 3 to 6 wk (Moorman & Reeves, 1979; Reeves et aI., 1979; Schwab & Reeves, 1981; Koide & Mooney, 1987; Abbott & Robson, 1991). The amount of colonization is assumed to be proportional to the total number of VAM propagules in the soil. The length of the assay is critical. If plants grow for too short a time, the full potential for colonization is not realized; however, plants grown for too long a time may become uniformly colonized despite differences in VAM populations-preliminary studies are needed to select the proper harvest time for a given plant, soil combination. Recently, an infection-unit method has been proposed to quantify mycorrhizal propagules (Franson & Bethlenfalvay, 1989). This study indicates that a count of infection units is a more reliable measure of the number of viable propagules than are other methods. However, this method is only applicable in short-term experiments because infection units are discern able only during the initial stages of colonization. 18-2 QUANTIFICATION OF VESICULAR-ARBUSCULAR MYCORRHIZAL COLONIZATION IN ROOTS 18-2.1 Visualizing Vesicular-Arbuscular Mycorrhizal Fungi in Roots The VAM fungi do not cause obvious morphological changes of roots; however, they produce arbuscules and, in many cases, vesicles in roots. To observe VAM structures within the root, it is necessary to clear cortical

SYLVIA

354

cells of cytoplasm and phenolic compounds, and then to differentially stain the fungal tissue. Phillips and Hayman (1970) published the oft-cited method to visualize VAM fungi in roots using trypan blue in lactophenol, but the use of phenol is now discouraged (Koske & Gemma, 1989). The clearing agent for nonpigmented roots is generally KOH, but H 20 2 (Phillips & Hayman, 1970) or NaOCI (Bevege, 1968; Graham et al., 1992) may be used for pigmented roots. Alternatives to trypan blue for staining are chlorazol black E (Brundrett et al., 1984) and acid fuchsin (Kormanik & McGraw, 1982). For nonpigmented roots, it is also possible to observe colonization non destructively by inducing autofluorescence (Ames et al., 1982). 18-2.1.1 Procedure for Clearing and Staining Roots 1. Place root samples (approximately 0.5 g) in perforated plastic holders (e.g., OmniSette tissue cassettes, Fisher Scientific, Pittsburgh, PA) and store in cold water until they are processed. 2. Place enough 1.8 M KOH into a beaker (without samples) to allow samples to be covered and heat to 80 °C in a fume hood. Goggles, gloves, and vinyl apron should be worn for protection. 3. Place samples in the heated KOH for the desired time-15 min for tender roots such as onion, 30 min for other roots. If samples are still pigmented after the initial treatment, rinse with at least three changes of water and then place them in a beaker with 30% (wt/wt) H 20 2 at 50 °C or 3% (wt/vol) NaOCI, acidified with several drops of 5 M HCI. Transfer roots to water as soon as samples are bleached white or become transparent. Times can vary from several seconds to several minutes. Check roots frequently to avoid destruction of the cortex. 4. Rinse with tap water using five changes of water. 5. Cover the samples with tap water and add 5 mL of conc. HCI for each 200 mL of water, stir, and drain. Repeat once. 6. Dispense enough trypan blue stain into a beaker (without samples) to cover samples and heat to 80 °C. To prepare the stain, add in order to a flask while stirring: 800 mL of glycerine, 800 mL of lactic acid, 800 mL of distilled water, and finally 1.2 g of trypan blue. 7. Place samples in the stain for 30 min, cool, and drain the stain into the large flask for reuse. Use a funnel and screen to remove debris. After several uses, additional trypan blue may be added to extend the use of the stain. 8. Rinse samples with one change of tap water to destain. Additional de staining in water may be necessary for some roots. 9. Store samples in a plastic bag in a refrigerator. 18-2.2 Estimation of Colonized Root Length Various methods have been used to estimate root colonization by VAM fungi (Kormanik & McGraw, 1982; Hayman, 1984). The gridline-

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

355

intersect method has the advantage of providing an estimate of both the proportion of colonized root and total root length (Giovannetti & Mosse, 1980). This is important because some treatments affect root and fungal growth differently. For example, when P is applied total root length may increase more rapidly than colonized root length and thus the proportion of colonized root will decrease even though the actual length of colonized root is increasing. 18-2.2.1 Gridline-Intersect Method 1. Spread a cleared and stained root sample evenly on a scribed plastic petri dish. A grid of squares is scribed on the bottom of the dish prior to use. The lines should be arranged so that the edge of the container does not coincide with a gridline and the two partial squares at each edge together make up a complete square. If the lines are 1.27-cm (0.5 in.) apart, then the total number of intersections is equal to total root length in centimeters (see no. 4 below). 2. Scan the gridlines under a dissecting microscope and record the total number of root intersections with the grid as well as the number of intersects with colonized roots. 3. Verify any questionable colonization with a compound microscope. To do this, cut out a small portion of the root with a scalpel, place it in water on a microscope slide, and look for VAM structures at 100 to 400 x . Remember that the stains are not specific for VAM fungi-other fungi colonizing the root will also stain so it is important to verify the presence of arbuscules or vesicles in the root with a compound microscope. 4. Calculate total and colonized root lengths (R) using the following equation (Newman, 1966): R=rrAn12H

where A is the total area in which roots are distributed, n is the number of intersections between roots and the scribed lines, and H is the total length of the scribed lines. According to Marsh (1971), when the interline distance is 14/11 of the measuring unit (14/11 cm = 0.5 in.) then nA = 2H and total or colonized root length in centimeter equals the number of total or colonized root intersections with the gridlines. 20-2.2.2 Modification of the Gridline-Intersect Method McGonigle et al. (1990) argued that the gridline-intersect method is somewhat subjective because arbuscules may be difficult to distinguish with a dissecting microscope. They proposed use of a magnified-intersect method where roots are observed at 200 x and arbuscules are quantified separately from vesicles and hyphae. Another limitation of the gridlineintersect method is that the intensity of colonization at each location is not estimated. To obtain an estimate of intensity a morphometric technique

SYLVIA

356

(Toth & Toth, 1982) can be used where a grid of dots is placed over an image of squashed roots and colonized cortical cells are counted. 18-2.3 Chemical Determinations Chitin determinations have been used to estimate fungal biomass in roots under controlled experimental conditions (Hepper, 1977; Bethlenfalvay et aI., 1981). However, its utility in native soil is limited because chitin is ubiquitous in nature, where it is found in the cell walls of many fungi and the exoskeletons of insects, and certain soils exhibit physical and chemical properties which prevent the technique from working properly (Jarstfer & Miller, 1985). Recently, assays for ergosterol have been used to quantify ectomycorrhizal fungal biomass (Salmanowicz et aI., 1989; Johnson & McGill, 1990; Martin et al., 1990) and this method should also be useful for VAM fungi. In addition, J.H. Graham (personal communication) has found that fatty acid analysis is applicable to the quantification of VAM fungal colonization. 18-2.3.1 Chitin Determinations 1. Oven-dried and ground roots (10 mg) are mixed with 4 mL of conc. KOH (120 g KOHl100 mL H 20) and autoclaved for 1 h in 15 mL screw-cap centrifuge tubes to degrade chitin to chitosan. For an internal control, root material from noncolonized plants should be assayed to correct for hexosamines in plant tissue and organisms other than VAM fungi. In addition, a standard curve using purified chitin should be included to correct for the incomplete hydrolysis of chitin. 2. Add and mix with a vortex mixer, 8 mL of 75% (voVvol) ice-cold ethanol (75%) to the 4 mL of autoclaved sample. Then layer 0.9 mL of a celite suspension (diatomaceous silica, suspended in 75% ethanol, 1 g per 20 mL) over the first ethanol wash. 3. The samples are centrifuged at approx. 5000 x gat 4°C for 10 min. The pellets are then resuspended and washed sequentially in 8 mL each of ice-cold, 40% ethanol, 0.01 M HCL, and distilled water. 4. The pellet is suspended in 1.5 mL of water and assayed colorimetrically as follows (Ride & Drysdale, 1971). Equal volumes (1.5 mL) of the washed chitosan suspension, 5% (wtlvol) NaN0 2 , and 5% (wtlvol) KHS0 4 are transferred to a centrifuge tube, shaken for 15 min, and centrifuged at 1500 x g for 2 min at 2°C. Two, 1.5-mL subsamples are removed from the tube and 0.5 mL of 12.5% (wtl vol) ammonium sulfamate is added to each, shaken for 5 min, and then 0.5 mL of 0.5% (wtlvol) 3-methyl-2-benzothiazolone hydrochloride (prepared daily) is added. This mixture is heated in a boiling water bath for 3 min, cooled, and 0.5 mL of 0.5% (wtlvol) FeCl3 (100 mL contained 0.83 g FeCI3 ·6H2 0, stored at 4°C and discarded after 3 d) is added. After 30 min, the absorbance is read at 650 nm.

VESICULAR·ARBUSCULAR MYCORRHIZAL FUNGI

357

18-3 QUANTIFICATION OF VESICULAR-ARBUSCULAR MYCORRHIZAL EXTERNAL HYPHAE 18-3.1 Introdnction Even though the hyphae that grow into the soil matrix from the root are the functional organs for nutrient uptake and translocation, few researchers have obtained quantitative data on their growth and distribution. This is largely due to the technical difficulties in obtaining reliable datathere is no completely satisfactory method to quantify external hyphae of VAM fungi in soil. Three major problems have yet to be overcome: (i) there is no reliable method to distinguish VAM fungal hyphae from the myriad of other fungal hyphae in the soil, (ii) assessment of the viability and activity of hyphae is problematic, and (iii) meaningful quantification is time consuming. Nonetheless, clarification of the growth dynamics of ex· ternal hyphae is essential to further understanding of their function in soil. Below are brief descriptions of some useful methods for quantification of external hyphae. Further detailed methods have been reviewed (Sylvia, 1992).

18-3.2 Indirect Methods for Total Hyphae 18-3.2.1 Colonization of "Receiver" Plants Several researchers have used colonization of a "receiver" plant, sep· arated from an inoculated "donor" plant by root·free soil, to estimate the rate of hyphal growth in soil (Warner & Mosse, 1983; Schiiepp et aI., 1987; Miller et aI., 1989; Camel et aI., 1991). The essence of this method is that a volume of soil is maintained free of roots by using non biodegradable fabric with a pore size (50-100 fim) that excludes roots, but which allows free passage of fungal hyphae. A tripartite system is constructed where one compartment contains the inoculated donor plant, the middle compartment contains root-free soil, and the third compartment contains the reo ceiver plant. With such a system, the distance that hyphae grow through the soil can be determined; however, it will underestimate the rate of hyphal growth since colonization is not instantaneous upon contact of a hypha with a root.

18-3.2.2 Soil Aggregation Graham et al. (1982) estimated relative hyphal development by weighing plant roots with adhering soil. They reasoned that the weight of the root ball would be proportional to the amount of hyphae that bound the unit together. Using this method, however, it is not possible to compare hyphal development among different soils or plant species since many factors affect the quantity of soil in the root ball.

SYLVIA

358

18-3.2.2.1 Method 1. The soil mass is air dried and then roots are shaken vigorously to remove air-dry soil that is not firmly attached to the root system. 2. The soil that remains attached is washed from the root system into a beaker of water and after it settles, the water is decanted and the soil residue dried and weighed. 3. The roots are blotted dry and their masses determined. The amount of soil adhering to roots is expressed as the amount of dry soil attached in milligrams per gram of root. 18-3.2.3 Chitin Determination Chitin determinations have also been used to estimate hyphal biomass in soil; however, the same limitations discussed for this method under root colonization apply here. For this method, dried soil samples are mixed with conc. KOH and autoclaved for 1 h to degrade chitin to chitosan. SubsampIes of the soil-KOH suspension are transferred to centrifuge tubes and assayed for chitin as described in section 18-2.3.1. By subtracting the chitin content of control soil without VAM fungi from soil containing VAM fungi, the biomass of VAM fungi can be estimated.

18-3.3 Direct Methods for Total Hyphae 18-3.3.1 Filtration-Gridline Method Attempts have been made to estimate hyphal development directly by extracting hyphae from soil and quantifying lengths by a modified gridlineintersect method (Abbott et aI., 1984; Miller et aI., 1987; Sylvia, 1988). The major problem with the filtration-gridline method is that hyphae of nonmycorrhizal fungi often are not distinguishable from those of VAM fungi. Hyphal diameter has been used to distinguish VAM fungi from other fungi in the soil. Abbott and Robson (1985) found that most hyphae in VAM fungal-inoculated pot cultures had diameters between 1 and 5 !AID. They concluded that it was not possible to distinguish hyphae of VAM fungi from nonmycorrhizal fungi by morphological or staining criteria. One approach has been to subtract the length of hyphae in control treatments from YAM-inoculated treatments; however, this assumes that no interactions occur among the hyphae in the soil.

18-3.3.1.1. Method 1. Soil cores of known volume are collected at random, thoroughly mixed, subsamples (e.g., 10 g) removed, suspended in water (e.g., 500 mL), and passed through a sieve with 250-llm diam. pores. 2. The filtrate is blended for 15 s and a portion (e.g., 25 mL) passed through a membrane with pores < 5 Ilm diam. The exact quantity of soil, water, and suspension should be determined empirically so

VESICULAR·ARBUSCULAR MYCORRHIZAL FUNGI

359

that a thin layer of soil is deposited on the filter membrane without excessive clogging. 3. The membrane is briefly flooded with a trypan blue solution (0.5 g of trypan blue, 500 mL of deionized water, 170 mL of lactic acid, and 330 mL of glycerin) and rinsed with deionized water. 4. The membrane can be cut to fit on a microscope slide and observed at 300 x through an eyepiece whipple disc that has a 10 by 10 lined grid. The total length of hyphae can be estimated by the gridlineintersect method (see section 18-2.2.1). Due to the spatial variability of hyphae on the membranes, it is advisable to quantify 20 to 60 fields per membrane. 18-3.3.2 Immunofluorescence Assay Serological techniques have the potential for specific detection of mycorrhizal hyphae in soil. Kough and Linderman (1986) used an immunofluorescence assay (IFA) to detect hyphae of VAM fungi in soil and provided detailed protocols for preparation of antigens and fluorescein labelling; however, cross reactivity has limited their usefulness. The development of hybridoma technology has opened the potential for produc· tion of large quantities of monoclonal antibodies specific for a single epitope (Halk & De Boer, 1985). Wright et al. (1987) were able to obtain highly specific monoclonal antibodies for spores and hyphae of VAM fungi. Procedures for the use of monoclonal antibodies with VAM fungi are presented in chapter 28 by Wright of this book. 18-3.4 Detection of Active Hyphae Stains that differentiate actively metabolizing cells from inactive cells have been used to estimate the "viability" of VAM hyphae. Sylvia (1988) used a solution containing iodonitrotetrazolium (INT) and NADH to locate active hyphae of two Glomus spp.; INT is reduced by the electrontransport system of living cells and results in formation of a red color. Schubert et al. (1987) used fluorescein diacetate (FDA) to identify functioning hyphae; FDA is hydrolyzed within living cells, releasing fluorescein which can be detected with UV illumination. 18-3.4.1 Details of the Iodonitrotetrazolium Method 1. A membrane with extracted hyphae (see section 18-3.3.1.1) is flooded with a solution containing INT and incubated for 8 h at 28°e. The INT solution consists of equal parts of INT (1 mg mL -1), NADH (3 mg mL -1), and 0.2 M Tris buffer (pH 7.4). 2. The membrane is counter-stained with trypan blue, rinsed with deionized water, and cut to fit on a microscope slide. 3. The proportions of active (hyphal contents stained reddish) and inactive hyphae are determined by the gridline-intersect method.

SYLVIA

360

18-4 RECOVERY OF VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGAL SPORES Hayman (1984) and Schenck and Perez (1990a) have reviewed several methods for extracting spores of VAM fungi from soil. Most of these techniques work best in sandy soils, and less well in clay or organic soils.

18-4.1 Wet Sieving and Decanting/Density Gradient Centrifugation 1. Place a 50 to 100 g sample into a 2-L container and add 1 L of water. Vigorously mix the suspension to free spores from soil. For clay soils, samples may be placed in a sodium hexametaphosphate solution instead of water (Porter et aI., 1987). 2. For fungal species that form spores in roots (e.g., Glomus intraradix and G. clarum), blend the soil-root sample for 1 min in 300 mL of water to free spores from roots. 3. Let the suspension settle for 15 to 30 s (times vary depending on soil texture) and decant the supernatant through standard sieves. Sieves should be selected so as to capture the spores of interest. Use a 425-!A.m sieve over a 45-!A.m sieve for unknown field samples. Examine the contents of the top sieve for sporocarps that may be up to 1 mm in diameter. For clay soils, it is advisable to repeat the decanting and sieving procedures on the settled soil. 4. Roots can be collected from the larger-mesh sieves for evaluation of internal colonization (see section 18-2.1.1). 5. Transfer sievings to 50-mL centrifuge tubes with a steady stream of water from a wash bottle and balance opposing tubes. 6. Centrifuge at 1200 to 1300 x g in a swinging-bucket rotor for 3 min, allowing the centrifuge to stop without braking. Remove the supernatant carefully to avoid disturbing the pellet. Also remove the organic debris that adheres to the side of the tube. 7. Suspend soil particles in chilled, 1.17 M sucrose, mix contents with spatula and centrifuge immediately at 1200 to 1300 x g for 1.5 min, applying the brake to stop the centrifuge. 8. Pour the supernatant through a small-mesh sieve, rinse spores held on the sieve carefully with tap water and wash spores into a plastic Petri dish scribed with parallel lines spaced 0.5-cm apart. Spores may be counted by scanning the dish under a dissecting microscope.

18-5 IDENTIFICATION OF VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI The identification of VAM fungi can be a frustrating process. Unless taxonomy is the major objective, it is recommended that the spores be keyed initially only to genus. Isolates of special interest can be classified to

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

361

species at a later date. A description of the taxonomy of the six genera known to form VAM is present in Fig. 18-1. Careful notes should be taken on the size, color, surface characteristics, and wall morphology of the spore types recovered. Initiate pot cultures with each spore type (see section 18-7.1.5.1), keeping detailed records on the origin and subsequent pot culture history of the isolate. It is imperative that each isolate be given a unique code and that this code be used in all reports about the isolate. Currently, the taxonomy of VAM fungi is based on a limited number of morphological characteristics of the spore. Detailed steps for identification of spores have been presented elsewhere (Morton, 1988; Schenck & Perez, 1990b). Especially useful features of the manual prepared by Schenck and Perez (1994) are a worksheet for recording spore characteristics useful in taxonomy, a key to genera, and a species guide or quick key to rapidly reduce the number of possible species. A portion of the worksheet is presented in Fig. 18-2. With experience, some fungi can be distinguished on the basis of their morphology within roots. Abbott (1982) was able to differential 10 species of fungi by their characteristic morphology in roots; however, the differences among genera were always greater than those among species within a genera. Although this method requires careful initial observations from pot cultures, it offers promise for distinguishing fungi in plant roots. Maintenance of good germplasm should be an essential part of any YAM-research program. The International Culture Collection of VAM Fungi (INVAM) maintains a collection of VAM isolates. Samples of VAM fungi may be submitted to INVAM for verification of classification and possible inclusion in their collection. For additional information contact Dr. J. Morton, Division of Plant and Soil Sciences, West Virginia University, Morgantown, WV 26506-6057, USA. Biochemical and molecular methods promise to make VAM taxonomy a more precise science. Protein profiles (Rosendahl et al., 1989). serology (Wright et al., 1987), gene amplification (Simon et al., 1992), and fatty-acid analysis (J.H. Graham, 1992, personal communication) all hold great potential, but none have been sufficiently developed to be included in this chapter.

18-6 ASSESSMENT OF GROWTH RESPONSE AND SELECTION OF EFFECTIVE ISOLATES 18-6.1 Phosphorus-Response Curves and Mycorrhizal Dependency Much of the interest in mycorrhiza has focused upon their ability to enhance P uptake and subsequently the growth of plants in soils of low to moderate P availability. The quantitative assessment of plant response to VAM fungi is affected by all the factors involved in the symbiosis. including the genotypes of both the plant and fungus, and the soil environment.

362

SYLVIA

Order Glomales Soil-borne fungi characterized by transient dichotomously-branched arbuscules in cortical cells of plant roots after establishing an obligate mutualistic symbiosis with many plant species. Spores appear to be obligately asexual, forming within or outside roots. Most species diversity is manifested in spore size, color, and microscopic features of subce"ular spore walls distinct in structural properties and histochemical properties (usua"y in Melzer's reagent). Suborder Glomlneae Arbuscular fungi forming intraradical vesicles in mycorrhizal roots; "chlamydospores" borne terminally, intercalarily, and laterally from one or more subtending hyphae. Members are referred to as "vesicular-arbuscular mycorrhizal (VAM) fungi." Family Glomaceae "Chlamydospores" (i) are produced singly, in aggregates, in an unorganized hyphal matrix, or in a highly ordered hyphal matrix, with the structural wall continuous with a wall of the subtending hypha; (ii) show morphological diversification mostly in the number and types of walls formed outside the structural wall (e.g., evanescent, expanding, mucliagenous, and unit walls); (iii) form flexible inner walls so far limited to those which are membranous, usually one or two in number, and which rarely stain positive in Melzer's reagent; (iv) seal off contents from that of the subtending hypha by different mechanisms, such as an amorphous plug, a septum, an inner membranous wall, or thickening of the structural wall; and (v) germinate usually by emergence of the germ tube through the subtending hypha. Genus Glomus Have a" familiar characters except that spores are not formed in a highly organized matrix originating from a columnar base. Family Acaulosporaceae "Chlamydospores" (i) are born latera"y from or within the neck of a sporiferous saccule that is formed terminally on a fertile hypha; usually singly but occasionally in aggregates; (ii) have an outer wall which is continuous with the subtending hyphal wall and which sloughs with age; (iii) are sessile following extraction from soil because of sloughing of attached hyphae, (iv) have a smooth to highly ornamented structural wall that does not originate from the wall of the subtending hypha; (v) form at least one flexible inner wall, but usually two or more; (vi) show most diversification in number and types of inner walls (e.g. the semi-rigid unit wall, beaded or smooth membranous wall, and amorphous wall), with the innermost wall types often producing a dextrinoid to dark red-purple reaction in Melzer's reagent; (vii) seal off contents from that of the subtending saccule neck by a plug indistinguishable from the structural wall; and (viii) germinate between a semi-rigid unit wall and the innermost pair of inner flexible walls when such walls are synthesized. Genus Acaulospora Spores borne latera"y on the neck of the sporiferous saccule in a continuous transition series from those with a glomus-like hyphal attachment to those borne on a pedicel and finally to those borne on a short collar. Genus Entrophospora Spores are formed within the neck of the sporiferous saccule. Spore ontogenesis and spore wall diversity mirror that in Acaulospora. Suborder Gigasporineae Arbuscular fungi which also form extraradical auxilIary cells Singly or in clusters; globose to subglobose "azygospores" often exceeding 200 JLm in diameter, forming on a bulbous sporogenous cell. Members of the group do not form vesicles and hence are "arbuscular mycorrhizal fungi." Fig. 18-1. Continued on next page.

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

363

Genus Gigaspora Spores do not differentiate any flexible innerwalls; germ tubes arise from a warty germinal wall which rarely separates from the laminated wall; auxiliary cells usually echinulate.

Genus Scutellospora Spores (i) always differentiate two or more flexible inner walls which often form in adherent pairs, (ii) show most diverSification in the number and types of inner walls (e.g., membranous walls of different thicknesses, coriaceous wall, amorphous wall), with the innermost flexible wall often produCing adextrinoid to dark red-purple reaction in Melzer's reagent, (iii) germinate via germ tubes arising from a persistent germination shield of variable shape and margin which always forms on the innermost pair of flexible inner walls. Auxiliary cells smooth to knobby. Fig. 18-1. Description of the major taxa forming vesicular-arbuscular and arbuscular mycorrhizal fungi. Adapted from Morton (1988). Used with permission.

Since growth response is directly related to the P content of the plant, P-response curves should be constructed for each plant-soil combination (Abbott & Robson, 1984). Mycorrhizal and nonmycorrhizal plants are grown over a range of applied-P levels and shoot biomass is plotted against P level. The benefits of constructing these curves are: (i) ensuring that evaluations of mycorrhizal responsiveness are at soil-P levels that are limiting the growth of nonmycorrhizal plants, (ii) estimates of the amount of nutrient required for the same yield of mycorrhizal and nonmycorrhizal plants can be made, and (iii) claims for non-P effects of mycorrhizae on plant growth can be tested on plants with the same percentage of maximum growth and nutrient content. The mycorrhizal dependency or responsiveness of a plant is usually assessed in terms of the ratio of the shoot dry mass of mycorrhizal (M) vs. nonmycorrhizal (NM) plants in a P-deficient soil as follows (Plenchette et aI., 1983): 100 x «M - NM) INM) While this method for determining mycorrhizal dependency is useful, it can give misleading information since the method does not take into account differences among plant species in their responsiveness to P. When only one P level is used in the assessment of mycorrhizal dependency, there is the possibility that one plant species will respond to VAM fungi and the other will not. However, at a different P level the second plant may be quite responsive. Controls for YAM-plant growth experiments are critical since other soil microorganisms may affect plant growth. Abbott and Robson (1984) discussed various strategies for obtaining suitable controls. Ames et al. (1987) reported that similar functional groups of microorganisms could be established if a YAM-inoculum wash is applied to both mycorrhizal and nonmycorrhizal treatments. This can be accomplished by making a

364

SYLVIA

I. Many of these observations can be made with a dissecting microscope but measurements should be made with a compound microscope. A. Spore color: In water _ _ . In mountant _ _ . Mountant used: _ _. B. Spore diameter: For globose spores: Range _ ~m Mean -/Lm. For irregularly-shaped spores: Length; Range _ ~m Mean _ ~m. Width; Range _ ~m Mean _ ~m. C. Composite spore wall thickness (determined on intact spores if possible: _ _ ~m. D. Attachment present? Yes _ No. _. If no, go to E. Sporiferous saccule present? Yes _ No _. E. Spore contents: Globular _ ; Reticulate _ ; Granular _ ; Other: _ . F. Spore with mantle or other surface hyphae? Yes _ No _. If no, go to G. Width of hyphae _ ~m; Color of hyphae _ _ ; Hyphae sinuous? Yes _ No_ G. Spores formed within the root? Yes _ No _. H. Auxiliary cells present? Yes _ No _. If no, go to I. If yes, Knobby _; Digitate _; Coralloid _; Echinulate _; Spiny _; Pigmented _ if yes, color: _ _ . I. Sporocarp present? Yes _ No _. If no, go to J. Sporocarp diameter _ _; Peridium present? Yes _ No _ - if yes, indicate color _ _ . J. Determine the genus of your specimen. (see Fig. 18-1); Genus _ . Go to 1,2, or 3, then II. 1) For Gigaspora or Scutellispora: a. Bulbous sporogenous cell dimensions: Width _ ~m Length _ ~m. b. Subtending hyphae septate? Yes _ No _ c. Surface ornamentation on spore present? Yes _ No _. Description _ _ _ _ _ __ d. Germination shield present? Yes _ No _. 2) For Acaulospora or Entrophospora: a. Sporiferous saccule present? Yes _ No _. If no, go to II. b. Sporiferous saccule collapsed? Yes _ No _. If yes, go to II. c. Sporiferous saccule dimensions (diameter or length I width) __ ~m. d. Description of sporiferous saccule contents (color; content appearance, e.g., granular; reticulate, globular; texture, e.g., smooth, rough, flaky): _ _ _ _ _ _ _ _ _ _ _ _ _ _ _ __ e. Hyphal length between spore and sporiferous saccule _ ~m. f. Hyphal diam. at spore attachment _ ~m. g. Pore diameter on spore at pOint of attachment _ ~m. h. Hyphal attachment scar present? Yes _ No _. If yes, indicate the number present _ . 3) For Glomus or Sclerocystis: a. Pore occluded? Yes _ No _. b. Pore diameter _ ~m. c. Presence of a septum at the pore Yes _ No _. Protruding septum? Yes _ No _. d. Hyphal width adjacent to spore wall _ ~m. e. Number of attach· ments per spore _ _. f. Outer wall of hypha contiguous with outer wall of spore? Yes _ No

g. Type of attachment: (Check all that apply) straight _; recurved _; funnel-shaped _; branched _; septate _; constricted _; swollen _; other: ____ , II. Make these observations on broken spores with a compound microscope. A. Number of wall groups _ . B. Width of each wall group: A = _ B = _ C=_D=_E=_. C. Number of walls within each group: A = _ B = _ C = _ D = _ E = _. Fig. 18-2. Continued on next page.

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

365

D. Type of wall(s) within each group: Wall Group: A _; B _; C _; D _; E _. A = Amorphous; C = Coriaceous; E = Evanescent (ephemeral), G = Germinal; L = Laminate; M = Membranous; P = Hyphal peridium; U = Unit; X = Expanding. Wall reaction to Meltzer's reagent. Indicate positive (+) or negative (-) for each wall group. If positive, also indicate wall color and wall type ef· fected. If other reagents or stains are used, record the spore wall reaction as for Meltzer's reagent.

Fig. 18-2. Worksheet for collecting important characters for the identification of vesiculararbuscular mycorrhizal fungi. Adapted from Schenck and Perez (1994). Used with permission.

suspension of the inoculum in water, passing it through a fine sieve ( < 10 !-lm) to exclude VAM propagules, and then applying equal amounts of the sieved suspension to all treatments. 18-6_2 Screening for Effective Isolates Various isolates of VAM fungi differ in their ability to colonize host plants and promote plant growth; therefore, mycorrhizal dependency can vary widely with fungal genotype. Before embarking on an inoculation program, indigenous and exotic isolates of VAM fungi should be screened for their ability to promote the growth of the target host plant in the soil where the host will grow. For expediency, screening trials are often conducted in the greenhouse; however, reliable field data which show consistent growth improvements as a result of inoculation of plants subject to routine commercial propagative practices will do far more to convince growers of the usefulness of mycorrhizae than a plethora of carefully controlled trials where conditions have been optimized for mycorrhizal performance (Jeffries, 1987).

Often, the most effective isolates are those that colonize the plant most rapidly (Abbott & Robson, 1982; Abbott & Robson, 1984), but not necessarily those that have the greatest amount of colonization at harvest (Hung et al., 1990). For this reason, sequential harvests and assessments of colonization should be conducted. Alternatively, a nondestructive (leaf-P status) assessment of effectiveness has been proposed (Aziz & Habte, 1987). When conducting screening trials, care must be taken to ensure that inoculum density is not a limiting factor. Ideally, inoculum potential of all isolates should be equalized (Daniels et aI., 1981). However, the time required to conduct these assays is sufficient for significant changes in propagule density to occur (Daft et aI., 1987; O'Donnell et al., 1992). Furthermore, when inoculum densities vary widely among isolates, standardization of the inoculum is not practical and effective isolates do not always produce large quantities of inoculum in culture (Sylvia & Burks,

SYLVIA

366

1988). Nonetheless,MPN or similar assays should be conducted on inoculum used in screening trials since the results are useful for interpretation of the results of YAM-inoculation studies.

18-7 PRODUCTION AND USE OF VESICULAR-ARBUSCULAR MYCORRHIZAL INOCULA

Jarstfer and Sylvia (1992a) recently reviewed methods for the production and use of VAM inocula and this section is largely adapted from that work. 18-7.1 Soil-Based Pot Cultures

The culture of VAM fungi on plants in disinfested soil, using spores, roots or infested soil as inocula, has been the most frequently used technique for increasing propagule numbers (Menge, 1983; Menge, 1984; Schenck & Perez, 1990a). 18-7.1.1 Selection of Host Plant

Many host plants have been used under a variety of conditions (Thompson, 1986; Sreenivasa & Bagyaraj, 1988; Liyanage, 1989). Examples of plants that have been used successfully are alfalfa, maize, onion, sudan grass, and wheat. Generally, the host selected should become well colonized (> 50% of the root length), produce root mass quickly, and be able to tolerate the high-light conditions required for the fungus to reproduce rapidly. Hosts that can be propagated from seed are preferable to cuttings since they can be more easily disinfested. Most seeds may be disinfested with 10% household bleach (0.525% NaOel) for 5 to 15 min followed with five washes of water (Tuite, 1969). Washing with water may also remove fungicides and other agrichemicals which may adversely affect VAM fungi (Tommerup & Briggs, 1981). 18-7.1.2 Soil Disinfestation

All components of the culture system should be disinfested prior to initiation of a pot culture. The method of soil disinfestation is especially important-the objective is to kill existing VAM fungi, pathogenic organisms, and weed seeds while preserving a portion of the nonpathogenic microbial community. The use of aerated steam or a commercial soil pasteurizer will eliminate VAM fungi from soils (Menge, 1983; Sylvia & Schenck, 1984). We commonly pasteurize soil by heating it to 85°C for two, 8-h periods with 48 h between treatments. Fumigation with a methylbromide/chloropicrin mixture or other soil-applied biocides is effective, but requires several days for the chemical to diffuse out of the soil. Ionizing

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

367

radiation (0.8-1.0 Mrad) has also been used (Jakobsen & Anderson, 1982; Jensen, 1983; Thompson, 1990). Autoc1aving should be avoided because it can result in the release of toxic inorganic and organic compounds (Wolf et al., 1989). 18-7.1.3 Light, Moisture, and Temperature Good light quality and high irradiance are necessary for maximum inoculum production (Ferguson & Menge, 1982a; Furlan & Fortin, 1977). Where natural light conditions are poor (PPFD < 500 !lmol m- 2 S-1), high-intensity lamps should be used. Soil moisture affects VAM sporulation, with nonsaturated and nonstressed water conditions providing maximum sporulation (Nelsen & Safir, 1982). Excessive moisture may encourage problems with hyperparasites in the culture (Paulitz & Menge, 1986; Daniels & Menge, 1980). The best strategy is to apply water regularly to well-drained soil. Temperature is also important for pot cultures. Sporulation is positively correlated with temperature from 15 to near 30°C for many VAM fungi (Schenck & Schroder, 1974; Schenck & Smith, 1982); however, at higher temperatures sporulation may decrease. For maximum sporulation, the temperature optimum for each isolate should be determined. 18-7.1.4 Fertilizers, Pesticides, and Pot Size Responses to P and N fertilization are also strain dependent (Sylvia & Schenck, 1983; Johnson et al., 1984; Thompson, 1986; Liyanage, 1989; Douds & Schenck, 1990a) and are affected by the relative amounts of N and P supplied (Sylvia & Neal, 1990). The usual approach is to supply the plants with adequate nutrition, except for P, the concentration of which should be limiting for growth of nonmycorrhizal plants. For routine maintenance of pot cultures a P-free fertilizer such as Peter's (Peter Fertilizer Products, Allentown, PA) 25-0-25 can be applied; however, to determine the optimal level of P fertility, P-response curves should be developed (Abbott & Robson, 1984). The optimal level ofP has been shown to differ for some species of VAM fungi (Thomson et al., 1986). Pesticides can affect VAM colonization and sporulation. Fungicides should be carefully selected for use in pot cultures (Nemec, 1980; Menge, 1982; Jabaji-Hare & Kendrick, 1987; Dodd & Jeffries, 1989). Ants and other crawling insects also cause cross contamination of pot cultures. Insecticides and nematicides may be used effectively in VAM pot cultures (Sreenivasa & Bagyaraj, 1989). Pot size should match the potential volume of the root system within practical space constraints. Ferguson and Menge (1982b) assessed pot volumes ranging from 750 to 15 000 cm and reported that larger containers resulted in higher spore concentrations. However, when using large volumes, any contamination will result in greater loss. In the greenhouse, pot

SYLVIA

368

cultures should be isolated from contaminated soil, splashing water, and crawling insects. In addition, specific isolates of VAM fungi should be kept well separated from each other.

18-7.1.5 General Procedures To initiate pot cultures, place a layer of inoculum 1 to 2 cm below the seed or cutting. Inoculum may consist of spores, colonized roots or infested soil. Infested soil is often used to initially obtain isolates from the field; however, these "mixed-species" cultures should rapidly progress to "single-species" cultures initiated from 20 to 100 healthy, uniform spores. For critical taxonomic studies, "single-spore" cultures should be produced (Fang et aI., 1983; Schenck & Perez, 1990a).

18-7.1.5.1 Method for Single-Spore Culture 1. Plastic tubes (such as available from Stuewe and Sons, Inc., Corvallis, OR) are plugged with polyester fiber batting (available at fabric stores). 2. Place moist, pasteurized soil-similar to that from which the spores were originally isolated-into the tubes to within 3 cm of the top. 3. Place a moist filter paper disk on the soil surface and transfer a single spore to it. Choose clean, bright, and nonparasitized spores. Spores can be transferred from water in petri dishes with micropipettes. 4. Add 1 to 2 cm of additional soil over the spore and plant seed of a suitable host on the surface. Place tubes in a well-lighted growth chamber or greenhouse and sample for colonization and sporulation after 12 to 16 wk. 5. Another method is to place a single spore directly on the surface of seedling root (approximately 2 cm below the crown) prior to transplanting it into a growth tube. The placement of the spore should be confirmed under a dissecting microscope before the growth medium is carefully poured around the root system. 6. The probability of success with one spore is relatively low so it is important that several tubes are established for each isolate. 18-7.2 Soil-less Media Culturing VAM fungi in soil-less media avoids the detrimental organisms in nonsterile soil and allows control over many of the physical and chemical characteristics of the growth medium. Soil-less media are more uniform in composition, weigh less, and provide aeration better than do soil media. Growth media that have been used include bark, peat, perlite, and vermiculite (Biermann & Linderman, 1983); sand (Ojala & Jarrell, 1980), calcined montmorillonite clay (Plenchette et aI., 1982), and expanded clay aggregates (Dehne & Backhaus, 1986). Moist soil-less media do not buffer P concentration so care must be taken to avoid high solution levels of P in the root zone. Frequent addition

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

369

of dilute, soluble nutrient solutions (Waterer & Coltman, 1988a; Liyanage, 1989; Douds & Schenck, 1990a), mixing of time-release fertilizer (Waterer & Coltman, 1988b) or the use of less-available forms of P (Thompson, 1986) should provide excellent cultures of VAM fungi in soil-less media. 18-7.3 Nutrient Flow and Aeroponic Systems 18-7.3.1 Introduction

At least seven species of VAM fungi have been grown in various nutrient-solution systems on hosts representing at least 21 genera (Jarstfer & Sylvia, 1992a). The primary benefit of these systems is that colonized roots and spores are produced free of any substrate, permitting more efficient production and distribution of inocula. Usually plants are inoculated with VAM fungi and grown in sand or vermiculite before they are transferred into a culture system. The plants are grown for a period of 4 to 6 wk under conditions conducive for colonization, after which they are washed and nondestructively checked for colonization (Ames et aI., 1982); however, it is also possible to inoculate plants directly in the culture system (Hung et aI., 1991). The P concentrations which have been reported to support VAM growth in solution cultures range from < 1 to 24 !tmol. A system that applies a fine nutrient mist to roots of intact plants (aeroponic culture) produces excellent inoculum (Sylvia & Hubbell, 1986; Hung & Sylvia, 1988). Greater concentrations of spores have been produced in aeroponic cultures when compared to soil-based pot cultures of the same age. Because the colonized-root inoculum produced in this system is free of any substrate, it can be sheared resulting in high propagule numbers per gram of root (Sylvia & Jarstfer, 1992). 18-7.3.2 Method for Producing Sheared-Root Inocula from Aeroponic Cultures

This method is modified from Jarstfer and Sylvia (1992b). 1. To inoculate the plants, distribute a spore suspension over 2 L of clean vermiculite in a disinfested pot and cover with more vermiculite. Either plant disinfested seed of the culture plant or push unrooted stem cuttings (also disinfected) into the vermiculite so the bud is level with the vermiculite surface. Sweet potato [Ipomoea halalas (L. ) Lam] has worked well for this purpose. 2. After the culture plants have grown for at least 4 wk, remove them from the vermiculite and check for VAM colonization. Displace the plants from their pots onto a disinfected surface and separate individual plants carefully to minimize root injury. Wash the roots in several changes of water to remove as much vermiculite as possible. 3. The roots of intact plants can be nondestructively examined with epifluorescence microscopy for intracellular granular fluores-

SYLVIA

370

4.

5.

6.

7.

cence,cence, indicating arbuscular formation (Ames et aI., 1982). If colonization is satisfactory, the plants are transferred into an aeroponic chamber by inserting roots through holes in the chamber cover. Only colonized plants should be placed in the aeroponic chamber. The aeroponic chamber should be of dimensions suitable to the desired quantity of inoculum or the space available. For sanitation reasons, an acrylic-lined chamber is desirable, but acrylic enamel paint over finished fiberglass will also work. The chamber should have a lid with 2.S-cm-diam. holes spaced 15 cm apart in a regular pattern to allow adequate distribution of the nutrient mist. Two systems of misting may be used to deliver the nutrient solution to the roots of the culture plants. The first uses an atomizing disk similar to those used in cold-water humidifiers (Zobel et aI., 1976). This system is limited to relatively small chambers with capacities from 60 to 80 L of nutrient solution. The second system uses a centrifugal pump to spray the nutrient solution at the upper portion of the root mass allowing the solution to flow down the roots. Multiple micro-irrigation nozzles may be used to cover the entire root system regardless of the number of plants or depth of the chamber. The pump should run intermittently to deliver a spray for 7 s every min. The nutrients for aeroponic culture are dilute as the roots are in constant contact with them. Deionized or distilled water should be used to make the nutrient solutions to avoid problems with pH or toxic levels of plant micronutrients. For convenience, nutrient solutions are made by diluting stock solutions (Table 18-1). When initiating a culture or routinely changing the nutrient solution, first fill the chamber with the desired amount of water then

Table 18-1. Nutrient solution for aeroponic culture of VAM fungi, based on a modified Hoagland's solution (Hoagland & Arnon, 1950). Stock solution 0.01 1.00 1.00 1.00 0.10 0.10

M KH2 P0 4 M KN0 3 M Ca(N0 3 )z4H2 0 M MgS0 4 7H2 0t MNaCI M NaFeEDTA

Micronutrient stock 46.20 mM H 3B0 3 9.53 mM MnCl 2 ·4H2 0 0.765 mM ZnS0 4 ·7H2 0 0.32 mM CuS0 4 ·5H 2 0 0.066 mM Na2 Mo04 ·2H2 0

t Keep this solution in a separate container.

Quantity of stock solution in final nutrient solution mLL-l 0.3 1.5 1.5 0.3 0.3 0.45 1.0

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

371

add the mixture of concentrated nutrients measured from the stock solutions. Mix well and check the pH after at least 15 min. The pH should be adjusted to 6.5 for most culture plants. 8. On a biweekly schedule, the roots of the culture plants should be cut back to 2 cm above the highest nutrient solution level and the root debris removed from the chamber, the inside walls scrubbed to remove any algae, and the nutrient solution drained quickly from the chamber and replaced with new solution. 9. To produce sheared-root inoculum, well-colonized roots harvested from aeroponic cultures (grown for 10-12 wk) are mixed 1 to 10 (wt/vol) with water and sheared with a food processor for 40 s (e.g., "Little Pro," Cuisinart, Inc., Norwich, CN). The resulting root fragments, vesicles, and free spores are collected over a fine sieve (45 !tm). The sheared-root material may be mixed directly with growing medium or may be added to hydrogels such as Natrosol to make a ftowable inoculum (Hung et al., 1991). 10. Spores may also be collected from fresh roots or from roots stored air dry by washing over standard sieves. For clean preparations, spores may be separated from root fragments using density-gradient centrifugation (see section 18-4.1).

18-7.4 Storage of Inoculum Spores of VAM fungi are commonly stored at 4 °C in dried pot-culture soil (Ferguson & Woodhead, 1982); however, many VAM species do not survive this treatment. Cryopreservation of spores at -60 to -70°C appears to be much more reliable (Douds & Schenck, 1990b). Cultures of VAM fungi should be dried slowly with the host plants and frozen in situ. The viability of colonized-root inoculum from aeroponic culture declines with storage. Root inoculum, air-dried prior to storage at 4°C, retains a greater density of VAM fungal propagules than roots stored moist. However, once roots are dried they cannot be sheared. For shortterm storage of active roots, continue to maintain the aeroponic culture and remove plants as needed to make inoculum. When plants are harvested, excess moisture should be removed from the roots to prevent development of anaerobic conditions. Roots may be stored for short times in this moist state.

18-7.5 Application of Vesicular-Arbuscular Mycorrhizal Inocula 18-7.5.1 Introduction Any method by which viable propagules of VAM fungi can be delivered efficiently to the rhizosphere (or potential rhizosphere) should produce the desired colonization. The composition of an inoculum should mesh economically with the method of application. In addition, to reduce

SYLVIA

372

the risk of spreading pathogens, the host plant used for inoculum production should not be related to the crop plant to be inoculated (Menge, 1983). For sanitation reasons, logistics, and cost, soil-less cultures or inoculum from hydroponic or aeroponic systems should be used. 18-7.5.2 General Methods

The most common application method is to place the inoculum below the seed or seedling prior to planting (Jackson et aI., 1972; Hayman et aI., 1981; Hayman, 1987). Hall (1979, 1980) successfully used infested soil pellets to inoculate plants in the field. To produce pellets, moistened soilclay-sand inoculum is sandwiched between aluminum foil, cut into squares, dried, and seed is glued to the pellets with gum arabic (Hall & Kelson, 1981). Seed coating (Hattingh & Gerdemann, 1975) and somatic embryo encasement (Strullu et aI., 1989) with inoculum are also methods worthy of further investigation. Sheared-roots from aeroponic cultures can be suspended in hydrogel for application as a flow able inoculum (Sylvia & Jarstfer, 1992). 18-8 MONOXENIC CULTURES FOR BASIC RESEARCH

The growth of VAM fungi in pure culture in the absence of a host has not been achieved. However, it is possible to colonize roots of intact plants or root-organ cultures to achieve monoxenic cultures that are useful for basic research on the symbiosis (Hepper, 1984). More recently, Ri T-DNA

transformed roots have been used to obtain colonized root cultures (Mugnier & Mosse, 1987; Becard & Fortin, 1988). It should be noted, however, that colonization rates in these systems are slow and only limited amounts of colonized roots have been produced. Transformed-root culture offers the most efficient method to grow colonized roots as no plant growth regulators are required for sustained growth. Becard and Fortin (1988) provide procedures for initiating transformed root cultures. A critical step is to obtain aseptically germinated spores of the VAM fungus or, alternatively, colonized root pieces (Williams, 1990). Hepper (1984) reviewed procedures for disinfesting and germinating spores. The most effective methods use chlorine containing compound such as hypochlorite, a surfactant, and antibacterial agents. REFERENCES Abbott, L.K. 1982. Comparative anatomy of vesicular-arbuscular mycorrhizas formed on subterranean clover. Aust. J. Bot. 30:485-499. Abbott, L.K., and A.D. Robson. 1982. The role of vesicular-arbuscular mycorrhizal fungi in agriculture and the selection of fungi for inoculation. Aust. J. Agric. Res. 33:389-408. Abbott, L.K., and A.D. Robson. 1984. The effect ofmycorrhizae on plant growth. p.113-130. In C.LI. Powell and D.J. Bagyaraj (ed.) VA mycorrhiza. CRC Press, Boca Raton, FL.

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

373

Abbott, L.K., and A.D. Robson. 1985. Formation of external hyphae in soil by four species of vesicular-arbuscular mycorrhizal fungi. New Phytol. 99:245-255. Abbott, L.K., and A.D. Robson. 1991. Factors influencing the occurrence of vesiculararbuscular mycorrhizas. Agric. Ecosystem Environ. 35:121-150. Abbott, L.K., A.D. Robson, and G. De Boer. 1984. The effect of phosphorus on the formation of hyphae in soil by the vesicular-arbuscular mycorrhizal fungus. Glomus Jasciculatum. New Phytol. 97:437-446. Adelman, M.J., and J .B. Morton. 1986. Infectivity of vesicular-arbuscular mycorrhizal fungi: Influence of host-soil diluent combinations on MPN estimates and percentage colonization. Soil. BioI. Biochem. 18:77-83. Alexander, M. 1982. Most probable number method for microbial populations. p. 815-810. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA. Madison, WI. Ames, R.N., E.R. Ingham, and c.P.P. Reid. 1982. Ultraviolet-induced autofluorescence of arbuscular mycorrhizal root infections: an alternative to clearing and staining methods for assessing infections. Can. J. Microbiol. 28:351-355. Ames, R.N., K.L. Mihara, and G.J. Bethlenfalvay. 1987. The establishment of microorganisms in vesicular-arbuscular mycorrhizal and control systems. BioI. Fert. Soils 3:217223. An, Z.-Q., J.W. Hendrix, D.E. Hershman, and G.T. Henson. 1990. Evaluation of the "Most probable number" (MPN) and wet-sieving methods for determining soil-borne populations of endogonaceous mycorrhizal fungi. Mycologia 82:576-581. Aziz, T., and M. Habte. 1987. Determining vesicular mycorrhizal effectiveness by monitoring P status of leaf disks. Can. J. Microbiol. 33:1097-1101. Becard, G., and J .A. Fortin. 1988. Early events of vesicular-arbuscular mycorrhiza formation on Ri T-DNA transformed roots. New Phytol. 108:211-218. Bethlenfalvay, G.J., R.S. Pacovsky, and M.S. Brown. 1981. Measurement of mycorrhizal infection in soybeans. Soil Sci. Soc. Am. J. 45:871-874. Bevege, D.l. 1968. A rapid technique for clearing tannins and staining intact roots for detection of mycorrhlzas caused by Endogone spp., and some records of infection in Australian plants. Trans. Br. Mycol. Soc. 51:808-810. Biermann, B.J., and R.G. Linderman. 1983. Effect of container plant growth medium and fertilizer phosphorus on establishment and host growth response to vesicular-arbuscular mycorrhizae. J. Am. Soc. Hortic. Sci. 108:962-971. Brundrett, M.C., Y. Piche, and R.L. Peterson. 1984. A new method for observing the morphology of vesicular-arbuscular mycorrhizae. Can. J. Bot. 62:2128-2134. Camel. S.B., M.G. Reyes-Solis, R. Ferrera-Cerrato, R.L. Franson, M.S. Brown, and G.J. Bethlenfalvay. 1991. Growth of vesicular-arbuscular mycorrhizal mycelium through bulk soil. Soil Sci. Soc. Am. J. 55:389-393. Cochran, W.G. 1950. Estimation of bacterial densities by means of the "most probable number." Biometrics 6:105-116. Daft, M.J., D. Spencer, and G .E. Thomas. 1987. Infectivity of vesicular-arbuscular mycorrhizal inocula after storage under various environmental conditions. Trans. Br. Mycol. Soc. 88:21-27. Daniels. B.A., P.M. McCool, and J.A. Menge. 1981. Comparative inoculum potential of spores of six vesicular-arbuscular mycorrhizal fungi. New Phytol. 89:385-392. Daniels, B.A., and J.A. Menge. 1980. Hyperparasitism of vesicular-arbuscular mycorrhizal fungi. Phytopathology 70:584-588. de Man. J.C. 1975. The probability of most probable numbers. Eur. J. Appl. Microbiol. 1:67-78. Dehne, H.W., and G .F. Backhaus. 1986. The use of vesicular-arbuscular mycorrhizal fungi in plant production. I. Inoculum production. Z. Pflanzenkr. Pflanzenschutz 93:415-424. Dodd, J.C., and P. Jeffries. 1989. Effect of fungicides on three vesicular-arbuscular mycorrhizal fungi associated with winter wheat (Triticum aestivum L.) BioI. Fert. Soils 7: 120128. Douds, D.D., and N.C. Schenck. 1990a. Increased sporulation of vesicular-arbuscular mycorrhizal fungi by manipulation of nutrient regimes. Appl. Environ. Microbiol. 56:413418. Douds, D.O., and N.C. Schenck. 1990b. Cryopreservation of spores of vesicular-arbuscular mycorrhizal fungi. New Phytol. 115:667-674.

374

SYLVIA

Fang, Y.c., A.c. McGraw, H. Modjo, and J.W. Hendrix. 1983. A procedure for isolation of single-spore cultures of certain endomycorrhizal fungi. New Phytol. 95:107-114. Ferguson, J.J., and J.A. Menge. 1982a. The influence of light intensity and artificially extended photoperiod upon infection and sporulation of Glomus Jasciculatum on sudan grass and on root exudation of sudan grass. New Phytol. 92:183-19l. Ferguson, J.J., and J.A. Menge. 1982b. Factors that affect production of endomycorrhizal inoculum. Proc. Fla. State Hortic. Soc. 95:39-39. Ferguson, J.J., and S.H. Woodhead. 1982. Increase and maintenance ofvesicular-arbuscular mycorrhizal fungi. p. 47-54. In N.C. Schenck (ed.) Methods and principles of mycorrhizal research. Am. Phytopathol. Soc., St. Paul. Fisher, R.A., and F. Yates. 1963. Statistical tables for biological, agricultural and medical research. Oliver and Boyd, Edinburgh. Franson, R.L., and G.J. Bethlenfalvay. 1989. Infection unit method of vesicular-arbuscular mycorrhizal propagule determination. Soil Sci. Soc. Am. J 53:754-756. Furlan, v., and J.A. Fortin. 1977. Effects of light intensity on the formation of vesiculararbuscular endomycorrhizas on Allium cepa by Gigaspora calospora. New Phytol. 70:335-340. Giovannetti, M., and B. Mosse. 1980. An evaluation of techniques for measuring vesicular arbuscular mycorrhizal infections in roots. New Phytol. 84:489-500. Graham, J.H., D.M. Eissenstat, and E.L. Drouillard. 1992. On the relationship between a rlants mycorrhizal ~ependency and rate of vesicular-arbuscular mycorrhizal colonization. Funct. Ecol. 5.773-779. Graham, J .H., and D. Fardelmann. 1986. Inoculation of citrus with root fragments containing chlamydospores of the mycorrhizal fungus Glomus intraradices. Can. J. Bot. 64:17391744. Graham, J.H., RG. Linderman, and J.A. Menge. 1982. Development of external hyphae by different isolates of mycorrhizal Glomus spp. in relation to root colonization and growth of Troyer citrange. New Phytol. 91:183-189. Halk, E.L., and S.H. De Boer. 1985. Monoclonal antibodies in plant-disease research. Annu. Rev. Phytopathol. 23:321-350. Hall, I.R. 1979. Soil pellets to introduce vesicular-arbuscular mycorrhizal fungi into soil. Soil BioI. Biochem. 11:85-86. Hall, I.R 1980. Growth of Lotus pedunculatus Cav. in an eroded soil containing soil pellets infested with endomycorrhizal fungi. N. Z. J. Agric. Res. 23:103-105. Hall, I.R., and A. Kelson. 1981. An improved technique for the production of endomycorrhizal infested soil pellets. N. Z. J. Agric. Res. 24:221-222. Halvorson, H.O., and N.R. Ziegler. 1933. Applications of statistics to problems in bacteriology. I. A means of determining bacterial populations by the dilution method. J. Bacteriol. 25: 101-12l. Hattingh, M.J., and J.W. Gerdemann. 1975. Inoculation of Brazilian sour orange seed with an endomycorrhizal fungus. Phytopathology 65:1013-1016. Hayman, D.S. 1984. Methods for evaluating and manipulating vesicular-arbuscular mycorrhiza. p. 95-117. In J.M. Lynch and J.M. Grainger (ed.) Microbiological methods for environmental microbiology. Academic Press, London. Hayman, D.S. 1987. VA mycorrhizas in field crop systems. p. 171-192. In G. Safir (ed.) Ecophysiology of VA mycorrhizal plants. CRC Press, Boca Raton, FL. Hayman, D.S., E.J. Morris, and RJ. Page. 1981. Methods for inoculating field crops with mycorrhizal fungi. Ann. Appl. BioI. 99:247-253. Hepper, C.M. 1977. A colorimetric method for estimating vesicular-arbuscular mycorrhizal infection in roots. Soil BioI. Biochem. 9:15-18. Hepper, C.M. 1984. Isolation and culture of VA mycorrhizal (VAM) fungi. p. 95-112. In C.L. Powell and D.J. Bagyaraj (ed.) VA mycorrhiza. CRC Press, Boca Raton, FL. Hoagland, D.R, and D.1. Arnon. 1950. The water-culture method for growing plants without soil. California Agric. Exp. Stn. Cir. 347 (revised edition), Berkeley. Hung, L.L., D.M. O'Keefe, and D.M. Sylvia. 1991. Use of a hydrogel as a sticking agent and carrier of vesicular-arbuscular mycorrhizal fungi. Mycol. Res. 95:427-429. Hung, L.L., and D.M. Sylvia. 1988. Production of vesicular-arbuscular mycorrhizal fungus inoculum in aeroponic culture. App!. Environ. Microbio!. 54:353-357. Hung, L.L., D.M. Sylvia, and D.M. O'Keefe. 1990. Isolate selection and phosphorus interaction of vesicular-arbuscular mycorrhizal fungi in biomass crops. Soil Sci. Soc. Am. J. 54:762-768.

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

375

Jabaji-Hare, S.H., and W.B. Kendrick. 1987. Response of an endomycorrhizal fungus in Allium porrum L. to different concentration of systemic fungicides, metalaxyl and fosetyl-AI. Soil BioI. Biochem. 19:95-99. Jackson, N.E., R.E. Franklin, and R.H. Miller. 1972. Effects of vesicular-arbuscular mycorrhizae on growth and phosphorus content of three agronomic crops. Soil Sci. Soc. Am. Proc. 36:64-67. Jakobsen, I., and A.J. Anderson. 1982. Vesicular-arbuscular mycorrhiza and growth in barley: effects of irradiation and heating of soil. Soil BioI. Biochem. 14:171-178. Jarstfer, A.G., and R.M. Miller. 1985. Progress in the development of a chitin assay technique for measuring extraradical soilborne mycelium of V-A mycorrhizal fungi. p. 410. In R. Molina (ed.) Proc. 6th North American Conf. Mycorrhizae. Forest Res. Lab, Oregon State Univ., Corvallis. Jarstfer, AG., and D.M. Sylvia. 1992a. Inoculum production and inoculation strategies for vesicular-arbuscular mycorrhizal fungi. p. 349-377. In B. Metting (ed.) Soil microbial technologies: Applications in agriculture, forestry and environmental management. Marcel Dekker, New York. Jarstfer, A.G., and D.M. Sylvia. 1992b. The production and use of aeroponically grown inocula of VAM fungi in the nursery. Florida Sea Grant Ext. Bull. 22, Gainesville. Jeffries, P. 1987. Use of mycorrhizae in agriculture. CRC Crit. Rev. Biotechnol. 5:319-357. Jeffries, P., and J.C. Dodd. 1991. The use of mycorrhizal inoculants in forestry and agriculture. p. 155-185. In D.K. Arora et al. (ed.) Handbook of applied mycology. Vol. I: Soil and plants. Marcel Dekker, New York. Jensen, A. 1983. The effect of indigenous vesicular-arbuscular mycorrhizal fungi on nutrient uptake and growth of barley in two Danish soils. Plant Soil 70:155-163. Johnson, B.N., and W.B. McGill. 1990. Comparison of ergosterol and chitin as quantitative estimates of mycorrhizal infection and Pinus contorta seedling response to inoculation. Can. J. For. Res. 20:1125-1131. Johnson, C.R., W.M. Jarrell, and J.A Menge. 1984. Influence of ammonium-nitrate ratio and solution pH on mycorrhizal infection, growth and nutrient composition of Chrysanthemum morifolium var circus. Plant Soil 77:151-157. Kendrick, B., and S. Berch. 1985. Mycorrhizae: Applications in agricultuie and forestry. p. 109-152. In c.w. Robinson (ed.) Comprehensive biotechnology. Vol. 4. Pergamon Press, Oxford. Koide, R.T., and H.A Mooney. 1987. Spatial variation in inoculum potential of vesiculararbuscular mycorrhizal fungi caused by formation of gopher mounds. New Phytol. 107: 173-182. Kormanik, P.P., and A-C. McGraw. 1982. Quantification of vesicular-arbuscular mycorrhizae in plant roots. p. 37-45. In N.C. Schenck (ed.) Methods and principles of mycorrhizal research. Am. Phytopathol. Soc., St. Paul. Koske. R.E., and J.N. Gemma. 1989. A modified procedure for staining roots to detect VA mycorrhizas. Mycol. Res. 92:486-505. Kough. J.L.. and R.G. Linderman. 1986. Monitoring extra-matrical hyphae of a vesiculararbuscular mycorrhizal fungus with an immunofluorescence assay and the soil aggregation technique. Soil BioI. Biochem. 18:309-313. Liyanage. H.D. 1989. Effects of phosphorus nutrition and host species on root colonization and sporulation by vesicular-arbuscular (VA) mycorrhizal fungi in sand-vermiculite medium. M.S. thesis, Univ. of Florida. Gainesville. Marsh, B. 1971. Measurement of length in random arrangements of lines. J. Appl. Ecol. 8:265-267. Martin, F., C. Delaruelle, and J.L. Hilbert. 1990. An improved ergosterol assay to estimate fungal biomass in ectomycorrhizas. Mycol. Res. 94:1059-1064. McGonigle, T.P., M.H. Miller. D.G. Evans, G.S. Fairchild, and J.A. Swan. 1990. A new method which gives an objective measure of colonization of roots by vesicular-arbuscular mycorrhizal fungi. New Phytol. 115:495-501. Menge, J.A. 1982. Effect of soil fumigants and fungicides on vesicular-arbuscular fungi. Phytopathology 72:1125-1132. Menge, J.A. 1983. Utilization of vesicular-arbuscular mycorrhizal fungi in agriculture. Can. J. Bot. 61:1015-1024. Menge, J.A. 1984. Inoculum production. p. 187-203. In C.L. Powell and D.J. Bagyaraj (ed.) VA mycorrhiza. CRC Press, Boca Raton, FL. Miller, D.D., M. Bodmer, and H. Schuepp. 1989. Spread of endomycorrhizal colonization and effects on growth of apple seedlings. New Phytol. 111:51-59.

376

SYLVIA

Miller, R.M., A.G. Jarstfer, and J.K. Pillai. 1987. Biomass allocation in an Agropyron smithii-Glomus symbiosis. Am. J. Bot. 74:114-122. Moorman, T., and F.B. Reeves. 1979. The role of endomycorrhizae in revegetation practices in the semi-arid West. II. A bioassay to determine the effect of land disturbance on endomycorrhizal populations. Am. J. Bot. 66:14-18. Morton, J.B. 1985. Underestimation of most probable numbers of vesicular-arbuscular mycorrhizae endophytes because of non-staining mycorrhizae. Soil BioI. Biochem. 17:383384. Morton, J.B. 1988. Taxonomy of VA mycorrhizal fungi: Classification, nomenclature, and identification. Mycotaxon 32:267-324. Morton, J.B., and G.L. Benny. 1990. Revised classification of arbuscular mycorrhizal fungi (Zygomycetes): A new order, Glomales, two new suborders, Glomineae and Gigasporineae, and two new families, Acaulosporaceae and Gigasporaceae, with an emendation of Glomaceae. Mycotaxon 37:471-491. Mugnier, J., and B. Mosse. 1987. Vesicular-arbuscular mycorrhizal infection in transformed root-inducing T-DNA roots grown axenically. Phytopathology 77:1045-1050. Nelsen, C.E., and G.R. Safir. 1982. Increased drought tolerance of mycorrhizal onion plants caused by improved phosphorus nutrition. Plant a 154:407-413. Nemec, S. 1980. Effects of eleven fungicides on endomycorrhizal development in sour oranges. Can. J. Bot. 58:522-527. Newman, E.I. 1966. A method for estimating the total length of root in a sample. J. Appl. Ecol. 3:139-145. O'Donnell, J.J., D.M. Sylvia, W.D. Pitman, and J.E. Rechcigl. 1992. Inoculation of Vigna parkeri with mycorrhizal fungi in an acid Florida spodosol. Trop. Grassl. 26:120-129. O'Keefe, D.M., and D.M. Sylvia. 1991. Mechanisms of the vesicular-arbuscular mycorrhizal plant-growth response. p. 35-54. In D.K. Arora et al. (ed.) Handbook of applied mycology. Marcel Dekker, New York. Ojala, J.C., and W.M. Jarrell. 1980. Hydroponic sand culture systems for mycorrhizal research. Plant Soil 57:297-303. Paulitz, T.C., and J.A. Menge. 1986. The effect of a mycoparasite on the mycorrhizal fungus, Glomus deserticola. Phytopathology 76:351-354. Phillips, J.M., and D.S. Hayman. 1970. Improved procedures for clearing roots and staining parasitic and vesicular-arbuscular mycorrhizal fungi for rapid assessment of infection. Trans. Br. Mycol. Soc. 55:158-161. Plenchette, C., J.A. Fortin, and V. Furlan. 1983. Growth response of several plant species to mycorrhizae in a soil of moderate P-fertility. I. Mycorrhizal dependency under field conditions. Plant Soil 70:199-209. Plenchette, C., V. Furlan, and J.A. Fortin. 1982. Effects of different endomycorrhizal fungi on five host plants grown on calcined montmorillonite clay. J. Am. Soc. Hortic. Sci. 107:535-538. Porter, W.M. 1979. The 'most probable number' method for enumerating infective prop agules of vesicular-arbuscular mycorrhizal fungi in soil. Aust. J. Soil Res. 17:515-519. Porter, W.M., A.D. Robson, and L.K. Abbott. 1987. Factors controlling the distribution of VAM fungi in relation to soil pH. J. Appl. Ecol. 24:663-672. Powell, C. LJ., and D.J. Bagyaraj (ed.) 1984. VA mycorrhiza. CRC Press, Boca Raton, FL. Reeves, F.B., D. Wagner, T. Moorman, and J. Kiel. 1979. Role of endomycorrhizae in revegetation practices in the semi-arid west. 1. Comparison of incidence of mycorrhizae in severely dIsturbed versus natural environments. Am. J. Bot. 66:6-13. Ride, J.P., and R.B. Drysdale. 1971. A chemical method for estimating Fusarium oxysporum f. lycopersici in infected tomato plants. Physiol. Plant Pathol. 1:409-420. Rosendahl, S., R. Sen. C.M. Hepper, and C. Azcon-Aguilar. 1989. Quantification of three vesicular-arbuscular mycorrhIzal fungi (Glomus spp.) in roots of leek (Allium porrum) on the basis of activity of diagnostic enzymes after polyacrylamide gel electrophoresis. Soil BioI. Biochem. 21:519-522. Salmanowicz, B., I.-E. Nylund, and H. Wallander. 1989. High performance liquid chromatography-assay of ergosterol: A technique to estimate fungal biomass in roots with ectomycorrhiza. Agric. Ecosystem Environ. 28:437-440. Schenck, N.C. (ed.). 1982. Methods and principles of mycorrhizal research. The Am. Phytopathol. Soc., St. Paul.

VESICULAR-ARBUSCULAR MYCORRHIZAL FUNGI

377

Schenck, N.e., and Y. Perez. 1990. Isolation and culture of VA mycorrhizal fungi. p. 237258. In D.P. Labeda (ed.) Isolation of biotechnological organisms from nature. McGraw Hill Publ. Co., New York. Schenck, N. e., and Y. Perez. 1994. Identification procedures. Chapter 4. In Methods manual for VA mycorrhizae Synergistic Publ., Gainesville, FL. (In press.) Schenck, N.C., and Y.N. Schroder. 1974. Temperature response of Endogone mycorrhiza on soybean roots. Mycologia 66:600-605. Schenck, N.e., and G.S. Smith. 1982. Responses of six species of vesicular-arbuscular mycorrhizal fungi and their effects on soybean at four soil temperatures. New Phytol. 92:193-201. Schubert, A., C. Marzachi, M. Mazzitelli, M.e. Cravero, and P. Bonfante-Fasolo. 1987. Development of total and viable extraradical mycelium in the vesicular-arbuscular mycorrhizal fungus Glomus clarum Nicol. & Schenck. New Phytol. 107:183-190. Schiiepp, H., D.D. Miller, and M. Bodmer. 1987. A new technique for monitoring hyphal growth of vesicular-arbuscular mycorrhizal fungi through soil. Trans. Br. Mycol. Soc. 89:429-435. Schwab, S.M., and F.B. Reeves. 1981. The role of endomycorrhizae in revegetation practices in the semi-arid west. III. Vertical distribution of vesicular-arbuscular mycorrhizal inoculum potential. Am. J. Bot. 68:1293-1297. Simon, L., M. Lalonde, and T.D. Bruns. 1992. Specific amplification of 18S fungal ribosomal genes from vesicular-arbuscular endomycorrhizal fungi colonizing roots. Appl. Environ. Microbiol. 58:291-295. Sreenivasa, M.N., and D.J. Bagyaraj. 1988. Selection of a suitable host for mass multiplication of Glomus fasciculatum. Plant Soil 109:125-127. Sreenivasa, M.N., and D.J. Bagyaraj. 1989. Use of pesticides for mass production of vesicular-arbuscular mycorrhizal inoculum. Plant Soil 119:127-132. Strullu, D.G., e. Romand, P. Callac, E. Teoule, and Y. Demarly. 1989. Mycorrhizal synthesis in vitro between Glomus spp. and artificial seeds of alfalfa. New Phytol. 113:545548. Sylvia, D.M. 1988. Activity of external hyphae of vesicular-arbuscular mycorrhizal fungi. Soil BioI. Biochem. 20:39-43. Sylvia, D.M. 1992. Quantification of external hyphae of vesicular-arbuscular mycorrhizal fungi. p. 53-65. In A.K. Varma et al. (ed.) Methods in microbiology: Experiments with mycorrhizae. Academic Press, New York. Sylvia, D .M., and J.N. Burks. 1988. Selection of a vesicular-arbuscular mycorrhizal fungus for practical inoculation of Uniola paniculata. Mycologia 80:565-568. Sylvia, D.M., and D.H. Hubbell. 1986. Growth and sporulation of vesicular-arbuscular mycorrhizal fungi in aeroponic and membrane systems. Symbiosis 1:259-267. Sylvia, D.M., and A.G. Jarstfer. 1992. Sheared-root inocula of vesicular-arbuscular mycorrhizal fungi. Appl. Environ. Microbiol. 58:229-232. Sylvia, D.M., and L.H. Neal. 1990. Nitrogen affects the phosphorus response of VA mycorrhiza. New Phytol. 115:303-310. Sylvia, D.M., and N.C. Schenck. 1983. Application of superphosphate to mycorrhizal plants stimulates sporulation of phosphorus-tolerant vesicular-arbuscular mycorrhizal fungi. New Phytol. 95:655-661. Sylvia, D.M., and N.C. Schenck. 1984. Aerated-steam treatment to eliminate VA mycorrhizal fungi from soil. Soil BioI. Biochem. 16:675-676. Thompson, J.P. 1986. Soilless cultures of vesicular-arbuscular mycorrhizae of cereals: Effects of nutrient concentration and nitrogen source. Can. J. Bot. 64:2282-2294. Thompson, J.P. 1990. Soil sterilization methods to show VA-mycorrhizae aid P and Zn nutrition of wheat in vertisols. Soil BioI. Biochem. 22:229-240. Thomson, B.D., A.D. Robson, and L.K. Abbott. 1986. Effects of phosphorus on the formation of mycorrhizas by Gigaspora calospora and Glomus fasciculatum in relation to root carbohydrates. New Phytol. 103:751-765. Tommerup, I.e., and G.G. Briggs. 1981. Influence of agricultural chemicals on germination of vesicular-arbuscular endophyte spores. Trans. Br. Mycol. Soc. 76:326-328. Toth, R., and D. Toth. 1982. Quantifying vesicular-arbuscular mycorrhizae using a morphometric technique. Mycologia 74:182-187. Tuite, J. 1969. Plant pathological methods, fungi and bacteria. Burgess Publ. Co., Minneapolis, MN.

378

SYLVIA

Varma, A.K., J.R. Norris, and D.J. Read (ed.). 1992. Methods in microbiology: Experiments with mycorrhizae. Vol. 24. Academic Press Limited, London. Warner, A., and B. Mosse. 1983. Spread of vesieular-arbuscular mycorrhizal fungi between separate root systems. Trans. Br. Mycol. Soc. 80:353-354. Waterer, D.R., and R.R. Coltman. 1988a. Phosphorus concentration and application interval influence growth and mycorrhizal infection of tomato and onion transplants. J. Am. Soc. Hortie. Sci. 5:704-708. Waterer, D.R., and R.R. Coltman. 1988b. Effects of controlled-release phosphorus and inoculum density on the growth and mycorrhizal infection of pepper and leek transplants. HortScience 23:620-622. Williams, P.G. 1990. Disinfecting vesicular-arbuscular mycorrhizas. Mycol. Res. 94:995-997. Wilson, J.M., and M.J. Trinick. 1982. Factors affecting the estimation of numbers of infective propagules of VAMF by the most probable number method. Aust. J. Soil Res. 21:7381. Wolf, D.C., T.H. Dao, H.D. Scott, and T.L. Lavy. 1989. Influence of sterilization methods on selected soil microbiological, physical, and chemical properties. J. Environ. Qual. 18:39-44. Wright, S.F., J.B. Morton, and J.E. Sworobuk. 1987. Identification of a vesieular-arbuscular mycorrhizal fungus by using monoclonal antibodies in an enzyme-linked immunosorbent assay. Appl. Environ. Microbiol. 53:2222-2225. . Zobel, R.W., P. Del Tredici, and J.G. Torrey. 1976. Method for growing plants aeroponically. Plant Physiol. 57:344-346.

Published 1994

Chapter 19 Isolation of Microorganisms Producing Antibiotics JEFFRY J. FUHRMANN, University of Delaware, Newark, Delaware

The soil has long been recognized as a reservoir for microorganisms capable of producing a variety of antimicrobial substances. The pioneering work of Selman A. Waksman unequivocally established soil actinomycetes as prominent producers of antibiotics (Waksman, 1967). Voluminous research has revealed that many other microbial groups produce compounds having a variety of negative effects on genetically similar or dissimilar microorganisms. The types of substances represented are diverse, and include "classical" antibiotics having low molecular weights, proteinaceous bacteriocins, iron-sequestering siderophores, lytic enzymes, and volatile compounds. Although some of these compounds are of interest for their potential therapeutic value, essentially all warrant study for their possible ecological roles in soil-plant systems. With respect to the latter, particular attention has been directed toward the modifying influences of antagonistic microorganisms on phytopathogens (Fravel, 1988; Leong, 1986; Weller, 1988) and members of the Rhizobiaceae (Barnet et aI., 1988; Parker et aI., 1977). Given the diversity of microorganisms and compounds represented, the development of standard assays for the detection of antibiosis is a difficult task. Numerous types of assays have been described in the literature, each accompanied by variations developed to meet unique experimental objectives and conditions. The methods described in this chapter convey the basic principles and procedures appropriate to each general approach, and draw attention to modifications having widespread application. Emphasis will be placed on procedures applicable to bacteria (including actinomycetes) and fungi. The reader is encouraged to consult the literature for additional information related to the particular organisms and experimental systems under consideration.

Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 379

FUHRMANN

380

19-1 GENERAL PRINCIPLES 19-1.1 Preface

It is generally conceded that in vitro studies for antibiosis do not reliably predict microbial interactions in natural soil environments (Fravel, 1988; Weller, 1988; Whipps, 1987). It is, therefore, recommended that the rationale for conducting such studies be clearly established prior to their initiation. Circumstances commonly cited for initiating attempts to isolate antibiotic-producing microorganisms include pre-existing studies establishing the presence of antagonists within defined taxonomic or physiological groups of microorganisms, evidence for high frequencies of antagonistic organisms within specific niches or soil types (e.g., suppressive soils), and coincidental observations suggesting antagonistic interactions are operative within a particular experimental system. 19-1.2 Sampling Strategies

No definitive guidelines can be given for obtaining environmental samples containing a high proportion of antagonistic microorganisms. This is largely due to a persistent lack of knowledge regarding the functional distribution of such organisms in the soil environment (Labeda & Schearer, 1990; Williams & Vickers, 1986). However, there are instances in which research objectives logically suggest certain sampling strategies. Such a situation exists when one wishes to identify microorganisms for the biocontrol of certain phytopathogens or which are particularly suited to a given ecological niche. In the former case, successes have been obtained by isolating from disease suppressive soils known to contain the pathogen (Weller et al., 1988), or from the surface of naturally occurring or introduced fungal tissues such as sclerotia (Utkhede, 1984). Similarly, in searching for antagonistic rhizobacteria suited to colonizing a particular plant species, it is logical to obtain isolations from roots of that species taken from various geographical locations (Lambert et aI., 1987). In many instances, however, the successful approach has simply been to sample from diverse soils or vegetative types. Potent antagonists have also been isolated from several unusual habitats, including soil insects (Attafuah & Bradbury, 1989). 19-1.3 Selection of Isolation Method

Microorganisms to be examined for production of antimicrobial substances are commonly referred to as antagonists or producers; these terms will be applied to both potential and proven antagonists. Conversely, those microorganisms being examined for sensitivity to antagonists are referred to as test organisms or indicator organisms. Antagonists can be detected by either (i) first randomly isolating microorganisms into pure culture and subsequently testing each for activity

MICROORGANISMS PRODUCING ANTIBIOTICS

381

against indicator organisms in a systematic manner (sections 19-4 and 19-5), or (ii) using a screening technique designed to detect putative antagonists prior to isolation and subsequent retesting (section 19-6). The choice between these two approaches in dictated primarily by the experimental objectives and the cultural characteristics of the organisms involved (Johnson & Curl, 1972). Use of previously isolated organisms is the more flexible approach and necessary when the culture media or conditions appropriate for isolating the potential antagonists do not permit proper growth of the test organism(s). Strategies and procedures for the isolation of specific types of microorganisms are described elsewhere in chapter 8 of this volume. Screening techniques have the potential for relatively high isolation efficiencies for antagonists. This benefit is best realized in focused studies, such as when only one test organism is under consideration or where there is preexisting information regarding the optimal conditions for production of specific antimicrobial compounds of interest (e.g., siderophores under iron-limited conditions). Instances where this strategy has been successful are described below (section 19-6).

19-1.4 Selection of Indicator Organisms In many cases, the choice of indicator organism(s) to be used will be dictated by the objectives of the particular study at hand. In the remaining instances, however, some thought must be given to the selection of organisms to be used as indicators. Microorganisms differ in their susceptibilities to various antibiotic substances, and this can considerably influence the experimental results (Fravel, 1988). Assays should be conducted with more than one indicator organism, or with a known, highly susceptible genotype, if the maximum number of putative antagonists are desired. Alternatively, use of a resistant indicator genotype will enhance identification of strong antagonists and possibly reduce the number of false positives detected.

19-1.5 Assay Standardization An assay must be carefully standardized if it is to provide reproducible and unambiguous results (Piddock, 1990). In addition to the environmental conditions used to incubate the assay cultures, it is important to control the methods employed to prepare media and inocula. Many studies have shown that small variations in media composition can influence microbial interactions (Fravel, 1988). Agar plates should be of uniform depth and age, particularly if quantitative measurements of inhibition will be made (Vidaver et al., 1972). Inocula should be cultured and processed under defined conditions to ensure that they are of a reasonably similar physiological state when used. Standardization of cell numbers is especially important, because inhibition of the test organism is generally inversely related to its initial population density (Piddock, 1990). Additionally, the growth stage of the putative producer should be carefully monitored

382

FUHRMANN

because production of antibiotic substances generally peaks following logarithmic growth (Demain et aI., 1983; Katz & Demain, 1977), although exceptions to this general rule are well documented (Katz & Demain, 1977; Tagg et aI., 1976).

19-2 MICROBIOLOGICAL MEDIA 19-2.1 General Comments Media composition strongly affects antibiosis by modifying microbial growth, differentially altering antibiotic synthesis and inactivation, and affecting the physiological sensitivity of microorganisms to particular antibiotics (Tagg et aI., 1976). This situation is made more complex by the fact that one is often trying to detect antibiotics of unknown identity and biosynthetic requirements (Fravel, 1988). Given the above uncertainties, it is strongly recommended that more than one medium be used in assaying for antibiosis. Use of several media may also help identify artifacts caused by such factors as growth-induced media acidification or nutrient depletion (Tagg et aI., 1976). Described below are media that have commonly been employed in antibiosis assays, and which are recommended in cases where guidance regarding media composition is unavailable. However, the reader is encouraged to consult the literature for additional media that may be better suited to particular applications. Additionally, new media formulations may be appropriately used in certain situations, although it is recommended that a common standard medium also be included for comparison. Media components that have been identified as enhancing antibiosis include glycerol (Axelrood et aI., 1988) and other soluble C sources (Gross & Vidaver, 1978), insoluble C sources (e.g., carboxymethyl cellulose) (Brewer et aI., 1987), plant extracts (Fuhrmann & Wollum, 1989; Weinhold & Bowman, 1968), and various N sources (Kanner et aI., 1978) including amino acids and peptones (Brewer et aI., 1987). Glucose concentration in media has been shown to differentially regulate production of antibiotic substances produced by Pseudomonas fluorescens (James & Gutterson, 1986). 19-2.2 Common Media for Antibiosis Assays Unless otherwise noted, all media constituents are combined and sterilized by autoclaving at 121°C for 15 min. Agar is omitted for preparing liquid media. Dehydrated preparations of many of these media are available commercially and often used for antibiosis assays. Other media of a more specialized nature are described separately in subsequent sections. See Dhingra and Sinclair (1985) and Johnson and Curl (1972) for additional media.

MICROORGANISMS PRODUCING ANTIBIOTICS

383

1. Corn meal agar: Used for the culture of fungi and co-culture of bacteria and fungi. Place 40 g of corn meal in 1000 mL of distilled water and keep at 58°C for 1 h. Clarify by filtration or centrifugation. Add 15 g agar. 2. Czapek Dox agar: Used for the culture of fungi and the co-culture of fungi and bacteria. Combine 3 g sodium nitrate (NaN0 3), 1 g potassium monohydrogen phosphate (K2HP0 4), 0.5 g potassium chloride (KCl) , 0.5 g magnesium sulfate heptahydrate (MgS0 4 ·7H20), and 1000 mL water. Once all components are dissolved, add 0.1 g ferrous sulfate (FeS0 4 ) and 15 g agar. Add 30 g sucrose just prior to sterilization. 3. Glucose nutrient agar: Used for the culture of bacteria and coculture of bacteria and fungi. Combine 5 g peptone, 5 g glucose, 3 g beef extract, 1 g yeast extract, and 15 g agar with 1000 mL distilled water. 4. Glycerol arginine agar (Porter et al., 1960): Used for the culture of actinomycetes. Combine 20 g glycerol, 2.5 g L-arginine, 1 g sodium chloride (NaCl), 0.1 g calcium carbonate (CaC0 3), 0.1 ferrous sulfate heptahydrate (FeS0 4 ·7H2 0), 0.1 g magnesium sulfate heptahydrate (MgS0 4 ·7H2 0), 20 g agar, and 1000 mL distilled water. Adjust pH to 7.0. 5. King's B agar (King et al., 1954): An iron-deficient medium used for the culture of bacteria and the co-culture of bacteria and fungi. Commonly used for the detection of bacterial siderophores (see section 19-7.3). Combine 20 g proteose peptone No.3 (Difco), 10 g glycerol, 2.5 g potassium monohydrogen phosphate (K2 HP0 4 ), 6 g magnesium sulfate heptahydrate (MgS0 4 ·7H20), 15 g agar, and 1000 mL water. Adjust pH to 7.2. Some investigators reduce the inorganic salts to 1.5 g each (e.g., Misaghi et al., 1982). 6. Malt extract agar: Used for the culture of fungi and co-culture of bacteria and fungi. Dissolve 20 g of malt extract in 1000 mL of distilled water with heating, adjust pH to 6.5 (optional), and add 20 g agar. 7. Potato dextrose agar: Used for the culture of fungi and co-culture of bacteria and fungi. Boil 200 g of peeled and sliced potatoes in 500 mL distilled water for 1 h. Clarify by filtration or centrifugation. To the filtrate, add 20 g glucose, distilled water to make 1000 mL, and 15 g agar. 8. Soybean-Casein Digest Agar: Used for the culture of bacteria and the co-culture of bacteria and fungi. Also known as Tryptic Soy Agar (Difco) and Trypticase Soy Agar (BBL). Combine 17 g pancreatic digest of casein, 3 g papaic digest of soy meal, 5 g of sodium chloride (NaCl) , 2.5 g potassium monohydrogen phosphate (K2 HP0 4), 2.5 g glucose, 15 g agar, and 1000 mL distilled water. 9. Yeast extract-dextrose-carbonate agar (Wilson et al., 1967): Used for the culture of bacteria (especially for bacteriocin production).

FUHRMANN

384

Combine 10 g yeast extract, 15 g agar, 20 g CaC03 powder, and 800 mL water. Autoclave and add 200 mL of an autoclaved solution containing 100 g glucose L -1. Cool the mixture to 50°C, stir to disperse the CaC03 • 10. Yeast extract-mannitol agar (Weaver & Frederick, 1982): Used for the culture of rhizobia. Combine 10 g mannitol, 0.5 g potassium monohydrogen phosphate (K2HP0 4), 0.5 g yeast extract, 0.2 g magnesium sulfate heptahydrate (MgS04 ·7H20), 0.01 g calcium carbonate (CaC03) (optional for Bradyrhizobium) , 15 g agar, and 1000 mL water. Adjust pH to 7.0.

19-3 PREPARATION OF INOCULA 19-3.1 General Comments Inocula should ideally be cultured in the same medium to be used for the antibiosis assay, although this is probably not necessary for routine tests. For critical studies or when the inocula will be tested with several different media, it is recommended that broth cultures be washed by centrifugation and resuspended in sterile water or other diluent of choice prior to standardization and use. 19-3.2 Bacterial Inocula Inocula of non-filamentous bacteria are generally most efficiently and reproducibly prepared in broth culture. Cultures are grown to late-logarithmic or early stationary growth phase, which for most bacteria will produce a cell density of 109 mL -1 (absorbance at 600 nm "'" 0.5 [1 cm light path]). Depending upon the intended application, these suspensions may be used directly but are generally diluted lO-fold with sterile distilled water or other diluent. If growth differs greatly among isolates, cell concentrations can be standardized using either direct microscopic counts or spectrophotometric methods. Actinomycetes or other bacteria producing particulate or filamentous growth in broth culture may require specialized procedures. If cultured in broth media, the bacteria can generally be dispersed by vigorous agitation or treatment in a blender for approximately 30 s using low speed. Alternatively, conidia can be collected from sporulating lawns on agar plates by adding sterile water and dispersing the spores with a sterile rubber spatula. Remaining large clumps may be removed by sieving through sterile cheesecloth or screens. Suspensions should be standardized as described for non filamentous bacteria. Actinomycetes can also be transferred to test plates as disks cut from agar lawns by using a sterile cork hole borer (generally 5-10 mm diam.) (Patel, 1974).

MICROORGANISMS PRODUCING ANTIBIOTICS

385

19-3.3 Fungal Inocula Depending on the assay procedure to be used, fungal inocula are applied either as propagule suspensions (spores or mycelial fragments) or as agar disks. Propagule suspensions are prepared using either the broth culture or agar lawn procedure described for actinomycetes. A final suspension density of 106 to 107 colony forming units (CFU) mL -1 is suitable for most assays. Agar disks are prepared by centrally inoculating an agar plate with a fungus and subsequently cutting 5- to 10-mm diam. disks from the periphery of the developing colony (when approximately 6-8 cm diam.) with a sterile cork hole borer. 19-4 DUAL CULTURE DETECTION METHODS 19-4.1 Introduction The unifying characteristic of these techniques is that the indicator organism(s) and potential antagonist( s) are cultured simultaneously, at least for part of the duration of the assay, on an agar medium. Furthermore, it is assumed that pure cultures of the microorganisms of interest have been obtained and inocula have been prepared as described above. 19-4.2 Microbial Lawn Technique 19-4.2.1 Principles This method is suited to the study of all combinations of bacteria and fungi as indicator organisms and potential antagonists. Inoculum of the indicator organism is evenly distributed on or within an agar medium contained in a petri dish, such that a uniformly dense lawn of the organism develops in the absence of antagonism. Potential antagonists are spotted onto localized areas of the plate surface so as to produce large colonies upon subsequent growth (hence, the "giant colony" technique of Robison, 1945). Antagonism is apparent as a zone of reduced or absent growth immediately surrounding the antagonistic colony. The primary method described below involves spotting or streaking the potential antagonist(s) on the agar medium and then uniformly spraying a suspension of the test organism on the agar surface, generally following a suitable incubation period (e.g., Axelrood et aI., 1988; Misaghi et aI.,1982). 19-4.2.2 Materials 1. Petri dishes (90 mm diam.) containing 15 mL of agar medium suitable for the growth of all organisms under consideration (larger volumes may obscure inhibition zones; the agar surface should be dry).

FUHRMANN

386

2. Inocula of indicator and potential antagonists. 3. Apparatus for spraying indicator inoculum (glass units for spraying chromatography reagents are well suited, autoclavable, but relatively expensive; surface-sterilized plastic squeeze- or pump-type sprayers are also suitable for many applications; see Stansly (1947) to construct a specialized sprayer). 4. Sterile micropipet or inoculation loop capable of reproducibly delivering small volumes (::515 !lL). 19-4.2.3 Procedures Inocula of potential antagonists are applied to the surface of duplicate agar plates as localized spots or steaks. Generally, each application consists of about 10 !lL of a suspension containing a total of approximately 106 bacterial or 104 fungal propagules. Fungi to be tested as potential antagonists are sometimes applied as agar disks and are commonly limited to one per plate, whereas one to several bacterial cultures are often tested as antagonists on a single plate. Once applied to the agar plate, the potential antagonists are usually incubated for a period to permit colony development and antibiotic production prior to being oversprayed with the indicator organism. The optimum incubation time depends on the growth rates of the microorganisms involved. Fast- and slow-growing antagonists are generally incubated for 24 to 96 hand 7 to 14 d, respectively. It may be desirable to spray the plates immediately after applying the potential antagonists in cases where their growth is particularly rapid or the test organism develops slowly. Bacterial and fungal densities in spray suspensions are typically about 108 and 106 CFU mL -1, respectively. Heavier applications of the test organism should be avoided as these may partially obscure zones of inhibition; in some cases, reduced cell densities of the indicator organism may enhance antibiotic detection (Piddock, 1990). Following a suitable incubation period, the plates are examined for colonies surrounded by inhibition zones. Each inhibition zone should be rated for the presence of microbial growth (i.e., clear or turbid). The presence of any resistant clones within the inhibition zone should also be noted. Measurements of inhibition zones are made from the edge of the antagonistic colony to the point of normal growth of the indicator organism. 19-4.2.4 Variations In situations where aerosol production must be minimized or strict control of microbial numbers is needed, measured amounts of indicator inocula may be incorporated directly into the agar medium. Described below are the more common modifications of this technique. 1. The indicator organism is added to 15 mL of molten (about 45°C) sterile agar to a density of approximately 106 (bacteria) or 104

MICROORGANISMS PRODUCING ANTIBIOTICS

387

(fungi) per milliliter. The agar is poured into a petri dish, allowed to solidify, and potential antagonists are applied as described above (e.g., Angle et aI., 1981; Trinick & Parker, 1982). 2. Petri plates containing 10 mL of noninoculated agar medium are prepared in the usual manner. The solidified base agar is then overlaid with 5 mL of molten (about 45°C) sterile agar containing indicator inocula at a density of approximately 106 bacteria or 1()4 fungi per milliliter. Potential antagonists are applied as described above. Although requiring one more operation than the first variation, this modification often produces more distinct inhibition zones due to the thinner indicator layer employed (e.g., Patel, 1974; Schwinghamer, 1971). 3. In cases where the potential antagonist displays an extremely rapid growth rate relative to the indicator organism, it may be helpful to use a deferred method in which growth of the antagonist is prevented beyond what is necessary for colony development. This can be accomplished by using a modification of the chloroform fumigation technique described for the detection of bacteriocins (section 19-7.2) (e.g., Fuhrmann & Wollum, 1989). Briefly, agar plates are prepared and spotted with potential antagonists as described for the primary procedure. Following a suitable incubation period, the colonies are killed by exposure to chloroform vapor and then overlaid with 5 mL of a agar medium seeded with indicator inocula. Inhibition zones are described and measured following an additional incubation. 19-4.3 Fungal Disk Technique 19-4.3.1 Principles This technique exploits the ability of filamentous fungi to spread and completely colonize an agar plate in the absence of antagonists. It is commonly used for assessing bacterial inhibition of fungi as well as interactions between fungal genotypes. Antagonism is observed as absent, reduced, or abnormal (e.g., matted or prostrate) fungal growth in the vicinity of the co-cultured organism. 19-4.3.2 Materials 1. Petri dishes (90 mm diam.) containing 15 mL of agar medium suitable for the growth of all organisms under consideration. 2. Inocula of indicator organisms and potential antagonists (bacteria as suspensions; fungi as agar disks). 3. Sterile micropipet or inoculation loop capable of reproducibly delivering small volumes (::515 ilL).

FUHRMANN

388

19-4.3.3 Procedures For studies of bacterial inhibition of fungi, aliquots of the potential antagonists containing approximately 106 CFU are uniformly spotted near the periphery of the plate (generally four or fewer spots per plate). The spotted organisms are incubated according to the guidelines described for the Microbial Lawn Technique. A fungal disk is then placed in the center of the plate and incubation is continued. Inhibition zones are described and measured once the fungal colony is sufficiently developed for observations to be made (e.g., Howell & Stipanovic, 1980; Thomashow & Weller, 1988). Inhibition among fungi is examined by simultaneously placing disks of two or more genotypes on separate areas of an agar plate. The distances separating the disks should be standardized; 5 cm is a commonly used value. The plates are then incubated and observed periodically for interactions among the developing colonies (e.g., Manandhar et aI., 1987). 19-4.4 Cross Streak Technique 19-4.4.1 Principles This technique is generally limited to the study of antagonistic interactions among bacteria (including actinomycetes). Potential antagonists are streaked in one direction on an agar medium. The test organism is subsequently streaked perpendicularly to the antagonist. An incubation period generally separates the two operations. Antagonism is apparent as absent or reduced growth of the indicator organism near the antagonist after a final incubation period. When compared with preceding methods, the cross streak technique is labor intensive and possibly more subject to variations in inoculation application rates. However, this method has the compensating advantage of maximum control over the relative timing of inoculation between antagonists and test organisms. Additionally, it permits more than one indicator organism to be tested with a potential antagonist on a single culture plate. 19-4.4.2 Materials 1. Petri dishes (90 mm diam.) containing 15 mL of agar medium suitable for the growth of all organisms under consideration (the agar surface should be dry). 2. Inocula of indicator organisms and potential antagonists (both as cell suspensions). 3. Inoculation loops capable of delivering reproducible volumes of inocula. 19-4.4.3 Procedures A suspension of the potential antagonist (about 108 CFU mL -1) is streaked in one direction on a agar medium either along one side of the

MICROORGANISMS PRODUCING ANTIBIOTICS

389

plate or so as to divide the plate into two halves. The applied organism is incubated as described for the Microbial Lawn Technique. A cell suspension of the test organism is streaked perpendicularly to the potential antagonist beginning at a standard distance from the former streak. Streaking away from the suspected producer ensures that an apparently antagonistic interaction is not an artifact resulting from a thinning of the test inoculum near the antagonist during the streaking process. Inhibition zones are described and measured following a suitable incubation period (e.g., Smith & Miller, 1974; Trinick & Parker, 1982).

19-5 CULTURE FILTRATE METHODS 19-5.1 Introduction These methods differ from the previous techniques in that culture of the potential antagonist and test organism are separated in time and space. The potential antagonist is cultured in a broth medium, and a cell-free filtrate of the culture supernatant is prepared. The filtrate is subsequently tested for its ability to inhibit the growth of the test organism on either liquid or agar media. Although cumbersome when compared with the Dual Culture Detection Techniques (section 19-4), these methods have the advantage of minimizing the possibility of artifacts arising from factors such as media acidification and nutrient depletion. 19-5.2 Preparation of Culture Filtrates 19-5.2.1 Materials l. Broth media prepared in erlenmeyer flasks (media additives inhibi-

2. 3. 4. 5.

tory to the test organism must be avoided). Pure cultures of potential antagonists. Centrifuge. Sterile membrane filter apparatus (0.2 or 0.45 ftm). Rotary evaporator or freeze dryer (optional).

19-5.2.2 Procedures In the simplest case, potential antagonists are cultured in one or more broth media, generally well into the stationary phase of growth. This is because many antibiotic compounds are secondary growth products that are excreted into the medium only after the culture enters stationary growth. Most of the cells are removed by centrifugation and the acidity of supernatant is checked and adjusted to the desired level (typically neutrality). The supernatant is then sterilized by passage through a membrane filter and tested for inhibitory activity.

FUHRMANN

390

In some instances, it may be desirable to concentrate the supernatant obtained by centrifugation prior to sterilization and testing. A rotary evaporator operated at moderate temperatures (40-60 0c) is commonly used for this purpose, although freeze drying is better suited for use with thermolabile compounds. The supernatant may be either simply reduced in volume by some constant amount or brought to dryness and redissolved in a solvent of choice. Various extraction procedures have also been employed but will not be discussed here (e.g., Cubeta et aI., 1985; Dunlop et aI., 1989; Howell & Stipanovic, 1980; James & Gutterson, 1986; Thomashow & Weller, 1988). 19-5.3 Paper Disk Technique 19-5.3.1 Principles

This method is conducted as a modified Microbial Lawn or Fungal Disk Technique in which sterile paper disks, moistened with culture filtrates, substitute for colonies of potential antagonists. It is suited to the study of inhibitions between all combinations of bacteria and fungi. 19-5.3.2 Materials 1. Petri dishes (90 mm. diam.) contammg 15 mL of agar medium suitable for the growth of the test organism (the agar surface should be dry). 2. Inocula of test organism(s). 3. Sterile culture filtrates of potential antagonists prepared as described above. 4. Sterile, dry paper disks (I-em diam. filter paper disks are commonly used; blank paper disks used for antibiotic sensitivity tests are also available; acceptable disks may be cut from filter paper using a paper punch). 19-5.3.3 Procedures

The paper disks are moistened with the culture filtrates either by dipping the dry disks in the filtrate and then draining or by applying a standard volume to each disk with a sterile micropipet. Using sterile forceps, the impregnated disks are arranged on the noninoculated agar surface. Alternatively, agar plates seeded with the indicator organism may be employed (section 19-4.2.4). Control disks moistened with fresh/concentrated medium should also be included. Nonseeded plates are subsequently sprayed with the test organism or a fungal disk is placed on the plate. It is generally not necessary to delay application of the indicator organism. Inhibition zones are described and measured following incubation (e.g., Angle et aI., 1981; Anusuya & Sullia, 1984; Robison, 1945).

MICROORGANISMS PRODUCING ANTIBIOTICS

391

19-5.4 Agar Well Technique 19-5.4.1 Principles This method is analogous to the Paper Disk Technique except that the culture filtrates are applied directly to wells cut in the agar medium with a sterile cork hole borer. 19-5.4.2 Materials 1. Petri dishes (90 mm diam.) containing a standard amount (15-25 mL) of agar medium suitable for the growth of the test organism. 2. Inocula of test organism(s). 3. Sterile culture filtrates of potential antagonists prepared as described above. 4. Sterile micropipet. 5. Cork hole borer (5-10 mm diam.). 6. Molten agar media (optional).

19-5.4.3 Procedures Indicator lawns are initiated by seeding the agar medium or spraying with the test organism. Wells are then formed in the agar medium by removing disks cut with a flamed cork hole borer. Some investigators seal the bottom of the wells with a drop of molten agar medium (e.g., Barefoot & Klaenhammer, 1983). Measured volumes of culture filtrates or appropriate control solutions are pipeted into the agar wells. If fungal disks are to be used, these are then placed on the plate. Inhibition zones around the wells are described and measured in the usual manner (e.g., Holland & Parker, 1966; Howell & Stipanovic, 1983). 19-5.5 Radial Growth Technique 19-5.5.1 Principles This method is used for assessing inhibition of filamentous fungi. Culture filtrates are added to the agar media prior to pouring assay plates. The radial growth of a fungus on medium amended with the culture filtrate is compared with that for the same fungus on unamended plates. Any inhibition of growth on the amended plates is assumed to result from antifungal substances released into the original broth culture by the antagonist. 19-5.5.2 Materials 1. Sterile petri dishes. 2. Sterile molten agar medium (concentration of the medium should be increased to compensate for any significant dilution caused by subsequent addition of culture filtrate). 3. Sterile culture filtrates. 4. Inocula of the indicator organism (fungal disks).

FUHRMANN

392

19-5.5.3 Procedures

Culture filtrates are incorporated into fresh molten agar medium prior to pouring into petri dishes. If nonconcentrated filtrates are used, these are commonly mixed in a 1:1 ratio with fresh agar medium prepared at double the normal concentration. Alternatively, fresh medium and filtrates can be mixed together in a range of ratios. In all cases, the concentration of fresh medium in the final mixture should be standardized. Small volumes of concentrated filtrates can be added without adjusting the concentration of the fresh medium. Control plates are amended with medium that has not been used to culture potential antagonists. Once the agar has gelled, a fungal disk is placed centrally on the plate and the culture is examined on a daily basis. More frequent observations may be necessary when working with fungi having high growth rates. Measurements of fungal colony size are made once the control colony has grown to approximately 6 cm in diameter (e.g., Dunlop et aI., 1989; Fravel et aI., 1987). 19-5.6 Biomass Technique 19-5.6.1 Principles

This method is conceptually similar to the Radial Growth Technique but is conducted using liquid rather than agar media. Growth in filtrateamended and nonamended broth is assessed by either a direct or indirect estimate of biomass production. It is adaptable to the study of both bacterial and fungal inhibition. 19-5.6.2 Materials 1. Flasks containing sterile liquid medium suited to the growth of the test organism (concentration of the medium should be increased to compensate for any significant dilution caused by subsequent addition of culture filtrate). 2. Sterile culture filtrates. 3. Inocula of the indicator organism (cell suspensions or fungal disks). 4. Buchner filter apparatus or centrifuge (for fungi). 5. Spectrophotometer/turbidimeter or centrifuge (for bacteria). 6. Microbalance and oven (65°C) (for direct biomass measurements). 19-5.6.3 Procedures

Culture filtrates are combined aseptically with fresh broth medium in a manner similar to that described for the Radial Growth Technique. Amended and non-amended broth are inoculated with the test organism using a standard volume of cell suspension or a standard number of fungal disks. Bacterial cell numbers in the initial inoculum should be limited to a total of approximately 105 CFU (see 19-5.6.4).

MICROORGANISMS PRODUCING ANTIBIOTICS

393

For direct biomass measurements, the cultures are incubated for a period sufficient to allow for substantial growth of the control organism at its maximum rate of biomass accumulation. Prolonged incubation should be avoided as it may mask differences among treatments. Fungal mycelia are recovered either by filtration or centrifugation. Bacteria are recovered by centrifugation. The recovered biomass is dried to constant mass at 65°C and weighed. Inhibition is reflected in reduced biomass in the treatments amended with culture filtrates relative to the nonamended control (e.g., Elad & Chet, 1987). The reader is directed to Gerhardt (1981) for additional discussion concerning the direct measurement of bacterial biomass produced in pure cultures. Bacterial biomass can also be monitored indirectly by turbidimetry. Absorbance at a given wavelength (generally 540-640 nm) is monitored with a spectrophotometer. Specially constructed flasks equipped with cuvets (e.g., Nephlo flasks manufactured by BellCo Glass, Inc., Vineland, NJ) simplify the taking of periodic measurements. See Koch (1981) for detailed information regarding determination of microbial biomass by turbidimetry. 19-5.6.4 Comments Particular attention to inoculation rates is important when using bacteria as indicator organisms in liquid culture because of the possible presence of spontaneous antibiotic-resistant mutants in the inoculum population. Growth of such mutants can lead to an underestimation of antibiotic production by the potential antagonists (Piddock, 1990). 19-6 SCREENING METHODS 19-6.1 Introduction Screening procedures are essentially modifications of the Microbial Lawn Technique that are designed to permit detection of putative antagonists prior to their isolation into pure culture, thereby potentially yielding high isolation efficiencies. They can often be performed quantitatively so as to permit an estimation of antagonistic populations. These techniques are primarily limited by the feasibility of developing a system that is sufficiently selective and yet suited to the growth of all organisms of interest. This requirement is particularly limiting in cases where conditions suitable for the selective culture of the desired group of antagonists are detrimental to growth of the test organism. For example, use of an antifungal agent such as cycloheximide for the selective culture of bacteria producing antifungal substances would be inappropriate unless a means can be found to minimize the effect of cycloheximide on the test fungi. Strategies for limiting problems of media incompatibility include the use of nutritionally selective base agar (as opposed to one containing

FUHRMANN

394

inhibitory compounds) in combination with an overlay agar medium specifically suited to the growth of the test organism (Freeman & Tims, 1955; Herr, 1959). Some investigators have successfully used reduced concentrations of inhibitory compounds in base agar by providing a third agar layer that is used to separate the base from the indicator layer (Pugashetti et aI., 1982). Treatment of soil samples prior to dilution plating to select for certain microbial groups may also have utility (Broadbent et aI., 1971; Panthier et aI., 1979). In many cases, however, the only recourse may be to limit an experiment to the study of physiologically similar organisms or to use only previously isolated organisms. 19-6.2 Single Agar Layer Technique 19-6.2.1 Principles This approach is inflexible when compared with the Multiple Agar Layer Technique described later. Its primary limitation is that the medium must be suited to the growth of both the antagonist(s) and indicator organism(s) of interest while preventing excessive growth of other undesirable microorganisms present in the environmental sample. However, provided this requirement is not prohibitive, the Single Agar Layer Technique is both simple and direct. Instances where this method have been used successfully are for the isolation of bacteria antagonistic to Streptomyces scabies (Weinhold & Bowman, 1968) and of fluorescent Pseudomonas spp. antagonistic to Erwinia (Burr et aI., 1978). 19-6.2.2 Materials

1. Petri dishes (90 mm diam.) containing 15 mL of a suitable agar medium (larger volumes may observe inhibition zones; the agar surface should be dry). 2. Inocula of indicator organism (cell suspension). 3. Apparatus for spraying indicator inoculum (see section 19-4.2.2). 4. Pipets, dilution blanks, and angled glass rod for preparing spread plates. 19-6.2.3 Procedures Dilutions of environmental samples containing antagonists are plated in the manner used for spread plates. Once colony development is apparent, the plates are sprayed with a suspension of the indicator organism and incubated as described for the Microbial Lawn Technique. Colonies surrounded by zones of inhibition are isolated into pure culture and subsequently retested for antagonistic activity using one of the previously described pure culture methods (sections 19-4 or 19-5) (Burr et aI., 1978).

MICROORGANISMS PRODUCING ANTIBIOTICS

395

19-6.2.4 Variations Cases may arise in which the agar medium used supports only minimal growth of the indicator organism but is not actively inhibitory. In such instances, it may be helpful to prepare the spray suspension in a broth medium that encourages growth of the indicator organism and, thereby, increases the visibility of inhibition zones. Indicator inocula may be incorporated into the agar medium prior to pouring, thereby avoiding the spraying operation (e.g., Weinhold & Bowman, 1968). However, success of this approach depends upon the antagonists displaying high growth rates relative to the indicator organism. Problems with spreading colonies may be minimized by using the pour plate method in which the diluted samples are incorporated directly into the agar medium. The indicator organism is then applied to the surface following an appropriate incubation period. The disadvantage of this approach is the increased difficulty of extracting and isolating antagonists from within the agar into pure culture. 19-6.3 Multiple Agar Layer Techniques 19-6.3.1 Principles Separate layers of agar are used to afford a suitable growth environment for the antagonistic and indicator organisms, respectively. In some cases, additional layers are employed to modulate the diffusion of inhibitory media additives or microbially produced antagonistic compounds from the base agar into the indicator layer. This approach was first described by Kelner (1948) and has since been modified many times. The original method employed up to four agar layers, but two (e.g., Freeman & Tims, 1955) and three (e.g., Herr, 1959; Panthier et al., 1979) layers have also been used successfully. In general, the procedure appears better suited to the isolation of antagonistic bacteria than fungi (Pugashetti et aI., 1982). The primary method described below is modified from that of Panthier et ai. (1979). 19-6.3.2 Materials

1. Sterile, molten (40-45 0c) agar medium suitable for the growth of the antagonists of interest. 2. Sterile, molten water agar (20 g agar L -1). 3. Dilutions of environmental samples. 4. Pipets. 5. Sterile, molten agar medium suitable for the growth of the indicator organisms. 6. Inocula of indicator organisms (cell suspension). 7. Apparatus for spraying indicator inocula (see section 19-4.2.2).

396

FUHRMANN

19-6.3.3 Procedures Sample dilutions are quantitatively incorporated into the medium chosen for growth of the antagonists. The mixture is distributed into petri dishes (10 mL plate-I) and allowed to solidify (base layer). The base layer is immediately covered with 5 mL of water agar (intermediate layer) which serves to confine colony development. After a suitable incubation period, 10 mL of the indicator medium is applied to the plate (indicator layer), cooled, and sprayed with a suspension of the indicator organism. After further incubation, any inhibition zones are described and, if desired, counted. Antagonistic organisms are recovered from the base layer, purified, and retested under pure culture conditions (section 19-4 or 19-5). 19-6.3.4 Variations Sample dilutions can be incorporated into the intermediate layer or can be applied as a completely separate layer situated between the base and intermediate layer (Kelner, 1948). Similarly, indicator inocula may be mixed with the indicator layer prior to pouring or simply spread over the indicator layer, thereby avoiding the spraying operation. 19-6.3.5 Comments The original method of Panthier et al. (1979) was for the isolation of actinomycetes antagonistic to various Rhizobium and Bradyrhizobium spp. For this objective, the sample dilutions contained 7 g phenol L -1 to reduce the growth of undesirable bacteria. Similarly, the base layer contained cycloheximide (200 mg L -1) to inhibit fungal growth. The reader is referred to Pugashetti et al. (1982) and Kelner (1948) for other strategies for using selective agents with this technique.

19-7 METHODS FOR SELECTED CLASSES OF COMPOUNDS 19-7.1 Introduction It is beyond the scope of this chapter to provide detailed experimental procedures for studying the entire range of known antimicrobial substances. Nevertheless, certain classes of compounds have received special treatment by investigators or have elicited development of unique and reasonably standardized protocols for their study. These compounds generally exhibit unique physical properties or modes of action, and this may, in part, explain their emphasis in the literature.

MICROORGANISMS PRODUCING ANTIBIOTICS

397

19-7.2 Bacteriocins 19-7.2.1 Principles Bacteriocins may be defined as nonreplicating, proteinaceous biocides whose antagonistic activity is limited to genetically similar organisms (Vidaver, 1976). There is, however, uncertainty regarding how strictly this definition should be interpreted (Hardy, 1982; Vidaver, 1983). The term is usually restricted to compounds produced by prokaryotic organisms, although the term has also been applied to substances produced by protozoa and yeasts (Vidaver, 1983). Genetically similar organisms are generally defined as those belonging to different species of the same genus or similar genera within a family. As is the case with other antimicrobial substances, bacteriocin production is strongly affected by media composition and other environmental conditions. Assays are typically conducted using agar media as these have generally been shown to give more reproducible results and greater sensitivity than liquid media. Although some organisms produce bacteriocins freely, it is sometimes necessary to induce production by treating cultures with ultraviolet radiation or chemical agents. The double agar layer method described below is modified from that of Vidaver et al. (1972). 19-7.2.2 Materials 1. Glass petri dishes containing 25 mL of agar medium suitable for the growth of the potential producers (partially dried plates [e.g., 28°C for 4 d] have been reported to enhance clarity of inhibition zones). 2. Inocula of producer and indicator organisms as cell suspensions (generally several microorganisms genetically similar and dissimilar to the producers are used as indicator organisms). 3. Sterile micropipet or multipoint replicator. 4. Chloroform. 5. Sterile, molten (45°C) soft agar medium (agar concentration reduced to 7.5 g L -1). 19-7.2.3 Procedures One to several potential producers are spotted onto the agar medium by micropipet (10 ~L) or multipoint inoculator. Following an incubation period to allow for colony development, the spotted organisms are killed by exposure to chloroform vapor (hence the need for glass petri dishes). This is conveniently done by inverting the petri dishes, transferring 3 mL of chloroform to the dish lid, replacing the inverted bottom, and allowing the chloroform to slowly evaporate in a fume hood (1-2 h). Any residual chloroform vapor is allowed to dissipate for one to several hours (overnight). Some investigators prefer to remove the colonies with a glass

FUHRMANN

398

microscope slide prior to treatment with chloroform (Tapia-Hernandez et aI., 1990). It should be noted that some bacteriocins are inactivated by chloroform (Tagg et aI., 1976). The chloroform-treated base agar is then overlaid with 3 to 5 mL of the molten soft agar inoculated with the indicator organisms to a density of approximately 106 cells mL -1. Inhibition zones are described and measured following an additional incubation period. Induction of bacteriocin production can be attempted by using either or both of the following procedures (Mayr-Harting et aI., 1972): 1. Ultraviolet radiation-Exponentially growing broth cultures are irradiated (254 nm) in a 2-mm layer in a dish for periods ranging from 30 to 300 s at a distance of 30 to 50 cm. The proper conditions are determined empirically. The irradiated culture is incubated to late logarithmic or early stationary growth phase in the dark and is then used to prepare inocula in the usual manner. 2. Chemical treatment-Exponentially growing cultures are treated with mitomycin C to a final concentration ranging from 0.1 to 1.0 ~g mL -1; occasionally concentrations up to 10 ~g mL -1 are used. The optimal concentration is determined empirically. The treated culture is incubated to late logarithmic or early stationary growth phase, centrifuged, and the cells are then used to prepare inocula. 19-7.2.4 Supplemental Procedures

Once putative bacteriocin producers have been identified, several supplemental tests are often performed to verify and characterize the suspected bacteriocins. Two common assays are summarized below. Not discussed here are assays for biocidal activity and for sensitivity to proteolytic enzymes, catalase, chloroform, heat, and acidity. The reader is referred to Mayr-Harting et al. (1972), Gross and Vidaver (1978), and TapiaHernandez et al. (1990) for further information and alternative protocols. 1. Assay for Phage-To verify that inhibition did not result from phage activity, agar disks are removed from inhibition zones, homogenized, serially diluted, and aliquots are spotted onto agar lawns of the indicator organism (see chapter 7 by Angle in this book). Phage produce decreasing numbers of discrete plaques with increasing dilution, whereas bacteriocins produce a single inhibition zone that gradually decreases in intensity with increasing dilution. It should be realized that a positive assay for phage does not rule out simultaneous bacteriocin production. 2. Assay for Dializability- Most bacteriocins are large molecules due to their proteinaceous nature, generally having molecular weights > 12 000 (Hardy, 1982). Estimates of the minimum molecular weight of inhibitory substances can be obtained by culturing the putative producer on an autoclaved dialysis membrane placed on an agar medium. Membranes having molecular weight exclusions ranging from 6000 to 12 000 are commonly employed. After a suit-

MICROORGANISMS PRODUCING ANTIBIOTICS

399

able incubation period, the membrane and associated colony are aseptically removed from the plate, and the base layer is overlaid with seeded soft agar in the normal manner. Development of inhibition zones indicates that the associated antimicrobial substance is smaller than the corresponding exclusion size of the membrane. 19-7.3 Siderophores 19-7.3.1 Principles Siderophores are iron (III) chelating compounds produced by many microorganisms under Fe-limited growth conditions that function to scavenge Fe needed for growth. Siderophores produced by one organism may reduce Fe availability to other microorganisms that either do not produce siderophores or compatible siderophore receptors or which produce siderophores with relatively weak chelating abilities (Leong, 1986). Therefore, antagonism results from competition for an essential nutrient rather than from an actively harmful compound. It is essential that Fe contamination of media and glassware be minimized so as to avoid repression of siderophore production. Media which are inherently deficient in Fe are most commonly used, although procedures for removing contaminating Fe from media are available (Waring & Werkman, 1942). For bacteria, and particularly for the fluorescent Pseudomonas spp., King's B medium has become the standard for most applications. Glassware should be scrupulously cleaned and treated with 6 M hydrochloric acid (Hel) to remove traces of Fe. Water used for making media and rinsing glassware should be deionized or distilled at minimum, preferably double glass-distilled. Many of the previously discussed detection and screening procedures may be used to identify microorganisms that exhibit siderophore-mediated antagonism. The procedure described below is essentially a specialized version of the Microbial Lawn Technique. Screening procedures for siderophore production by fluorescent Pseudomonas spp. have also been described (e.g., Burr et aI., 1978). 19-7.3.2 Materials 1. Petri dishes containing 15 mL of King's B agar or other Fe deficient medium. 2. Inocula of potential antagonists and indicator organisms (cell suspensions). 3. Sterile micropipet. 4. Spray apparatus for applying indicator organisms.

19-7.3.3 Procedures The assay is conducted as described for the Microbial Lawn Technique. Antagonistic microorganisms should be retested using both iron

FUHRMANN

400

deficient and iron sufficient media, the latter amended to 50 !-lm of Fe as FeCI3 . Putative siderophore producers are identified as exhibiting antagonistic activity in the Fe deficient medium only. 19-7.3.4 Comments Fluorescent Pseudomonas spp. often cause strong siderophore-mediated antagonisms and are characterized by the production of diffusible yellow-green pigments (siderophores) that exhibit a blue-green fluorescence under longwave (365 nm) ultraviolet radiation. This fluorescence is eliminated by the addition of ferric Fe to growth media containing the siderophores (Misaghi et aI., 1982). Many other microorganisms produce siderophores which mayor may not fluoresce. Additional confirmation of siderophore production may be obtained using the colorimetric assay of Schwyn and Neilands (1987). Loss of inhibitory activity on iron sufficient media is not considered proof that an antagonism is siderophore-induced. Gill and Warren (1988) characterized a fluorescent pseudomonad that produced an antagonistic substance in iron limited culture that did not function in Fe assimilation. A fluorescent siderophore was co-produced in Fe deficient media, whereas neither compound was produced in media amended with Fe. 19-7.4 Mycolytic Enzymes 19-7.4.1 Principles Microorganisms may antagonize fungi by producing enzymes that lyse fungal cell walls. One procedure for assessing mycolytic activity, and the one described here, is to apply dilutions of environmental samples to myceliallawns and subsequently examine the mycelia for zones of lysis (Carter & Lockwood, 1957). Alternatively, previously isolated microorganisms are grown in media amended with compounds which induce mycolytic enzymes, and the resulting culture supernatants are tested for specific lytic enzymes by using appropriate colorimetric assays (e.g., Elad et aI., 1982). 19-7.4.2 Materials 1. Petri dishes containing 15 mL of peptone agar (L -1: 5 g peptone, 20 g agar) seeded throughout with propagules (about 105 mL -1) of the test fungus. 2. Diluted environmental samples. 3. Spray apparatus for applying sample dilutions.

19-7.4.3 Procedures The seeded agar plates are incubated until a uniform mycelial lawn is obtained. The plates are then sprayed with the sample dilutions and incubated for an additional 2 to 21 d. Lytic colonies will be surrounded by clear

MICROORGANISMS PRODUCING ANTIBIOTICS

401

zones free of fungal hyphae. The associated organisms are isolated into pure culture and retested for lytic activity. 19-7.4.4 Comments

Carter and Lockwood (1957) identified a number of factors that affect the results of this screening procedure. Seeding of the agar with fungal propagules by incorporation was found superior to surface application. Prostrate mycelial growth was necessary for detection of lysis, and this depended on the nature of the test fungus and medium. Glomerella cingulata (Ston.) proved to be superior to the other indicator fungi tested, although positive results were obtained for most species examined. In general, media which supported sparse fungal growth, such as peptone agar, gave the greatest isolation efficiencies. 19-7.5 Volatile Compounds 19-7.5.1 Principles

Volatile compounds have been examined primarily for antagonistic effects towards fungi (Dennis & Webster, 1971; Keel et al., 1989). Certainly one factor contributing to this emphasis is the radial growth habit of fungal mycelia that can be easily monitored under conditions appropriate to the study of volatile compounds. Antagonism is observed as a reduced rate of radial enlargement of a fungal colony in the presence of the inhibitory compound(s) when compared to negative control cultures. Recent research has focused on the role of cyanide as a volatile inhibitor of microbial growth (e.g., Ahl et al., 1986; Keel et al., 1989). Described below are both a general procedure and one designed specifically for the detection of cyanide production. 19-7.5.2 Materials (General) (Dennis & Webster, 1971) 1. Separate petri dishes containing agar media suitable for growth of

the antagonists and test organisms, respectively. 2. Inocula of the antagonists (cell suspensions or agar disks). 3. Inocula of the test fungus (agar disks). 19-7.5.3 Procedures (General)

The antagonists are applied individually to plates of the proper agar medium. Inocula should be applied so as to produce abundant growth. Depending on the relative growth rates of the organisms being tested, the subsequent operation is either performed immediately or following an appropriate incubation period. The petri dish lids are replaced by an inverted dish bottom containing an agar disk of the test fungus centrally located on a suitable agar medium. To provide controls, the bottom containing the test fungus is placed on a non-inoculated plate of the antagonist medium.

FUHRMANN

402

The two dish bottoms are sealed together with parafilm or similar material. Radial growth of the test fungus is monitored and all treatments are measured once the control colonies are approximately 6 cm in diameter. If preferred, this and the following procedure may be conducted by using divided petri dishes rather than by joining two undivided dishes. 19-7.5.4 Materials (Cyanide) (Bakker & Schippers, 1987) 1. Petri dishes containing an agar medium suitable for the antagonists that has been amended with 4.4 g glycine L -1 (glycine has been shown to stimulate cyanide production; soybean-casein digest agar and King's B agar have both been used successfully as basal media). 2. Inocula of the potential cyanogenic antagonists. 3. Filter paper. 4. Cyanide detection solution (5 g picric acid and 20 g of sodium carbonate [NaC0 3] dissolved in 1000 mL distilled water). 19-7.5.5 Procedures (Cyanide) Single antagonists are streaked onto the glycine-containing medium and the plates are inverted. A piece of filter paper impregnated with the cyanide detection solution is placed in the inverted lid of the petri dish, and the plate is sealed with parafilm or similar material. Upon incubation, cyanogenic organisms will cause the filter paper to change color from yellow to orange-brown. Organisms rated positive for cyanide production are subsequently tested for antagonism toward test fungi according to the general procedure described above (19-7.5.3); the glycine-amended medium should be retained as the antagonist medium. Cyanide production can be verified using procedures described by Bakker and Schippers (1987). 19-8 CONCLUDING COMMENTS Efforts to isolate microorganisms capable of producing antimicrobial compounds is clearly hampered by a lack of information regarding their spatial distribution and ecological relevance in soil environments (Labeda & Schearer, 1990; Williams & Vickers, 1986). This deficiency is reflected in the inability of in vitro assays for antagonism to reliably predict microbial interactions in natural soil environments (Fravel, 1988; Weller, 1988; Whipps, 1987). It is particularly remarkable that, despite the rich history of research concerning antibiotic production by soil microorganisms, there are only intermittent reports giving direct evidence for production of antibiotics in situ (Thomashow et aI., 1990). Nevertheless, these infrequent reports, combined with the common observation that many soil isolates produce antimicrobial compounds in vitro, will undoubtedly sustain interest in this area of research. The increasing efforts to develop rational strategies for using biological agents to enhance plant productivity will certainly intensify this interest.

MICROORGANISMS PRODUCING ANTIBIOTICS

403

REFERENCES Ahl, P., C. Voisard, and G. Defago. 1986. Iron bound-siderophores, cyanic acid, and antibiotics involved in suppression of Thielaviopsis basicola by a Pseudomonas fiuorescens strain. J. Phytopathol. 116: 121-134. Angle, J.S., B.K. Pugashetti, and G.H. Wagner. 1981. Fungal effects on Rhizobiumjaponicum-soybean symbiosis. Agron. J 73:301-306. Anusuya, D., and S.B. SulIia. 1984. The antibiotic effect of culture filtrates of some soil fungi on rhizobial growth in cultures. Plant Soil 77:387-390. Attafuah, A., and J.F. Bradbury. 1989. Pseudomonas antimicrobica, a new species strongly antagonistic to plant pathogens. J. Appl. Bacteriol. 67:567-573. Axelrood, P.E., M. Rella, and M.N. Schroth. 1988. Role of antibiosis in competition of Erwinia strains in potato infection courts. Appl. Environ. Microbiol. 54:1222-1229. Bakker, A.W., and B. Schippers. 1987. Microbial cyanide production in the rhizosphere in relation to potato yield reduction and Pseudomonas spp-mediated plant growth-stimulation. Soil BioI. Biochem. 19:451-457. Barefoot, S.F., and T.R. Klaenhammer. 1983. Detection and activity of lactacin B, a bacteriocin produced by Lactobacillus acidophilus. Appl. Environ. Microbiol. 45: 1808-1815. Barnet, Y.M., M.J. Trinick, R.A. Date, and R.J. Roughley. 1988. Ecology of the root-nodule bacteria. p. 1-22. In w.G. Murrell and 1.R. Kennedy (ed.) Microbiology in action. John Wiley and Sons, New York. Brewer, D., F.G. Mason, and A. Taylor. 1987. The production of alamethicins by Trichoderma spp. Can. J. Microbiol. 33:619-625. Broadbent, P., K.F. Baker, and Y. Waterworth. 1971. Bacteria and actinomycetes antagonistic to fungal root pathogens in Australian soils. Aust. J. BioI. Sci. 24:925-944. Burr, T.J., M.N. Schroth, and T. Suslow. 1978. Increased potato yields by treatment of seedpieces with specific strains of Pseudomonas fiuorescens and P. putida. Phytopathology 68:1377-1383. Carter, H.P., and J.L. Lockwood. 1957. Methods for estimating numbers of soil microorganisms lytic to fungi. Phytopathology 47:151-154. Cubeta, M.A., G.L. Harman, and J.B. Sinclair. 1985. Interaction between Bacillus subtilis and fungi associated with soybean seeds. Plant Dis. 69:506-509. Demain, A.L., Y. Aharonowitz, and J.-F. Martin. 1983. Metabolic control of secondary biosynthetic pathways. p. 49-72. In L.c. Vining (ed.) Biochemistry and genetic regulation of commercially important antibiotics. Addison-Wesley Publ. Co., London. Dennis, C., and J. Webster. 1971. Antagonistic properties of species-groups of Trichoderma. II. Production of volatile antibiotics. Trans. Br. Mycol. Soc. 57:41-48. Dhingra, O.D., and J.B. Sinclair. 1985. Basic plant pathology methods. CRC Press, Boca Raton, FL. Dunlop, R.W., A. Simon, K. Sivasithamparam, and E.L. Ghisalberti. 1989. An antibiotic from Trichderma koningii active against soilborne plant pathogens. J. Natl. Prod. 52:67-74. Elad, Y., and 1. Chet. 1987. Possible role of competition for nutrients in biocontrol of Pythium damping-off by bacteria. Phytopathology 77:190-195. Elad, y', 1. Chet, and Y. Henis. 1982. Degradation of plant pathogenic fungi by Trichoderma harzianum, Can. J. Microbiol. 28:719-725. Fravel, D.R. 1988. Role of antibiosis in the biocontrol of plant diseases. Annu. Rev. Phytopathol. 26:75-91. Fravel, D.R., K.K. Kim, and G.c. Papavizas. 1987. Viability of microsclerotia of Verticillium dahliae reduced by a metabolite produced by Talaromyces fiavus. Phytopathology 77:616-619. Freeman, T.E., and E.C. Tims. 1955. Antibiosis in relation to pink root of shallots. Phytopathology 45:440-442. Fuhrmann, J., and A.G. Wollum, II. 1989. In vitro growth responses of Bradyrhizobium japonicum to soybean rhizosphere bacteria. Soil BioI. Biochem. 21:131-135. Gerhardt, P. 1981. Diluents and biomass measurement. p. 504-507. In P. Gerhardt (ed.) Manual of methods for general bacteriology. Am. Soc. Microbiol., Washington, DC. Gill, P.R., Jr., and G.J. Warren. 1988. An iron-antagonized fungistatic agent that is not required for iron assimilation from a fluorescent rhizosphere pseudomonad. J. Bacteriol. 170: 163-170.

404

FUHRMANN

Gross, D.C., and A.K. Vidaver. 1978. Bacteriocin-like substances produced by Rhizobium japonicum and other slow-growing rhizobia. Appl. Environ. Microbiol. 36:936-943. Hardy, K.G. 1982. Bacteriocins. p. 368-378. In R.G. Burns and J.H. Slater (ed.) Experimental microbial ecology. Blackwell Sci. Publ., Oxford. Herr, L.J. 1959. A method of assaying soils for numbers of actinomycetes antagonistic to fungal pathogens. Phytopathology 49:270-273. Holland, A.A., and C.A. Parker. 1966. Studies on microbial antagonism in the establishment of clover pasture. II. The effect of saprophytic fungi upon Rhizobium tri/olii and the growth of subterranean clover. Plant Soil 25:329-340. Howell, c.R., and R.D. Stipanovic. 1980. Suppression of Pythium ultimum-induced damping-off of cotton seedlings by Pseudomonas fluorescens and its antibiotic, pyoluteorin. Phytopathology 70:712-715. Howell, C.R., and R.D. Stipanovic. 1983. Gliovirin, a new antibiotic from Gliocladium virens, and its role in the biological control of Pythium ultimum. Can. J. Microbiol. 29:321-324. James, D.W., Jr., and N.!. Gutterson. 1986. Multiple antibiotics produced by Pseudomonas fluorescens HV37a and their differential regulation by glucose. Appl. Environ. MicrobioI. 52:1183-1189. Johnson, L.F., and E.A. Curl. 1972. Methods for research on the ecology of soil-borne plant pathogens. Burgess Publ. Co., Minneapolis. Kanner, D., N.N. Gerber, and R. Bartha. 1978. Pattern of phenazine pigment production by a strain of Pseudomonas aeruginosa. J. Bacteriol. 134:690-692. Katz, E., and A.L. Demain. 1977. The peptide antibiotics of Bacillus: chemistry, biogenesis, and possible functions. Bacteriol. Rev. 41:449-474. Keel, C., C. Voisard, C.H. Berling, G. Kahr, and G. Defago. 1989. Iron sufficiency, a prerequisite for the suppression of tobacco black root rot by Pseudomonas fluorescens strain CHAO under gnotobiotic conditions. Phytopathology 79:584-589. Kelner, A. 1948. A method for investigating large microbial populations for antibiotic activity. J. Bacteriol. 56:157-162. King, E.O., M.K. Ward, and D.E. Raney. 1954. Two simple media for the demonstration of pyocyanin and fluorescin. J. Lab. Clin. Med. 44:301-307. Koch, A.L. 1981. Growth measurement. p. 179-207. In P. Gerhardt (ed.) Manual of methods for general bacteriology. Am. Soc. Microbiol., Washington, DC. Labeda, D.P., and M.C. Shearer. 1990. Isolation of actinomycetes for biotechnological applications. p. 1-19. In D.P. Labeda (ed.) Isolation of biotechnological organisms from nature. McGraw-Hill Publ. Co., New York. Lambert, B., F. Leyns, L. van Rooyen, F. Gossele, Y. Papon, and J. Swings. 1987. Rhizobacteria of maize and their antifungal activities. Appl. Environ. Microbiol. 53:18661871. Leong, J. 1986. Siderophores: Their biochemistry and possible role in the biocontrol of plant pathogens. Ann. Rev. Phytopathol. 24:187-209. Manandhar, J.B., P.N. Thapliyal, K.J. Cavanaugh, and J.B. Sinclair. 1987. Interaction between pathogenic and saprobic fungi isolated from soybean roots and seeds. Mycopathology 98:69-75. Mayr-Harting, A., A.J. Hedges, and R.C.W. Berkeley. 1972. Methods for studying bacteriocins. Meth. Microbiol. 7A:315-422. Misaghi, I.I., L.J. Stowell, R.G. Grogan, and L.c. Spearman. 1982. Fungistatic activity of water-soluble fluorescent pigments of fluorescent pseudomonads. Phytopathology 72:33-36. Panthier, J.J., H.G. Diem, and Y. Dommergues. 1979. Rapid method to enumerate and isolate soil actinomycetes antagonistic toward rhizobia. Soil BioI. Biochem. 11:443-445. Parker, C.A., M.J. Trinick, and D.L. Chatel. 1977. Rhizobia as soil and rhizosphere inhabitants. p. 311-352. In R.W.F. Hardy and A.H. Gibson (ed.) A treatise on dinitrogen fixation. Section IV: Agronomy and ecology. John Wiley and Sons, New York. Patel, J.J. 1974. Antagonism of actinomycetes against rhizobia. Plant Soil 41:395-402. Piddock, L.J.Y. 1990. Techniques used for the determination of antimicrobial resistance and sensitivity in bacteria. J. Appl. Bacteriol. 68:307-318. Porter, J.N., J.J. Wilhen, and H.D. Tresner. 1960. Method for preferential isolation of actinomycetes from soil. Appl. Microbiol. 8:174-178. Pugashetti, B.K., J.S. Angle, and G.H. Wagner. 1982. Soil microorganisms antagonistic toward Rhizobium japonicum. Soil BioI. Biochem. 14:45-49.

MICROORGANISMS PRODUCING ANTIBIOTICS

405

Robison, R.S. 1945. The antagonistic action of the by-products of several soil microorganisms on the activities of the legume bacteria. Soil Sci. Soc. Am. Proc. 10:206-210. Schwinghamer, E.A. 1971. Antagonism between strains of Rhizobium tri/olii in culture. Soil BioI. Biochem. 3:355-363. Schwyn, B., and J.B. Neilands. 1987. Universal chemical assay for the detection and determination of siderophores. Anal. Biochem. 160:47-56. Smith, R.S., and R.H. Miller. 1974. Interactions between Rhizobium japonicum and soybean rhizosphere bacteria. Agron. J. 66:564-567. Stansly, P.G. 1947. A bacterial spray apparatus useful in searching for antibiotic-producing microorganisms. J. Bacteriol. 54:443-445. Tagg, J.R., A.S. Dajani, and L.W. Wannamaker. 1976. Bacteriocins of gram-positive bacteria. Bacteriol. Rev. 40:722-756. Tapia-Hernandez, A., M.A. Mascarua-Esparza, and J. Caballero-Mellado. 1990. Production of bacteriocins and siderophore-like activity by Azospirillum brasilense. Microbios 64:73-83. Thomashow, L.S., and D.M. Weller. 1988. Role of a phenazine antibiotic from Pseudomonas fluorescens in biological control of Gaeumannomyces graminis var. tritid. J. Bacteriol. 170:3499-3508. Thomashow, L.S., D.M. Weller, R.F. Bonsall, and L.S. Pierson, III. 1990. Production of the antibiotic phenazine-1-carboxylic acid by fluorescent Pseudomonas species in the rhizosphere of wheat. Appl. Environ. Microbiol. 56:908-912. Trinink, M.J., and C.A. Parker. 1982. Self-inhibition ofrhizobial strains and the influence of cultural conditions on microbial interactions. Soil BioI. Biochem. 14:79-86. Utkhede, R.S. 1984. Antagonism of isolates of Bacillus subtilis to Phytophthora cactorum. Can. J. Bot. 62:1032-1035. Vidaver, A.K. 1976. Prospects for control of phytopathogenic bacteria by bacteriophages and bacteriocins. Annu. Rev. Phytopathol. 14:451-465. Vidaver, A.K. 1983. Bacteriocins: The Lure and the reality. Plant Dis. 67:471-475. Vidaver, A.K., M.L. Mathys, M.E. Thomas, and M.L. Schuster. 1972. Bacteriocins of the phytopathogens Pseudomonas syringae, P. glycinea, and P. phaseolicola. Can. J. Microbiol. 18:705-713. Waksman, S.A. 1967. The actinomycetes. Ronald Press Co., New York. Waring, W.S., and C.H. Werkman. 1942. Growth of bacteria in an iron-free medium. Arch. Biochem. 1:303-310. Weaver, R.W., and L.R Frederick. 1982. Rhizobium. p. 1043-1070. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Weinhold, A.R., and T. Bowman. 1968. Selective inhibition of the potato scab pathogen by antagonistic bacteria and substrate influence on antibiotic production. Plant Soil 28:1224. Weller, D.M., W.J. Howie, and R.J. Cook. 1988. Relationship between in vitro inhibition of Gaeumannomyces graminis var. tritid and suppression of take-all of wheat by fluorescent pseudomonads. Phytopathology 78:1094-1100. Weller, D.M. 1988. Biological control of soilborne plant pathogens in the rhizosphere with bacteria. Ann. Rev. Phytopathol. 26:379-407. Whipps, J.M. 1987. Effect of media on growth and interactions between a range of soil-borne glasshouse pathogens and antagonistic fungi. New Phytol. 107:127-142. Williams, S.T., and J.C. Vickers. 1986. The ecology of antibiotic production. Microb. Ecol. 12:43-52. Wilson, E.E., F.M. Zeitoun, and D.L. Frederickson. 1967. Bacterial phloen canker, a new disease of Persian walnut trees. Phytopathology 57:618-621.

Published 1994

Chapter 20 Microbiological Procedures for Biodegradation Research DENNIS D. FOCHT, University of California, Riverside, California

The geochemical cycling of organic wastes through the metabolism of microorganisms was considered as fait accompli during the first 75 yr that the discipline of microbiology had existed. Although the phenomenal growth in the production of synthetic organic chemicals following the end of World War II should have raised questions about the limitation of microbial metabolic diversity, soil was still considered as the great cleansing agent. Gale (1952) best summarized the concept of microbial infallibility by suggesting that there existed in nature a microorganism capable of metabolizing any conceivable compound that the organic chemist might choose to synthesize. The opposing doctrine of molecular recalcitrance was advanced by Alexander (1965), who noted that many synthetic chemicals-particularly the chlorinated hydrocarbon insecticides-were persistent in soil. He questioned the validity of expecting that microbial enzymes, which were a product of 1 to 2 billion years of evolution, should attack substrates that were introduced recently into the environment. To keep the argument in perspective, it should be noted that there is nothing mystical about xenobiotic or synthetic chemicals ("xenos" der. Greek "foreign"). Some are biodegradable (e.g., organophosphate insecticides) and some are very recalcitrant (e.g., chlorinated aromatic hydrocarbons). Similarly, the same can be noted for natural organics (e.g., amino acids and carbohydrates vs. lignin and humic polymers). As biodegradation research has come into its own as a discipline over the last 25 yr, it is apparent that most recalcitrant compounds-particularly the chlorinated aromatic hydrocarbons (CAHs) - are neither used as growth substrates nor mineralized by a single microbial species. Rather, they are metabolized by the action of two or more physiologically different microorganisms. In most of these cases, the organism that carries out the initial oxidation is unable to use the recalcitrant compound as a growth substrate, and must rely on an exogenous substrate to sustain its activity Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

407

408

FOCHT

and survival. The phenomenon whereby an organism can transform a compound, but not use it as a growth substrate has been termed cometabolism (Horvath, 1972). Perhaps the term fortuitous metabolism (Dagley, 1984) better explains the underlying phenomenon: i.e., enzymes, which have resulted from the evolutionary selection of a microorganism to a natural substrate (e.g., benzene), fortuitously catalyze the metabolism of a structurally similar xenobiotic substrate (e.g., chlorobenzene) because of low enzyme specificity. However, at some point in the catabolic pathway, a particular enzyme with high substrate specificity will not attack the xenobiotic product, which then accumulates in pure culture. Thus, methods designed to select for a single organism able to use the target compound as a growth substrate will fail with recalcitrant compounds. This failure, however, cannot be taken as an indication that the target compound is inherently recalcitrant to microbial attack because the xenobiotic metabolite may be metabolized by another microorganism, having enzymes with different substrate specificity. The enrichment culture procedure, which is the molecular backbone of biodegradation research, has been successful when its limitations regarding growth are taken into consideration. The use of structurally similar growth-promoting analogs has greatly enhanced the use of the enrichment culture procedure in understanding how recalcitrant compounds are metabolized in culture and in soil. This process, referred to as analog enrichment (Horvath, 1972; Brunner et al., 1985), has proven useful in the isolation of bacteria capable of cometabolizing many recalcitrant compounds, particularly DDT, PCBs, and other chlorinated aromatic hydrocarbons that are not used as growth substrates by any single microorganism. Cyclohexane is an excellent example of a compound that is biodegradable in soil, yet does not select for isolates able to use it as a growth substrate (Perry, 1984). This enigma was solved in independent studies by Beam and Perry (1973, 1974) and deKlerk and van der Linden (1974), who showed that two species were involved in complete catabolism of cyclohexane. Bacteria isolated from enrichment culture on n-alkanes, were able to fortuitously oxidize cyclohexane to cyclohexanol by a monooxygenase, yet were unable to further metabolize the product. Another strain, obtained from enrichment culture on cyclohexanol, was able to complete the process. These elegant studies show why it would not be possible to isolate either organism by selective enrichment with cyclohexane because the first member, the cometabolizer, gets insufficient energy from the substrate and is, therefore, unable to grow and provide sufficient levels of cyclohexanol for enrichment of the commensal. A similar situation exists for the cometabolic-commensal mineralization of PCBs in soil, in which the first member obtains no energy for its growth (Focht & Brunner, 1985). The intent of this chapter is to focus on the isolation, growth, and physiology of the microorganisms involved in biodegradation. Soil biodegradation studies will not be covered here since the methods and procedures (e.g., respiration, CO2 evolution, extraction and chromatographic analysis of substrates and metabolites) can be found in other chapters of this book.

PROCEDURES FOR BIODEGRADATION RESEARCH

409

20-1 THE ENRICHMENT CULTURE 20-1.1 Principles The enrichment culture, also referred to as selective or elective culture, is based on the Darwinian concept of natural selection, namely, that the organism best able to exploit the niche (i.e., use the substrate for growth and energy) under all other environmental constraints (temperature, pH, O 2 , and essential nutrients) will be the one that is selected. It is not the intent of this procedure to use inorganic nutrient conditions similar to those in soil (Angle et aI., 1991) or in any other environment because these conditions are suboptimal for microbial growth. The objective of the enrichment culture procedure is generally to provide conditions for the rapid isolation of the desired catabolic phenotype. This procedure will be focused only on aerobic selection as this is the oldest, fastest, and most commonly used condition for selection of xenobiotic-degraders. Moreover, the isolation and significance of anaerobic bacteria that degrade xenobiotic compounds is covered in chapter 13 by Tiedje in this book. Methods for selection and isolation will be confined to mesophilic conditions, although the procedures herein could be readily adapted to thermophilic or psychrophilic conditions for selection from environments likely to have a different microbial flora, such as high-temperature composts or antarctic brine pools. The pH at which enrichment cultures are established may influence the predominance of one microbial group over another: for example, the procedures described herein will invariably lead to the selection of bacteria over fungi. Whether this is due exclusively to a pH effect or the greater metabolic diversity of bacteria is not clear, although the latter would appear to be the case. No eucaryotic organism capable of using an aromatic hydrocarbon as a growth substrate has ever been reported since noted by Gibson and Subramanian (1984). A major limitation of the enrichment culture is that it will work only for those organisms that are prototrophic: i.e., those that are able to synthesize all amino acids, nucleotides, vitamins, and cofactors from the inorganic N, P, and S sources and carbonaceous substrate. A defined medium generally selects against fungi and gram-positive bacteria, which are more fastidious. These limitations can be overcome to a small degree by including trace quantities of yeast extract (YE) or some other vitamin and amino acid source. Growth supplements, however, should not be used in quantities large enough to support growth as a C source. Generally, a YE concentration of 50 mg L -1 will support a population density of pseudomonad bacteria of 108 cells mL -1, which will effect faint turbidity. Thus, a concentration of 25 mg L -1 of YE is acceptable as a cofactor supplement. High concentrations of 500 mg L -1 YE have led to erroneous conclusions being made about the utilization of chlorinated aromatic hydrocarbons as "sole carbon sources" (Vandenbergh et aI., 1981). Use of high concentration of YE in the enrichment process merely selects for organisms that are tolerant to the xenobiotic chemical.

FOCHT

410

20-1.2 Materials

1. Platform shaker. 2. Erlenmeyer flasks (125 or 250 mL), screw-cap needed for volatile substrates. 3. Pipets. 4. Eppendorf tubes, 1 mL. 5. Wire. 6. Aluminum foil or cotton. 7. Mineral salts medium (below). Stock solutions K2HP04, 1 M NaH 2 P0 4, 1 M (NH4)2S04' 1 M MgS0 4 ,lM Ca(N03)z' 1 M Fe(N0 3h, 1 M Trace minerals (below) MnS04 ZnS0 4 CuS0 4 NiS0 4 CoS0 4 Na 2Mo0 4

Additions, mL

10 3

10

1

0.1 0.01 1

Final concentration, mM

10 3 10

1 0.1 0.01

0.001 0.001 0.001 0.0001 0.0001 0.0001

Final pH = 7.25. The second column represents the amount of each stock solution to be added to a final volume of 1 L, and the third column represents the final concentration. Add about 0.9 L of distilled water before adding any of the solutions above, or precipitates will form, and then fill to volume. Stock solutions are given without water of hydration to make it easier to use any hydrated form that is available in the laboratory. The trace mineral solution is made up with all the compounds listed. The addition of 1.0 mL to the media above will give a final concentration of each compound in column 3. For multichlorinated compounds or high concentrations of chlorinated compounds, it may be necessary to double the concentrations of K2HP0 4 and NaH2P0 4 to buffer the acidity from the production of HeI. Do not be concerned about the MgHPO 4 precipitate that is formed with doublestrength phosphate additions after autoclaving as it will disappear upon cooling. This salt is unusual in having a lower solubility in hot water than in cold water. 20-1.3 Procedure

There is generally no need for aseptic technique, sterile media, or sterile glassware during the enrichment process unless the source of the

PROCEDURES FOR BIODEGRADATION RESEARCH

411

+-rHREADED NECK WITH CAP

250ml ERLENMEYER FLASK

Fig. 20-1. Nephelometry flask used for growing pure cultures and measuring optical density without removing sample. Volatile substrates such as benzene are placed in a 1.0 ml Eppendorf tube that is suspended above with a wire fastened over the neck of the flask. If the Eppendorf tube is positioned in a manner that might cause spillage of solvent when reading the optical density of the tube, the spectrophotometer should be raised at the appropriate end to prevent this occurrence . Drawn to 0.4 scale.

inoculum is important. Aerial dispersal of xenobiotic-degrading bacteria is not likely to be as important in laboratories not previously engaged in biodegradation research, as in those where many isolates have been obtained. An example of the latter point was the isolation of a 2,6-dinitrophenol-degrading bacterium, Alcaligenes eutrophus JMP 134 (Ecker et aI., 1992), which probably originated from contaminated glassware according to H.-J. Knackmuss (personal communication). The original isolate, obtained by enrichment on 2,4-dichlorophenoxyacetic acid (Pemberton et al., 1979), was found to grow on 2,6-dinitrophenol, and was indistinguishable from the one isolated many years later. Thus, the use of sterile glassware is critical when evaluating the efficacy of inoculation on biodegradation in soil that may lack an indigenous population able to metabolize the xenobiotic chemical. Ample precedences for lack of biodegradation in soil, in lieu of inoculation, exist with 2,4,5-trichlorophenoxyacetic acid (Chatterjee et aI., 1982), pentachlorophenol (Edgehill & Finn, 1983; Crawford & Mohn, 1985), 3-chlorobenzoate (Pertsova et aI., 1984; Focht & Shelton, 1987) and parathion (Barles et aI., 1979). Use of nonsterile glassware could, therefore, give false positive results from the uninoculated control. Add 100 mL of mineral salts solution to a 250-mL Erlenmeyer flask and add the substrate. A concentration range of 100 to 500 mg L -1 is sufficient for nonvolatile substrates. Toxic substrates such as phenol should not exceed 300 mg L -1. Volatile compounds such as toluene and benzene should be added to the Eppendorf tube suspended at the top of the flask (Fig. 20-1). Less volatile compounds such as naphthalene, biphenyl, and diphenylmethane can be added directly to the medium or to the Eppendorf tube. Acidic substrates (e.g., benzoic acids) should be neutralized with an

FOCHT

412

equivalent amount of NaOH added to the media, or preferably made up previously as aqueous stock solutions of the Na or K neutralized salts since some of the acids dissolve slowly. The source and amount of inoculum is a matter of choice. Generally a 11100 mass basis is a good working approximation to minimize soil from obscuring growth, as noted by an increase in turbidity. All flasks can be covered with aluminum foil or cotton, except those containing volatile substrates, and placed on a platform shaker at a temperature not exceeding 30°C. (The commonly used temperature of 37 °C in clinical microbiology is too high for the growth of most gramnegative soil bacteria.) When growth is apparent, transfer 1 mL of the enrichment culture to fresh media containing the same composition as above and incubate. When this subculture shows evidence of growth by an increase in turbidity, it is time to isolate pure cultures by aseptic technique.

20-2 ISOLATION OF PURE CULTURES 20-2.1 Principles Questions may arise as to the necessity of making subcultures rather than directly plating or streaking out samples onto agar media containing the sole C source. If there is an abundant population of microorganisms present in the sample, then it is not necessary to enrich, and pure cultures can be isolated directly. However, in many cases, the numbers of organisms present may be too low for a direct isolation, and consequently they must be enriched to permit the population to increase to a density sufficient for isolation. Also, the presence of exogenous substrates in the sample may effect growth on agar plates without any metabolism of the desired substrates. Finally, for those very odd, yet common, cases requiring 7 to 13 mo for successful enrichment and isolation of chlorobenzene-utilizers (Reineke & Knackmuss, 1984; DeBont et aI., 1986; Shraa et aI., 1986; Spain & Nishino, 1987; Haigler et aI., 1988) or dichlorobenzoate-utilizers (Hernandez et aI., 1991), a long adaptive phase appears to be a necessity. Whether this is due to physiological adaptation, de novo enzyme synthesis, enzyme modification, mutation, or genetic exchange is unknown. Generally, a compound is readily biodegradable if an enrichment culture is obtained quickly and with little effort. This is certainly predictable with simple aromatic acids and hydrocarbons. However, failure to obtain enrichment cultures could either be indicative of recalcitrance (e.g., CAHs) or action by a consortium (e.g., cyclohexane). On occasions, utilization of a C source in liquid media after several transfers may be apparent, yet selection and growth of isolated colonies on agar plates cannot be demonstrated. In all such cases, these microbial communities involve a syntrophic or cross-feeding effect by which the interaction is obligatory and mutually symbiotic to both.

PROCEDURES FOR BIODEGRADATION RESEARCH

413

20-2.2 Materials 1. 2. 3. 4. 5. 6.

Mineral salts agar (1.5%) for petri plates. Mineral salts agar (1.5%) for slants. Desiccator or sealed container. Incubator set at 28°C. Inoculating loop. MS media (50 mL) containing desired substrate in 250 mL Erlenmeyer flasks, sterile. 20-2.3 Procedures

Nonvolatile substrates should be added to the agar before autoclaving. Do not attempt to autoclave volatile substrates! When it is apparent that the substrate is supporting bacterial growth, remove a sample with an inoculating loop and streak the suspension according to standard microbiological procedures onto sterile mineral salts agar (1.5%) plates containing the appropriate substrate. For volatile substrates, use mineral salts agar plates containing no substrate. After streaking these plates, there are two methods that can be used for ensuring aseptic contact of the volatile substrate with the culture. Petri plates are inverted and placed on the raised floor of a desiccator with the substrate placed in a small beaker or petri plate lid underneath the raised floor at the bottom of the desiccator. The other method is to add one drop of the liquid or a few crystals of the solid (as the case may be) to the bottom inverted lid of the petri plate, place the agar lid above, and seal the two lids together with parafilm or some other temporary but effective sealer. It is advisable to use glass, and not plastic, petri plates when isolating microorganisms on volatile organic compounds. The plates should be incubated at 28°C or ambient temperature for several days before the presence of single isolated colonies is noted. It is not unusual to wait 7 to 10 d before colonies of sufficient size are noted with some substrates. Generally, hydrocarbon substrates select for faster (2-3 d) and more luxuriant growth than chlorinated aromatic acids. The final proof that the compound is used for growth as a sole C source is to aseptically transfer a single isolated colony to sterile liquid media, incubate, and observe growth by the appearance of turbidity.

20-3 MAINTENANCE OF CULTURES 20-3.1 Principles Xenobiotic-degrading bacteria frequently lose their degradative ability by spontaneous loss of plasmids, which code for many of the degradative enzymes. The influence of substrate on genetic rearrangement of a TOL-like plasmid in Pseudomonas putida has been recently documented

414

FOCHT

(Carney & Leary, 1989). The Darwinian dogma regarding the environment solely as a selective agent that is independent of influencing mutational events has recently come under question (Hall, 1990; Cairns et aI., 1988). Whether or not the spontaneous loss of degradative plasmids or their rearrangement in response to substrates is suggestive of neo-Lamarckianism is not likely to be resolved before the role of environment and mutation is settled. It is important for microbiologists to recognize that transpositional changes between plasmid and chromosomal DNA occur frequently with xenobiotic-degrading bacteria, which necessitates that the selection pressure be maintained to eliminate undesirable mutants. Consequently, continuous growth on complex media is discouraged-particularly when it is apparent that reversion from the desired phenotype is high. On the other hand, there are instances in which bacteria are "refreshed" by occasional transfer to rich media. Sylvestre and Fauteux (1982) noted that a facultative anaerobe, strain B206, lost the ability to grow on 4-chlorobiphenyl or biphenyl after a series of transfers: When propagated on nutrient agar, it regained the ability to grow again on either biphenyl or 4-chlorobipheny1. Complex media may be useful to "refresh" problematic cultures, to check for purity by observing the presence of more than one colony type, and for quickly growing up large batches of cells, but they should be used judiciously. 20-3.2 Materials The same materials used in the previous section. 20-3.3 Procedure The previous section described the growth of bacteria on agar plates with volatile substrates. The same basic principle applies to maintaining stock cultures of bacteria on agar slants. A closed container (e.g., desiccator containing the substrate) can be used for incubation of slants. However, once the slants are removed, they should be sealed and placed in a refrigerator to retard metabolism and conserve the small amount of substrate remaining. Alternative substrates for benzene, toluene, ethylbenze and xylene (BTEX) are toluates (methyl benzoates), which are suitable for maintaining the selection pressure for BTEX utilization (Carney & Leary, 1989). Cultures using less volatile substrates such as biphenyl or naphthalene can be maintained on slants as described above or on slants containing the substrate, which is added immediately after the tubes have been removed from the autoclave. This can be done most rapidly by the addition of 100 !JL of an acetone solution containing 70 g L -1 of material prior to slanting the tubes in a hood and allowing them to cool with the cap fastened loosely to permit the loss of solvent. As this procedure violates prima facie the concept of sterilization, it is necessary to frequently ascertain purity by streaking onto plates as described above. Slants, nevertheless, are more

PROCEDURES FOR BIODEGRADATION RESEARCH

415

manageable than plates for sending requests of cultures and maintaining isolates over several weeks. Stock cultures are best maintained pure by removing a loopful of culture from an isolated colony on a plate and mixing it in 0.1 mL of sterile phosphate buffer (see section 20-6.2), which is then added to a sterile vial containing 30 to 50% glycerol in water, and stored at -80°C. 20-4 GROWTH IN LIQUID CULTURES 20-4.1 Principles

In most cases, strains that are isolated on solid media will grow in liquid media. Occasionally, however, there will be strains that grow poorly in liquid media. Solid insoluble substrates, such as biphenyl, generally give low cell densities when the substrate is added to the medium prior to autoclaving, presumably because of the small surface area that is caused when the material cools into a solid lump. (As stated before, autoclaving of these substrates is discouraged in lieu of proper ventilation designs for public health reasons.) The addition of substrate by a volatile solvent sometimes facilitates this problem. However, miscible solvents such as acetone form azeotropic mixtures, such that residual solvent may be toxic to some strains. The best growth with biphenyl is achieved by powdering the substrate with a mortar and pestle and adding it to sterilized media immediately before inoculation. This nonaseptic microbiological procedure can be justified if a sufficient inoculum density is used to give a starting concentration in excess of 107 cells mL -1 on kinetic grounds as it would be difficult to argue that the introduction of an air-borne contaminant would have a density sufficient to outgrow the inoculant. The same argument would apply regarding the likelihood that contaminants could be introduced from the nonsterile substrate. As a matter of satisfaction, an uninoculated control could be easily run to verify the source of growth. However, it cannot be stressed too strongly that this nonaseptic method should never be used for continual subculturing or maintenance of stock cultures. Once the experiment has been completed, the cells should be discarded in accordance with standard microbiological procedures. To ensure purity, stock cultures should always be maintained axenically on sterilized slants and verified on plates. Although utilization of a growth substrate may seem rather obvious with fast-growing strains, the same cannot be said with substrates that are toxic at moderate concentrations and must be added in small quantities. Pentachlorophenol is such a case in point since it is used specifically to inhibit microbial growth. Moreover, the maximum concentration (300 mg L -1) that can be used for growth of Flavobacterium spp. (Crawford & Mohn, 1985) does not provide much C (27% by mass) or energy, and the organism must be grown with an auxiliary C source. Thus, growth curves are required to conclusively demonstrate that an organism uses the target

FOCHT

416

compound in the presence of an auxiliary growth substrate, as opposed to merely being tolerant to it. With chlorinated substrates, production of inorganic chloride would further confirm that the substrate was mineralized at the expense of growth, particularly if the release was close to complete stoichiometry of what was added. It would not be possible to demonstrate this on solid media. This section is not intended to provide a detailed kinetic examination of growth, but rather to provide a general understanding of how fast bacteria grow on xenobiotic substrates and their efficacy of utilization. 20-4.2 Materials 1. Shaking platform. 2. Erlenmeyer side-arm flasks (250 mL; Fig. 20-1) containing 30 mL media. 3. Spectrophotometer for measuring cell density turbidimetrically (525 nm). 4. Centrifuge. 5. Media for growing culture. 6. Culture.

20-4.3 Procedure

To obtain the most reproducible results, and to reduce the length of the lag phase from inoculation to the first observed increase in turbidity, it is best to use a fresh culture that has previously been growing in the same liquid media that is under investigation. The density of the inoculum should be about 109 cells mL-l, and the flasks should be inoculated with an amount sufficient to give slight turbidity: the absorbance or optical density (OD) should be about 0.02 to 0.04 with an 18-mm thick test tube at 525 nm at the beginning. The use of dilute cell suspensions or inoculations from slants usually result in an increased apparent lag because cell densities below this concentration cannot be detected turbidimetrically. The term apparent lag is used here to distinguish from a real biological lag phase, which constitutes an actual failure of cells to immediately divide and begin the exponential growth phase. The time at which measurements are taken will depend on how fast the culture grows. Cultures that grow fast (3 h or less doubling time) can be completed in a day, while those that grow slowly (12 h doubling time or more) will take several days, although they can be measured conveniently twice a day. Those strains in between will require more dedication and sleepless nights. The use of nephthelometry flasks (Fig. 20-1) obviates the risk of contamination that could occur by repeated sampling; it also maintains a constant volume. To minimize light interference, the flask should be

PROCEDURES FOR BIODEGRADATION RESEARCH

417

covered completely with a thick dark plastic cover that does not let light pass through. Portfolios that are distributed at scientific meetings, work well for this purpose. The two important kinetic parameters that can be determined from growth curves are the doubling time (tD ) and the yield coefficient (Y). The former can be determined from a semi-log plot of the optical density vs. time from a straight line of those points representing the exponential (or log) phase of growth. The latter can be approximated by the difference of the maximal turbidity (stationary growth phase) and the initial turbidity. The method for the latter is applicable only if all the substrate has gone into cell growth and not into maintenance metabolism. Thus, high substrate concentrations in which sufficient substrate remains after cells have reached stationary growth are unsuitable for determining yield coefficients. These methods are intended as approximations only. For more detailed and exacting analysis of the kinetics of growth, the reader is referred elsewhere (Simkins & Alexander, 1984; Focht & Shelton, 1987). Absorbance measurements are more useful when related to cell mass. This relationship is established by growing cells in about 1.0 L of media to obtain sufficient mass of cells for weighing (100-500 mg). A good general rule of thumb for pseudomonad bacteria is the following relationship: 1010 cells mL -1 = 1 mg dry mass mL -1 = OD 4.0. Cell densities this high are rarely observed with batch cultures except with rich media under conditions of high aeration. Cells should be harvested from the late exponential growth phase to avoid the production of lytic products and slimes that may add to the weight. It is desirable to use a refrigerated centrifuge so that autolysis will be minimized. Harvest cells by centrifugation at 6000 g for 10 min. Discard the supernatant, and wash the cells by resuspending them in about 300 mL of deionized water or about the amount that a single centrifuge bucket will hold. Centrifuge, wash, and resuspend two more times to remove the salts from the medium, which would otherwise add to the weight. Resuspend the last (third) cell pellet into duplicate volumes of 50 mL and mix thoroughly. Remove 1 mL from each sample, dilute with 9 mL of water, mix thoroughly and record the absorbance. Add the duplicate cell suspensions to two tared breakers, and wash the remaining cells into the beaker with deionized water. Evaporate the water at 90°C or below. Avoid boiling temperatures since this will cause bumping and loss from boil over and oxidation of organic mater. When the samples are dry, weigh the beakers and take the average value. For better precision, more samples can be taken, but it is generally unnecessary to make a standard curve over several concentration ranges. Yield coefficients are usually expressed as g cells g-l substrate or g cells mol- 1 substrate. When it is desirable to do a total C balance, the total C content of the cell mass can be determined by methods mentioned elsewhere in this book. A working approximation is that 55% of the total dry mass of the cell is comprised of C.

FOCHT

418

20-5 PREPARATION OF WASHED CELL SUSPENSIONS 20-5.1 Principles Xenobiotic substrates that are cometabolized can be studied in the presence of the growth substrate during batch culture growth. However, this method is problematical when the growth substrate and xenobiotic compound have properties that do not distinguish them by common analytical methods. Moreover, comparisons among metabolism of several xenobiotic compounds would require a separate culture. A culture study could involve several days and a degree of uncertainty regarding sampling time and frequency. The preparation of washed, resting cells eliminates all of the problems mentioned above. Moreover, it enables the investigator to choose what substrate the organisms should be grown on and to make comparisons between them. It is particularly advantageous to know if the metabolism of a xenobiotic compound is constitutive or inducible since this would have relevance to the performance of inoculants in soil. Thus, comparisons can be made in a single experiment, not only between xenobiotic compounds, but among those cells grown on different substrates, e.g., glucose and chlorobenzoates. The use of washed cells, which are unable to grow because of N limitations, enables the investigator to concentrate the cell density, which thereby enhances the rate of metabolism and saves time. Finally, the preparation of washed cells is a prelude to the preparation of cell-free extracts for use in enzymatic studies. 20-5.2 Materials 1. 2 L flask containing 1 L of growth medium. (Depending on the quantity of cells needed, smaller flasks and less media could be used.) 2. Platform shaker. 3. Pipets. 4. Centrifuge. 5. Phosphate buffer (0.05 M, either Na or K or both, pH 7.2). 6. Spectrophotometer.

20-5.3 Procedure The substrate concentration of the medium should not be limiting; i.e., it should equal or exceed that necessary to achieve maximal turbidity or cell density. Cells should be harvested from the late exponential growth phase. To know all these parameters generally requires that growth curves be performed previously as in section 4. Prior to harvesting, remove an aliquot for spectrophotometric analysis of the cell density. Follow the same procedure in section 4 for harvesting cells except for using phosphate buffer

PROCEDURES FOR BIODEGRADATION RESEARCH

419

instead of deionized water for washing. After the third and final centrifugation, suspend the cells in buffer to 25 mL. Make a 11100 dilution of the suspension and measure the 00. Adjust the final 00 to 4.0 with phosphate buffer according to the formula below, where Vb is the volume of buffer to be added, Vs is the volume of the cell suspension (24.9 mL), ODs is the 00 of the cell suspension (let us use 10 as an example), and 00 4 .0 is the desired 00 (4.0).

Thus, Vb = 37.35 mL in the example above to give a total washed cell volume of 62.25 mL at 00 4.0. There is nothing fixed in stone about a washed cell suspension being 1 mg dry mass mL -1. Higher or lower cell densities can be used. Many pseudomonad bacteria tend to undergo autolysis at higher cell densities. This is particularly more pronounced for experiments that may be conducted for several hours. The use of lower cell densities simply means that more time may be needed to complete the experiment. The cell suspension can then be used in any reaction mixture to assess O 2 consumption, disappearance of substrate, or analysis of metabolites. Oxygen consumption measurements are generally completed in a few minutes (see section 7), while the other experiments may be conducted over a few hours. Contamination or demise of the culture is generally not a problem over a 24-h period. If the cell suspension is not going to be used right away, it can be kept refrigerated or on ice for several hours prior to its use.

20-6 PREPARATION AND USE OF CELL-FREE EXTRACTS 20-6.1 Principles Although the use of washed cell suspensions can give valuable information about the rates of substrate disappearance and qualitative identification of metabolites, it is sometimes desirable to obtain information about an important, single reaction-step in the catabolic pathway. A few pertinent examples are the dehalogenation of 4-chlorobenzoate to 4-hydroxybenzoate (Elsner et aI., 1991), the hydrolysis of parathion to p-nitrophenol (Munnecke, 1979), the dehalogenation of halo me thanes by methane monooxygenase (Tsien & Hanson, 1992), and the dioxygenation of naphthalene (Haigler & Gibson, 1990). The best characterized enzymes in the catabolism of aromatic compounds are the catechol dioxygenases, which incorporate both atoms of O 2 into the splitting of the aromatic ring. ortho-Pyrocatechase (EC 1.13.11.1), splits catechol between the two hydroxyl groups to give cis, cis-muconate, which has Amax = 260 nm. meta-Pyrocatechase (EC 1.13.11.2) splits the ring adjacent to one of the hydroxyl groups to give muconic semialdehyde, a bright yellow-product that has Amax = 375 nm at neutral to alkaline

FOCHT

420

pH: the color is reversibly abolished by addition of acid. Bacteria normally possess constitutive levels of the ortho fission enzyme, presumably as a result of tryptophan catabolism. Both catechol dioxygenases can be distinguished from each other by measuring product disappearance at the two different wave lengths or by inactivating ortho activity at 55°C for 15 min and inactivating meta activity with a 3% final solution of H 20 2 • The analysis of the two catechol dioxygenases presented below are given as an illustration for the preparation and use of cell-free extracts. 20-6.2 Materials 1. 2. 3. 4.

Same as for growing cells. Sonic probe or French pressurized cell. UV-visible spectrophotometer. Methanolic or N,N-Dimethyl formamide solutions of catechol (10 mg mL -1). 20-6.3 Procedure

Follow the same procedure above for growing up cells except that the final suspension should be more concentrated in about 10 mL buffer since this will facilitate breakage. Cool the cell suspension in an ice bath, and disrupt the cells by sonic bursts of 15 s duration followed by 15 s of swirling the tube in the ice bath. About 8 to 12 bursts will be necessary to get good breakage for gram-negative bacteria. When the cells have been broken, as indicated by a peculiar burnt-like odor and a glue-like appearance, clarify the extracts by centrifugation at 4 °C at 40 000 g for 30 min. Remove a small aliquot, and determine the protein concentration by one of the standard methods commonly used (Colowick & Kaplan, 1957; Bradford, 1976). The protein concentration should be between 5 to 15 mg mL -1. Since gram-positive bacteria are more resistant to mechanical breakage, the addition of an equal volume of glass beads ( < 10 !lm) is recommended when using sonication. Alternatively, the cell suspension can be passed through a French pressurized cell at 25 MPa (20000 lb in. -2). Cell-free extracts can be purified for isolation of pure enzymes or they can be used for crude enzyme assays as is common in the determination of catechol dioxygenase activity. The measurement of this activity can be done respirometrically, as described in the following section, or spectrophotometrically, as described herein. Catechol dioxygenase activity is an extremely rapid reaction, which begins immediately upon addition of substrate. Thus, everything should be in place before substrate and cell-free extract are added. The procedures that follow are based on a curvette volume of 1.0 mL. All additions should be changed accordingly to the volume of the curvette. Dilute the CFE with 0.067 M phosphate buffer to give a concentration of 0.1 mg protein mL -1. Add 1.0 mL of the diluted cell-free extract to the

PROCEDURES FOR BIODEGRADATION RESEARCH

421

curvette and allow 1 to 2 min for temperature equilibration. When the temperature is constant, add 5 !lL of the catechol solution, plug the end of the curvette, invert it rapidly to insure mixing, and place it immediately into the spectrophotometer, which is hooked up to a chart recorder, or some other appropriate signal device. Metapyrocatechase activity is measured at 375 nm, while orthopyrocatechase activity is measured at 260 nm. Product formation will stop in a few minutes as noted by a plateau. This position on the chart recorder will then represent the complete stoichiometric conversion to product. Thus, the specific activity can be determined from the rate (slope of product formation) per unit of protein added. Occasionally, stoichiometry of the meta ring fission product may be < 100%. Instead of a plateau, there will be a peak due to hydrolysis of the ring fission product. Should this happen, it will be necessary to preincubate the CFE at 55°C for 15 min, which will inactivate hydrolase activity, but not metapyrocatechase activity (Dagley et aI., 1960). 20-7 OXYGEN CONSUMPI'ION 20-7.1 Principles Respirometry studies are rapid, provide information about a catabolic pathway, and compare relative transformation rates among different xenobiotic compounds. In the case of CFE, the stoichiometry of O 2 for an oxygenase-catalyzed reaction can be confirmed precisely since there is virtually no respiratory activity because of the absence of the particulatelinked respiratory cytochrome system. Respirometry with washed cells provides rejection or presumptory acceptance of what the intermediate products might be. However, the failure to oxidize a presumptive intermediate may also be caused by its inability to be transported into the cell. A classic example of a cell being unable to induce for a transport protein is the inability of E. coli to metabolize and use citrate, even though it has all the components of an active TCA cycle. Many years ago, Warburg and Gilson manometers were commonly used in physiological work. These instruments are rarely, if ever used anymore in biodegradation research. A more rapid and less cumbersome method is the measurement of O 2 uptake in solution by an O 2 probe. Depending on the extent of information needed, each analysis takes only a few minutes, in contrast to hours by manometry. There is also little cleaning that accompanies the end of a run. 20-7.2 Materials 1. 2. 3. 4.

Oxygen meter with probe. Recirculating water bath with tubing. Chart recorder or other event marker. Magnetic stirring platform.

FOCHT

422 REMOVABLE GROUND . . - GLASS PLUG WITH Jir HOLLOW CENTER

TO

WATER~

BATH

........"..;--. FROM +-WATER BATH

MAGNETIC STIRRER

Fig. 20-2. Apparatus used for measuring consumption of dissolved 02' Outer jacket maintains constant temperature of cell (inside) by a recirculating water bath. Drawn to half scale.

5. 6. 7. 8.

Magnetic stirring bar (3 by 12 mm). Incubation cell and assembly (Fig. 20-2). Cell suspension or cell-free extract. Substrate concentrations (10-100 mg mL -1 in methanol or dimethyl formamide). 9. Vacuum flask and sipper. 20-7.3 Procedure

The example given below is for a system having a 2.0-mL reaction chamber. Amounts of materials can be adjusted accordingly. Higher concentrations of substrate are more suitable for washed cell suspensions where comparisons of rates are desirable. If the total stoichiometry with relation to O 2 uptake is desired, the concentration of the substrate should not exceed the oxygen demand available. For example, 1 !-lmol toluene (92 !-lg) would require 9 !-lmol O 2 for complete combustion to CO2 and H 20, yet only 0.5 !-lmol O 2 would be available in the entire 2.0 mL of the cell because of its low solubility (0.25 mM at 25 0q. Cell suspensions or cell-free extracts, particularly metapyrocatechase, may lose activity with time if they are not kept on ice. However, they must first be equilibrated to the temperature of the water bath (25-30 0q prior to adding the substrate. Since this may require several minutes between samples, it is a good idea to incubate a small batch of cells or cell-free

PROCEDURES FOR BIODEGRADATION RESEARCH

423

extracts in the water bath to accommodate material for the next two runs. The first run should be with cells alone, to determine the base rate of O 2 consumption. With washed cells, this will represent primarily endogenous respiration. In the case of cell-free extracts, consumption of O 2 will be caused primarily by the electrode. Thus, it is important to take several baseline samples interspersed between the experimental samples, to obtain an accurate control level of consumption that will be subtracted from the actual experimental values. When the system is ready to go, add 10 flL of the solution substrate to the reaction chamber and record the downscale pen drift as O 2 is consumed. This rate when subtracted from the basal rate will represent the activity, which can be expressed in specific activity units per mass of cells or mass of protein. When a run is finished, the material is removed with a sip per and the reaction chamber is rinsed several times with water. The sipper can be made from glass or stainless steel tubing that is connected to flexible tubing in line with a vacuum flask and vacuum line. 20-8 CHLORIDE DETERMINATION 20-8.1 Principles Because chlorinated organic compounds represent the greatest contribution to soil and water pollution, there is considerable interest in the nature of halogen removal and in their catabolic pathways. There is a strong current interest in anaerobic de halogenations as well (see chapter 13 in this book). Generation of inorganic chloride, moreover, is strong evidence for the metabolism to the compound in question. The following method is designed for pure cultures that grow in a chloride-free medium. The method is not intended for use in soil where other anions (e .g., nitrate) and ligands interfere. The method is based on a simple straightforward precipitation of inorganic chloride with AgN03 under acidic conditions that do not cause formation of insoluble phosphate or sulfate salts. The method has its drawbacks when insoluble precipitates, particularly dichlorobenzoic acids, are formed upon the addition of phosphoric acid. In some cases, the precipitate can be removed by centrifugation, while in others, it floats to the top, which necessitates removal by extraction with ethyl acetate or other suitable solvent prior to addition of silver nitrate. 20-8.2 Materials 1. 2. 3. 4. 5. 6.

10 mM NaCl, in 100 mL of mineral salts solution. 100 mL of a mineral salts solution. Concentrated (15 M) H 3P0 4 . 1.0 M AgN0 3. Test tubes. Spectrophotometer.

FOCHT

424

20-8.3 Procedure

Prepare at least five standard concentrations to give a range of chloride from 0.2 to 3.2 mM by using the appropriate volumes of NaCl solution (0.2-3.2 mL) and the corresponding volume of mineral salts solution without NaCl (9.8-6.8 mL) to give the same volume of 10 mL in each tube. When the standards have been prepared, add 0.06 mL concentrated H 3P04 to all the tubes and mix thoroughly. Add 1O!lL of AgN0 3 to one standard, mix thoroughly and measure immediately at 525 nm. Make a standard curve or calculate the regression equation. Use this for determining chloride in the culture. From culture supernatants, add 10 mL to the spectrophotometer tube followed by 0.06 mL of concentrated H 3P0 4 . If a precipitate forms, remove it by centrifugation or extraction as mentioned above. Add 10 !lL AgN0 3 solution, and measure under the same conditions given above.

20-9 CONCLUSION

The real challenge for microbiologists is to identify and isolate the members of catabolic consortia that together, but not singly, can effect total destruction of the target molecule. This can only be done by logically envisioning a plausible catabolic pathway to establish separate enrichment cultures using the hypothetical degradation products for isolation of the desired microftora. Only when all members of the catabolic consortia can be identified, will it be possible to devise strategies for bioremediation (i.e., the biologically mediated detoxification of contaminated soil) of recalcitrant compounds. The isolation and identification of all members also makes for an attractive strategy for combining all the genes involved in the complementary catabolic pathway into a single organism. It is no longer relevant to think of biodegradation in terms of single organisms per se but rather in their contribution to the total gene pool that exists scattered in nature for the catabolism of any conceivable organic molecule.

REFERENCES Angle, J. Scott, S.P. McGrath, and R.H. Chaney. 1991. New culture medium containing ionic concentrations of nutrients similar to concentrations found in soil solution. Appl. Environ. Microbiol. 57:3674-3676. Alexander, M. 1965. Biodegradation: Problems of molecular recalcitrance and microbial fallability. Adv. Appl. Microbiol. 7:35-76. Beam, H.W., and J.J. Perry. 1973. Co-metabolism as a factor in microbial degradation of cycloparaffinic hydrocarbons. Arch. Microbiol. 91:87-90. Beam, H.W., and J.J. Perry. 1974. Microbial degradation of cycloparaffinic hydrocarbons via co-metabolism and commensalism. Arch. Microbiol. 91:87-90. Bradford, M.M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principles of protein dye-binding. Anal. Biochem. 72:248-254.

PROCEDURES FOR BIODEGRADATION RESEARCH

425

Brunner, W., F.H. Sutherland, and D.D. Focht. 1985. Enhanced biodegradation of polychlorinated biphenyls in soil by analog enrichment. J. Environ. Qual. 14:324-328. Cairns, J., J. Overbaugh, and S. Miller. 1988. The origin of mutants. Nature (London) 335:142-145. Carney, B.F., and J.v. Leary. 1989. Novel alterations in plasmid DNA associated with aromatic hydrocarbon utilization by Pseudomonas putida strain R5-3. Appl. Environ. Microbiol. 55: 1523-1530. Chatterjee, D.K., J.J. Kilbane, and A.M. Chakrabarty. 1982. Biodegradation of 2,4,5trichlorophenoxyacetic acid in soil by a pure culture of Pseudomonas cepacia. Appl. Environ. Microbiol. 44:514-516. Colowick, S.P., and N.O. Kaplan. 1957. Methods in enzymology III. Academic Press, New York. Crawford, R.L., and W.W. Mohn. 1985. Microbiological removal of pentachlorophenol from soil using a Flavobacterium. Enzyme Microb. Technol. 7:617-620. Dagley, S. 1984. Introduction. 1-11. In D.T. Gibson (ed.) Microbial degradation of organic compounds. Marce Dekker, New York. Dagley, S., W.C. Evans, and D.W. Ribbons. 1960. New pathways in the oxidative metabolism of aromatic compounds by micro-organisms. Nature (London) 188:560-566. DeBont, J.A.M., M.J.A.W. Vorage, S. Hartmans, and W.J.J. van den Tweel. 1986. Microbial degradation of 1,3-dichlorobenzene. Appl. Environ. Microbiol. 52:677-680. deKlerk, H., and A.e. van der Linden. 1974. Bacterial degradation of cyclohexane: Participation of a co-oxidation reaction. Antonie van Leeeuwenhoek 40:7-15. Ecker, S., T. Widmann, H. Lenke, O. Dickel, P. Fischer, e. Bruhn, and H.-J. Knackmuss. 1992. Catabolism of 2,6-dinitrophenol by Alcaligenes eutrophus JMP 134 and JMP 222. Arch. Microbiol. 158: 149-154. Edgehill, R. U., and R.K. Finn. 1983. Microbial treatment of soil to remove pentachlorophenol. Appl. Environ. Microbiol. 45:1122-1125. Elsner, A., F. Loffier, K. Miyashita, R. Miiller, and F. Lingens. 1991. Resolution of 4-chlorobenzoate dehalogenase from Pseudomonas sp. strain CBS3 into 3 components. Appl. Environ. Microbiol. 57:324-326. Focht, D.D., and W. Brunner. 1985. Kinetics of biphenyl and polychlorinated biphenyl metabolism in soil. Appl. Environ. Microbiol. 50:1058-1063. Focht, D.D., and D. Shelton. 1987. Growth kinetics of Pseudomonas alcaligenes C-O relative to inoculation and 3-chlorobenzoate metabolism in soil. Appl. Environ. Microbiol. 53: 1846-1849. Gale, E.F. 1952. The chemical activities of bacteria. Academic Press, London. Gibson, D.T., and V. Subramanian. 1984. Microbial degradation of aromatic hydrocarbons. p. 181-252. In D.T. Gibson (ed.) Microbial degradation of organic compounds. Marcel Dekker, New York. Haigler, B.E., and D.T. Gibson. 1990. Purification and properties of NADH-ferrodoxinNAP Reductase, a component of naphthalene dioxygenase from Pseudomonas sp. strain NCIB 9816. J. Bacteriol. 172:457-464. Haigler, B.E., S.F. Nishino, and J.e. Spain. 1988. Degradation of 1,2-dichlorobenzene by a Pseudomonas sp. Appl. Environ. Microbiol. 54:294-301. Hall, B.G. 1990. Spontaneous point mutations that occur more often when advantageous than when neutral. Genetics 126:5-16. Hernandez, B.S., F.K. Higson, R. Kondrat, and D.D. Focht. 1991. Metabolism of and inhibition by chlorobenzoates in Pseudomonas putida PIll. Appl. Environ. Microbiol. 57:3361-3366. Horvath, R.S. 1972. Microbial cometabolism and the degradation of organic compounds in nature. Bacteriol. Rev. 36: 146-155. Munnecke, D.M. 1979. Hydrolysis of organophosphate insecticides by an immobilized-enzyme system. Biotechnol. Bioeng. 21:2247. Pemberton, J.M., B. Corney, and R.H. Don. 1979. Evolution and spread of pesticide degrading ability among soil micro-organisms. p. 287-299. In K.N. Timmis and P.A. Piihler (ed.), Plasmids of medical, environmental and commercial importance. Elsevier/ North Holland Biomedical Press, Amsterdam. Perry, J.J. 1984. Microbial metabolism of cyclic alkanes. p. 61-97. In R.M. Atlas (ed.) Petroleum microbiology. Macmillan, New York. Pertsova, R.N., F. Kunc, and L.A. Golavleta. 1984. Degradation of 3-chlorobenzoate in soil by pseudomonads carrying biodegradative plasmids. Foila Microbiol. 29:242-247.

r'

426

FOCHT

Reineke, W., and H.-J. Knackmuss. 1984. Microbial metabolism of haloaromatics: Isolation an~ properties of a chlorobenzene-degrading bacterium. Appl. Environ. Microbiol. 47.395-402. Shraa, G., M.L. Boone, M.S.M. Jetten, A.R.W. van Neerven, P.J. Colberg, and A.J.B. Zehnder. 1986. Degradation of 1,4-dichlorobenzene by Alcaligenes sp. strain A175. Appl. Environ. Microbiol. 52:1374-1381. Simkins, S., and M. Alexander. 1984. Models for mineralization kinetics with the variables of substrate concentration and population density. Appl. Environ. Microbiol. 47:12991306. Sylvestre, M., and J. Fauteux. 1982. A new facultative anaerobe capable of growth on chlorobiphenyls. J. Gen. Appl. Microbiol. 28:61-72. Spain, J.C., and S.F. Nishino. 1987. Degradation of 1,2-dicholorobenzene by a Pseudomonas sp. Appl. Environ. Microbiol. 53:1010-1019. ' Tsien, H.C., and R.S. Hanson. 1992. Soluble methane monooxygenase component-B gene probe for identification of methanotrophs that rapidly degrade trichloroethylene. Appl. Environ. Microbiol. 58:953-960. Vandenbergh, P.A., R.H. Olsen, and J.F. Colaruotolo. 1981. Isolation and genetic characterization of bacteria that degrade chloroaromatic compounds. Appl. Environ. MicrobioI. 42:737-739.

Published 1994

Chapter 21 Algae and Cyanobacteria F. BLAINE METTING, JR., Battelle, Pacific Northwest Laboratories, Battelle Boulevard, Box 999, P7-54, Richland, Washington

Algae are Oz-evolving photosynthetic organisms that contain plant-like chlorophylls but are distinct from higher plants in that every gametangial cell is fertile-sexual structures with differentiated tissues are not produced,1 Thus the term algae encompasses seaweeds (macro algae) as well as microscopic forms (microalgae). Phycology, the study of algae, has evolved historically to include the prokaryotic cyanobacteria (blue-green algae) because of their morphological, physiological, and ecological similarities to eukaryotic micro algae (Bold & Wynne, 1985). This chapter will present methods for studying microalgae and cyanobacteria that inhabit soil and rocks (endolithic habitats). General methods for studying algae in the laboratory and field can be found in a four volume series of monographs sponsored by the Phycological Society of America (Stein, 1973; Hellebust & Craigie, 1978; Gantt, 1980; Littler & Littler, 1985). Microalgae inhabit all terrestrial surfaces. Growth rates and the complexity of community development depend primarily on available light and moisture, but are also influenced by temperature and properties of the substrate, such as pH. Numbers and biomass of microalgae vary greatly among soils, commonly ranging, respectively, between 109 and lO lD colonyforming units (CFU) per mZ (103 -106 per g), and from < 1 to 1500 kg wet weight ha- 1 (Metting, 1993). The largest communities (in terms of biomass) are found in flooded rice (Oryza sativa L.) soils (Roger & Kulasooriya, 1980). Successively smaller mixed populations occur on arid and semiarid soils with minimal plant cover, temperate agricultural soils, and grasslands, forests, and other habitats where plant canopies and litter limit the flux of sunlight at the soil surface. On and in rocks, populations of IThe gametangium is the sexual organ that produces gametes. Individual cells of many single-celled soil microalgae (e.g., Chlamydomonas) act as gametes. In algae with multicellular gametangia, each cell is a potential gamete. Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

427

METTING

428

micro algae form in cracks and fissures (chasmolithic communities) and in the fabric of the rock itself (cryptoendolithic communities). Cryptoendolithic communities are remarkably similar in both hot and cold deserts throughout the world. Biomass estimates for these communities range from about 2 to 200 mg (as chlorophyll a) per mZ (Metting, 1991). Organic matter production by photosynthesis (CO z fixation), Nz fixation, and surface consolidation are significant ecological attributes of the soil algal community. All of these activities are particularly important in semiarid and arid habitats (steppes and deserts) characterized by minimal vascular plant cover. In these situations, microalgae, together with lichens and mosses, make up what are known as cryptogamic crusts. In many situations, these crusts are the major source of biologically fixed N and can be a major factor acting to minimize erosive loss of topsoil by water runoff and wind (Metting, 1991). There is evidence that increased rates of desertification, such as in sub-Saharan Africa, can result from excessive grazing of fragile landscapes that destroys cryptogamic crusts by the trampling of livestock (Knutsen & Metting, 1991). The most important influence that microalgae have in agriculture is biological Nz fixation by cyanobacteria in wetland rice cultivation. Cyanobacterial Nz fixation can provide all of the N requirements in some traditional farming systems (China, India, and Vietnam) and can contribute to the fertility of soils in which high-yielding hybrid rice varieties are grown. Estimates of 100 kg of N fixed ha- I ye l or more have been reported, although values between 10 and 30 kg are more common. Both free-living species and cyanobacteria symbiotic with the water fern Azalia are important, although grazing by arthropods and competition from non Nz-fixing micro algae limits their overall contribution to rice production (Roger et al. , 1993). On pivot-irrigated farm ground in the USA, mass-cultured green micro algae that produce copious quantities of extracellular mucilages (mostly Chlamydomonas spp.) have been used on a small scale as soilconditioning agents to control erosion of sandy soils and improve infiltration of water into heavy soils (Metting et al., 1988). Because of their ecological importance and because they are physiologically similar to plants, soil micro algae and cyanobacteria have also proven to be valuable as ecological indicators for studying non-target pesticide effects (Pipe, 1992). 21-1 IDENTIFICATION OF SOIL ALGAE AND CYANOBACTERIA 21-1.1 Principles Algae and cyanobacteria are readily recognized as distinct from other soil microorganisms by their chlorophyll a content. Using standard light microscopic methods (see chapter 6 by Bottomley in this book), cultured algae or algal cells and filaments in soil samples appear as various shades of green, blue-green, and yellow-green depending on the content of acces-

ALGAE AND CYANOBACTERIA

429

sory pigments that can include various combinations of different chIorophylls, carotenoids, xanthophylls, and phycobilins. 2 Table 21-1 shows the major algal classes that include terrestrial micro algae and lists references to principal taxonomic treatises with descriptions of many of the species. Identification of microscopic algae at the genus level is based on morphological properties. The major distinction among soil algae is between the prokaryotic cyanobacteria and the eukaryotic groups. It is apparent upon microscopic examination that cyanobacterial cells lack an internal organization that is homologous to the system of organelles, particularly chloroplasts and nuclei, that are readily distinguishable in eukaryotic microalgae. The distinctive coloration imparted by blue-green and red phycobilins also set the cyanobacteria apart from nearly all other soil algae. Most cyanobacteria are blue-green because of a preponderance of phycocyanins. Most unicellular cyanobacteria and many species within the common filamentous N2-fixing genera Anabaena and Nostoc are good examples. Other cyanobacteria appear brown or black due to a large content of red phycoerythrins together with the blue-green phycocyanins. Examples include numerous filamentous N2-fixing genera inhabiting rice soils, such as Aulosira and Tolypothrix (Roger & Kulasooriya, 1980). Chlorophyceae (green algae) and Xanthophyceae (yellow-green algae) are not reliably differentiated from one another on the basis of color. Green algae are usually grass-green, but many soil-inhabiting species are shades of olive- or yellow-green. Mature cells and resting spores can accumulate large quantities of carotenoids and will appear various shades of orange or red. Yellow-green algae can also be grass-green or yellow-green. Two features are used to distinguish these groups, the nature of the chloroplast(s) and the presence or absence of starch. Chloroplasts in green micro algae vary widely in number (usually one), shape (cup-shaped, reticulate, axial, and variously lobed) and location (parietal, polar, and central) within the cell. In contrast, most yellow-green micro algae harbor many small, uniformly disc-shaped, parietal chloroplasts (Metting, 1981). Green algae accumulate starch as the storage product of photosynthesis; yellow-green algae store chrysolaminarin, oils, or fats (Bold & Wynne, 1985). Presence or absence of starch is readily determined by staining with a dilute solution of iodine in potassium iodide. A black staining reaction indicates the presence of starch. Diatoms are common soil inhabitants and easily recognized by the distinct nature of the cell wall, which is known as the frustule. The frustule is made of silicon and has two halves, known as valves, that fit together like the top and bottom of a petri dish. Ornamentation of the valves is the principal criterion for taxonomic treatment at the species and subspecies levels (see section 21-2.6). Soil diatoms are usually colored various shades of brown or golden-brown.

2 Algal divisions and classes are based, in part, on pigmentation together with the chemical nature of the cell wall and of the principal storage product (photosynthate).

430

METTING

Morphological variation among species within all classes of cyanobacteria and eukaryotic micro algae includes unicellular (single-celled), colonial, siphonous, and filamentous forms. Cell diameters range from 1 to 2 !l (e.g., the unicellular cyanobacterium Anacystis) to 50 !l or more (e.g., the green coccoid alga Neospongiococcum). Coccoid species are round unicellular forms that do not aggregate (except during cell division). Coccoid green microalgae are extremely common soil inhabitants (e.g., Chlorococcum and related genera). Non-coccoid unicellular species have ellipsoid, fusiform, ovoid, or cylindrical cells (Metting, 1981). Two types of colonial algae are common in soil. In the first, cellular aggregates are indefinite in size, cell number, and relative cellular position; cell division is continuous under optimal growth conditions, and the aggregates readily fragment. Species exhibiting this morphological habit are often aggregated within a common, palmelloid matrix of extracellular polysaccharide. Examples include commonly encountered cyanobacteria (e.g., Gloeocapsa) and green micro algae (e.g., Chlamydomonas and PalmeUa). The second colonial type, known as a coenobium, has a fixed number and arrangement of cells. Sarcinoid aggregates are packets with cell numbers in multiples of four (rarely more than 16 or 32 cells). Common soil genera are the green micro algae Chlorosarcinopsis, Tetracystis, and Fasiculochloris (Metting, 1981). Siphonous microalgae include all bulbous and tubular-shaped eukaryotic genera and species that are multinucleate (coenocytic). These genera are distinguished morphologically by the conspicuous lack of transverse walls or septa. The yellow-green genera Botrydium and Vaucheria are common soil algae that inhabit garden paths and bench tops in greenhouses. On soil or agar, Botrydium appears as a collection of bubble-like sacs that can be visible to the naked eye. Vaucheria is a branching filament whose oogamous sexual production of zygospores is commonly observed by microscopic examination of samples collected from soil surfaces. Other siphonous microalgae are much less common in soil. Filamentous algae and cyanobacteria are common in soil. Filaments can be branched or unbranched and, particularly for cyanobacteria, are often enclosed together as bundles within a comm~n gelatinous matrix. Filamentous cyanobacteria include individual species with the greatest degree of cellular differentiation within the prokaryotic kingdom. Many species have barrel-shaped vegetative cells that house photosynthetic functions, heterocysts (rounded, anaerobic Nz-fixing cells), and thick-walled resistant spores known as akinetes on a single filament whose length can be as long as a few millimeters. Anabaena is perhaps the best example of a genus with this morphology. Nostoc is closely related to Anabaena and is also common on soil. Nostoc filaments can be linear or they can be organized into variously shaped globular aggregates, the diameters of which can reach several centimeters on soil and in flooded rice fields. Many genera of filamentous green and yellow-green algae are also common soil

ALGAE AND CYANOBACTERIA

431

inhabitants. Ubiquitous genera of green micro algae include Stichococcus, short filaments with 10 to 15 ovoid cells, and Klebshormidium, whose filaments can be many dozens of cells in length. Bumilleria and Tribonema are representative filamentous yellow-green algae (Metting, 1981). Most microalgae inhabiting soil and rocks are not taxonomically distinct from planktonic forms, although certain groups are proportionately more common in terrestrial habitats and many planktonic groups have never been detected in soil. However, many coccoid, sarcinoid, and pseudofilamentous green and yellow-green microalgae may be unique to the soil habitat. Hundreds of species in at least 185 algal and cyanobacterial genera have been identified in or isolated from soil or rocks (Metting, 1981, 1991). 21-1.2 Materials and Procedure Use any brand of light microscope together with glass slides and other standard materials (see chapter 6 by Bottomley) for identification of soil algae and cyanobacteria. No special materials are required. Detailed (Prescott, 1978) and diagrammatic (Metting, 1981) keys are recommended to help in the preliminary identification of soil algae to the genus level. The references in Table 21-1 are recommended for confirmation of generic identity and identification of individual species. Mount the soil sample or algal biomass collected from a culture vessel onto a glass microscope slide and examine it under the microscope beginning with scanning power. To distinguish anatomical features of eukaryotic micro algae (chloroplast number, size, and shape) or to view small cells (1-5 !!), it may be necessary to use the oil emersion with the highest power lens.

Table 21-1. Algal classes that include soil algae. Algal classt

Common name of the group

References to taxonomic treatment of species

Chlorophyceae

Green algae

Euglenophyceae Xanthophyceae Bacillariophyceae

Euglenoids Yellow-green algae Diatoms

Cyanophyceae

Cyanobacteria (blue-green algae)

Ett!. 1983 Komarek & Fott, 1983 Starmach, 1972 Leedale, 1967 Ettl,1978 Bock,1963 Krammer & Lange-Bertalot, 1986, 1988 Castenholtz et aI., 1989 Geit!er, 1930-32 (reprinted, 1985)

t Algal classes with species that are encountered only rarely in soil include Rhodophyceae (red algae) and Eustigmatophyceae (eustigmatophytes).

METTING

432

21-2 DIRECT METHODS FOR ENUMERATION 21-2.1 Principles Direct methods include techniques that permit or attempt some measurement or estimation of quantitative or qualitative (taxonomic) properties of the soil microalgal community at the time of sampling by direct examination of the soil sample. Special methods for obtaining soil samples are not necessary if the samples are subjected to microscopic examination soon after collection or if quantitative data are not the objective. For quantitative or comparative study, however, the following method is recommended for obtaining samples of the surface veneer of soil. Microalgal communities are only commonly abundant or play important roles in the upper few millimeters to centimeters of soil where light is available for phototrophic growth. 21-2.2 Materials and Procedure for Collecting Soil Veneers A special method is required for collecting samples for studying soil algae. Quantitative data for soil algae should be reported on the basis of area. Thus, values per square centimeter should be reported. For dense populations in rice soils, values per square meter are common. It is also acceptable to report values per gram of soil, but not in lieu of reporting on the basis of area. For most soils, 1 cm2 by 1 cm in depth is approximately equal to 3 g (air-dry) soil. Sampling devices (samplers) for collecting soil surface veneers are not readily available from scientific supply houses. Therefore, it is necessary to have the samplers fabricated. Choose a material that is strong enough to permit the use of force in penetrating hard dry soils and that will withstand repeated autoclaving. The preferred sampler is crafted from hollow, stainless steel tubing that is cylindrical or square in cross-section, 20 cm in length, and beveled at a 45 angle on one end to facilitate vertical penetration of the soil. Round samplers are preferred although square ones can be used. A loose-fitting stainless steel cap is made to fit the non-beveled end of the tube. For most soils, a cross-sectional area of 1 or 3.33 cm2 is ideal. The larger area is used for algal populations that are not readily visible. In this situation, it is common to pool three samples giving a total of 102 cm with which to work. A solid round (or square) dowel or other implement for use in pushing soil cores through the samplers should be fabricated to exactly fit within the sampler. For repeated collection of soil samples for studying algae, at least 20 to 30 samplers and two solid dowels should be available (Rayburn et aI., 1982). Samplers should be capped, packaged in brown paper sacks (three, four, or five per sack), and sterilized by autoclaving or baking in an oven prior to sampling. Orient the samplers in the sacks so that the capped end 0

ALGAE AND CYANOBACTERIA

433

is grasped when opening the sack in the field and the beveled (open) ends are not exposed. The choice of a random or systematic sampling pattern depends on the objectives of the study and limitations imposed by the site (see chapter 1 in this book). Collect samples by pushing the beveled end of a sampler into the soil to a depth of at least half the length of the tube. If necessary, use a rubber mallet to facilitate penetration of a hard soil. Place the sampler with its cap intact and containing the fresh core, beveled end first, into a clean sack, plastic bag, or other suitable container. Repeat the procedure with a new sampler and as many times as needed to collect the required number of samples. Return the samples to the laboratory, keeping them dark and at ambient temperature. Samples may be stored on ice if it will be more than a day between sampling and analysis. Depending on the study objectives and any preliminary information or assumptions regarding the structure or density of the surface algal communities, soil veneers from the individual sample tubes can be analyzed separately (as 1- or 3.33-cm2 samples) or they can be pooled. Alternatively, a single 1- or 3. 33-cm2 sample will often yield sufficient chlorophyll when pigmentation is used as the algal biomass index. Process the samples at the laboratory bench and have ready the solid dowel(s), a supply of alcohol, a forceps, a supply of single-edged razor blades, and suitable borosilicate flasks, beakers, or other containers for collecting the soil samples. It is also recommended that a lined garbage can and plastic bucket or tub filled with detergent and hot water be close by for convenient disposition of used sample tubes. Holding a sample tube in one hand, use the other hand to remove the cap and then to force the soil core through the tube by pushing with the solid dowel. When the surface of the soil core is pushed beyond the lip of the tube, use a flame-sterilized razor blade to carefully slice off a measured thickness of the veneer into a waiting flask or beaker. Continue to push the entire core through the tube into the garbage can, put the tube and cap into the wash tub, and repeat the procedure with the remaining sampling tubes. 21-2.3 Microscopy and Enumeration by Cell Counting

General methods for the use of light microscopy in the qualitative and quantitative examination of soil microalgae are the same as those described for bacteria and fungi (see chapter 6 in this book). However, because they are often much larger than heterotrophic eubacteria and are pigmented, the direct observation of microalgae and cyanobacteria in soil is less difficult. Any light microscope equipped with a high-intensity light source, a condenser, and a stage that is capable of accommodating a cell-counting chamber (haemacytometer) can be used for standard microscopic examination (Waaland in Gantt, 1980). A special apparatus and procedure,

METTING

434

described below (21.2.4.2) is required for fluorescence microscopy. The limitation to optical methods is interference from soil particles, particularly when cell densities are less than about 104 per cm2 • Special cytological methods for cultured algae, including staining procedures, are detailed in Gantt (1980). Two general methods for counting micro algal cells in soil or soil dilutions include the use of a standard cell-counting chamber (see chapter 6 in this book) or counting directly on glass slides that can be calibrated with the ocular and stage micrometers. Except for the larger, filamentous forms, the use of the counting chambers, with either transmission or epifluorescence microscopy, for enumeration of microalgae in soil samples is usually practical only for population densities greater than or equal to about 104 to 105 per cm2 •

21-2.4 Enumeration by Chlorophyll Autofluorescence 21-2.4.1 Principles Chlorophyll strongly fluoresces when excited by blue and UV wavelengths. This is the basis for estimating micro algal biomass by extraction of chlorophyll from soil (21-3.3). It also permits the direct microscopic observation of algae in soil and allows distinction between living and nonliving cells (Shields, 1982). Although a transmission fluorescence microscope is suitable, epifluorescence is preferable because of its greater versatility and ease of use. For example, epifluorescence microscopy permits nearly simultaneous viewing under fluorescence and bright-field or phasecontrast images. Also, cell-counting chambers can be used with epifluorescence microscopy but, because of their thickness, cannot be used for transmission microscopy. For transmission microscopy, a special condenser illuminates the specimen so that light from the condenser does not directly enter the objective. Only light reflected, scattered, or fluoresced by the specimen enter the objective. The object thus appears to be self-luminous in a surrounding dark field.

21-2.4.2 Materials and Procedure If a standard fluorescence microscope is not available, it is possible to convert a standard compound microscope for the purpose. Packages to upgrade a standard system for fluorescence applications are available from most microscope manufacturers. Components of a package will include a UV light source (100 W Mercury arc lamp), a dark-field condenser, and appropriate excitation and barrier filters. Some microscopes may also require modified objectives (J.R. Johansen, 1992, personal communication). Prepare an appropriate dilution of soil, taking care to avoid transferring particles 2: 0.1 mm into the cell-counting chamber. Focus the condenser of the light source to produce a parallel beam. Insert a blue filter between the light source and microscope. Introduce one drop of soil sus-

ALGAE AND CYANOBACTERIA

435

pension into the counting chamber. The best magnification for epiftuorescence enumeration is 100 to 400 x . Count the algal cells in several fields or along several transects calculated to be statistically appropriate (see chapter 2 by Parkin and Robinson in this book). Living cells appear red against a black background, while dead cells, although sometimes difficult to distinguish, appear greenish gray. Soil particles will sometimes fluoresce yellow or green. Work in a darkened room for best results. 21-2.5 Implanted Slide Method 21-2.5.1 Principles The procedure involves the implantation of a microscope slide vertically in the soil, leaving the top 1.5 cm ofthe slide projecting above the soil surface. Light attenuates downward from the exposed portion of the slide, encouraging differential micro algal growth on the surface of the slide in contact with the soil, thus providing a means by which to study the vertical distribution of the soil algal community (Pipe & Cullimore, 1980). 21-2.5.2 Materials and Procedure Mark two glass microscope slides with small scratches at 2-mm intervals along their entire lengths and near both edges of one face on each slide. Place the two marked faces together, fastening pairs of slides together at each end with an adhesive tape. Push slide pairs into the soil until only the upper 1.5 cm is exposed. To hasten colonization of the slides or for comparative purposes, the soil may be moistened with deionized glassdistilled water or with an appropriate algal medium (section 21-5.1.3). After incubation for the desired length of time, carefully retrieve the paired slides. The best way to remove the paired slides is in a block of soil, rather than by pulling the slides from the soil. Carefully chip the soil away from the exposed surfaces of each slide and then separate each pair. Allow the slides to air dry. Remove large soil aggregates by gentle tapping against a hard surface. Spray the colonized surfaces with a molten 2% agar solution in water. Dehydrate the slides in the air stream provided by a laminar flow bench or a hair dryer. The result will be a thin covering of agar that acts as an adhesive covering the soil particles and micro algae and sealing them onto the surface of the glass. The slides can be microscopically examined immediately (or following storage) with the addition of a few drops of water. Wrapped in aluminum foil and kept dry in the dark at room temperature, slides can be stored for up to 2 yr. Individual slides can be alternately wetted and dried through several cycles without consequence, so long as they are not kept moist for longer than a few hours at anyone time. Reference to the scratch marks permits quantification of spatial distribution on the surface as a function of the original vertical distribution in soil.

436

METTING

21-2.6 Method for Diatom Frustules 21-2.6.1 Principles

Frustule is the term used to designate the cell wall of diatoms. Each frustule is composed of two halves, or valves, that fit together much like the halves of a petri dish. Diatoms are unique among algae in that their cell walls are composed of silicon. Because the taxonomy of diatoms is based on frustule ornamentation and because silicon is resistant to harsh treatment, an important aspect of the study of these micro algae is the use of chemical methods for the preparation of samples for qualitative and quantitative microscopic study. The following quantitative method has been used successfully in several studies of soil diatoms (e.g., Johansen et aI., 1982, 1984). 21-2.6.2 Materials and Procedure Prepare a composite sample of 1- or 3.33-cm2 samples as described above (section 21-2.2). Dry the composite sample for 1 hat 105°C in an oven. Pass the dry sample through a 1-mm mesh screen. Disperse a 1-g subsample in 20 mL of deionized glass-distilled water in a 100-mL beaker. Add 10 mL of concentrated HN0 3 and bring the mixture to a boil on a hotplate placed behind the protective sash in a forced-air fume hood. If the soil has a moderate to high organic matter content, additional oxidation is beneficial and can be achieved by the addition of about 0.5 g (or more) K2Cr207 after the first 10 min of boiling in concentrated HN0 3 . Be prepared to avoid spattering (bumping) as the mixture boils by repeatedly removing the beaker from the hotplate. Discontinue the heating and allow the beaker and contents to cool to room temperature when the volume has been reduced by boiling to < 20 mL. Divide the 20-mL sample into two equal portions in 15-mL glass or plastic centrifuge tubes. Centrifuge the contents for a few seconds in a tabletop clinical centrifuge. Decant the liquid supernatants. Bring the volume back to 10 mL with water and resuspend the soil with a vortexer. Repeat the centrifugation, decanting, and resuspension six or more times. After the final rinse, bring the volume of cleaned soil to 10 mL. Repeated rinsing with deionized glass-distilled water is necessary to remove all of the nitric acid, to prevent the appearance of acid halos that otherwise interfere with microscopic examination of the details of frustule morphology. Make a 10-fold dilution of this mixture in deionized glass-distilled water for preparing slides for microscopic examination. Semi-permanent slides for repeated study can be made in a suitable mounting medium (Hyrax and Naphrax are recommended; J.R. Johansen, 1992, personal communication). Place 0.5 mL of the diluted soil mixture on a clean 18-mm square glass coverslip. Allow to air dry overnight. The next day, add two to three drops of mounting medium on a glass slide and place it on a hotplate. The heat should be sufficient to boil the mount ant within 5 to 20 s. Boil the mountant until the boiling slows but does not stop (a few

ALGAE AND CYANOBACTERIA

437

seconds). Remove the slide from the heat and place the 18-mm square coverslip, diatom side down, onto the heated mountant. Place the slide back on the hotplate allowing it to boil again while tapping the coverslip lightly with a teasing needle to spread the medium to its edges. Tapping the slide several times onto the edge of the work bench by dropping it from a height of about 3 cm will help rid the mount of entrapped air bubbles. Enumerate the diatom frustules at 1000 x . Determine the width of the field of view and convert to millimeters. This value times 18 is the area of the slide covered by one transect. To obtain quantitative counts, start at the top or bottom of the slide at the nearest even whole number on the vernier scale of the microscope stage. Because each diatom frustule is composed of two halves (valves), count the number of valves and divide by two to obtain an estimate of the original number of frustules and, thus, diatom cells. Count the total number in nine transects, each I-mm apart. The number of cells per gram of dry soil is calculated by: FCrrAM, where F = number of frustules C = area of coverslip (324 mm2) T = number of transects scanned A = area of one transect (18 mm x width of field), and M = mass of air dry soil on coverslip (0.0025 g). Identification of soil diatom species is performed by comparing the characteristic valve morphology with taxonomic descriptions given by Anderson and Rushforth (1976), Ashley et ai. (1985), Johansen et ai. (1981), and the authorities cited in Table 21-1. 21-2.7 Methods for Cyanobacteria in Rice Fields 21-2.7.1 Principles

Cyanobacteria are ubiquitous in rice fields. With minor modifications, the same direct and indirect methods described in other sections of this chapter are also used for sampling and analysis of rice fields for studying cyanobacteria and other microalgae. Modifications reflect the nature of the rice field environment and the fact that most of the important cyanobacterial species in rice fields are filamentous and often form dense mats. The methods described in this section are also suited for studying the algal component of non-agricultural wetlands, such as intermittently flooded marshes and estuaries (Roger et aI., 1991). Rice fields are usually flooded for some part or all of the cropping cycle. Thus, portions of the associated microalgal communities float and are subject to distribution patterns that are influenced by water movement, wind, and the spacing of individual rice plants. Individual filaments or portions of cyanobacterial aggregates can also move vertically in the water column and will adhere to submerged and emergent portions of the rice shoot (Roger et aI., 1993).

438

METTING

Cyanobacteria are usually irregularly distributed in rice fields. Correlations between means and variances of different indices of cyanobacterial abundance (most probable numbers [MPNs], plate counts, wet biomass) and photodependent N2 fixation (as acetylene reduction) in nearly all studies have shown in most cases that the variables are distributed in a lognormal pattern. This has been observed for single-locus samples collected in the same plot and for composite samples collected in replicated plots. For this reason, random sampling is discouraged. Sampling strategies should be selected only after careful examination of the conditions at the site (Roger et aI., 1991). 21-2.7.2 Modified Procedure for Sample Collection, Serial Dilution, and Plating The use of beveled sampling tubes (section 21-2.2) is adequate for non- flooded rice soils. A modification of the sampling tube is recommended for collecting samples from flooded fields. When obtaining samples, take care to minimize disturbance. To avoid muddying the waters, the person collecting the sample should stand or kneel on a dike or on a wooden board placed across the study area. In place of a beveled tube, use one with two blunt open ends. Instead of a cap at one end, close both ends with rubber stoppers. Gently push the tube down through the water column into the soil with a circular twisting motion. When the desired depth is reached: (i) plug the top, (ii) very carefully remove the tube, and (iii) plug the bottom while it is still submerged (Roger et aI., 1991). As has been mentioned, any of the direct or indirect cell counting, plating, MPN, or pigment extraction methods can be used when studying cyanobacteria in rice fields. However, because of the filamentous nature of many species and the fact that mucilagenous sheaths surrounding individual filaments or entire mats are common, it is necessary to use more forceful methods and longer periods of time when homogenizing samples prior to serial dilution for plating or inoculating MPN tubes. To facilitate disruption of filaments, Roger et aI. (1991) recommend that soil samples be ground in a mortar and pestle and sieved to a particle size;::: 0.25 mm and that composites of at least 10 samples that include the upper 5 mm of soil be stirred at 400 rpm for 15 min prior to serial dilution. For plating, agar concentrations > 1% are not recommended because they have been shown to inhibit the growth of some unicellular Nz-fixing species (Van Baalen, 1965). BG-ll medium is recommended (section 21-5.1.3.1). To discourage the growth of eukaryotic microalgae, add 20 to 50 mg cycloheximide per liter of medium. To enrich for N2-fixing species, do not include fixed N in the medium (section 21-5.1.3). 21-2.7.3 Procedure for Visual Estimation of Coverage For observational purposes, visual estimates often are sufficient to follow the development of algal communities during a cropping cycle when

ALGAE AND CYANOBACTERIA

439

Table 21-2. Indices of abundance for visual estimates of cyanobacteria in rice fields. Relative abundance

Code

No visible growth Few colonies present More colonies present More colonies present More colonies present Growth covers one-fourth of the area Growth covers one-third of the area Growth covers one-half of the area Growth covers two-thirds of the area Growth covers three-fourths of the area Growth covers the entire area

0

+ ++ +++ ++++

114 113 112 2/3 3/4

All

Index

Index percentage

0 1 2 3 4 5 6 7 8 9 10

0 5 10 15 20 25 33

50 67 75 100

it is either logistically difficult or not convenient to use more accurate methods. Roger et al. (1991) describe two methods. The square count method is convenient when algae are not abundant and when the rice plants have been transplanted in a grid pattern. The index of abundance is easier and more accurate when algal growth is denser. These techniques can also be used to estimate coverage by Azolla in fields where this water fern is used as a biofertilizer. Where farming practices do not include transplantation of seedlings into grid patterns, such as in the USA, where common practices include aerial seeding, it is necessary to construct a grid using spaced wooden stakes or steel rods. 21-2.7.3.1 Square Count Method. Individual rice plants in transplanted fields provide a grid that can facilitate making visual estimates of algal coverage. The square count method is used when algae are not abundant. The observer records the presence or absence (+ or - ) of growth in as many squares as possible formed by individual rice plants. If desired as a preliminary measure, the percentage of squares with visible growth can be correlated to kg fresh weight ha -1. Calculation of a regression equation for a representative area yields a useful tool for larger areas. 21-2.7.3.2 Index of Abundance Method. This is a semiquantitative index based on how much of the floodwater surface area is covered with algal growth. Use the subjective codes and indices shown in Table 21-2 to determine the average index percentage. This index is most accurate when standardized beforehand. 21-2.8 Procedure for Endolithic Microalgae

Except for collecting the rock specimens themselves, no special methods are used for the study of endolithic microalgae (E.I. Friedmann, 1992, personal communication). In the field, samples are chipped off of rocks and collected in Whirl-Pak plastic bags. Samples from cold deserts should be frozen for transport to the laboratory. Whole rocks, fragments, or crushed

METTING

440

rock are used for the laboratory study of photosynthesis (02/C02 uptake/ evolution) and N2 fixation using standard methods (see chapters 38 by Zibilske and 43 by Weaver and Danso, respectively). Microalgae are isolated from rocks with standard plating techniques and are cultured in standard algal media. For extraction of chlorophyll (21-3.3.3), dimethyl sulfoxide is preferred (Bell & Sommerfeld, 1987; E.!. Friedmann, 1992, personal communication).

21-3 INDIRECT METHODS FOR ENUMERATION 21-3.1 Enumeration of Colony-forming Units on Solid Media Standard soil dilution and plating methods (see chapter 8 by Zuberer in this book) are recommended for use in enumerating populations of soil algae as CFU on agar plates. 3 Simple modifications are used to compensate for the slower growth of micro algae and cyanobacteria relative to heterotrophic eubacteria and fungi. Modifications of the standard plate count method for use with microalgae include: 1. The use of antibiotics to control the growth of bacteria. At lower soil dilutions (10- and 100-fold dilutions), soil particles are a source of organic compounds that support growth of heterotrophs. To discourage the growth of unwanted bacteria, supplement any algal medium with penicillin and streptomycin (21-5.1.3). 2. Pour the agar plates nearly full to reduce the likelihood of desiccation during the 1- to 3-wk (sometimes longer) incubation period. To prevent condensation from smearing the colonies, plates should be inverted and incubated on shelves under standard conditions (section 21-5.1.2). Wrapping the edges with parafilm will retard desiccation.

21-3.2 Most Probable Number Method for Enumeration Prepare a soil dilution series as described in chapter 5. From each of six successive dilutions (1()2 through 106 ), transfer 1 mL to each of five 15-mL culture tubes containing one of the algal media described below (21-5.1.3). If the objective is the enumeration of specific groups, the appropriate medium and recommended supplements should be used (215.1.3). If, on the other hand, total numbers of microalgae (including cyanobacteria) are desired, soil water tubes using soil from the study site should be used (21-5.1.3.5). Incubate the tubes on culture tube racks under standard conditions (21-5.1.2). After 3 wk, record the number of 3For microalgae, colonies on agar are sometimes referred to as plant masses (Archibald & Bold, 1970).

ALGAE AND CYANOBACTERIA

441

tubes with visible algal growth. Find the MPN of micro algae in the original sample using the information provided in chapter 5 by Woomer. 21-3.3 Chlorophyll Extraction and Quantification 21-3.3.1 Principles Extraction and spectrophotofluorometric quantification of chlorophyll is a useful index of soil algal biomass. Usefulness derives from the facts that (i) all groups of micro algae and cyanobacteria produce chlorophyll, (ii) the procedure is easy to perform and somewhat less labor and equipment intensive than the MPN or plate count methods, and (iii) data are available within hours of sampling, compared to the days or weeks required to incubate agar plates or culture tubes. The major disadvantages of the method are that (i) it fails to distinguish among microalgal species or higher taxonomic groupings (e.g., among cyanobacteria, eukaryotic microalgae, and lichen phycobionts), and (ii) as an index of biomass or metabolic activity, chlorophyll content can vary markedly among different micro algae, at various stages of growth within individual populations, and in response to environmental conditions (Metting & Barry, 1986; Sharabi & Pramer, 1973). Two methods are described. The first is an ethanol extraction procedure developed for routine use with cultivated and irrigated silt loam and sandy loam soils (Metting & Barry, 1986). The second is a dimethyl sulfoxide (DMSO) procedure widely used for chlorophyll extraction from soils and rocks (Bell & Sommerfeld, 1987; J.R. Johansen, 1992, personal communication) . 21-3.3.2 Procedure for Estimating Cell DensitylPigment Correlation in Soil The chlorophyll method is most useful in controlled studies of the influence of experimental variables on unialgal populations on soil or when studying relatively homogeneous populations in the field. In both cases, a standard curve is prepared prior to the experiment by extracting and measuring chlorophyll from soil to which a known number of cells of a selected species with a known biomass has been added. To prepare a curve, produce a unialgal population of the desired species in liquid culture under standard conditions (section 21-5.1.2). At the desired stage of log or stationary phase population growth, quantify the cell densities by direct microscopic enumeration (section 21-2.3). Depending on the species, culture densities can reach 106 per mL for filamentous cyanobacteria and as many as 108 or more per millimeter of unicellular eukaryotic species. Prepare a cell concentration and dilution series from the liquid culture by centrifugation and lO-fold serial dilution to yield 10 mL each of 103 through 10 10 cells per mL for use in experiments in which cell densities on soil cannot be predicted. Add the lO-mL aliquots to 10 g (3 cm2) of air-dry soil, mix thoroughly, and proceed to quantify the soil

442

METTING

chlorophyll content as described below (section 21-3.3.3.2). The procedure is easily modified to produce a narrow range or one with more correlation points between log concentrations of cells. 21-3.3.3 Ethanol Extraction Method for Quantification of Chlorophyll in Soil 21-3.3.3.1 Materials. Have available: 1. Twice as many clean 50- or 100-mL flasks as there are samples to be processed. To half of them, add a pinch of Na2S04 (approximately 1 g). 2. A Turner model 111 Filter Fluorometer or similar instrument with blanking rod and quartz cuvettes. A spectrophotometer, such as the Bausch & Lomb Spectronic 20 or 21 models, can be used in the absence of a fluorometer. However, fluorometric measurement is preferred because of its greater sensitivity, and hence usefulness, for detecting small quantities of pigment. 3. Two or more hot plates. 4. 95% ethanol, petroleum ether, and deionized glass-distilled water. Heat about 200 mL of 95% ethanol in a separate 500-mL flask and have it at hand for sample filtrates. 5. Two or more 250-mL side-arm flasks with Buchner funnels and no. 1 Whatman filter paper. Two or more 250-mL separatory funnels. Jars, jugs, or beakers for waste liquids. Canisters of 5- and lO-mL volumetric glass pipets. Two or more 25-mL graduated cylinders. 21-3.3.3.2 Procedure. Collect individual or pooled samples of the surface veneer ofthe soil to be studied, as outlined above (section 21-2.2). Perform the chlorophyll extraction procedure as soon as possible after collecting the samples to minimize changes in pigment concentration. There is no objective way by which to determine how much soil (how many cm2) to use for a single extraction. In general, if algal growth is not visible to the naked eye, as much as 10 cm2 may have to be extracted to yield sufficient chlorophyll for fluorometric quantification. If there is visible algal growth, then as little as 1 cm2 may yield sufficient pigment. Luxuriant growth will yield an excess of chlorophyll-necessitating dilution prior to fluorometric measurement. Perform all extraction and separation steps in a fume hood. Once familiar with the procedure, it is possible to simultaneously process a small number of flasks. However, it is recommended that the researcher first become familiar with the procedure by subjecting a single sample to the following steps. Using the method described in section 21-2.2, add the surface veneer from the desired number of sample tubes to a clean dry flask and repeat for each desired sampling unit. Add 5 mL of cold 95% ethanol to each flask. Cover the mouth of each flask with aluminum foil. Perforate the foil with two or three small holes with a dissecting needle, safety pin, or the pointed

ALGAE AND CYANOBACTERIA

443

end of a paper clip. While occasionally swirling the flask, bring the combined contents to a boil over low to moderate heat on a hotplate. To avoid bumping, remove the flask(s) from the hotplate as soon as they boil. Filter the contents through no. 1 Whatman filter paper into a 250-mL side-arm flask. Rinse the flasks with an additional 5 mL of hot 95% ethanol. Pour the leachate into a 25-mL graduated cylinder and note the volume (which should be :::; 10 mL). Transfer the leachate to a 250-mL separatory funnel with the stopcock closed. Add an aliquot of petroleum ether equal to about twice the volume of the leachate. Gently swirl the contents for 3 to 4 min, holding the funnel at a 45° angle. Add a volume of deionized glass-distilled water equal to the total volume of leachate plus the petroleum ether, thus doubling the total liquid in the funnel. Swirl the contents for an additional 3 to 4 min. Drain the lower ethanol/water mixture and a small portion of the upper layer. Drain the remaining petroleum ether with the dissolved chlorophyll into one of the 50- or 100-mL flasks into which was placed the Na2S04. Swirl the contents and fill a quartz cuvette with a representative fraction. Excite each sample from the petroleum ether fraction at 430 nm (chlorophyll a lambda maximum) and pass to the detector mostly 669 nm fluorescence (chlorophyll a fluorescence maximum). Use the 1 x slit arrangement. Dilute the sample in petroleum ether if the reading is off the scale. The density of chlorophyll in !!g per cm2 soil is equal to the fluorescence reading multiplied by 0.66 (times any appropriate dilution factors). When using a spectrophotometer, the density of chlorophyll is given by the following relationship (Jensen in Hellebust & Craigie, 1978): mg chlorophyll a mL -1

=

[COD at 666 nm) - (OD at 730 nm)] x mL of sample x 10 890 21-3.3.4 DMSO Extraction Method for Quantification of Chlorophyll in Soil 21-3.3.4.1 Materials and Procedure. The DMSO extraction procedure obviates the need for heating the samples and using separatory funnels, as described above (21-3.3.3.2). If rock samples are being extracted, it is recommended that they first be homogenized by crushing with a mortar and pestle or a rolling pin. Some benefit may also be derived from crushing and mixing soil samples. Working in dim light to minimize degradation of the phaeophytin fraction, add about 1.5 g of rock or soil to 5 mL of DMSO in a screw-cap centrifuge tube. Mix the contents of the tube for 20 s on a vortexer and incubate for 45 to 60 min at 65°C. Vortex the tubes a second time about half way through the incubation. Centrifuge at top speed in a clinical centrifuge for about 1 min or until the particulate matter has collected in a pellet. Record the absorbance of the supernatant solution at 665 and 750 nm. After each reading, pour the liquid contents back into the tube used

METTING

444

for the extraction and clean the cuvette with DMSO. Acidify the sample with 10 !AL (or two drops) of 1 N HCI. Wait 10 min and proceed to again record absorbances at 665 and 750 nm. For each value at 665 nm, subtract the reading at 750 nm to correct for turbidity. Calculate the concentrations of chlorophyll a and phaeophytin from the following formulas (Lorenzen, 1967). mg chlorophyll a g-l = A x K x (665 t =0 - 665 t =a) x V L mg phaeophytin g-l = A x K x (R[665 t =al - 665 t =0) x V L where A = absorption coefficient of chlorophyll a (= 11.0) K = factor to equate reduction in absorbancy to initial chlorophyll a concentration (= 1.7,0.7, or 2.43) t = 0 = absorbance after initial 45 to 60 min incubation t = a = absorbance after acidification for 10 min L = path length of cuvette in centimeters R = maximum ratio of 665 t =0 to 665 t =a in the absence of phaeophytin pigments 21-4 METHODS FOR ISOLATION AND PURIFICATION OF MICROALGAL CULTURES

Microalgae and cyanobacteria enriched from soil can easily be isolated into unialgal culture and subsequently purified into axenic culture using traditional bacteriological and traditional phycological methods developed for planktonic species from marine and freshwater habitats (Hoshaw and Rosowski in Stein, 1973). Use sterile inoculating loops or flame-drawn glass Pasteur pipets to remove individual microalgal colonies (or portions of colonies) from agar surfaces (or liquid culture tubes) on (or in) which multiple colony types have arisen following inoculation with aliquots from serial soil dilutions. The colony material can then be inoculated onto the surface of a fresh agar plate or into a culture tube containing a suitable liquid medium for subsequent purification into clonal unialgal or axenic culture. 21-4.1 Procedure for Preparing Fine Capillary Pipets 21-4.1.1 Principle

The procedure for preparing fine bored glass capillaries from individual glass Pasteur pipets is a standard phycological technique for collecting and transferring single cells or short pieces of filaments from one tube, spot

ALGAE AND CYANOBACTERIA

445

dish, or petri dish to another. Individual capillaries are prepared one at a time as they are needed. To minimize contamination, always use presterilized equipment and work at a laminar flow bench or in a clean room. 21-4.1.2 Materials and Procedure Ahead of time, pack several borosilicate glass Pasteur pipets into a glass pipet canister (Belleo) with an aluminum closure after having first stuffed the blunt end of each pipet with a small piece of absorbent cotton. Autoclave the canisters. Dry them in an oven, and allow them to cool to room temperature before proceeding. To make a capillary pipet, work at a laminar flow bench. Holding the blunt end of a pipet in one hand, grasp the tip of the opposite end with a pair of forceps. Hold a portion (between 3-8 to 10 cm) of the fine end of the pipet over a flame for 3 to 5 s, depending on distance from the flame until a short (1-2 cm) region begins to glow. When the glass is fluid, remove the pipet from the flame and, in one motion, pull evenly on both ends until the molten piece is reduced in thickness to the desired bore. Snap off the tip using the tweezers. 4 To use a pipet, squeeze a 1- to 2-mL rubber bulb, attach it to the end of a pipet, and release the pressure on the bulb to create a vacuum. Touching the fine tip of the capillary to a portion of a colony on an agar surface or a cell( s) in solution is sufficient to draw the desired cells into the pipet. The cell(s) can then be ejected into a fresh culture tube or onto a fresh agar plate by compressing the rubber bulb. 21-4.2 Isolation and Purification by Repeated Washing 12-4.2.1 Principle This traditional phycological method is used to isolate individual cells into clonal culture from mixed planktonic communities. It is also used to isolate single cells from mixed cultures in MPN tubes prepared from soil dilutions and homogenizations prepared from colonies on agar plates (Hoshaw and Rosowski in Stein, 1973). 21-4.2.2 Materials and Procedure Prepare ahead of time 1. Sterile canisters of glass Pasteur pipets (21-4.1.2) and 7- x 22-mm glass spot dishes (Corning 7223) placed inside 100- x 20-mm glass petri dishes. 2. Sufficient volumes of the appropriate sterile algal medium (215.1), tap water, or deionized glass-distilled water. 3. Binocular dissecting microscope, forceps, and rubber bulbs. 4It is necessary to practice this technique to consistently prepare capillaries of desired thickness.

METIING

446

Line up a series of three (or more) dishes for a total of nine (or more) spot dishes. Add six to eight drops of the desired medium to all but one of the wells in the spot dishes. Pipet six to eight drops of the mixed algal culture to the remaining well. Perform all subsequent manipulations with the aid of a binocular dissecting microscope. Use a flame-drawn glass pipet (section 21-4.1) to isolate individual cells or pieces of filaments from the first well and to squirt them into the adjacent well. While moving the tip of the pipet through the liquid, maintain sufficient pressure on the rubber bulb to prevent the premature uptake of liquid. Repeat until at least 12 to 15 cells have been isolated into the second well, using a fresh pipet for each cell. Repeat the procedure until as many cells as possible have been passed through as many wells as needed to ensure that at least one cell is available to inoculate fresh culture tubes (or agar surfaces) to give rise to clonal cultures. 21-4.3 Isolation and Purification by the Streak Plate Method Prepare a suitable number of petri dishes containing the appropriate medium solidified with agar and supplemented with antibiotics to discourage bacterial contamination (section 21-5.1.3). Fill the petri plate nearly full with agar to delay desiccation during the time required for slow-growing isolates to develop adequately sized colonies from single cells. Place a small inoculum from the original spread plate or MPN tube in one comer by touching the agar surface with the inoculating loop or Pasteur pipet and streak for isolation. Incubate the plates upside down on shelves under standard conditions (section 21-5.1.2). Keeping the dishes upside down, scan the agar surface after 2 to 3 d at regular intervals at 30 x power with a binocular dissecting microscope to locate individual cells or groups of cells that are distinctly separate from others. Mark the location of the cells by drawing a circle on the bottom of the closed petri dish with a marking pen. Tum the dish right side up and, using a higher power of magnification, proceed to collect the cell(s) with a sterile flame-drawn pipet. Deposit the cell(s) in a culture tube with the appropriate liquid medium. Repeat the procedure to collect as many individual cells as are reasonable. Incubate the tubes under standard conditions (section 21-5.1.2). 21-4.4 Purification by the Centrifugation Technique Microalgae can usually be separated from adhering bacteria by repeated centrifugation and washing in sterile water with or without detergent. This method is based on the fact that microalgae settle out more readily than the much smaller bacteria. Work with sterile equipment in a clean setting. Fill a sterile 5-, 10- or 15-mL glass tissue grinder with an appropriate aliquot from a log-phase culture of a unialgal culture. For best results with mucilagenous species, add a small drop of Tween 20 or Triton X-l00 detergent. Homogenize the

ALGAE AND CYANOBACTERIA

447

contents with a few short, twisting strokes, taking care not to use so much force that the contents of the reservoir are lost when plunging the barrel. Fill a sterile lS-mL centrifuge tube with about 10 mL of the homogenate. Hold the tube on a vortexer for a few seconds. Centrifuge the tube at full speed in a tabletop clinical centrifuge (about 2000 rpm) for about 30 s. The micro algal cells will form a pellet. Pour off the supernatant. Vortex again for a few seconds. Quickly and forcefully add 10 mL of fresh sterile water to disperse the pellet. Repeat the centrifugation, decanting, vortexing, and resuspension steps. Do not add detergent after the first repetition. Place a drop from the pellet onto an agar surface after 12 repetitions (or sooner if there is a danger of losing the algal pellet altogether). Choose the appropriate algal medium (section 21-S.1.3). Supplement the medium with O.S g of a protein extract or O.S g of yeast extract to facilitate the identification of micro algal colonies that remain associated with bacteria. Streak the pellet to separate the individual cells. Incubate under standard conditions (section 21-S.1.2) and examine under a dissecting microscope, as described above (section 21-4.3). 21-4.5 Purification by the Zoospore Technique Zoospores are the single-celled flagellated stage produced by many green and some yellow-green microalgae. A simple purification procedure depends on their motility and phototactic response (Kessler, 1984). Inoculate a 30 mL culture tube containing about S mL of an appropriate sterile medium (section 21-S.1.3) with a loop full of algal mass taken from an agar surface or another liquid culture. Aseptically place a wad of sterile cotton above the level of liquid in the tube and then add enough additional sterile medium to saturate and totally immerse the cotton. Place the tubes below a source of light. If the inoculum subsequently produces zoospores, they will swim upward through the cotton barrier where they can be captured with a capillary pipet and transferred to fresh medium. Zoospores are axenic when first liberated from the parental cell wall. In traversing the cotton, many of the zoospores that were subsequently contaminated will have the few contaminating bacterial cells removed in the process. 21-5 METHODS FOR GROWTH AND STORAGE OF MICROALGAL CULTURES 21-5.1 Culture Methods and Growth Media 21-5.1.1 Principles Most terrestrial micro algae are obligate photoautotrophs or facultative photoheterotrophs. Many are obligate photoheterotrophs requiring the provision of vitamins or other growth factors from the environment. Most facultative heterotrophs use only simple sugars or organic acids,

448

METTING

although some are capable of growth on polysaccharides or proteins (e.g., starch and gelatin). Many are also capable of dark heterotrophic growth. A few soil microalgae are obligate heterotrophs, including some euglenoids whose saprophytic or predatory nutritional requirements make them ecologically equivalent to protozoa (Metting, 1981). 21-5.1.2 Standard Methods for Culturing Microalgae and Cyanobacteria

Microalgal cultures are maintained in liquid media or on agar slants. A wide range of conditions of irradiance, photoperiod, and temperature are employed for different experimental purposes. A dedicated room with precise temperature control, such as a fully contained, refrigerated walk-in room, is preferred. At a minimum, the room should be refrigerated and have sufficient circulation of cooled air to counteract the heat generated by the lighting fixtures. Mounting the lighting fixture ballasts outside the room is recommended to aid temperature control. Provision of central pressurized air or placement of a dedicated air compressor is recommended for aeration and agitation of liquid cultures. The following are standard culture conditions: 1. Constant air temperature of 20 or 25°C. 2. Light provided by a bank of 40-W cool-white fluorescent bulbs of the standard type involving two lamps per ballast and providing about 3700 lux light intensity. Pairs of lamps are spaced at lO-cm intervals and placed vertically (against a wall) 30 cm behind the culture vessels or, alternatively, placed horizontally 30 cm above horizontal shelves for petri dishes or culture vessels. These spacings provide an irradiance of between 200 and 300 !J.E per cm2 of photosynthetically active radiation per second at the surface of the culture vessel or petri dish, depending on the age of the bulbs. 3. Two standard photoperiods are employed about equally often. These are 12-h light/12-h dark and 16-h lightl8-h dark. Special culture methods, including continuous fermentor culture and light-temperature gradient plate culture, are given by Starr (in Stein, 1973). Large-scale, outdoor mass culture is described by Knutsen and Metting (1991). 21-5.1.3 Culture Media

Selected basal culture media are described below for cultivation of terrestrial (and freshwater planktonic) microalgae. Many additional media are in common use as preferred by individual researchers. The recipes for many of these are listed by Borowitzka (1988), Castenholtz et al. (1989), Starr (1978), and Stein (1973). Any of the following media can be used as the base for an agar (solid) medium. Additionally, certain compounds and antibiotics can be added to discourage the growth of various groups of microorganisms, as follows:

449

ALGAE AND CYANOBACTERIA

1. To discourage the growth of heterotrophic bacteria in enrichment of soil samples, add 1 mL of the following filter-sterilized antibiotic stock solution per 100 mL of algal medium: 0.6 g of penicillin (1625 units mg- 1) and 1.0 g of streptomycin sulfate in 200 mL of deionized-glass distilled water. 2. To suppress diatom growth, add 1 to 10 mg Ge 20 L -1 of medium. 3. To suppress the growth of eukaryotic microalgae when enriching for cyanobacteria, add 20 to 50 mg cycloheximide L -1 of medium. 4. To avoid discouraging the growth of unicellular, N2 -fixing cyanobacteria, solidify the medium with no more than 1% agar. 21-5.1.3.1 BG-ll Medium for Cyanobacteria (Allen, 1968). To 1 L of deionized, glass-distilled water, add: NaN0 3 K2 HP0 4 MgS0 4·7H20 CaCI2·2H20 Citric acid Ferric ammonium citrate Na2-EDTA Na2C03 Trace element solution

1.5 g 0.04 g 0.075 g 0.036 g 0.006 g 0.006 g 0.001 g 0.02 g 1.0 mL

Trace element solution (to make 1 L of stock solution): H 3B0 3 MnCI 2·4H20 ZnS04·7H20 Na 2 Mo0 4·2H2 0 CuS04·5H20 Co(N0 3 h6H 20

2.86 1.81 0.222 0.39 0.079 0.494

g g g g g g

Adjust to pH 7.1 after autoclaving. Some species of cyanobacteria require vitamin B12 which can be added after autoclaving to a concentration of 1 ~g L -1. Partial enrichment for N2-fixing cyanobacteria is possible by eliminating NaN0 3 from the medium. It is then known as BG-ll o medium. Solid BG-ll medium is prepared with 10 to 15 g of agar per liter. 21-5.1.3.2 Bold's Basal Medium for Green and Yellow-green Microalgae (Starr, 1978). This medium is also suitable for many cyanobacteria. Diatoms do not grow well in this medium (J.R. Johansen, 1992, personal communication). Prepare the five individual macro nutrient solutions (listed in a), one chelate solution (either of those in b), and three minor element stock solutions (c, d, and e). To 940 mL of deionized, glass-distilled water, add 10 mL of each of the macro nutrient solutions and 1 mL of each of the EDTA and minor element solutions. Final pH is 6.6.

METTING

450

a. Macronutrient solutions: NaN0 3 10 g 400 mL -1 K2HP0 4 3.0 g 400 mL -1 KH2P04 7.0 g 400 mL-1 3.0 g 400 mL-1 MgS0 4·7H20 CaCI2·2H20 1.0 g 400 mL -1 b. EDTA solution 50 g L-1 Na2-EDTA 31 g L-1 KOH c. Iron solution 4.98 g L-1 FeS04·7H20 1 mL L-1 conc. H 2S04 d. Boron solution 11.42 g L-1 H 3B0 3 e. Micronutrients 1.44 g L-1 MnCl 2 ·4H2 0 8.82 g L-1 ZnS04·7H20 0.71 g L-1 Mo03 1.57gL-1 CuS04·5H20 0.49 g L-1 Co(N0 3h·6H20 Solid Bold's basal medium is prepared with 10 to 15 g agar L -1. 21-5.1.3.3 Medium for Diatoms: Guillard's (We) Freshwater Enrichment Basal Salt Mixture (Guillard, 1975). This all-purpose medium for freshwater algae can be modified to partially enrich for diatoms by (i) providing silicon for cell wall (frustule) formation, and (ii) providing a wide NP ratio to take advantage of the fact that many (though not all) diatoms have efficient P uptake systems compared to other microalgae. The basal medium and the vitamin mixture are both available commercially from Sigma Chemical Company, St. Louis, MO (Sigma Plant Cell Culture Catalogue). To enrich for diatoms, eliminate the NaHC03 and reduce the concentration of K2HP0 4 lO-fold (S.S. Kilham, 1992, personal communication). Prepare the following six individual macronutrient stock solutions and seven minor element solutions. Add 1 mL of each of the macronutrient and micronutrient solutions to 1000 mL of deionized, glass-distilled water. Vitamin stocks should be membrane-filtered (or autoclaved) and stored frozen. Substitution of 200 mg L -1 of TES or PIPES for TRIS is recommended for diatom culture as is the addition of 10 mg H 3B0 3 L -1 (S.S. Kilham, 1992, personal communication).

a. Macronutrient solutions NaN03 K2HP0 4 MgS0 4·7H20 CaCl2 ·2H2 0 NaHC03 Na2Si03·9H20

85.01 8.71 36.97 36.76 12.60 28.42

g L-1 gL-1 g L-l g L-1 g L-1 g L-1

ALGAE AND CYANOBACTERIA

451

e. Micronutrients 0.18 g L-I MnCI 2·4H20 0.022 g L-I ZnS0 4·7H20 0.006 g L-I Na2Mo0 4·2H20 0.01 g L-I CuS04·5H20 0.01 gL-I CoCl2·6H20 3.15 gL-I FeCI3 ·6H20 4.36 g L-I Na2·EDTA c. Vitamins Thiamine·HCl 0.01 mg L-I Biotin 0.5 [.tg L-I Cyanocobalamin 0.5 [.tg L-I d. TRIS solution-use 2 mL L -I of medium. 50 g 200 mL-I Tris(hydroxymethyl)-aminomethane 21-5.1.3.4 Soil Extract Medium (Starr, 1978). A soil extract can be used in place of the defined microelement, chelate, and organic constituents in the media described above. It is often used to improve the likelihood of enriching for micro algal species with undefined growth factor requirements that are supplied by soil. Soil extract is prepared as follows: 1. Place about 2 g of CaC0 3 on the bottom of a 2-L flask and add a layer of a fertile air-dried garden soil to a depth of about 3 to 5 cm. 2. Slowly add deionized glass-distilled water down the inside wall of the flask so as not to disturb the soil, until the flask is about twothirds to three-fourths full. 3. Submit the flask to fractional.sterilization (tyndallization) for 2 h in a steam chamber on two successive days. Store the flasks under refrigeration and allow the supernatant to clear for 7 d before use. To use, add 40 mL of the soil water supernatant to 960 mL of any defined medium. The supernatant may be added before the medium is autoclaved or as a filter-sterilized supplement after autoclaving. Solid soil extract medium is prepared with 10 to 15 g agar L -1. 21-5.1.3.5 Soil Water Tubes (Starr, 1978). Soil water tubes are used with the MPN method (section 21-3.2) when a quantitative index of the complete microalgal community is desired in place of enrichment for selected components. Soil water tubes are also suitable for the short-term maintenance of many microalgal species. Soil water tubes are prepared as a simple modification of the procedure outlined immediately above (section 21-5.1.3.4). Place 1 to 2 cm of a fertile air-dried garden soil over a pinch of CaC03 in the bottom of a series of 15-mL culture tubes. Slowly add deionized glass-distilled water down the inside wall to within 4 to 5 cm of the top of the individual tubes so as not to disturb the soil. Submit racks of the tubes to fractional sterilization (tyndallization) for 2 h in a steam chamber on two successive days. Store the tubes under refrigeration, allowing the supernatants to clear for 7 d before use.

452

METTING

21-5.1.3.6 Organic Carbon and Energy Sources. It is often possible to isolate obligately or facultatively photoheterotrophic or heterotrophic microalgae in liquid or on agar plates where sufficient concentrations of suitable organic compounds remain following serial dilution of the soil. Microalgal cultures that grow poorly (or not at all) when inoculated into defined mineral salts media may require the provision of one or more organic compounds from the environment. For further determination of the nutritional status of these cultures, it is recommended that a subjective comparison be made between growth in one or more defined mineral media without provision of vitamins, the same media with vitamins, the same media with undefined complex organic compound mixtures (0.2% yeast extract or casamino acids), and with growth in a soil extract medium or in soil water tubes. Microalgae and cyanobacteria grow much slower than bacteria or fungi under photoheterotrophic and heterotrophic conditions. For this reason, it is imperative to use only axenic cultures to study patterns of utilization of organic C and energy sources in the light or in the dark. Organic compounds of interest should be dissolved in deionized glass-distilled water and filter-sterilized through sterile 0.22- or 0.45-!! membrane filters. A wide range of concentrations or organic compounds are used to study heterotrophy in microalgae and cyanobacteria (Neilson, Blankley, and Lewin in Stein, 1973). Any defined organic compound can be tested as a potential source of organic C or energy. Standard methods for the taxonomic treatment of eukaryotic microalgae from soil include screening for utilization of 0.25 M glucose, fructose, ribose, xylose, mannose, Na-acetate, and Napyruvate in Bold's basal medium in the light and in the dark. Tests for the production of extracellular amylase and gelatinase are also standard for taxonomic determinations at the species level (Archibald & Bold, 1970; Metting, 1980). 21-5.2 Storage and Preservation Methods 21-5.2.1 Short-Term Storage Methods

Methods for storage and preservation of micro algal and cyanobacterial cultures are similar to those used for other microorganisms. For shortterm storage (between 4 and 10 wk), cultures should be streaked onto agar slants in screw-cap culture tubes. The length of storage time possible with this method will vary depending on storage conditions and rate of desiccation of the agar medium. The time between transfers of cultures to fresh agar slants can be lengthened by: 1. Storage at reduced levels of light. This is accomplished within the

walk-in culture room by placing racks of cultures on agar slants on a separate set of shelves that are removed from the light source(s). A sheet of thin, translucent polyethylene plastic sheeting can be

ALGAE AND CYANOBACTERIA

453

draped in front of the shelves to further reduce irradiance and minimize airflow that would otherwise hasten desiccation of the agar. 2. Wrapping a piece of domestic plastic wrap or laboratory wax paper (Para-Film) tightly around the screw cap. 3. Storage at reduced temperature. Most microalgae will tolerate long periods of refrigeration (3-6 mo or more) when stored as agar slants or in a liquid medium. Soil water tubes are useful for this purpose. Some filamentous green and yellow-green microalgae (e.g., Zygnema and Vaucheria) prefer the medium if the CaC0 3 is omitted. Euglena and other euglenoids prefer soil-water tubes to which half of a pea (Pisum sativum L.) cotyledon is added (Starr, 1978). 21-5.2.2 Long-Term Preservation Methods Many micro algae and cyanobacteria can be preserved for indefinite periods by drying, freezing, or freeze-drying (lyophilization) (HolmHansen in Stein, 1973). Drying and storage at room temperature is not recommended, but it should be noted that Nostoc and other filamentous cyanobacteria have been revived following many decades of storage as specimens on herbarium paper in botanical collections (Metting, 1981). Freezing with cryoprotective agents such as glycerol or dimethyl sulfide is appropriate for microalgae. For long-term preservation by freezing, a deep freeze at -25°C is satisfactory for some species. For other cultures, cryogenic freezing at or below -40 °C is necessary. Individual cultures must be tested to determine what conditions are necessary. Species that can successfully be frozen without excessive cellular damage or death can be stored for many months. Methods used to freeze-dry bacteria are appropriate for microalgae. Not all species, however, are equally tolerant of the procedure and each must be tested to determine suitability. In some cases, iterative modification of the standard method can successfully identify conditions specific for a given species. Species that can successfully be freeze-dried can be stored for many years. Indeed, there is evidence that cryptoendolithic communities in the dry interior valleys of Antarctica undergo cycles of natural lyophilization and may be many thousands of years old (Johnston & Vestal, 1991). 21-6 METHODS FOR ESTIMATING PHOTOSYNTHESIS Photosynthesis (C0 2 fixation) by communities of soil algae is estimated by the same methods outlined for soil respiration described in chapter 38 in this book. The contribution of algal photosynthetic production to soil respiration is estimated by comparing CO 2 or O 2 uptake and evolution in the light and dark and assuming that any difference is due to photosynthesis. The value of estimates from in situ (field) measurements depends

454

METTING

largely on the size of the algal community. Algal contribution is easily masked by soil respiration except in situations where a large, visible crust is present. Better estimates of potential photosynthetic contributions by algal communities are achieved by studying thin (millimeters to centimeters) surface veneers from soil cores brought into the laboratory (Shimmel & Darley, 1985). 21-7 METHODS FOR MEASURING CYANOBACTERIAL DINITROGEN FIXATION Most cyanobacteria are capable of biological N2 fixation. Species that form specialized cells called heterocysts are capable of N2 fixation in aerobic environments. Also, many filamentous and aggregate-forming species that do not have heterocysts can fix N in aerobic environments, but only in localized microaerophilic microsites within aggregates of cells that are protected from excessive O 2 diffusion by encapsulating mucilages. Other nonheterocystous cyanobacteria with the capacity to fix N2 can do so only under strictly anaerobic conditions (Castenholtz et al., 1989). The methods outlined in chapter 43 for direct (15N) and indirect (C2H 2 reduction) quantification of bacterial N2 fixation should also be used for cyanobacteria. 21-8 METHODS FOR STUDYING ENDOSYMBIOTIC CYANOBACTERIA IN CYCAD ROOTS 21-8.1 Principles Cyanobacteria form symbiotic relationships and fix N2 in association with liverworts, fungi (lichens), Gunnera, the aquatic fern Azolla, and cycads. Gunnera is a genus of tropical flowering herbs mostly occurring at high elevations in which cyanobacteria inhabit glandular stem cells. As mentioned in the introduction, the Azolla-Anabaena symbiosis is important to flooded rice culture. Lumpkin and Plucknett (1980) have reviewed methods for studying this symbiosis. Most cyanobacteria that form intercellular bands of filaments in coralloid roots of cycads are in the genera Anabaena and Nostoc. The association is common to the cycad genera Zamia, Macrozamia, Encephalartos, and Cycas. In some symbioses, the cyanobacteria are localized in root tissue at or near the surface where they are exposed to light and conduct photosynthesis. In others, they are excluded from light and are accordingly dependent on other mechanisms for provision of reductant and ATP from the host. Unlike nodules formed in rhizobial and Frankia symbioses, endosymbiotic cyanobacteria reside in normally differentiated host tissue (with the exception of Gunnera). In common with other bacterial symbionts, however, endosymbiotic cyanobacteria do display several structural, ultrastructural, and physiological alterations from the free-living form (Grilli-Caiola, 1975).

ALGAE AND CYANOBACTERIA

455

Methods for studying the symbiosis are fundamentally similar to those outlined in chapter 12 for legume symbionts. Although whole plant experimentation is complicated by the relatively larger size, woody nature, and much slower growth rates of cycads compared to herbaceous legumes, excision of roots or of root segments and isolation and cultivation of cyanobacteria from root nodules is relatively simple and routinely accomplished. Methods for in situ direct (Nl5) or indirect (C 2H 2 reduction) measurement of N2 fixation are identical to those described in chapter 43 for N2-fixing legumes. 21-8.2 Procedure for CoraIloid Roots and Root Sections Actively growing coralloid roots or root sections are collected from natural cycad populations by digging one or a series of shallow excavations around mature plants and cutting off desired segments (Lindblad et al., 1991). Whole roots or sections can also be collected from smaller plants in the field or from a greenhouse collection by removing the entire root mass from the soil and selectively cutting desired whole roots or segments. Transport the roots to the laboratory, preferably in an inert N2 or Ar atmosphere. Prepare a series of I-mm-thick sections under N2 or Ar in degassed 10 mM HEPES buffer (pH 8.0). Determine the wet weights of the sections. Analyze the sections for N2-fixation potential using the methods outlined in chapter 43. 21-8.3 Procedure for Isolating Endosymbiotic Cyanobacteria Wash healthy coralloid roots or root sections under running tap water. Blot the roots dry, and cut them transversely into l-cm lengths. Surface sterilize the sections by immersion in 80% ethanol (2 min), then rinse in deionized glass-distilled water (2 min), 2.5% Na-hypochloride (2 min), and again in deionized glass-distilled water (7 min). Cut the l-cm sections into I-mm sections and suspend them in BG 11 medium (section 21-5.1.3.1) in a petri dish in the light. The cyanobacterial filaments will migrate into the medium from the roots (Grobbelaar et al., 1987). Individual filaments can then be isolated and purified into clonal culture by repeated washing (section 21-4.2), streaking on agar (section 21-4.3), or centrifugation (section 21-4.4).

ACKNOWLEDGMENT The following people made important contributions to this chapter or reviewed and commented on the manuscript: Jeffrey R. Johansen, John Carroll University (diatom frustule preparation, fluorescence microscopy). Susan S. Kilham, Drexel University (diatom culture). William J. Zimmerman, University of Michigan-Dearborn (symbiotic cyanobacteria). Pierre

METTING

456

Roger, ORSTOM, University of Marseilles (methods for rice soils). E. Imre Friedmann, Florida State University (methods for endolithic habitats). Annette E. Pipe, Lakeland College (implanted slide method). REFERENCES Allen, M.M. 1968. Simple conditions for growth of unicellular blue-green algae on plates. J. Phycol. 4:1-3. Anderson, D.C., and S.R. Rushforth. 1976. The cryptogamic flora of desert soil crusts in southern Utah, U.S.A. Nova Hedwigia 28:691-729. Archibald, P.A., and H.C. Bold. 1970. Phycological studies XI. The genus Chlorococcum Meneghini. University of Texas Publ. no. 7015, Austin. Ashley, J., S.R. Rushforth, and J.R. Johansen. 1985. Soil algae of cryptogamic crusts from the Uintah Basin, Utah, U.S.A. Great Basin Nat. 45:432-442. Bell, R.A., and M.R. Sommerfeld. 1987. Algal biomass and primary production within a temperate zone sandstone. Am. J. Bot. 74:294-297. Bock, W. 1963. Diatomeen extrem trockener Standorte. Nova Hedwigia 5:199-254. Bold, H.C., and M.J. Wynne. 1985. Introduction to the algae. 2nd ed. Prentice-Hall, Englewood Cliffs, NJ. Borowitzka, M.A. 1988. Algal growth media and sources of algal cultures. p. 456-465. In M.A. Borowitzka and L.J. Borowitzka (ed.) Micro-algal biotechnology. Cambridge University Press, Cambridge. Castenholtz, R., G.L. Gherna, R. Lewin, R. Rippka, J.B. Waterbury, and B.A. Whitton, 1989. Section 19. Oxygenic photosynthetic bacteria. Group 1. Cyanobacteria. p. 17101799. In J.T. Staley led.) Bergey's manual of systematic bacteriology. Vol. 3. Williams and Wilkins, Baltimore. Ettl, H. 1983. Chlorophyta I. Phytomonadina. 9 Teil. In H. Ettl et al. (ed.) SiiBwasserflora von Mitteleuropa. Gustav Fischer Verlag, Stuttgart, Germany. Ettl, H. 1978. Xanthophyceae. 1 Teil. In H. Ettl et al. (ed.) SiiBwasserflora von Mitteleuropa. Gustav Fischer Verlag, Stuttgart, Germany. Gantt, E. 1980. Handbook of phycological methods. Vol. 3. Developmental and cytological methods. Cambridge Umversity Press, Cambridge. Geitler, L. 1930-32. Cyanorhyceae. In L. Rabenhortst's Kryptogammenflora. Reprinted in 1985 by Koeltz Scientific Books, Koenigstein, Germany. Grilli-Caiola, M. 1975. A light and electron microscopic study of blue-green algae growing in the coralloid-roots of Encephlartos altensteinii and in culture. Phycologia 14:25-33. Grobbelaar, N., W.E. Scott, W. Hattingh, and J. Marshall. 1987. The identification of the coralloid root endophytes of the southern African cycads and the ability of the isolates to fix dinitrogen. S. Afr. J. Bot. 53:111-118. Gui\lard, R.R.L. 1975. Culture of phytoplankton for feeding marine invertebrates. p. 29-68. In W.e. Smith and M.H. Chanley led.) Culture of marine invertebrate animals. Plenum Press, New York. Hellebust, J.A., and J.S. Craigie. 1978. Handbook of phycological methods. Vol. 2. Physiological and biochemical methods. Cambridge University Press, Cambridge. Johansen, J.R., A. Javakul, and S.R. Rushforth. 1982. Effects of burning on the algal communities of a high desert soil near Wallsburg, Utah. J. Range Manage. 35:598-600. Johansen, J.R., S.R. Rushforth, and J.D. Brotherson. 1981. Subaerial algae of Navajo National Monument, Arizona. Great Basin Nat. 41:433-439. Johansen, J.R., L.L. St. Clair, B.L. Webb, and G.T. Nebeker. 1984. Recovery patterns of cr~ptogamic soil crusts in desert rangelands following fire disturbance. Bryologist 87.238-243. Johnston, C.G., and J.R. Vestal. 1991. Photosynthetic carbon incorporation and turnover in Antarctic cryptoendolithic communities: are they the slowest-growing communities on earth? Appl. Environ. Microbiol. 57:2308-2311. Kessler, J.O. 1984. A new method for concentrating and purifying swimming algae. Appl. Phycol. Forum 1:2-4. Komarek, J., and N. Fott. 1983. Chlorophyceae (Griienalgen) Ordung: Chlorococcales. In G. Huber-Pestalozzi (ed.) Das Phytoplankton des SiiBwassers, Band 26, Teil 7, Heft. 1. E. Schweizerbart'sche Verlagsbuchhandlung, Stuttgart, Germany.

ALGAE AND CYANOBACTERIA

457

Knutsen, G., and F.B. Metting. 1991. Microalgal mass culture and forced development of biological crusts in arid lands. p. 487-506. In J. Skujins (ed.) Semiarid lands and deserts. Soil resource and reclamation. Marcel Dekker, New York. Krammer, K., and H. Lange-Bertalot. 1986. Bacillariophyceae. 1 Teil. Naviculaceae. In H. Ett!. et a!. (ed.) SiiBwasserflora von Mitteleuropa 9. Gustav Fischer Verlag, Stuttgart, Germany. Krammer, K., and H. Lange-Bertalot. 1988. Bacillariophyceae. 2 Tei!. Bacillariaceae, Epithemiaceae, Surirellaceae. In H. Ettl et a!. (ed.) SiiBwasserflora von Mitteleuropa. Gustav Fischer Verlag, Stuttgart, Germany. Leedale, G.F. 1967. Euglenoid flagellates. Prentice-Hall, Englewood Cliffs, NJ. Lindblad, P., C.A. Atkins, and J.S. Pate. 1991. N2 -fixation by freshly isolated Nostoc from coralloid roots of the cycad Macrozamia riedlei (Fisch. ex Gaud.) Gardn. Plant Physio!. 95:753-759. Littler, D., and M. Littler. 1985. Handbook of phycological methods. Vo!. 4. Ecological field methods: Macroalgae. Cambridge University Press, Cambridge. Lorenzen, C.F. 1967. Determination of chlorophyll and pheo-pigments: septrophotometric equations. Limno!. Oceanogr. 12:343-346. Lumpkin, T.A., and D.L. Plucknett. 1980. Awlla: botany, physiology, and use as a green manure. Econ. Bot. 34:111-153. Metting, F.B. 1980. New species of green microalgae (Chlorophycophyta) from an eastern Washington silt loam. Phycologia 19:296-306. Metting, F.B. 1981. The systematics and ecology of soil algae. Bot. Rev. 41:195-312. Metting, F.B. 1991. Biological surface features of semiarid lands and deserts. p. 257-293. In J. Skujins (ed.) Semiarid lands and deserts. Soil Resource and Reclamation. Marcel Dekker, New York. Metting, F.B. 1993. Structure and physiological ecology of soil microbial communities. p. 3-25. In F.B. Metting (ed.) Soil microbial ecology. Applications in agriculture and environmental management. Marcel Dekker, New York. Metting, F.B., and A.D. Barry. 1986. Palmelloid microalgae as soil-conditioning agents. Phase I Final Project Report 85-SBIR-8-0067. USDA, Washington, DC. Metting, F.B., WR. Rayburn, and P.A. Reynaud. 1988. Algae and agriculture. p. 335-370. In C.A. Lembi and J.R. Waaland (ed.) Algae and human affairs. Cambridge University Press, Cambridge. Pipe, A.E. 1992. Pesticide effects on soil algae and cyanobacteria. Rev. Environ. Contamination Toxico!. 127:95-170. Pipe, A.E., and D.R. Cullimore. 1980. An implanted slide technique for examining the effects of the herbicide diuron on soil algae. Bull. Environ. Contamination Toxico!. 24:306-312. Prescott, G.W.1978. How to know the freshwater algae. 3rd ed. Wm. C. Brown, Dubuque, IA. Rayburn, W.R., R.N. Mack, and F.B. Metting. 1982. Conspicuous algal colonization of the ash from Mount St. Helens. J. Phyco!. 18:537-543. Roger, P.A., Jimenez, R., and Santiago-Ardales, S. 1991. Methods for studyinS blue-green algae in ricefields: distributional ecology, sampling strategies, and estimatIOn of abundance. IRRI Res. Paper Ser., No. 150. Int. Rice Res. Inst., Los Banos. Manila, Philippines. Roger, P.A., and S.A. Kulasooriya. 1980. Blue-green algae and rice. Int. Rice Res. Inst., Los Banos, Manila, Philippines. Roger, P.A., W.J. Zimmerman, and T.A. Lumpkin. 1993. Microbiological management of wetland rice fields. p. 417-455. In F.B. Metting (ed.) Soil microbial ecology. Applications in agriculture and environmental management. Marcel Dekker, New York. Sharabi, N.E.-D., and D. Pramer. 1973. A spectrophotometric method for studying algae in soi!. Bull. Eco!. Res. Commun. NFR Sweden 17:224-227. Shields, L.M. 1982. Algae. p. 1093-1101. In A.L. Page et a!. (ed.) Methods of soil analysis. Part 2. Chemical and microbiological properties. 2nd ed. ASA and SSSA, Madison, WI. Shimmel, S.M., and W.M. Darley. 1985. Productivity and density of soil algae in an agricultural system. Ecology 66:1439-1447. Starmach, K. 1972. Chlorophyta III. Zielenice nitlowate. In K. Starmach and J. Sieminska (ed.) Flora Slodkowodna Polski, Tom. 10. Polska Akad. Nauk, Inst. Bot., Warsaw, Poland. Starr, R.C. 1978. The culture collection of algae at the University of Texas at Austin. J. Phyco!. 14 (supplement):47-100.

458

METTING

Stein, J.R. 1973. Handbook of phycological methods. Vol. 1. Culture methods and growth measurements. Cambridge University Press, Cambridge. Tchan, Y.T. 1952. Study of soil algae. I. Fluorescence microscopy for the study of soil algae. Proc. Linn. Soc., London 77:265-269. Van Baalen, C. 1965. Quantitative surface plating of coccoid blue-green algae. J. Phycol. 1:19-22.

Published 1994

Chapter 22 Nematodes RUSSELL E. INGHAM, Oregon State University, Corvallis, Oregon

Nematodes are unsegmented roundworms and one of the most ecologically diverse groups of animals on earth. They exist in nearly every habitat known from the tops of mountains to the depths of the ocean and from hot deserts to the cold of the Antarctic. Nematodes eat bacteria, fungi, algae, yeasts, diatoms, and may be predators of several small invertebrate animals, including other nematodes. In addition, they may be parasites of invertebrates, vertebrates (including humans) and all above and belowground portions of plants. Nematodes range in length from 82!Lm (marine) to 9 m (whale parasite) but most species in soil are between 0.25- and 5.5-mm long. Nematodes may constitute as much as 90% of all multicellular animals in soil (Oostrinbrink, 1971) and often exceed several million m- 2 (Table 21-1). While the total biomass of nematodes may often be less than that for other faunal groups (Coleman, 1976; Kusmin, 1976; MacLean, 1974) nematode metabolic activity is often higher (Kusmin, 1976; Reichle, 1977; Sohlenius, 1977). Although nematodes are recognized as a major consumer group in soils, the exact feeding habits of most soil nematodes are not known. They are generally grouped into four to five trophic categories based on the nature of their food, the structure of the stoma and esophagus and method of feeding (Yeates, 1971). Plant-feeding nematodes possess stylets with a wide diversity of size and structure and are the most extensively studied group of soil nematodes because of their ability to cause plant disease and reduce crop yield. Fungal-feeding nematodes have slender stylets but are often difficult to categorize and have been included with plant feeders in many ecological studies. Bacterial-feeding nematodes are a diverse group and usually have a simple stoma in the form of a cylindrical or triangular tube, terminating in a valve-like apparatus that may bear minute teeth (Nicholas, 1975). Predatory nematodes are usually large species possessing either a large stylet or a wide cup-shaped cuticular-lined stoma armed with powerful teeth (Nicholas, 1975). Omnivores are sometimes considered as Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 459

INGHAM

460

Table 22-1. Density, biomass, and respiration estimates for nematode communities in selected ecosystems. (After Sohlenius, 1980.) Ecosystem Desert, USA Tundra, Sweden Beech forest, England Tropical forest, Africa Pine forest, Sweden Mixed forest, Poland Pasture, Poland Grassland, Denmark Potato field, Poland Rye field, Poland

Numbers

Biomass

Respiration

millions/m2 0.4 4.1 1.4 1.7 4.1 7.0 3.5 10.0 5.5 8.6

g fresh wtlm 2 0.1 1.1 0.4

Kcallm2 .y 7.6 7.3 5.2

0.6 0.7 2.2 14.0 0.7 1.1

8.2 33.0 50.0 339.0 14.3 6.0

a fifth trophic category of soil nematodes. These nematodes may fit into one of the categories above but also ingest other food sources. For example, some bacterial feeders may also eat protozoa or algae and some styletbearing nematodes may pierce and suck algae as well as fungi or higher plants. Stages of animal-parasitic nematodes, such as hookworms, may also be found in soils but generally are not common in most soil samples. Nematodes appear to have several important functions in soil ecosystems. Most notably, they are important pests of agricultural crops. However, even in native ecosystems, plant consumption by nematodes can be substantial, exceeding that of other herbivores (Ingham & Detling, 1984). The importance of plant consumption by nematodes on system productivity has not been adequately studied in most native ecosystems. However, nematicide studies in native North American grasslands reduced populations of plant-feeding nematodes and increased plant growth (Ingham & Detling 1990; Stanton et al., 1981). Nutrient cycling dynamics may be influenced by nematodes feeding on bacteria and fungi, regulating substrate utilization and the mineralization of nutrients (Ingham et al., 1985b). Nematodes are generally most abundant in the rhizosphere (Ingham & Coleman, 1983; Ingham et aI., 1985a) and thus may interact with other rhizosphere organisms such as plant pathogens (Sikora & Carter 1987; Riedel, 1988), rhizobia (Huang, 1987), mycorrhizae (Ingham, 1988), and other nematodes (Eisenback & Griffin, 1987). Nematodes also fill an important link in the soil food web, transferring plant and microbial-based energy and nutrients to higher trophic categories in the soil food web (Hunt et aI., 1987). Thus, nematodes may be studied by soil biologists from several different disciplines. The most important methods for adequate study of soil nematodes in the field relate to: (i) taking appropriate samples, (ii) reliable extraction procedures for recovering nematodes from soil or plant tissues, and (iii) enumeration and identification of the nematodes recovered.

461

NEMATODES

22-1 NEMATODE SAMPLING

The first consideration in sampling for nematodes is to design a sampling plan that is appropriate for the current objectives. For example, the sampling procedure for describing the nematode community from a particular species of plant may be quite different from that for determining if a fallow field must be treated with nematicide before a crop is planted. Available resources may also influence sampling intensity. Researchers may be able to afford to process more samples than a grower trying to diagnose the cause of plant disease. Most studies to evaluate procedures for sampling nematodes have only been concerned with estimating populations of plant-parasitic nematodes. The protocols for collecting nematode samples have been thoroughly reviewed (Barker et al., 1985a; McSorley, 1987) and are beyond the scope of this discussion. Several principles are necessary to mention, however, and many apply to sampling for other soil organisms and abiotic properties as well. 22-1.1 Nematode Distribution

Nematode species are not uniformly distributed throughout the soil but generally have a patchy distribution with areas of high densities mixed among areas of low densities (Fig. 22-1). Nematode species are often distributed differently within the same field so that areas of high density of one species may not correspond with high densities of another species (Goodell & Ferris, 1980, 1981). Nematode distribution and density may be related to soil type (e. g., high in sandy areas and lower in heavier soil), some other edaphic factor, past cropping histories or be unrelated to any apparent parameter. Inadequate sampling can result in unreliable estimates of nematode density and distribution. Inaccurate estimates can lead to expensive errors if they are used as the basis for nematode management decisions, or result in the development of errant concepts if used to describe ecological principles. The structure of the root system influences distribution of nematode populations. In annual cropping systems, the structure of preceding crops, as well as the current crop, may influence nematode distribution. Patterns of spatial distribution can be described by taking a series of soil cores and computing the mean and variance. Most distributions can be described by the negative binomial functions indicating clumped distributions. Thus, data must be transformed by normalizing functions such as Y = log(x + c) [where Y is the transformed count, x is the original count and c is a constant, usually 1] before analysis (McSorley, 1987). The density of a nematode species in soil will also affect the precision of the population estimate made from any sampling plan. If a system is to be sampled frequently, initial extensive sampling can determine the index of dispersion (k) (Southwood, 1978) associated with negative binomial distributions and assist development of an optimum sampling design for future samplings (McSorley, 1987).

INGHAM

462

5] FINE

TEXTURED

~~:::-2~~~.:)-,.;T.;;JJ~~I~IIIIIIII.fll~C:OARSE

TEXTURED

100

o

Fig. 22-1. Three-dimensional depiction of the distribution of Helicotylenchus digonicus (B) and Meloidogyne arenaria (C) in relation to soil texture (A) in an alfalfa field. (After Goodell and Ferris, 1980.)

22-1.2 Sample Collection Nematode samples may be taken for many reasons. A bulk sample of several soil cores may be taken at one time to determine which nematode species are present. Many individual cores can be analyzed to map the population distribution in that area. Other areas may be sampled repeatedly throughout the year to determine the occurrence of temporal changes in the populations of one or several species. In all cases, however, the biology of the nematode species of interest will dictate the time, depth, and pattern of the sampling design. The primary objectives for taking nematode samples include: (i) detection and survey, (ii) diagnosis, (iii) advisory, and (iv) research (Barker & Campbell, 1981). When nematode samples are taken for general survey purposes, or, to make collections for taxonomic work, there may be little concern with the precision of the sampling design. However, if the sampling objective is for quantification of populations, quarantine, or other regulatory purposes, sampling may need to be thorough to assure detection of serious pathogens. Nematodes samples

NEMATODES

463

are generally taken with a cylindrical soil probe (generally 2.5 cm diam.) but shovels, trowels, or augers may be used as well. Sampling depth depends on the rooting pattern of the crop being sampled but is generally 30 to 45 cm. A common objective for nematode sampling is to aid diagnosis of poor plant growth. If a crop is present and patches of poorly growing plants are observed, it is advisable to take separate samples from the poorly growing patches and from areas that appear healthy. This may confirm that nematodes should be implicated as a cause for the poor plant productivity if densities are high in the patches of poor growth and less in the healthy appearing areas. It is often preferable to take samples from the periphery of the patch of poor plant growth than from the center of the patch. Often plant growth in the center of these patches may be so poor that nematode densities will have declined due to inadequate root production. Higher numbers may be obtained at the periphery where populations may be rapidly increasing. When sampling for diagnosis, adequate precision is usually obtained economically by taking several samples of composited soil cores from a relatively small affected area. Another reason for taking nematode samples is to determine if nematode populations are at a level that may damage a crop to be planted or are increasing towards damaging levels in an existing crop (advisory). If one is sampling a fallow field, or a field without visible differences in plant growth, the whole field (or subdivided portions of a large field) must be sampled. The most representative samples are obtained by taking several cores from a given area, compositing the cores into a single sample, mixing the soil thoroughly, and taking a sufficient subsample (for example 1 Lor 1 kg) for analysis. The pattern in which cores are collected is dependent on the cropping system being sampled as illustrated in Fig. 22-2. The number of samples taken from a field depends on the value of the crop and the relative nematode risk (e.g., risk to a high cash value perennial from a nematode that vectors plant viruses would be much greater than the risk to a low cash value annual crop from a nematode that reduces yield slightly). It has been estimated that 1% of crop production costs may be economical for nematode sampling. For example, if it costs $500/acre to produce a crop and $20/sample for nematode sampling and analysis, then $5/acre could be spent on sampling. Thus, each sample should represent four acres (Ferris et aI., 1981a). Southwood (1978) recommended that sampling for advisory purposes should be of sufficient intensity to estimate the population with a standard error within 25% of the mean. However, the level of precision that can be attained may be governed by economic factors such as the value of the crop threatened and the relative pathogenicity of the nematode species of interest. The chances of detecting nematodes and numbers of samples needed at different population densities is discussed by Barker (1985a). Taking nematode samples for research purposes generally requires estimates with a standard error of 10% of the mean (Southwood, 1978).

NEMATODES

465

rooted perennials, often to depths of 0.5 to 1 m. Ingham et al. (1985a) recovered nematodes to 1.5 m in a shortgrass prairie. Extreme climates often encourage vertical migration so sampling to 30 to 45 cm is recommended for regions with hot, dry summers (Barker, 1986). However, some nematodes, such as Meloidogyne chitwoodi, may migrate 1.5 m upward to cause significant crop damage (Mojtahedi et aI., 1991). In these situations, sampling to 90 or 120 cm may be necessary. Most nematode analysis laboratories request that 500 to 1000 cm3 of soil be submitted for analysis. Extraction techniques limit the amount of soil that can be processed so rarely can entire samples be extracted. Large samples should be thoroughly mixed before removing a subsample for extraction. Size of the subs ample depends on the particular procedure used (see below). If soil is thoroughly mixed, only one aliquot is necessary to estimate the population in the sample. Homogeneity of mixing can be tested by extracting replicate aliquots. Care must be taken in mixing, however, to prevent nematode mortality from mechanical damage during mixing (Barker & Campbell, 1981). It is essential that samples be protected from drying out or getting too hot. Soil samples should be sealed in plastic or heavy paper bags in the field and kept in the shade or in an ice chest. Samples should be submitted for analysis as soon as possible. Many extraction and identification procedures require nematodes to be alive and poor sample handling may kill nematodes before the soil reaches the laboratory. This results in an underestimate of the population that was actually present at the time of sampling. Samples should be labeled on the outside of the bag. Labels placed next to soil often disintegrate or become illegible. Samples should be extracted as soon as possible and stored at 10 to 15°C until extracted (Barker & Campbell, 1981). If endoparasites are suspected, roots (or aboveground plant parts for stem, foliar, or seed-gall nematodes) should be collected, sealed in plastic bags, and sent for analysis. Plant material containing nematodes should be kept cool and moist and examined as soon as possible. Whole plants are best stored free from soil. Since shoots often decompose more quickly than roots, they should be kept in separate bags if stored for more than a day or two. Polyethylene bags are excellent storage containers. 22-1.3 Sampling Pattern Several parameters must be considered before designing a pattern for collecting nematode samples. Goodell and Ferris (1981) examined sampling patterns that included random, stratified random and division of the field into north-south or east-west strips and evaluated each pattern with a predetermined time (cost) constraint. In this study, the optimum pattern was division of the field into north-south strips because the strips in this direction isolated a streak of fine-textured soil. Thus, if information about distribution of soil type, or other edaphic factors, is known it may be used to stratify sampling when designing a sampling pattern.

INGHAM

466

In most systems, the sampling pattern will be determined by the structure of the plant community. If the area is fallow or has a homogeneous plant community such as pasture or alfalfa, a simple random or stratified random pattern may be best. Samples are generally taken by walking in a "W" pattern and taking cores at intervals along the traversed path (Fig. 22-2). With row crops, samples should be randomly taken from within the row only and taking cores "across" different rows rather than "along" the same rows provides a more accurate estimate of the overall field mean (Noe & Barker, 1982). In systems where plants are grouped as discrete individuals, such as orchards, samples should be sampled from the drip line since it represents the area occupied by feeder roots and generally contains the highest population of nematodes (Barker, 1986). In a study of vineyards, Ferris and McKenry (1976) found that nematode densities were greatest and variation least when samples taken 30 to 45 cm from the vine and to a depth of 60 cm. When sampling for several nematode species the sampling design should be based on the most difficult species to estimate (Goodell & Ferris, 1981; McSorley & Parrado, 1982). If the same area is sampled repeatedly, selecting cores from the same location (e.g., same plants in an orchard, same distance from stem) each time will reduce the variance associated with nematode distribution (Fidler et al., 1959; Goodell, 1982). 22-1.4 Timing of Sampling Collections Densities of most nematodes reach a maximum and minimum at different times of the year. Nematodes in temperate climates tend to reach minimum population levels in winter or early spring while, in warmer climates, populations may be at their minimum in summer and early fall. Different species cycle in different patterns, however, and may require sampling at different times (Barker, 1985a; Barker & Campbell, 1981). Timing of sampling depends on the sampling objective. Most samples are best taken at time of maximum population to provide maximum probability of detection. This provides information that can be used in management decisions for the subsequent crop. In practice, however, samples are often taken shortly before planting or in the spring when populations may be low so then management options may be implemented before populations increase. In special instances, where nematode infection may damage the quality of a belowground crop, such as potato (Solanum tubarosum), samples may also be take during midseason to determine if the crop may need to be harvested early to escape damage. Sufficient knowledge of the nematode species of interest must be known to optimize the sampling time. If population dynamics are studied, sampling at a minimum of 2-wk intervals are generally necessary to monitor rapid population changes that may occur.

NEMATODES

467

22-1.5 Sample Size The most appropriate method for obtaining accurate nematode sample estimates is with sufficient sample sizes. If the study objectives include precise mapping of the distribution of a nematode community or specific associations between different nematodes or nematodes and other soil organisms, many soil cores may need to be taken and processed individually. However, many sampling objectives do not require this level of precision. Sampling effort can be improved for most objectives, particularly those for advisory purposes, by increasing the number of samples taken and by increasing the number of subsamples or soil cores that comprise a single sample. Although both approaches increase the cost of sampling, it is generally more cost effective to increase the number of cores/sample than to increase the number of samples processed, provided the area represented by a single sample is not too large (Goodell, 1981). The number of cores (subsamples) taken to represent an area depends on the size of the area sampled. For advisory purposes, 10 cores are taken to represent < 1 acre, 20 to 30 cores for 1 to 5 acres and 50 to 100 cores for areas 6 to 10 acres in size. However, it is not generally recommended that one attempt to represent more than 5 acres with a single sample because of the heterogeneous distribution of nematodes (Santo et aI., 1982). Large fields should be subdivided and different samples taken from the subdivisions. Stratification of sampling can minimize variance among repeated sampling from a single stratum while maximizing the differences in densities between strata (Goodell, 1982). Often fields are subdivided based on some natural stratification (Fig. 22-3). This practice allows identification of areas that need to be treated and other areas that may not need to be treated, thus limiting the amount of treatment necessary and reducing treatment costs to the grower. When the expense for more precise estimates may be permissible (e.g., high cash value crop and research studies), the relationship between sample size and level of precision may be used to determine optimum sampling effort if sufficient information about nematode distribution is available (McSorley, 1987). Estimates of the precision expected from multiple samples through subdivision or replicated sampling can be evaluated by computer simulation (Goodell & Ferris, 1981; McSorley, 1982). 22-1.6 Nematode Sampling for Ecological Studies Few quantitative studies have been made on the precision of various sampling plans for ecological studies in native ecosystems. Most principles associated with agroecosystems are likely to apply to native systems as well but the questions addressed may be more complex. While sampling of agroecosystems may be primarily concerned with the macro distribution of

INGHAM

468

1

2

3

5

6

7

ALFALFA

WHEAT

.. , 10

.'

.

".. ,,":


6.5 (Sparling & West, 1990). The manner in which COz is removed from the chamber during incubation has been shown to affect the rate at which it is evolved from neutral to high pH soils (Martens, 1987). This effect was attributed to the increased pCO z in the gas phase during static periods that influenced the aqueous carbonate equilibrium, causing more COz to be dissolved in the aqueous phase. The periodic flushing events of these semi-static systems were not long enough to shift the equilibrium toward the gaseous phase of COz. Constantly aerated systems appear to favor the removal of COz from the aqueous phase in neutral and high pH soils. Accumulating COz or depleting Oz in closed incubation systems may also directly affect metabolic activity. Macfadyen (1973) observed decreased respiration as COz concentrations increased. By controlling rates of aeration, it is possible to prevent conditions that may inhibit or stimulate respiration (Cheng & Coleman, 1989). Another problem that can affect results is the influence of COz trapping on autotrophic processes. Ammonium oxidation was reduced 30% when NaOH traps exceeded 50 mM in a static system (Kinsbursky & Saltzman, 1990). Keeney et al. (1985) reported increasing inhibition of nitrification as COz concentrations were increased from 0.3 to 100%, with no nitrification occurring in 100% COz. Gas exchange rates that affect the balance of COz and 0z in the atmosphere in contact with the soil may affect nitrification and other processes in incubation systems. Excessive air flow rates can intensify pressure leaks in the incubation system and cause siphoning of the base solution when the flow is interrupted during alkali trap changes (Weaver, 1974), both of which result in the loss of COz. Passing the COz-scrubbed air through water to rehydrate it should be monitored cautiously. Without adequate temperature control in the system, condensation of the water vapor may occur, increasing the water content of the soil. Incubation in the laboratory is usually designed to examine aspects of soil metabolism, with little emphasis on reproducing the horizontal characteristics of soil and in situ COz diffusion patterns. Therefore, soil can be distributed in the incubation chamber to minimize the constraints on gas exchange between the soil and atmosphere. This can be done by incubating

CARBON MINERALIZATION

855

taller vessels on their sides, which allows a given amount of soil to be spread out over a larger area (Gilmour & Gilmour, 1985). NaOH and KOH solutions are the most commonly used base solutions to trap respired CO 2 , but Ba(OH)z (Blet-Charaudeau et al., 1990; Witkamp, 1969) and Ba0 2 (Fine et al., 1986) have also been infrequently employed. This may be partly explained by the potential for producing films of BaC03 on the quiescent surface of the base solution that can interfere with further absorption of CO 2 (Blet-Charaudeau et al., 1990). This problem may not be as significant in dynamic systems as it is in static systems since the base is agitated by the bubbling action of the eluting air stream. Further, the use of KOH for 14C work may be compromised by the presence of naturally occurring 40K (D.D. Focht, 1991, personal communication). Methods for radiorespirometry are covered in chapter 39. In choosing combinations of sample size, vessel size, and the other conditions of incubation, the researcher should consider the potential consequences of those choices on the chemical and biological properties of the system that may affect C mineralization. For example, it may be necessary to cover the incubation flasks with an opaque material to preclude photosynthesis by algae and cyanobacteria.

38-4.2 Static Methods for Carbon Dioxide and Oxygen 38-4.2.1 Principles The use of sealed chambers for incubation of soil in the laboratory provides a simple means of determining CO 2 using base absorption, or CO 2 and O 2 if gas chromatography is used for analysis. This method differs from dynamic or flow systems in that a given volume of atmosphere is entrapped above the soil in a closed, non-aerated chamber. Evolved CO 2 is either allowed to accumulate in the headspace of the container for GC analysis (Christensen, 1987; Linn & Doran, 1984; West & Sparling, 1986) or is trapped in base solutions for titrimetric or conductimetric determination (Anderson & Ineson, 1982; Chapman, 1971; Gloser & Tesarova, 1978; Nordgren et al., 1988). Incubation chambers must be opened frequently for aeration to avoid potential problems with unnaturally high CO 2 accumulations or severely depleted O 2 supply in the incubating soil. The emphasis in the following closed-chamber methods is on setup and incubation strategies; the analytical methods for CO 2 and O 2 determinations by gas chromatography and titrimetry having been covered in previous sections.

38-4.2.2 Static Incubation-Gas Chromatographic Analysis \

38-4.2.2.1 Special Apparatus 1. Vessels suitable for incubation may be constructed from commercially available glass mason or preserving jars (about 1 L capacity) with

856

ZmILSKE

threaded rings that seal a rubber-gasketed lid tightly against the lip of the jar. Holes must be drilled through the lids so that they will accept rubber serum vial stoppers. Drill the holes somewhat smaller in diameter than that of the serum vial stopper. Remove defects in the holes by filing so that the rubber stopper is not cut as it is forced through the hole with a twisting motion. The stopper must form an airtight seal with the lid. 2. Rubber serum vial stoppers, 9 mm size; available in bulk from commercial sources. Larger stoppers generally provide longer service than smaller sizes. At least one stopper per vessel must be provided. 3. Gas chromatograph and calibration gases as described in section 38-3.2.1 and 38-3.2.2, respectively. 38-4.2.2.2 Procedure 1. Place 100 g of the soil into the jars; wipe the lip of the jar to remove any debris and close tightly with a threaded ring and lid. The jar may be laid on the side and shaken gently to distribute the soil over the larger surface area of the side. 2. After all the vessels have been prepared, take the first gas sample to establish baseline CO2 concentrations. Incubate the jars under the desired conditions and withdraw gas samples periodically for GC analysis of respired CO2 or O2 uptake. 3. Gas samples are taken by syringe and directly injected into the sample port of the GC (or into the sample loop if the GC is so equipped). Gas may be mixed prior to sampling by inserting the needle of a 50-cc syringe through the septum and alternately withdrawing and re-injecting container air. Remove the mixing syringe and take a 1 cc sample with a gas-tight syringe. Purge 0.5 cc from the syringe and immediately inject the remaining 0.5 cc into the Gc. Alternately, the gas sample may be injected into a vacutainer for later analysis (Neilson & Pepper, 1990). Follow procedures in 38-3.2.3 to generate the data to determine CO2 and O 2 in the injected sample. 4. Open the vessels after sampling as necessary to aerate the soil for 15 to 30 min. Close the vessels and continue the incubation. If vessels are aerated, another sample must be taken immediately after they are closed again to mark the new baseline amounts of respiratory gases in the headspace. 38-4.2.2.3 Calculation of Results. Follow the procedures in 38-3.2.4 for calculating the concentrations of CO 2 and O2 in the gas sample. Gas concentrations are adjusted for headspace gas volume in the vessel and the temperature at sampling time to express the data in mass units (i.e., J.tg CO 2 g soil- 1) or on a molar basis (Ross et aI., 1975). If the headspace volume is small, uncomplicated vessel volume can be estimated by the weight difference determined between an empty vessel and the same vessel filled with water (Nadelhoffer, 1990). Soil sample volume must be estimated as well. This may be done by including a replicate mass of the soil

CARBON MINERALIZATION

857

to be incubated during headspace determination. Soil sample volume may be estimated independently by calculation if the bulk density and water content are known. Data are converted to mass units by multiplying the temperaturecorrected gas concentration determined for the injected sample (0.5 mL) by 2 and then by the headspace volume of the container to obtain the total mass of CO2 or O 2 in the headspace. This is then divided by the mass of the soil in the container to express the value on a dry soil basis. The time interval is accounted for as required to determine production rates or total CO2 produced.

38-4.2.2.4 Comments. A manometric method (Kroeckel & Stolp, 1985) has been reported in which the vessel volume is calculated from the rise of the liquid level in a manometer in response to pressurization of the closed system by air injection. Another pressurization method for the determination of headspace volume involves the use of a pressure transducer with excitation and measurement circuitry (Myrold, 1988; Parkin et aI., 1984; Rice et aI., 1988). In this method, the pressure transducer is attached to stiff tubing ending in a syringe needle. This needle is inserted through a septum into the vessel. The vessel is pressurized with air by use of another syringe. The transducer produces a mV reading that stabilizes within a few seconds when the vessel is empty or within a minute when it contains soil (T.B. Parkin, 1991, personal communication). Measurement of several vessels of known volume produces a graph that can be used to estimate the unknown volume of another vessel. The circuitry required to operate a pressure transducer is found in such common laboratory equipment as multimeters and data loggers. A + 12V DC source is also required. The cost of a suitable pressure transducer is relatively small (approximately $50 U. S.). Pressure methods for determining vessel volume are also effective in determining vessel leaks. Vessel sample size relationships may be determined beforehand such that the CO2 level does not rise above 2% during the incubation period (Sparling, 1981). This would make the aeration procedure unnecessary. Harper and Lynch (1985) and Bowen (1990) flushed incubating vessels with sterile air when O 2 concentration fell below 10%. Both of these methods attempt to prevent the problems with microbial metabolism that can occur where significant alterations in respiratory gas concentrations arise during incubation. A ratio of 10:1 or greater, headspace volume (mL):soil weight (g), is often used for closed systems (cf. Bottner, 1985; deCatanzaro & Beauchamp, 1985; Jenkinson & Powlson, 1976; van Gestel et aI., 1991). Combining C mineralization determinations with estimations of other microbial activities is commonly desired. The following method is designed for laboratory experiments in which it is necessary to remove portions of the soil periodically during the incubation for other analyses, such as N or S mineralization or biomass determinations.

858

ZmILSKE

38-4.2.3 Static Incubation-Titrimetric Determination 38-4.2.3.1 Special Apparatus 1. Sealable, large-mouth glass containers about 4 L in volume make suitable incubation vessels. Mason or preserving jars, as described in section 38-4.2.2.1 (used without drilling holes into the lids), may be appropriate for smaller amounts of soil, or if only one sample is to be incubated in the jar. Larger jars allow easier access to the beakers. The lids normally provided with large jars obtained commercially may not seal properly. A few disks of aluminum foil cut to fit snugly inside the sealing side of the lid usually corrects this. If not, large rubber stoppers can be used to seal the jars. 2. Small beakers (50 mL). These will contain water to maintain the humidity within the vessel. Larger beakers (100 mL) to accommodate several, individual soil samples within the large incubation jar. 3. Burette (50 mL) clamped to a stand. 38-4.2.3.2 Reagents 1. Reagents needed are found in section 38-4.1.3; CO2 -free water, 1 N of NaOH, standardized 0.5 N of HCI, 3 N of BaCI2 , and the phenolphthalein indicator solution. 38-4.2.3.3 Procedure 1. Weigh 25 g replicates of the soil into l00-mL beakers. Carefully arrange the soil sample replicates and one beaker containing 50 mL of CO2-free water in the jar. 2. Quickly pipet 1 N NaOH into a 100 mL beaker. Use 10 mL of the base for each 25 g of soil in the jar. Place this beaker in a readily accessible location in the jar. Seal the jar tightly and incubate appropriately. Set up and incubate a blank containing only water and NaOH-containing beakers. 3. Periodically, open the jar and remove the NaOH beaker for CO 2 determination. Allow the jar to remain open during the titration procedure. This is done to replenish O 2 in the jar for the next incubation period. Gently mix the contents of beaker and transfer 25 mL to a 125-mL erlenmeyer flask. Add 25 mL of CO2-free water, an excess of 3 N BaCI2 , and three or four drops of phenolphthalein indicator. Titrate with 0.5 N of HCl. Titrate the NaOH from the blank in the same manner. An alternate method for titration of base solutions involves the use of carbonate dehydratase (EC 4.2.1.1) also called carbonic anhydrase (Underwood, 1961). In this method, the enzyme is added to the diluted NaOH trap and titrated sequentially to pH 10, 8.3, and 3.7. The volume of the dilute acid used between the last two points is used to calculate the amount of CO2 trapped. The advantages of this method are the rapidity with which titration can be carried out and that the determination of the endpoint is made with a pH meter. The latter eliminates the need for subjective determination of endpoints with chromogenic indicators. 4. If other procedures are to be performed on the soil, remove one of the replicate beakers of soil. Add water, if necessary, to the beaker in the

CARBON MINERALIZATION

859

jar. Place a fresh beaker of NaOH in the jar, reducing the amount of base used if a soil sample was removed. Seal the jar and continue the incubation. Replenish the NaOH trap in the blank and continue the incubation.

38-4.2.3.4 Calculation of Results. Results are calculated as described in section 38-4.1.5, adjusting the calculated results as necessary if soil samples were removed from the jar during incubation. 38-4.2.3.5 Comments. Placement of a small sponge (Yoda & Nishioka, 1982) or piece of filter paper (Klein et al., 1984) into liquid alkali has been suggested to improve CO2 absorption, presumably by increasing the surface area of the absorbing liquid. The only other commonly used acid for titrating the base solutions is H 2S04, As sulfuric acid is added in the presence of excess Ba2 +, BaS0 4 precipitates and increases the cloudiness of the solution, which may affect the visual detection of the endpoint. Carbon dioxide trapped in base solutions may be purged by acidification of the alkali. This is useful for purging 14C-C02 from base solutions and trapping it again in a scintillation fluor (Donnelly et al., 1990). REFERENCES Alexander, M. 1977. Introduction to soil microbiology. 2nd ed. John Wiley and Sons, New York. Anderson, J.M., and P. Ineson. 1982. A soil microcosm system and its application to measurements of respiration and nutrient leaching. Soil BioI. Biochem. 14:415-416. Anderson, J.P.E. 1982. Soil respiration. p. 831-871. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Annis, P.e., and G.R. Nicol. 1975. Respirometry system for small biological samples. J. Appl. Ecol. 12:137-141. Baldocchi, D.D., S.B. Verma, D.R. Matt, and D.E. Anderson. 1986. Eddy-correlation measurements of carbon dioxide efflux from the floor of a deciduous forest. J. Appl. Ecol. 23:967-975. Bartha, R., and D. Pramer. 1965. Features of a flask and method for measuring the persistence and biological effects of pesticides in soil. Soil Sci. 100:68-70. Blackmer, A.M., and J.M. Bremner. 1977. Gas chromatographic analysis of soil atmospheres. Soil Sci. Soc. Am. J. 41:908-912. Blet-Charaudeau, C., J. Muller, and H. Laudelout. 1990. Kinetics of carbon dioxide evolution in relation to microbial biomass and temperature. Soil Sci. Soc. Am. J. 54:13241328. Boddy, L. 1983. Carbon dioxide release from decomposing wood: Effect of water content and temperature. Soil BioI. Biochem. 15:501-510. Bottner, P. 1985. Response of microbial biomass to alternate moist and dry conditions in a soil incubated with 14C_ and 15N-labeled plant material. Soil BioI. Biochem. 17:329-337. Bowen, R.M. 1990. Decomposition of wheat straw by mixed cultures of fungi isolated from arable soils. Soil BioI. Biochem. 22:401-406. Brooks, P.D., and E.A. Paul. 1987. A new automated technique for measuring respiration in soil samples. Plant Soil 101:183-187. Buyanovsky, G.A., and G.H. Wagner. 1983. Annual cycles of carbon dioxide level in soil air. Soil Sci. Soc. Am. J. 47:1139-1145. Carlyle, J.C., and U Ba Than. 1988. Abiotic controls of soil respiration beneath an eighteenyear-old Pinus radiata stand in Southeastern Australia. J. Ecol. 76:654-662. Cary, J.W., and C. Holder. 1982. A method for measuring oxygen and carbon dioxide in soil. Soil Sci. Soc. Am. J. 46:1345-1347 Castelie, A.J., and J.N. Galloway. 1990. Carbon dioxide dynamics in acid forest soils in Shenandoah National Park, Virginia. Soil Sci. Soc. Am. J. 54:252-257

860

zmILSKE

Chapman, S.B. 1971. A simple conductimetric soil respirometer for field use. Oikos 22:348353 Cheng, W., and D.C. Coleman. 1989. A simple method for measuring CO 2 in a continuous air-flow system: Modifications to the substrate-induced respiration technique. Soil BioI. Biochem. 21:385-388 Cheng, W., and D.C. Coleman. 1990. Effect of living roots on soil organic matter decomposition. Soil BioI. Biochem. 22:781-787 Christensen, B. 1987. Decomposability of organic matter in particle size fractions from field soils with straw incorporation. Soil BioI. Biochem. 19:429-435. Clark, M.D., and J.T. Gilmour. 1983. The effect of temperature on decomposition at optimum and saturated soil water contents. Soil Sci. Soc. Am. J. 47:927-929. Cropper, W.P., Jr., K.C. Ewel, and J.W. Raich. 1985. The measurement of soil CO 2 evolution in situ. Pedobiologia 28:35-40. deCatanzaro, J.B., and E.G. Beauchamp. 1985. The effect of some carbon substrates on denitrification rates and carbon utilization in soil. BioI. Fert. Soils 1: 183-187. de Jong, E., R.E. Redmann, and E. A. Ripley. 1979. A comparison of methods to measure soil respiration. Soil Sci. 127:300-306. Dobbins, D.C., and F.K. Pfaender. 1988. Methodology for assessing respiration and cellular incorporation of radiolabeled substrates by soil microbial communities. Microb. Ecol. 15:257-273. Donnelly, P.K., J.A. Entry, D.L. Crawford, and K. Cromack, Jr. 1990. Cellulose and lignin degradation in forest soils: Response to moisture, temperature, and acidity. Microb. Ecol. 20:289-295. Edwards, N.T. 1982a. A timesaving technique for measuring respiration rates in incubated soil samples. Soil Sci. Soc. Am. J. 46:1114-1116. Edwards, N.T. 1982b. The use of soda-lime for measuring respiration rates in terrestrial systems. Pedobiologia 23:321-330. Edwards, N.T., and B.M. Ross-Todd. 1983. Soil carbon dynamics in a mixed deciduous forest following clear-cutting with and without residue removal. Soil Sci. Soc. Am. J. 47:10141021. Edwards, N.T., and P. Sollins. 1973. Continuous measurement of carbon dioxide evolution from partitioned forest floor components. Ecology 54:406-412. Fine, P., A. Feigin, and Y. Waisel. 1986. A closed, well-oxygenated system for the determination of the emission of carbon dioxide, nitrous oxide, and ammonia. Soil Sci. Soc. Am. J. 50:1489-1493. Gilmour, C.M., and J.T. Gilmour. 1985. Assimilation of carbon by the soil biomass. Plant Soil 86:101-112. Gloser, J., and M. Tesal'ova. 1978. Litter, soil, and root respiration measurement. An improved compartmental analysis method. Pedobiologia 18:76-81. Gordon, A.M., R.E. Schlentner, and K. van Cleve. 1987. Seasonal patterns of soil respiration and CO 2 evolution following harvesting in the white spruce forests of interior Alaska. Can. J. For. Res. 17:304-310. Gupta, S.R., and J.S. Singh. 1977. Effect of alkali concentration volume and absorption area on the measurement of soil respiration in a tropical sward. Pedobiologia 17:233-239. Gupta, S.R., and J.S. Singh. 1981. Soil respiration in a tropical grassland. Soil BioI. Biochem. 13:261-268. Harper, S.H.T., and J.M. Lynch. 1985. Colonization and decomposition of straw by fungi. Trans. Br. Mycol. Soc. 85:655-661. Heinemeyer, 0., H. Insam, E.A. Kaiser, and G. Walenzik. 1989. Soil microbial biomass and respiration measurements: An automated technique based on infra-red analysis. Plant Soil 116:191-195. Hendricks, C.W., E.A. Paul, and P.D. Brooks. 1987. Growth measurements of terrestrial microbial species by a continuous-flow technique. Plant Soil 101:189-195. Insam, H. 1990. Are the soil microbial biomass and basal respiration governed by the climatic regime? Soil BioI. Biochem. 22:525-532. Jawson, M.D., L.F. Elliott, R.I. Papendick, and G.S. Campbell. 1989. The decomposition of 14C-labeled wheat straw and 15N-labeled microbial material. Soil BioI. Biochem. 21:417-422. Jenkinson, D.S., and D.S. Powlson. 1976. The effects of biocidal treatments on metabolism in soil- V. A method for measuring soil biomass. Soil BioI. Biochem. 8:209-213.

CARBON MINERALIZATION

861

Kanemasu, E.T., M.L. Wesley, B.B. Hicks, and J.L. Heilman. 1979. Techniques of calculatin~ energy and mass fluxes. p. 156-182. In B.J. Barfield and J.F. Gerber (ed.) Modifications of the aerial environment of crops. Am. Soc. Agric. Eng., St. Joseph, MI. Kaspar, H.P. 1984. A simple method for the measurement of N 20 and CO 2 flux rates across undisturbed soil surfaces. N. Z. J. Sci. 27:243-246. Kaspar, H.P., and J.M. Tiedje. 1980. Response of electron-capture detector to hydrogen, oxygen, nitrogen, carbon dioxide, nitric oxide and nitrous oxide. J. Chromatogr. 193:142-147. Keeney, D.R, K.L. Sahrawat, and S.S. Adams. 1985. Carbon dioxide concentration in soil: Effects on nitrification, denitrification, and associated nitrous oxide production. Soil BioI. Biochem. 17:571-573. Kinsbursky, RS., and S. Saltzman. 1990. CO 2 -nitrification relationships in closed soil incubation vessels. Soil BioI. Biochem. 22:571-572. Klein, D.A., P.A. Mayeux, and S.L. Seaman. 1972. A simplified unit for evaluation of soil core respirometric activity. Plant Soil 36:177-183. Klein, T.M., N.J. Novick, J.P. Kreitinger, and M. Alexander. 1984. Simultaneous inhibition of carbon and nitrogen mineralization in a forest soil by simulated acid precipitation. Bull. Environ. Contam. Toxicol. 32:698-703. Knapp, E.B., L.P. Elliott, and G.S. Campbell. 1983. Microbial respiration and growth during the decomposition of wheat straw. Soil BioI. Biochem. 15:319-323. Kroeckel, L., and H. Stolp. 1985. Influence of oxygen on denitrification and aerobic respiration in soil. BioI. Fert. Soils 1:189-193. Kursar, T.A. 1989. Evaluation of soil respiration andsoil CO 2 concentration in a lowland moist forest in Panama. Plant Soil 113:21-29. Linn, D.M., and J.W. Doran. 1984. Effect of water-filled pore space on carbon dioxide and nitrous oxide production in tilled and nontilled soils. Soil Sci. Soc. Am. J. 48: 1267-1272. Loos, M.A., A. Kontson, and P.C. Kearney. 1980. Inexpensive soil flask for 14C-pesticide studies. Soil BioI. Biochem. 12:583-585. Lundegardh, M. 1927. Carbon dioxide evolution of soil and crop growth. Soil Sci. 23:417453. Macfadyen, A. 1973. Inhibitory effects of carbon dioxide on microbial activity in soil. Pedobiologia 13:140-149. Marinucci, A.C., and R Bartha. 1979. Apparatus for monitoring the mineralization of volatile 14C-labeled compounds. Appl. Environ. Microbiol. 38:1020-1022. Martens, R. 1985. Limitations in the application of the fumigation technique for biomass estimations in amended soil. Soil BIOI. Biochem. 17:57-63. . Martens, R. 1987. Estimation of microbial biomass in soil by the respiration method: importance of soil pH and flushing methods for the measurement of respired CO 2 • Soil BioI. Biochem. 19:77-81. Mathes, K., and T. Schriefer. 1985. Soil respiration during secondary succession: influence of temperature and moisture. Soil BioI. Biochem. 17:205-211. Minderman, G., and J.C. Vulto. 1973. Comparison of techniques for the measurement of carbon dioxide evolution from soil. Pedobiologia 13:73-80. Myrold, D.D. 1988. Denitrification in ryegrass and winter wheat cropping systems of western Oregon. Soil Sci. Soc. Am. J. 52:412-416. Nadelhoffer, K.J. 1990. Microlysimeter for measuring nitrogen mineralization and microbial respiration in aerobic soil incubations. Soil Sci. Soc. Am. J. 54:411-415. Naganawa, T., K. Kyuma, H. Yamamoto, Y. Yamamoto, H. Yokoi, and K. Tatsuyama. 1989. Measurement of soil respiration in the field: Influence of temperature, moisture level, and application of sewage sludge compost and agro-chemicals. Soil Sci. Plant Nutr. 35:509-516. Neilson, J.W., and I..L. Pepper. 1990. Soil respiration as an index of soil aeration. Soil Sci. Soc. Am. J. 54.428-432. Nordgren, A. 1988. Apparatus for the continuous, long-term monitoring of soil respiration rate in large numbers of samples. Soil BioI. Biochem. 20:955-957. Nordgren, A., E. Baath, and B. SOderstrom. 1988. Evaluation of soil respiration characteristics to assess heavy metal effects on soil microorganisms using glutamic acid as a substrate. Soil BioI. Biochem. 20:949-954. Parkin, T.B., H.P. Kaspar, A.J. Sextone, and J.M. Tiedje. 1984. A gas-flow soil core method to measure field denitrification rates. Soil BioI. Biochem. 16:323-330.

862

ZmILSKE

Parkinson, K.J. 1981. An improved method for measuring soil respiration in the field. J. Appl. Ecol. 18:221-228. Paul, E.A., and EE. Clark. 1989. Soil microbiology and biochemistry. Academic Press, San Diego. Redmann, R.E., and Z.M. Abouguendia. 1978. Partitioning of respiration from soil, litter and plants in a mixed-grassland ecosystem. Oecologia 36:69-79. Revsbech, N.P., and B.B. J!1lrgensen. 1986. Microelectrodes: Their use in microbial ecology. Adv. Microb. Ecol. 9:293-352. Rice, C.W., P.E. Sierzega, J.M. Tiedje, and L.W. Jacobs. 1988. Stimulated denitrification in the microenvironment of a biodegradable organic waste injected into soil. Soil Sci. Soc. Am. J. 52:102-108. Robards, K., V.R. Kelly, and E. Patsalides. 1992. Determination of dissolved gases in water by gas chromatography. p. 53-86. In J.C. Giddings et al. (ed.) Advances in chromatography. Marcel Dekker, New York. Robertson, G.P., and J.M. Tiedje. 1985. An automated technique for sampling the contents of stoppered gas-collection vials. Plant Soil 83:453-457. Ross, D.J., B.A. McNeilly, and L.E Molloy. 1975. Studies on a climosequence of soils in tussock grasslands 4. Respiratory activities and their relationships with temperature, moisture, and soil properties. N. Z. J. Sci. 18:377-389. Sakamoto, K., and T. Yoshida. 1988. In situ measurement of soil respiration rate by a dynamic method. Soil Sci. Plant Nutr. 34:195-202. Salonius, P.O. 1983. Effects of air drying on the respiration of forest soil microbial populations. Soil BioI. Biochem. 15:199-203. Schwartzkopf, S. 1978. An open chamber technique for the measurement of carbon dioxide evolution from soils. Ecology 59:1062-1068. Seto, M. 1982. A preliminary observation on CO 2 evolution from soil in situ measured by an air current method-An example in rainfall and plowing sequences. Jpn. J. Ecol. 32:535-538. Sextone, A.J., N.P. Revsbech, T.B. Parkin, and J.M. Tiedje. 1985. Direct measurement of oxygen profiles and denitrification rates in soil aggregates. Soil Sci. Soc. Am. J. 49:645-651. Shelton, D.R., and T.B. Parkin. 1989. A semiautomated instrument for measuring total and radiolabeled carbon dioxide evolution from soil. J. Environ. Qual. 18:550-554. Smith, K.A. 1983. Gas chromatographic analysis of the soil atmosphere. p. 407-454. In K.A. Smith (ed.) Soil analysis: Instrumental techniques and related procedures. Marcel Dekker, New York. Sparling, G.P. 1981. Microcalorimetry and other methods to assess biomass and activity in soil. Soil BioI. Biochem. 13:93-98. Sparling, G.P., and A.W. West. 1990. A comparison of gas chromatography and differential respirometer methods to measure soil respiration and to estimate the soil microbial biomass. Pedobiologia 34:103-112. Stotzky, G. 1965. Microbial respiration. p. 1550-1572. In C.A. Black (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Swift, M.J., O.W. Heal, and J.M. Anderson. 1979. Decomposition in terrestrial ecosystems. D.J. Anderson et al. (ed.) Studies in ecology. Vol. 5. Blackwell Scientific, Oxford, England. Umbreit, W.W., R.H. Burris, and J.E Stauffer. 1972. Manometric and biochemical techniques: A manual describing methods applicable to the study of tissue metabolism. 7th ed. Burgess Publ. Co., Minneapolis. Underwood, A.L. 1961. Carbonic anhydrase in the titration of carbon dioxide solutions. Anal. Chern. 33:955-956. van Cleve, K., Pol. Coyne, E. Goodwin, C. Johnson, and M. Kelley. 1979. A comparison of four methods for measuring respiration in organic material. Soil BioI. Biochem. 11:237246. van Gestel, M., J.N. Ladd, and M. Amato. 1991. Carbon and nitrogen mineralization from two soils of contrasting texture and microa~gregate stability: Influence of sequential fumigation, drying and storage. Soil BioI. BlOchem. 23:313-322. Vance, E.D., and N.M. Nadkarni. 1990. Microbial biomass and activity in canopy organic matter and the forest floor of a tropical cloud forest. Soil BioI. Biochem. 22:677-684. Wagner, G.H., and ~.A. .Buyanovsky. 198~. Use of gas sampling tubes for direct measurement of 14C02 III soil air. Int. J. Radlat. Isot. 34:645-648.

CARBON MINERALIZATION

863

Weaver, R.W. 1974. A simple, inexpensive apparatus for simultaneous collection of CO2 evolved from numerous soils. Soil Sci. Soc. Am. Proc. 38:853. Weber, M.G. 1985. Forest soil respiration in eastern Ontario jack pine ecosystems. Can J. For. Res. 15:1069-1073. West, A.W., and G.P. Sparling. 1986. Modifications to the substrate-induced respiration method to permit measurement of microbial biomass in soils of differing water contents. J. Microbiol. Methods 5:177-189. Wildung, R.E., T.R. Garland, and R.L. Buschborn. 1975. The interdependent effects of soil temperature and water content on soil respiration rate and plant root decomposition in arid grasslands soils. Soil BioI. Biochem. 7:373-378. Witkamp, M. 1969. Cycles of temperature and carbon dioxide evolution from litter and soil. Ecology 50:922-924. Yoda, K., and M. Nishioka. 1982. Soil respiration in dry and wet seasons in a tropical dry-evergreen forest in Sakaerat, NE Thailand. Jpn. J. Ecol. 32:539-541.

Published 1994

Chapter 39 Isotopic Methods for the Study of Soil Organic Matter Dynamics DUANE C. WOLF, University of Arkansas, Fayetteville, Arkansas

J. O. LEGG, University of Arkansas, Fayetteville, Arkansas THOMAS W. BOUTTON, Texas A & M University, College Station, Texas

The vast majority of all organic C and N in the world's terrestrial environment is present in the form of soil organic matter, which contains approximately 1.5 x 1018g of C (Post et aI., 1982) and 0.095 x 1018g of N (Post et aI., 1985). In addition to its importance as a reservoir of C, N, and other nutrients, this pool of soil organic matter ha3 many properties that define the structural and functional attributes of natural and agricultural ecosystems. The benefits of soil organic matter in crop production have been recognized for centuries (Allison, 1973), although the reasons for the beneficial effects have not been clearly understood. Organic matter is highly important in relation to soil aggregation, tilth, cation exchange capacity, nutrient supply, soil water, biological activity, and other soil characteristics. Repeated additions of organic matter to soils, normally occurring from plant and animal residues, and subsequent biological, chemical, and physical processes that occur, create a highly complex and dynamic system. This complex system attracted the attention of chemists as early as the 18th century when acid and alkali extraction procedures were first used to remove organic matter from soils (Russell, 1961). In the 1940s, isotopes of C and N came into use in studies of transformations of these elements during organic matter decomposition. Paul and van Veen (1978) reviewed these early studies and proposed a model to describe the rate of organic matter decomposition. It is important to emphasize that, in the decomposition of labeled compounds, other organic compounds may be synthesized simultaneously from the mineralized C and N. The position of the labeled atom

Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

WOLF ET AL.

866

in the material undergoing mineralization can influence the results from decomposition studies. Soil organic matter not only has fundamental importance at the ecosystem level, but also has considerable importance for global biogeochemistry due to the size of the pool and its linkages to atmospheric CO2 via primary production and decomposition. There is currently much interest and speculation regarding whether soil organic matter is a net carbon sink or source in the global carbon cycle under present environmental conditions as well as under conditions of elevated atmospheric CO2 concentration and altered climate (Prentice & Fung, 1990; Schlesinger, 1990). As a result of the controversial role of soil organic matter in global biogeochemistry, methodologies for quantifying fluxes of soil organic matter have become extremely important. In this chapter, we will address isotopic methods of assessing fluxes associated with soil organic matter. There are basically four approaches available: (i) use of organic matter labeled with 14C or 13C; (ii) use of organic matter labeled with 15N; (iii) use of natural variation in 13C in organic matter; and (iv) use of 14C injected into the atmosphere during nuclear weapons testing. Approaches 1 and 2 are most useful for shortterm studies of 1 to 10 yr, and approaches 3 and 4 are useful for examining fluxes on time scales ranging from tens to thousands of years. Since methods involving the use of "bomb" 14C have been reviewed recently (Harrison et ai., 1990; Goh, 1991), we will address some of the more common methods of using 13C, 14C, and 15N to study organic matter decomposition. The References section will provide citations for obtaining further information on pertinent isotopic methodology as well as related analytical techniques. 39-1 DECOMPOSITION OF 14C-LABELED ORGANIC MATTER IN SOILS 39-1.1 Introduction

Historically, the use of 14C-Iabeled materials to study soil organic matter decomposition has been accomplished by adding radiolabeled plant material, microorganisms, microbial products, or specific compounds to the soil and measuring the amount of 14C02 evolved during an aerobic incubation (Ladd & Martin, 1984; Wolf & Legg, 1984). Various methods are available to produce the radiolabeled materials and to collect the evolved 14C02 • The objective of this section is to present methods for the production of 14C-Iabeled materials and define several methods available for the collection and assay of the 14C02 evolved during aerobic laboratory incubation studies. It is not our intent to define all available techniques, but rather to provide details of the more commonly used procedures. Using even a weak beta-emitting radioactive isotope such as 14C requires particular at-

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

867

tention to safety considerations. The use of radioisotopes generally requires a permit or license and training in radiation safety. The aspects of health and safety will not be addressed in this section, but any researchers undertaking a project involving 14C should contact their local Radiation Safety Officer for details on licensing and permits. 39-1.2 Obtaining 14C-Labeled Organic Materials Obtaining 14C-Iabeled materials for studies on organic matter decomposition is the first and possibly the most difficult step. The choice of organic material may be dictated by the objective of the given research project and the availability of specific research facilities in which to carry out labeling experiments. Products that can be labeled range from the simplest of pure compounds purchased from several commercial sources (Table 39-1) to whole plants grown in a 14C02 environment or microorganisms and microbial products produced in the laboratory on a wide range of labeled organic substrates. Obviously the cost, time, and facilities will have an impact on the organic material available for any given experiment. 39-1.2.1 Labeling Plant Material One method used to produce 14C-Iabeled plant material is to grow plants in a 14C02 environment in a growth chamber and harvest the plants after a suitable growth period. Another approach is to inject a 14C-Iabeled precursor directly into the plant and let the metabolic activity of the plant incorporate the label into various biochemical fractions. Both methods require an appropriate containment facility to prevent contamination of the laboratory environment. 39-1.2.1.1 Materials 1. 14C02 or 14C-Iabeled precursor. 2. Suitable growth and containment facility. 39-1.2.1.2 Procedure. Details of construction and operation of 14C02 growth chambers have been presented by several researchers (Andersen et aI., 1961; Cheshire & Griffiths, 1989; Harris & Paul, 1991; Jenkinson, 1960; Scully et aI., 1956; Smith et aI., 1962; Warembourg & Kummerow, 1991). At harvest, the plant may be separated as desired into various components such as shoots, leaves, and roots. Once the plant material is harvested, it should be lyophilized. Oven drying the plant material has the potential to release 14C02 into the laboratory environment and is not recommended. After freeze-drying, the material can be ground and sieved to the required size. A second method of labeling plant material is to treat growing plants with 14C-Iabeled precursors of lignin or cellulose biosynthesis. Haider et ai. (1977) used a syringe to inject 14C-Iabeled p-coumaric acid, a precursor in the synthesis of lignin, into the base of young corn (Zea mays L.)

WOLFET AL.

868

Table 39-1. Partial listing of commercial sources of 14C-labeled materials or liquid scintillation counting cocktails. Company name and address American Radiolabeled Chemicals, Inc. 11624 Bowling Green Dr. St. Louis, MO 63146 Amersham Corp. 2636 S. Clearbrook Dr. Arlington Heights, IL 60005 Curtin Matheson Scientific, Inc. P.O. Box 1416 Houston, TX 77251-1416 Dupont Co., Biotechnology Systems 549 Albany St. Boston, MA 02118 Fisher Scientific 900 Stewart Ave. Plano, TX 75074 ICN Biomedicals, Inc. 3300 Hyland Ave. Costa Mesa, CA 92626 Isolab, Inc. Drawer 4350 Akron, OH 44321 Isotope Products Laboratories 1800 N. Keystone St. Burbank, CA 91504 Moravek Biochemicals, Inc. 577 Mercury Lane Brea, CA 92621 Packard Instrument Co. 800 Research Parkway Meriden, CT 06450 Research Products International Corp. 410 N. Business Center Dr. Mount Prospect, IL 60056 Sigma Chemical Co. 3050 Spruce St. St. Louis, MO 63103

Telephone number

14C-Labeled material

1-800-331-6661

X

1-800-323-9750

X

1-800-879-2670

X

X

1-800-551-2121

X

X

1-800-766-7000 1-800-854-0530

LSC cocktail

X X

1-800-321-9632

X X

1-818-843-7000

X

1-800-447-0100

X

X

1-203-238-2351

X

X

1-800-323-9814 1-800-325-3010

X X

X

plants. The plants were injected three times and allowed to grow an additional 3 wk before harvest. The plant tops were harvested and small molecular weight compounds extracted before the material was used for decomposition studies. Similarly, lignin components of white oak (Quercus albus), red maple (Acer rubrum), and cattail (Typha lati/ola) have been labeled by feeding plants aqueous solutions of L-[U-14C)phenylalanine or [2'-14C(side chain)] ferulic acid through their cut stems (Crawford, 1978; Crawford & Crawford, 1976). Cellulosic components of lignocelluloses were labeled by substituting solutions of D_[U-14C)glucose (Crawford et aI., 1977).

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

869

39-1.2.1.3 Comments. Safety is critical when using radioisotopes, and it is important to prevent contamination of the laboratory (Coleman & Corbin, 1991). Always wear a laboratory coat, chemical safety goggles, and disposable gloves. Work on absorbent paper and use a pipette bulb for all pipetting. Avoid being exposed to 14C-Iabeled aerosols, dust, and volatile material. All grinding and sieving procedures should be conducted in a properly vented fume hood. It is obvious that there should be no eating, drinking, or smoking in the laboratory at any time. The growth medium may be contaminated with 14C, and suitable disposal procedures must be followed. 39-1.2.2 Labeling Microbial Biomass and Microbial Products Most studies using 14C-Iabeled microbial biomass involve incubation of a pure culture of a given microorganism with a suitable labeled substrate such as uniformly 14C-Iabeled glucose. The microbial biomass is harvested and lyophilized following an appropriate incubation period. The dry biomass can be ground and sieved before it is added to the soil in an incubation experiment. 39-1.2.2.1 Materials 1. 14C-Iabeled substrate such as D-[U_14C]glucose. 2. Pure culture of microorganism (see American Type Culture Collection Catalog). 3. 250-mL Erlenmeyer flasks. 4. Orbital shaker. 5. System to collect evolved 14C02 (see Fig. 39-1,39-2,38-2). 6. Freeze dryer. 7. Autoclave. 39-1.2.2.2 Procedure. Prepare a growth medium suitable for the microorganism of interest. Add an appropriate amount of 14C-Iabeled substrate to the medium, add the inoculant, and connect the flask to the CO 2 collection system (see Fig. 39-1 and chapter 38 in this book). Place the 250-mL flask on the shaker and incubate until sufficient microbial growth has occurred. The microbial biomass can be harvested by centrifugation and then lyophilized, ground, and sieved before it is used in soil decomposition studies. Wagner and Krzywicka (1975) used 14C02 to label algal biomass and D_[U-14C]glucose was used by Reyes and Tiedje (1973) to produce labeled Saccharomyces cerevisiae. Labeled whole cells have been fractionated and their components used in soil decomposition studies (Hurst & Wagner, 1969; Nakas & Klein, 1979). Measurement and turnover of microbial biomass have been evaluated (Jenkinson & Powlson, 1976; Kassim et aI., 1981, 1982) and reviewed (Jenkinson & Ladd, 1981; Wagner, 1975). Additionally, 14C-Iabeled microbial products such as polysaccharides, polyphenols, and proteins have been isolated and used in decomposition studies. Oades and Wagner (1971) and Zunino et ai. (1982) used a mineral

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

871

salts medium supplemented with D_[U-14C]glucose to grow Leuconostoc spp. The polysaccharide material was collected and used in decomposition studies. Soil fungi have been grown on a mineral salts medium with D_[U-14C]glucose to produce melanins used in laboratory incubations (Martin & Haider, 1979; Martin et aI., 1972, 1982). The 14C-Iabeled proteins produced by Chlorella pyrenoidosa and Microcoleus spp. also have been obtained (Verma et aI., 1975).

39-1.2.2.3 Comments. The spent microbial growth medium containing 14C-Iabeled material must be disposed of in a suitable manner. It is important to realize that a radioactive waste container with spent medium can be a source of 14C02 in the laboratory if microbial growth occurs. 39-1.2.3 Specific 14C-Labeled Compounds In certain studies related to organic matter decomposition, it is desirable to amend the soil with specific compounds such as an amino acid, phenol, or sugar (Haider & Martin, 1975). These compounds may be labeled in a specific position in the molecule, or they may be labeled uniformly. In most cases, the compound can be obtained from commercial sources, a few of which are listed in Table 39-1. Some companies also have a service available to synthesize special compounds.

39-1.3 Methods and Approach to Incubations of 14C-Labeled Organic Materials It is not possible to present all of the methods that have been used to determine decomposition of 14C-Iabeled organic materials in soil. References should be consulted for additional methods or specific details of more specialized incubation systems (Anderson, 1982; Marvel et al., 1978; Stotzky, 1965). Some of the common systems are presented in Fig. 39-1. In general terms, the soil is placed in a flask and amended with the 14C-Iabeled organic material and attached to a CO 2 collection unit (see chapter 38 in this book). For aerobic incubation studies, O 2 must not be a limiting factor. For this reason, a continuous flow-through system has been used. The evolved 14C02 can be trapped in a base such as KOH or NaOH and the 14C activity assayed by liquid scintillation counting techniques. Several static systems have also been used and have the advantages of a simple design and conservation of space.

39-1.3.1 Materials 1. 2. 3. 4. 5.

Liquid scintillation spectrometer. Liquid scintillation counting (LSC) cocktail. Scintillation vials. Pipettes. KOH or NaOH containing 10 mgIL Tropaeolin 0 (Aldrich Chemical Co.).

WOLFET AL.

872

6. Soil, field moist and sieved. 7. 14C-Iabeled organic material. 8. 14C02 collection unit (see Fig. 39-1,39-2, and 38-2). 39-1.3.2 Procedures

The amount of 14C activity to add to the soil will depend upon the amount of 14C expected to be found in the fraction of interest or evolved as 14C02 , analytical efficiency of the laboratory procedure that determines whether the activity in the fraction of interest is diluted or concentrated, and the minimal activity in the LSC cocktail to give efficient counting (Voroney et aI., 1991). It is generally necessary to determine the counting efficiency for each procedure and LSC cocktail used and it may be necessary to do a preliminary or pilot study to determine the appropriate levels of activity to add. In an example given by Voroney et al. (1991), to detect a difference of 20 counts/min (cpm) at a 95% confidence level, one would have to count a sample containing lOOO cpm for lO min at an 85% counting efficiency. In many cases, it may not be possible to obtain a level of lOOO cpm in the LSC cocktail; using longer counting times and lower confidence intervals will enable the researcher to work with samples containing activity as low as 100 cpm in the LSC cocktail. The 14C-Iabeled organic material should be added at a rate not to exceed 2% of the soil weight (Jenkinson, 1971). Typically, a lOO-g sieved soil sample should be weighed onto nonabsorbent waxed paper or aluminum foil. If the 14C-Iabeled organic material to be added is a dry solid, it can be weighed and added directly to the soil and carefully mixed into the soil. Because of the potential for contamination, the operation should be completed in a fume hood. If the 14C-Iabeled material is in aqueous solution, it can be added to the soil and thoroughly mixed. Once the labeled material is added, the soil should be adjusted to the required soil water potential by adding distilled water and the soil should be carefully mixed to ensure uniform water distribution. Excessive mixing should be avoided as puddling of the soil could result that would reduce oxygen diffusion into the soil. The moist soil containing the 14C-Iabeled material should be transferred into a 250-mL Erlenmeyer flask and attached to the CO2 collection unit. Appropriate controls (soil without organic amendment) and blanks (no soil or organic amendment in the flask) should be prepared in the same manner and attached to the CO2 collection unit. It may also be appropriate to include a soil amended with unlabeled organic material. Several CO2 collection units that have been used are shown in Fig. 39-1. The unit shown in Fig. 39-2 is similar to the unit given in chapter 38, Fig. 38-2. The details of the reactions involved are given in chapter 38, and the same samples used to determine the total amount of CO2 evolved can and often are used to determine the amount of 14C02 evolved. One modification used for the 14C02 determination is that Tropaeolin 0 is added to the base to give it an orange color indicating a pH > 12.7. If the base

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

873

2mm POLYPROPYLENE TUBING

AIR--'

0

0

0

0

0

"

0

0

0 0

0

0

( "AQUARIUM" VALVE

1 MKOH Fig. 39-2. Apparatus for studying the decomposition of 14C-labeled organic materials in soil (Stevenson, 1986).

becomes partially neutralized and the pH declines to < 11, the indicator will turn yellow which indicates that the CO2 removal efficiency has been compromised. In studies using 14C-Iabeled materials, it is recommended that air be drawn through the system by a vacuum. Thus, if a leak develops in the system, there is little possibility of contamination of the laboratory atmosphere with 14C02 . A leak in the system would substantially reduce the specific activity of the collected CO2 , In experiments using 14C-Iabeled material, it is also desirable to include a secondary or backup tube to collect any 14C02 that might not be collected in the first tube. The air flow rate is adjusted to one bubble per second to provide adequate oxygen levels to maintain aerobic conditions. Faster rates may result in incomplete trapping of the CO 2 , Gas exchange rates of 20 volumes per hour have been used (Wagner & Chahal, 1966), but a level of one volume exchange per hour is adequate in most cases. Relatively economical "aquarium" valves can be obtained at local pet supply stores and have proved durable and satisfactory. At various intervals, the CO2 collection tubes are replaced, and the amount of both total CO 2 evolved and 14C02 evolved can be determined. To determine the total CO2 evolved, see chapter 38. To determine the 14C02 evolved, add an aliquot of the NaOH or KOH containing the evolved 14C02 to a scintillation vial containing a suitable LSC cocktail. A partial list of commercial suppliers of LSC cocktails is given in Table 39-1. New generation LSC cocktails do not contain aromatic solvents, are nonflammable, and do not result in toxic vapors. The cocktails are biodegradable and result in fewer disposal problems. The specific volumes of base

WOLFET AL.

874

and cocktail will be determined by the load capacity of the cocktail, the activity in the sample, and the base concentration. Generally, 1 mL of 0.5 M NaOH or KOH can be counted using 10 mL of LSC, but the manufacturer can provide specific guidelines. For determining the radioactivity in the sample, the liquid scintillation spectrophotometer should be used according to the manufacturer's specifications. Discussions of scintillation counting procedures (L' Annunziata, 1979) and measurement of radionuclides (L'Annunziata, 1984) should be consulted. An alternative approach is to determine the amount of activity remaining in the soil following a given incubation period. This approach generally requires combustion of the soil to convert the 14C-Iabeled organic material to 14C02 and collecting the 14C02 from the combustion and counting it. The 14C02 can be collected in base and the base added to a suitable cocktail, or the 14C02 can be collected directly in certain cocktails and counted. Specific details are provided by L' Annunziata (1979) and Voroney et al. (1991). 39-1.3.3 Calculation of Results Depending upon the specific scintillation spectrometer used, the results may be given as counts per unit time such as second (cps) or minute (cpm). The counts must be corrected for counting efficiency and background and historically have been given as disintegrations per unit time (dps or dpm). The SI base unit is becquerel (Bq). One becquerel is equivalent to a nuclear transformation per second, or dps. Useful conversions include 1 !A-Ci = 37 kBq = 3.7 x 104 dps = 2.22 X 106 dpm (Corbin & Swisher, 1986). Once the appropriate dilution factors are taken into account, the percentage of the added 14C evolved as 14C02 or remaining in the soil can be calculated (Eq. [1] and [2]). Modem-day scintillation counters allow the researcher to program the instrument to complete many of the calculations. 14C in sample

=

% added 14C evolved as 14C02

Sample cps - Background cps

(dilution factor) Counting efficiency (expressed as a decimal fraction) 14C evolved from sample 14C added to soil

(100)

[1]

[2]

39-1.3.4 Comments It is important to report percentage of the added 14C evolved as 14C02 or the percentage added 14C remaining in the soil rather than percentage decomposition. Historically, percentage decomposition was calculated from 14C02 evolution data, but the amount of 14C02 evolved may be substantially influenced by the amount of 14C incorporated into microbial

875

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

biomass or soil organic matter. The microbial products formed may be subsequently mineralized and thus influence the amount and rate of labeled substrate recovered as 14C02. If, in addition to 14C02, volatile organics are lost from the soil and absorbed by the base used to collect 14C02, the amount of 14C02 evolved will be overestimated. Kearney and Kontson (1976) placed a polyurethane filter preceding the base trap and were able to sorb the evolved volatile products for subsequent 14C determination. Another technique for collecting 14C02 uses chromatographic tubes (chapter 38) that can be counted by liquid scintillation techniques. Additionally, Mayaudon (1971) presents a detailed discussion of radiorespirometry techniques in soil systems. Also, a direct soil counting technique has been used for determining 14C remaining in soil treated with an herbicide (Lavy, 1975; Scott & Phillips, 1972). 39-2 13C NATURAL ABUNDANCE TECHNIQUE: BACKGROUND AND PRINCIPLES 39-2.1 Introduction Approximately 98.89% of all C in nature is 12C, and 1.11 % is 13e. The relative proportions of these two stable isotopes in nature vary slightly around these average values as a result of isotopic fractionation during physical, chemical, and biological processes (Boutton, 1991b). The 13C/12C ratio of organic C found in terrestrial environments is determined largely by the C isotope fractionation that occurs during photosynthesis. Plants with the C3 photosynthetic pathway exhibit greater discrimination against 13C than plants with the C4 pathway. These natural isotopic differences between plants can be used to study the dynamics of organic matter in soil. 39-2.1.1 Stable Isotope Terminology Because natural variation in the ratio of 13C/12C is small, stable C isotope ratios are expressed in relative terms as b 13C PDB values: b13C PDB e/oo)

=

[ Rsample -

R pDB

1 x 1()3

[3]

R pDB where Rsample is the mass 45 (13C 160 160) to mass 44 (12C 160 160) ratio of the sample and R pDB is the 13C/12C ratio of the international PDB limestone standard, which has a value of 0.0112372 (Craig, 1957). The PDB standard was a Belemnitella americana limestone fossil from the Cretaceous Pee Dee formation in South Carolina. Corrections are made for the presence of 180 and 17 0 in the CO2, Thus, b 13C PDB is a relative index that indicates the parts per thousand (per mil, or °/00) difference between the 13C/12C ratio of the sample and that of the PDB standard. For example,

876

WOLFET AL.

a b13CPDB value of -10 %0 indicates a sample with a 13C/12C ratio 10 parts per thousand lower than the PDB standard; a b13CPDB value of + 5 %0 indicates a sample with a 13C/12C ratio 5 parts per thousand greater than the PDB standard. 39-2.1.2 Stable Carbon Isotope Ratios of Plants and Soils The 13C natural abundance technique for studying soil organic matter dynamics uses natural differences in b 13 CPDB values between plants with the C3 and C4 pathways of photosynthesis. Atmospheric CO2 has a b13CPDB value of approximately -8 %0 (Levin et aI., 1987). During photosynthesis, plants with the C3 pathway discriminate against atmospheric 13C02 to a greater extent than C4 plants (O'Leary, 1988). The C 3 plants have b 13 CPDB values ranging from approximately -32 to -20 %0 (mean = -27 %0), while C4 plants have b13CPDB values ranging from -17 to -9 %0 (mean = -13 %0). Thus, C 3 and C4 plants have distinct stable C isotope ratios and differ from each other by approximately 14 %0 on average (Smith & Epstein, 1971). Plants with Crassulacean acid metabolism (CAM) usually have b13CPDB values typical of C4 plants; however, under certain environmental and developmental circumstances, some CAM species are able to switch to a C 3 mode of photosynthesis. These "facultative" CAM species will have b13CPDB values depending upon the relative proportions of C fixed via CAM and C3 modes. The CAM plants have b 13 CPDB values ranging from approximately -28 to -10 %0, but are most commonly -20 to -10 %0. Most terrestrial plant species are C 3. Most temperate zone and all forest communities are dominated by C 3 species. However, C 4 and CAM plants are significant components of many plant communities, particularly in warm, arid, or semiarid environments (Osmond et aI., 1982). For example, tropical and subtropical grasslands consist almost exclusively of C4 grasses, and CAM plants (e.g., Cactaceae, Euphorbiaceae) are important in many desert communities. In general, the proportion of C4 species in a flora increases as latitude and altitude decrease (e.g., Teeri & Stowe, 1976; Boutton et aI., 1980). Although there are small isotopic differences between different parts ofthe same plant (up to 2 %0 different from the whole plant) and between specific biochemical fractions within plants (up to 8 %0 different from the whole plant), the C isotopic signature of the whole plant is largely preserved as dead plant tissue decomposes and enters the soil organic matter pool (Nadelhoffer & Fry, 1988; Melillo et al., 1989). Thus, soil organic matter in C4 plant communities will have b13CPDB values near -13 %0, while organic matter from soils in C 3 communities will be near -27 %0. This natural isotopic "label" in the soil organic matter enables reconstruction of the prior history of plant communities (Dzurec et al., 1985) and also permits estimation of soil organic matter dynamics in situ over relatively long periods without any type of experimental disturbance.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

39-2.1.3 Natural

13C

877

and Measurement of Organic Matter Dynamics

If a community dominated by C3 plants has been compositionally stable for a relatively long time (e.g., 500-1000 yr), then the soil organic matter in that community is in isotopic equilibrium with that C3 vegetation; that is, they should have approximately the same b13 C values. If that C3 community (e.g., a forest) is converted to a C4 plant community (e.g., a corn field or tropical grass pasture), then the isotopic composition of the soil organic matter will begin to shift towards that of C4 vegetation as the C3 component decays out of the soil and is replaced by C4 organic matter inputs. The rate at which the original mass of C3-derived organic matter (which is uniquely and readily identifiable by its characteristic b 13 C value) decays out of the system through time is a direct measure of the turnover rate of organic matter in that system. It should be noted that cultivation of the soil will accelerate organic matter turnover. Thus, changing the b13 C value of the organic matter inputs (i.e., from C3 ~ C4 , or from C4 ~ C3 ) is equivalent to in situ labeling of the soil organic matter (Balesdent et al., 1987). Measurements of turnover rates using the 13C natural abundance technique are best suited to time periods of tens to thousands of years because: (i) the C isotopic difference between C3 and C4 plants is relatively small; and, (ii) the mass of existing soil organic C derived from the previous vegetation is large relative to the annual increments of organic C derived from the new vegetation (Balesdent et al., 1988). However, significant differences in b 13C have been detected in upper A horizons in as little as 3 mo following a single input of C4 litter into a C3 plant system (Insam et al.,1991). The use of the 13C natural abundance technique obviously will be limited to situations where there has been a change from C3 ~ C4 or C4 ~ C3 vegetation. However, these situations are common and have been used to study organic matter dynamics where: (i) C3 rain forest has been converted to C4 pasture or C4 crops; (ii) C4 grassland has been converted to C3 crops; (iii) C4 savanna has been converted to C3 woodland; and (iv) C3 cropland has been converted to C4 cropland (Balesdent et al., 1987, 1988, 1990; Vitorello et al., 1989; Martin et al., 1990; Skjemstad et al., 1990; Cerri et al., 1991).

39-2.2

13C

Natural Abundance Technique: Methodology

39-2.2.1 Special Apparatus 1. Vacuum manifold capable of achieving 10- 3 torr for evacuating and sealing combustion tubes. 2. Gas-oxygen torch for making quartz combustion tubes and sealing the tubes after being loaded with sample and evacuated. 3. Shade 8 or darker welding goggles to protect eyes while heating quartz with gas-oxygen torch.

WOLFET AL.

878

4. High vacuum system capable of 10- 3 torr for cryogenic separation and purification of CO2 produced during combustion of organic matter in sealed quartz tubes. 5. Analytical balance readable to 0.01 mg. 6. Programmable muffle furnace. 7. Dewar flasks for holding liquid N or dry ice slush during CO 2 isolation procedure. 8. Nier-type, dual inlet, triple collector gas isotope ratio mass spectrometer for measuring ratios of isotopic species of CO2 produced by combustion of organic matter. 39-2.2.2 Reagents 1. HCl (0.5 M) to volatilize carbonate C from soils containing pedogenic or lithogenic carbonates. 2. NaCl, NaI, ZnBr2' or CsCl to produce high density (1.2-1.8 g cm- 3 ) liquid for isolation of undecomposed particulate organic debris in soils. 3. Quartz or vycor tubing (9 mm o.d. x 7 mm i.d.) for combustion of soil organic matter samples (Quartz Scientific, Fairport Harbor, OH). 4. Wire-form CuO with a low C background suitable for microanalysis (Fisher Scientific C474-500 or equivalent). 5. Reduced Cu granules, -10 to +40 mesh (Aldrich Chemical Co. catalog no. 31,140-5 or equivalent). 6. Quartz wool. 7. Liquid N. 8. Ethanol-dry ice slush (-78 0e). 9. Carbon isotope standards calibrated relative to the international PDB standard (available from NIST, Gaithersburg, MD, or from IAEA, Vienna, Austria). 39-2.2.3 Procedure 39-2.2.3.1 Field Sampling. The study site must consist of an area known to have been converted from a C3-dominated to a C4-dominated plant community (or vice versa) at a precisely known time, and there should be a remnant of the original plant community nearby to provide baseline samples. Ideally, one would like to sample sites on the same soil and in close proximity to one another that have undergone the same type of conversion at different times. Sampling such a chronosequence would provide detailed kinetics of the turnover process. To characterize the isotopic composition of the organic matter inputs, live plant tissue as well as litter should be sampled from both the baseline site with the original vegetation and the derived site with the new plant community. Plant tissue should be dried at 70°C, ground to pass a O.4-mm (40-mesh) screen, and set aside for isotopic analysis.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

879

Soil samples can be obtained either by digging a pit or taking cores. Pit sampling reveals soil structure and horizonation more readily than cores, does not compress the profile, and enables more accurate sampling for bulk density. Sampling depth intervals according to horizonation enables 6 13 C to be related to pedogenesis, and bulk density measurements at several depths in the profile will allow C dynamics to be expressed on Mg Clba basis (1 Mg = 106g). Samples should be taken from several depth intervals to a depth of approximately 1 m. Samples from several pits or cores can be bulked to obtain representative 6 13C values of a large area or kept separate for analysis of spatial variability. 39-2.2.3.2 Preparation of Soil Samples. Soil samples should be passed through a 2-mm screen and rocks and large roots removed. Undecomposed particulate organic debris, or the "light fraction" (Stevenson & Elliott, 1989), is removed from the soil samples by flotation in high density (1.21.8 g cm- 3 ) inorganic solutions. Suitable inorganic chemicals for the preparation of high density liquids include NaCl, NaI, ZnBrz, CsCI, and Na metatungstate. Approximately 100 g of soil is added to a 600-mL beaker, and the beaker is filled with a saturated NaCI solution (density == 1.2 g cm- 3 ). The soil is stirred vigorously, and particulate organic debris floats to the surface. After the soil settles, the organic debris can be siphoned or strained off the surface of the liquid. By repeating this process five times, virtually all particulate organic debris is removed from the soil, and the removal can be verified by examining the soil with a dissecting microscope. The particulate organic debris, which is largely roots, can be pooled with the larger roots removed by sieving, treated with 0.5 M HCl to remove any adhering carbonate C, dried at 70°C, ground to pass a O.4-mm (40-mesh) screen, and set aside for isotopic analysis. The NaCl is then removed from the root-free soil by repeated washing in distilled water. Failure to remove residual salt may interfere with determination of organic C content later in the procedure. If soils have pedogenic or lithogenic carbonates present, this C must be destroyed. Although the 6 13 CPDB value of pedogenic carbonate is related to the C3 to C4 composition of the plant community, both pedogenic and lithogenic carbonate are significantly enriched in 13C and would seriously confound stable C isotope measurements of the soil organic matter. Approximately 100 g of soil is placed in a 600-mL beaker which is then filled with 0.5 M HCI and stirred. Soil is left in the 0.5 M HCI for 3 d, and the HCI solution is replaced daily. When carbonates have been removed, soils are washed repeatedly in distilled water to remove excess HCI. This acid pretreatment to eliminate carbonates has no effect on either the organic C content of the soil, or on the 6 13 C value of the soil organic C (Boutton et aI., unpublished data). If turnover of only the bulk soil organic matter is to be measured, then the soils can be dried, ground to pass a 0.5-mm screen, and set aside for isotopic analysis. However, because different organic matter fractions exhibit different turnover rates, most investigators now choose to process soil

880

WOLFET AL.

samples into more defined categories such as particle-size fractions, aggregate size classes, or the classical humic fractions. The most useful information is gained when biologically significant fractions are analyzed. In some studies, particle-size fractionation has been followed by extraction of specific humic substances from the particle-size fractions. Others have described in detail the methods for the isolation of particle-size fractions (Jackson, 1969; as modified by Tiessen & Stewart, 1983), aggregate size fractions (Kemper & Rosenau, 1986), and the humic fractions (Schnitzer, 1982). Following any of these more detailed procedures, the soil organic matter fractions must be dried and ground to pass a 0.5-mm screen prior to isotopic analysis. 39-2.2.3.3 Conversion of Organic Carbon to Carbon Dioxide for Mass Spectrometry. Due to instrumental requirements, C must be converted to CO 2 for stable isotope ratio measurements by mass spectrometry. The most common and simplest method to convert organic C to CO2 is by combustion with an excess of CuO in an evacuated, sealed quartz tube at 850°C (Boutton, 1991a). This method does not change the C isotope composition of the original sample, produces quantitative yields of C (which permit determination of percentage C in the sample), and is rapid and relatively inexpensive. Quartz or vycor tubing (9 mm o.d. x 7 mm i.d.) is cut to 20-cm lengths, and the tubes are sealed at one end with a gas-oxygen torch. When heating quartz or vycor to the softening point, always work under a fume hood to exhaust the resulting toxic gases and wear quartz-working goggles (Wale Apparatus, Hellertown, PA) or shade 8 or darker welding goggles to protect eyes from intense glare. The prepared tubes sealed on one end only, a porcelain crucible containing the wire-form CuO catalyst (e.g., Fisher Scientific, catalog no. C474-500), and another crucible containing quartz wool are then heated in a muffle furnace at 850°C for 1 h to remove potential organic contaminants. Upon cooling, the CuO and quartz wool can be stored separately in clean jars and the combustion tubes stored in a desiccator until ready for use. On a piece of weighing paper, weigh out 1.0 g of CuO catalyst. Then, tare the balance and weigh out enough sample to provide approximately 2 to 3 mg of C. If this step can be carried out on an analytical balance readable to 0.01 mg or on a microbalance, the percentage C in the sample can be determined later. The amount of soil required to provide 2 to 3 mg of C will vary mostly as a function of depth in the profile. Plant tissue, roots, and litter usually contain 40 to 50% C, and a 4- to 5-mg sample of these materials is adequate to provide 2 to 3 mg of C. Too much C can result in explosion of tubes during combustion. When the appropriate amount of sample has been weighed, mix the sample and CuO thoroughly, and use a long-stem funnel to deliver the sample/CuO mixture to the bottom of a pre-combusted quartz tube. To hold the soil in place during evacuation of the tubes later, a plug of pre-combusted quartz wool can be inserted into the combustion tube and positioned above the soil/CuO mix-

882

WOLF ET AL.

additional hours, and then cooled to room temperature. If a programmable muffle furnace is not available, it is acceptable to simply turn off the furnace after 2 h at 900 DC, and allow it to cool slowly to room temperature. While the furnace is at 650 DC, the reduced copper granules eliminate halogens, and catalyze the conversion of any CO to CO2, NO x to N2, and SOx to CUS0 4 (Frazer & Crawford, 1963). Since N0 2 (mass 46) and N20 (mass 44) have the same masses as isotopic species of CO2, it is critical that these gases be eliminated prior to mass spectrometric analysis. High resolution mass spectrometric analysis of the gases produced by combustion of several different organic compounds by this method have revealed the presence of only CO2, H 20, and N2 (Boutton et aI., 1983). Sealed tubes that have been combusted should not be stored for more than 5 d prior to isolation of CO2 because carbonate forms slowly and fj13CPDB of the CO2 decreases by 1 to 3 D/OO after 2 wk (Engel & Maynard, 1989). If tubes are stored for more than 5 d, the problem can be avoided simply by recombusting the tubes prior to CO2 isolation and purification (Engel & Maynard, 1989). Prior to isotopic analysis, the CO2 produced by combustion must be isolated from the other combustion products and purified by cryogenic distillation. A vacuum system for this purpose is shown in Fig. 39-3b. When operating this vacuum system or handling combusted quartz tubes, safety glasses should be worn to protect against explosions, implosions, or cryogenic liquids. The combustion tube is scored at one end and inserted into the tube cracker (Des Marais & Hayes, 1976), a sample bottle is attached to the manifold, all valves are opened, and the entire vacuum system is pumped down to < 10- 3 torr. Then, a Dewar flask containing liquid N (-196 DC) is placed around the purification trap, and valve E is closed. The top of the combustion tube is broken off by flexing the tube cracker, and the gases produced during combustion are released into the vacuum system; water and CO2 freeze into the purification trap. After 4 min, valve F is closed, valve E is opened, and all noncondensible gases (mostly N2) are pumped away. When the vacuum is restored to 10-3 torr, valve B is closed, and the liquid N Dewar flask is removed from the purification trap and replaced with a Dewar containing an ethanol-dry ice slush (-78 DC). The CO2 sublimes but water remains frozen in the trap. To measure the volume of CO2 produced from the organic matter, the CO2 is transferred into the manometer cold finger by cooling it with a Dewar of liquid N; 2 min should be allowed for the CO2 to freeze into the cold finger. When the transfer is complete, valve E is closed, the liquid N Dewar is removed from the cold finger, and the CO2 is allowed to expand into the mercury manometer calibrated previously with known volumes of CO2, The manometer reading is noted and, together with the weight of the sample combusted, is used to calculate the percentage C of the sample. Valve A is then closed, and valves B, C, and D are opened. The CO2 is transferred into the sample bulb by immersing it in a liquid N Dewar for 2 min. When the transfer is complete, valves C and D are closed, and the

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

883

sample bulb is detached from the vacuum system and attached to the inlet system of the mass spectrometer. An electronic capacitance manometer could be used in place of the mercury manometer shown in Fig. 39-3b. The capacitance manometer is more expensive but eliminates the need for mercury and is significantly more accurate for determining percentage C. Accuracy for electronic manometers ranges from 0.15 to 1% of reading, depending upon the model, while that for a mercury manometer ranges from approximately 2 to 5% of reading. Organic matter can be combusted to CO 2 in Pyrex tubing at 550 °C (Sofer, 1980; Vitorello et aI., 1989) with procedures identical to those outlined above. The primary advantage of this alternative is that Pyrex tubing is approximately 5 to 10% of the cost of quartz tubing. If Pyrex is used, great care should be taken to ensure good contact between sample and CuO, and combustion time should be increased to 12 h. Some reports indicate that accuracy and precision of l)13CPDB values obtained by combustion in Pyrex tubes at 550 °C are poorer than those obtained with quartz at 850 °C (Boutton et al., 1983; Le Feuvre & Jones, 1988; Swerhone et aI., 1991). Furthermore, combustion at 550 °C may not give quantitative yields of CO 2 , eliminating the possibility of determining percentage C during CO 2 isolation and purification (Boutton et aI., 1983). 39-2.2.3.4 Mass Spectrometric Analysis and Isotopic Indices. Stable C isotope ratios are measured on the CO 2 generated by the above procedure with a dual-inlet, triple-collector gas isotope ratio mass spectrometer. The high precision of these instruments is due to simultaneous collection of the ion beams (masses) of interest and to repeated measurements of sample and standard gases by alternate switching during a single isotope ratio determination. The theory and methods of determining the isotopic composition of CO 2 by mass spectrometry have been reviewed (Craig, 1957; Deines, 1970; Mook & Grootes, 1973; Gonfiantini, 1981; Santrock et aI., 1985). The b 13C PDB value is determined using Eq. [3]. Most of the error in isotopic measurements results from sample preparation. Mass spectrometer precision (1 SD), as determined by repeated analyses of the same gas sample, is often as low as 0.01 0/00 . By contrast, different preparations of aliquots of the same sample generally will have a precision (1 SD) of 0.1 0/00 for both plants and soils. The PDB standard was derived from a limestone of marine origin with an absolute 13C!12C ratio of 0.0112372 (Craig, 1957). As the basis of the PDB scale, it has a b 13C PDB value of 0 0/00 • The PDB standard no longer exists, but several other primary standards were calibrated against it before the supply was exhausted, so it is still possible to express l)13C values relative to PDB. Primary C isotope standards are available from the National Institute of Standards and Technology (NIST, formerly the National Bureau of Standards or NBS) or from the International Atomic Energy Agency (IAEA) in Vienna, Austria.

WOLFET AL.

884

For some mass balance calculations, it is more appropriate to use the absolute ratio (R) or the fractional abundance (F) of 13C in a sample. The absolute ratio is calculated by rearrangements of Eq. [3]:

1000

+1

1

X RPDB

[4]

where RpDB = 0.0112372. The fractional abundance is the fraction of total C in a sample that is 13C: Rsample

F=

Rsample

+1

[5]

Additional details on these indices and their relationship with 013C PDB are provided by Hayes (1983).

39-2.2.3.5 Calculating Sources of Soil Organic Matter. The relative proportions of soil C derived from C3 and C4 sources can be determined by simple mass balance calculations. Assuming that we have a situation where a C4 plant community has replaced a C3 community, then the proportion of C (p) derived from the C4 community at some later point in time (t) can be calculated as: F

=

(p)Fc4 + (l-p) FC3

[6]

where F is the fractional abundance (see Eq. [5]) of the soil organic matter fraction of interest at time t after the transition from C3 ~ C4, F C4 is the fractional abundance of the C4 organic matter inputs (often an average of shoot, roots, and litter), FC3 is the fractional abundance of the soil organic matter fraction of interest prior to the change in vegetation, and 1-p is the proportion of C3 plant-derived C still present in the soil at time t. Since the 013C PDB scale is not linear, it is technically more correct to use fractional abundance (F) as an isotopic index in mass balance calculations such as Eq. [6]; however, over the range of 013CPDB values encountered in plant-soil systems at natural abundance, 013C PDB is sufficiently linear that Eq. [6] can be rewritten as:

o = (p) OC4 + (1-p) OC3

[7]

where 013C PDB values have been substituted for the fractional abundances used in Eq. [6]. Equation [7] can be rearranged and simplified to:

p=

[8]

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

885

Since the percentage C of each of the organic matter fractions was measured above as part of the sample combustion procedure outlined earlier, the relative proportions of C from C 4 (p) and C3 (l-p) sources can be used to compute the actual masses of C from each organic matter source. If the total mass of C M (with units of mg C/g soil) is known, then the mass of C from C 4 vegetation (MC4) can be calculated as:

[9]

MC4=M(p)

Similarly, the mass of C from C 3 vegetation (MC3) can be calculated as: MC3 = M (l-p)

[10]

Thus, if the study site consists of plots that have been switched from C3 to C 4 at different times or if multiple times are available from the same site, the decay of the C3 C out of the soil system and the rate of entry of C 4 C into the soil system can be described with respect to time (Balesdent et aI., 1987; Skjemstad et aI., 1990; Andreux et aI., 1990; Cerri et aI., 1991). If soil bulk density measurements are taken, it is possible to convert the data acquired above into C content per unit area: Mg C/ha = (g C/g soil) x Qb x I X 104

[11 ]

where Qb is the bulk density of the soil layer under consideration (Mg soil/m3), and I is the thickness (meters) of the soil layer under consideration. Carbon dynamics for the entire soil can be determined by solving Eq. [11] for each depth interval (e.g., 0-0.1 m, 0.1-0.2 m, etc.) and then summing the results. Although we have assumed a situation where a C 4 plant community has replaced a C3 community, the calculations for the reverse situation (C 4 ~ C3) are directly analogous to those described above.

39-2.2.3.6 Kinetics of Soil Organic Matter Turnover. Although many complex processes are responsible for soil organic matter turnover, it is generally accepted that the overall process can be described reasonably well according to first order rate kinetics which assume a single homogeneous pool of soil organic C (Jenkinson & Rayner, 1977; Paul & van Veen, 1978; Paul & Clark, 1989). When studying organic matter dynamics where a C 4 community has replaced a C3 community, the decay of C 3 C out of the system can be approximated by the negative exponential or first order decay model: At = Ao e- kt orInt (AlA) 0

=

-kt

[12]

where At is the mass of C 3-derived C at some time t after the C 3 ~ C 4 switch, Ao is the mass of C 3-derived C at time 0, k is the fractional rate constant with units of time -1, and t is the length of time elapsed since the

886

WOLF ET AL.

C3 ~ C 4 switch. The value of the fractional rate constant k is equal to the slope of the line obtained by plotting In (A/Ao) against time. The half-life (t l12 ) of C3-derived C in the system is then equal to 0.693/k, and the mean residence time or turnover time is equal to -11k (Paul & Clark, 1989). Because of the poor fit of a single compartment model to most experimental results, more complex multiexponential (multicompartmental) models would permit representation of the kinetics of labile and recalcitrant pools of soil C (e.g., Andreux et aI., 1990). 39-2.2.4 Comments The 13C natural abundance technique for measuring soil organic matter dynamics is complementary to the tracer approaches using 14C_ or 15N-enriched organic matter. Natural 13 C allows work to be done on large spatial scales as opposed to small plots or pots, it involves minimal perturbation to the system, and it can elucidate kinetics over relatively long periods of tens to hundreds of years. Furthermore, this technique usually operates on time scales that allow all soil organic matter pools, even those that are recalcitrant, to become "labeled." In tracer experiments with 14C or 15N, the duration of the experiments (usually < 5 yr) is such that the more recalcitrant fractions may not become labeled. Because this technique involves only naturally occurring stable isotopes of C, there are no hazards, regulations, or disposal problems associated with its use. The technique also has some weaknesses that should be addressed. One of the major problems is that it is difficult to measure the exact b 13 C PDB value of the organic matter inputs following the vegetation change (e.g., 6 C4 in Eq. [8]). The 6 13 C PDB values of plant tissue vary slightly in response to environmental conditions and can show small differences between plant parts and between biochemical fractions. However, 6 13 C PDB values of litter samples integrated through time should serve as a reasonable estimate of the isotopic composition of the organic matter input. Another potential problem is that, in well-drained mineral soils that have supported a stable plant community for a long time, 613CPDB of the soil organic matter increases by 1 to 2 °/00 from the surface to approximately 1 m in depth (Stout et aI., 1981). It has been suggested that this 13C-enrichment is a consequence of microbial metabolism, with microbes typically being slightly more enriched in 13C than the substrate they grow on. Since organic matter increases in age with depth in the profile (Scharpenseel & Neue, 1984), the enrichment in the deeper layers may simply reflect the consequences of a longer history of microbial metabolism. While the natural abundance method assumes that the isotopic composition of the soil organic matter will equilibrate to the 6 13 C PDB value of the organic matter inputs of the new plant community, it is clear that the organic matter input alone does not determine the equilibrium 6 13 C PDB values of the soil. Further work is needed to elucidate the consequences of 13C enrichment with soil depth on this technique.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

887

39-3 DECOMPOSITION OF 15N-LABELED ORGANIC MATTER IN SOILS 39-3.1 Introduction A study of the decomposition of 15N-Iabeled organic matter primarily involves the processes of immobilization and mineralization. Major aspects of these processes are covered in chapter 42. Indigenous soil organic N occurs in many complex forms that have varying mineralization rates. Much of the organic N is in a "passive" phase (Jansson & Persson, 1982) which is essentially inert over short periods. Incorporation of 15N-Iabeled organic matter into soils permits an evaluation of the mineralization potential of newly incorporated organic N relative to that of the indigenous soil N, as well as determination of the possible size of the inert pool of soil N. Many studies of the nature and composition of soil organic matter have been carried out (Allison, 1973), but factors influencing the stability of the organic N constituents are still not clearly understood. The use of isotopic methods thus far has not enhanced our current knowledge of the stabilizing factors, but information has been obtained concerning the rate at which newly incorporated N becomes stabilized in forms similar to that of the indigenous soil organic N (Legg et aI., 1971). Studies of the transformations of labeled organic N incorporated into soils have been carried out in field, greenhouse, and laboratory experiments. Although field experiments provide useful information from soils under natural conditions (e.g., Ladd & Amato, 1986), time requirements are large compared to greenhouse and laboratory studies. For that reason, emphasis will be placed on the latter two modes of study. Many variations of the described methods exist, and reference should be made to the literature cited for additional material. Methods for the determination of mineral N are covered in chapters 41 and 42, organic N in chapter 40 and in section 39-4.3, and 15N measurements are given in chapter 40. Additional discussions of methods are presented by Bremner and Mulvaney (1982), Keeney and Nelson (1982), Hauck (1982), and Mulvaney (1993).

39-3.2 Labeling Organic Matter with 15N Generally, the method selected for labeling organic matter will depend upon the type and purpose of the experiment. The methods may be categorized as (i) labeling plants by supplying growing plants with 15N_ labeled fertilizer or (ii) labeling soil biomass by supplying soil microorganisms with 15N plus an energy source (see also chapter 40). In any case, the enrichment with 15N should be sufficient to be easily detectable in the total soil N. This depends upon the amount of 15N added, its enrichment, and the total N content of the soil. The minimum amount of N per sample for many mass spectrometers is about 0.5 to 1 mg. It is useful to have an excess 15N percentage of 0.500 or more if the total N of the sample is so small that

888

WOLF ET AL.

an addition of a measured amount of unlabeled (NH4hS04 is needed to meet the minimum N requirement. Calculation of results from this dilution technique has been covered by Hauck (1982). 39-3.2.1 Growing Plants with 15N-Labeled Fertilizer 39-3.2.1.1 Growing Plants in Soil to be Labeled. A greenhouse pot experiment may be set up in which 15N-Iabeled fertilizer is added to the soil, plants are grown, and harvested plant material is chopped or ground and returned to the soil. This process can be repeated as many times as desired, provided a short incubation period is allowed between crops so that some N is mineralized to reduce N deficiency in the succeeding crop. No further additions of N need be made, and additions from seed in succeeding crops can be eliminated by growing a crop, such as oat (Avena sativa L.), to sufficient maturity for the seed to be harvested and used for the next crop. A typical example will be given that can be modified as needed. 39-3.2.1.2 Procedure. A bulk sample of field soil (top 15-20 cm) is obtained, sieved through a 2-mm sieve to remove extraneous plant material, and mixed well. To 2-kg soil (oven-dry basis) contained in plastic bags within the pots, add 200 mg of enriched N in solution (5-10% excess 15N) and other nutrient elements as needed. These materials may be easily mixed throughout the soil in the plastic bags. The form of labeled N (nitrate, ammonium, or urea) is optional. Plant 15 oat seeds in each pot of soil, cover, and water to near field capacity. Other plants can be used, but the plant must be grown to maturity so the seeds produced can be used for subsequent planting. After germination, plants may be thinned to an equal number for each pot. Return the plant material to the soil surface. During plant growth, water should be added daily by weighing the pots and adjusting the soil to field capacity. When the seed heads are sufficiently mature for germination, plants are cut at the soil surface, seeds are collected, and the stems and leaves are cut into about 2-cm pieces on a sheet of paper or plastic. The soil is removed from the pot, broken up on a plastic sheet, and roots are cut into small pieces. Soil and plant material are replaced in the plastic bag and mixed. Water is added as required for field capacity, the tops of the bags are loosely closed to reduce evaporation and allow aeration, and incubation can proceed on the greenhouse bench for 2 to 3 wk. Seeds from the pots are kept separate, allowed to air dry during the incubation period, and then used to replant the pot from whence they came. This procedure can be repeated as many times as desired. At definite cropping intervals, triplicate pots may be removed from the experiment and analyzed. In this case, the plant material is oven dried (60°C), weighed, ground in a Wiley mill, and analyzed for total N and atom % 15N. Plant roots may be harvested, washed, and handled in the same manner. The 15N-Iabeled soil may be used immediately or air dried and reserved for further study.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

889

39-3.2.1.3 Growing Plants without Soil. An initial advantage of labeling plant material with 15N in the absence of soil is that the 15N is not diluted by the uptake of soil N. The labeled plant material can then be added to any number of soils for decomposition studies. One disadvantage is that the 15N has not had the opportunity to recycle into less readily mineralized forms as in the preceding method. Sand-culture systems generally have been used to grow the plants, although solution-culture with proper aeration should also be suitable. The procedure described is similar to that used by Ladd et al. (1981). 39-3.2.1.4 Procedure. Weigh 3-kg quantities of clean, washed sand (air-dry basis) in plastic pots without drainage holes and place in the greenhouse or growth chamber. Plant seed of the desired crop and moisten the sand. After germination and thinning of plants to a suitable number, apply a nutrient solution to maintain the sand at 10% gravimetric water content. According to Gauch (1972), the nutrient solution proven satisfactory for many types of plants contains (meq/L) Ca, 10; Mg, 4; K, 4; N0 3 , 10; HP0 4, 4; and S04' 4, with trace elements (mglL) Cu, 0.02; Zn, 0.05; Mn, 0.5; B, 0.5; and Fe, 3 (supplied as NaFe-EDTA). The labeled nitrate (5-10 atom % 15N) may be contained in the nutrient solution or added separately at intervals during the growth period. Harvest the plants at the desired stage of maturity and wash roots to remove sand. Dry the plant material and grind to the fineness needed for the soil to be labeled. For laboratory incubation studies, the plant material should be ground to pass a I-mm sieve. For greenhouse and field studies, where large volumes of soil are involved, coarser material may be used. 39-3.2.2 Laheling Soil Biomass Microbial biomass forms a highly important constituent of soils in that it provides for the transformation of all organic materials entering the soil, as well as being a small but labile pool of plant nutrients. Jenkinson and Ladd (1981) estimated that about 2 to 3% of the organic C in soils they examined was present as microbial biomass. Labeling soil biomass directly, rather than through the decomposition of 15N-Iabeled plant materials, averts any complications that might arise from the presence of labeled plant compounds resistant to decomposition. The time required for adequate labeling is also quite short (Chichester et al., 1975; Kelley & Stevenson, 1985). A simple incubation procedure with the soil to be labeled, a readily available C source, and a 15N-Iabeled inorganic compound results in a labeled soil within a week. The procedure described is similar to that of Kelley and Stevenson (1985) but with a higher CIN ratio.

39-3.2.2.1 Procedure. Weigh 100 g of soil (oven-dry basis) that has been air dried and passed through a 2-mm sieve, into a 250-mL Erlenmeyer flask. Wet the soil with a solution containing 1.0 g of glucose C and 10 mg

WOLFET AL.

890

of N as (15NH4hS04 (5 atom % 15N). Incubate the soil for 1 wk at 30 °e. Remove the soil from the incubator, air dry, and store for future use. Larger soil samples may be used, or several flasks may be included in the incubation to provide enough labeled soil for the experiment. 39-3.3 Determination of Mineralization Rates The decomposition of soil organic N is measured in terms of net mineralization rates (see chapter 42). Since the labeled organic N is not uniformly distributed throughout the organic matter of the soil, estimates of mineralized N derived from both the indigenous and labeled organic N are made. Both plant uptake studies and laboratory incubation tests have been used to determine mineralization rates. Similar results have been obtained by these two methods (e.g., Broadbent & Nakashima, 1967; Legg et aI., 1971). Since plant uptake studies are more laborious and time consuming, only laboratory procedures will be outlined. It is possible to use either aerobic or anaerobic incubation to determine N mineralization (see chapter 41). In most cases, aerobic incubation has been the method of choice for the long-term, consecutive incubations and extractions that are required for organic matter decomposition studies. The aerobic system is especially useful with soil organic matter labeled with both e and N (e.g., Broadbent & Nakashima, 1974). For these reasons the aerobic system is the preferred method. The time requirements for carrying out long-term incubations or plant uptake experiments have varied from several months to several years. Generally, it is necessary to continue tests until the relative amounts of mineralized N derived from indigenous and labeled organic sources become stabilized. The calculated "availability ratios" (Broadbent & Nakashima, 1967) vary among soils, but the reasons for such variations have not been elucidated. The described procedure essentially follows that of Broadbent and Nakashima (1967) and allows determinations of mineralization rates at regular intervals for extended periods. Size of the soil sample to be incubated may vary, depending upon the expected mineralization rate. 39-3.3.1 Procedure Weigh out triplicate 50-g samples of labeled soil (oven-dry basis) on paper sheets, moisten, mix, and transfer to leaching tubes with fritted glass bottoms. By moistening and mixing the soil, the finer particles will not segregate out and form layers in the tubes that may impede leaching. For clayey soils that are not easily leached, it may be necessary to mix the soil with sand or expanded vermiculite to facilitate leaching. A thin layer of glass wool at the top of the soil column will reduce the dispersive action of the leaching solution on the soil particles. The leaching tubes are fitted with rubber stoppers suitable for use on small suction flasks.

891

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

Before incubation, leach the soil tubes with 80 mL of saturated CaS04 solution, applied in four equal aliquots, to remove initial mineral N. Remove excess solution with suction, and incubate the samples at 35°C in a humid atmosphere for 2 wk. Transfer the leachate to semi-micro Kjeldahl flasks and determine the inorganic N by distillation with MgO and Devarda's alloy (see chapter 40). Alternatively, the leachate may be made up to 100-mL volume and aliquots taken for analysis. The ammonia from the distillation, after titration to determine the N content, is prepared for l5N analysis in the mass spectrometer (chapter 40). After the 2-wk incubation, leach the soil samples again in the prescribed manner and determine the inorganic Nand l5N content of the leachate. Any number of incubations may be carried out to obtain a pattern of the organic N decomposition that is occurring. If the amount of N mineralized in 2 wk becomes insufficient for l5N analysis, the incubation period may be extended, with precautions taken for maintaining the water content of the soil. At the conclusion of the incubation and leaching part of the experiment, remove soil from the leaching tubes and dry and prepare it for total N analysis (see chapters 40 and 41) as well as l5N content. A comparison of the data for the original soil and that for the residual soil plus mineralized inorganic N will indicate whether any appreciable undetermined loss of labeled N has occurred during the prolonged incubation period. The above procedure can be modified in many ways, such as size of soil sample and different leaching solutions. Quite often, adaptations are easily made to accommodate the laboratory equipment available without any sacrifice in precision of results. 39-3.4 Preparation of Samples for

lSN

Analysis

The general procedures involved in l5N analysis will not be given in this chapter. Refer to chapter 40, Hauck (1982), and Mulvaney (1993) for details. Each mass spectrometry laboratory has developed its own special apparatus for preparing and analyzing l5N samples; therefore, it is necessary for anyone unfamiliar with a particular mass spectrometry laboratory to determine the most appropriate form of the research samples. 39-3.5 Calculations The "availability ratio" concept developed by Broadbent and Nakashima (1967) has been found useful in both plant uptake and mineralization studies to determine the relative mineralization rates of indigenous and l5N-labeled organic N (e.g., Chichester et aI., 1975; Legg et aI., 1971). For extracts of mineral N after incubation, the equation is as follows: . Aval'1 ab'l' llty ratlO

Labeled N (extract) / Total N (extract) =-------------Labeled N (soil) / Total N (soil)

[13]

WOLFET AL.

892

If the 15N-Iabeled soil organic N has the same availability to microorganisms as the indigenous organic N, the availability ratio will be one. On the other hand, if the labeled N is more susceptible to mineralization than the indigenous N, the availability ratio will be greater than one. A ratio of less than one is conceivable, but not likely. The component values in the equation are those at the beginning of each incubation; therefore, in a succession of incubations, the values for labeled and total N in the soil must be corrected to account for the N mineralized and extracted in the previous incubation. This correction assumes that no other losses of any consequence have occurred. An analysis of soils after the experimental period will indicate whether such losses actually occurred. When the organic N of soils is first labeled, the initial availability ratios obtained either by cropping or mineralization are generally high but quickly decrease to a level much closer to unity. This decrease indicates that the more labile fraction of the organic N is soon mineralized, leaving a more resistant fraction that appears to become more stable with time. The relationship between availability ratios and the degree of stability of labeled organic N incorporated into the soil has not been completely elucidated. Other useful calculations can be made by employing the general equation for estimating quantities involved in isotopic equilibria:

A = B(1-y)/y

[14]

Fried and Dean (1952) used this equation to obtain a measure of the

availability of a soil nutrient, A, in terms of a given rate of a labeled fertilizer, B, where the proportion of the nutrient derived from the fertilizer, y, could be determined in the plant. The A value for N can be determined when a soil is being labeled with 15N, as in section 39-3.2.1.1, by determining the fraction of total N in the plants that was derived from the fertilizer. Jansson (1958) used the same equation in mineralization studies of recently incorporated labeled organic N. In this case, A is the amount of soil organic N in the active phase, B is the amount of newly immobilized labeled N, and y is the proportion of mineralized N after incubation that is derived from the labeled organic N. In long-term incubation or plant uptake experiments, increasing A values are generally observed. This increase reflects a stabilization of the labeled N and an eqUilibration with increasing amounts of soil N in a passive form. Understanding the dynamics of residue decomposition and soil organic N turnover can be enhanced by using reaction kinetics to describe the microbial transformations involved. Paul and Clark (1989) provide a clear explanation of the mathematical equations and the utilization of specific types of data for determining zero-order, first-order, and hyperbolic reactions. Such calculations for degradation rates require relatively short-time

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

893

intervals between measurements; otherwise, microbial synthesis and soil organic matter formation may become complicating factors. 39-3.6 Comments Studies of the decomposition of organic N in soils require several different approaches owing to the complexity of the problem, and it is not possible to cover all of them in a single chapter. Notable among these are the procedures that have been devised to extract and separate organic compounds from the soil (Schnitzer, 1982). Some of the early work with 15N described the changes taking place in the quantities of labeled N found in different extracts with time (e.g., Stewart et aI., 1963). Another means of separating recently incorporated 15N in soils is organomineral sedimentation fractionations (e.g., Chichester, 1970). Modeling also has become important to increased understanding of organic matter turnover and N cycle rates (e.g., Jenkinson & Rayner, 1977; Paul & van Veen, 1978; Myrold & Tiedje, 1986). Such procedures, along with the basic methods described, provide ample opportunities for even greater advances in studies of organic N decomposition.

39-4 EXTRACTION OF LABELED ORGANIC FRACTIONS IN STUDIES OF SOIL ORGANIC MATTER DYNAMICS 39-4.1 Introduction In the preceding sections, it has been shown how the soil organic matter is labeled and how subsequent biological transformations are followed by various procedures. Much of the labeled organic matter becomes difficultly mineralizable and apparently enters stable forms similar to indigenous organic matter. Chemical extractions of soil organic matter have been used to determine the movement of labeled compounds into the various extracted fractions, as well as unknown non hydrolyzable forms. Currently, there are no standardized procedures for separating the organic from the inorganic soil phase, and many variations in extractants, treatments, and conditions have evolved over the years. In general, the extractions involve hot mineral acids or bases, followed by separation into various fractions for analysis. Several excellent reviews cover the details of several methods for extraction and fractionation of soil organic matter (Bremner, 1965; Schnitzer, 1982; Stevenson, 1965; 1982b). As Stevenson (1965) stated: The great difficulty in all fractionation procedures is that the methods employed either separate out products which are not definite chemical entities, or they form artifacts which do not have the properties of the original material. Nevertheless, the various fractionation procedures have proved useful for

WOLFET AL.

894

studying soil organic matter, and they will probably continue to be used in the future. Quite often, different extraction procedures are used for C and N fractionations owing to differences in determinations of isotopic composition and interferences that may occur. For that reason, separate extraction procedures will be described for C and N in the following sections. 39-4.2 Extraction of Organic Matter Containing Labeled Carbon 39-4.2.1 Introduction Numerous methods have been used to extract C-containing components found in soil organic matter. Many of the procedures have been summarized by Stevenson (1982a). The classical procedure involves alkaline extraction of the soil and precipitation of the humic acid fraction by acidification. The fulvic acid fraction is soluble in both the base and acid. Recently, substantial research has focused on determination of the fraction of organic C found in the microbial biomass that is the more biologically active and dynamic fraction of organic C found in soil. Procedures for estimating C levels present in microbial biomass are given in chapter 36. The procedure given in the following section is a generalized scheme for the classical method of separation of humic and fulvic acid fractions found in soil organic matter. The more detailed discussions of the procedure given by Schnitzer (1982) and Stevenson (1982a) should be consulted. The procedure is easily adapted to determine the amount of 14C in humic and fulvic acids as long as appropriate safety precautions are followed when working with radioactive material. 39-4.2.2 Materials 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Soil containing 14C. Hydrochloric acid (HCI), 0.05 M, 2 M. Sodium hydroxide (NaOH), 0.5 M. N2 source. Horizontal or wrist-action shaker. Centrifuge. Polypropylene or polyethylene centrifuge tubes. pH meter. Dry combustion or wet oxidation unit. Liquid scintillation spectrometer. Liquid scintillation counting cocktail. Scintillation vials.

39-4.2.3 Procedures Weigh 40 g (dry weight equivalent) of labeled soil into a 250-mL centrifuge tube and add 200 mL of 0.05 M HCl. Stir the mixture with a

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

895

stirring rod. The dilute acid removes any free carbonates and polyvalent cations and increases organic matter extraction efficiency. Centrifuge and discard the supernatant. This and all subsequent centrifugations are carried out at 1500 x g. Add 200 mL of distilled water, stir, centrifuge, and discard the supernatant. Add 200 mL of 0.5 M NaOH and displace the air in the centrifuge tube with N2 • Shake the centrifuge tube for 12 to 24 h at room temperature. Separate the dark-colored supernatant that contains the humic and fulvic acids from the soil by centrifugation and decant the supernatant and filter it through glass wool to remove suspended organic solids. Collect the supernatant in a 1-L beaker. For maximum organic matter removal, repeated extraction with 0.5 MNaOH is required. Generally, two to three extractions are adequate for most soils. Following the alkaline extractions, add 200 mL of distilled water to the residual soil, shake for 10 min, centrifuge, and add the rinse water to the supernatant in the beaker. Add 2 M HCI to the alkaline extract and adjust to pH 1 using a pH meter to monitor the change. The dark-colored precipitate is humic acid and the straw-colored solution is the fulvic acid. Allow the mixture to set overnight at room temperature or refrigerate. Centrifuge the pH 1 suspension to separate the fractions. Once the humic acid has been isolated, add 200 mL of distilled water, mix, centrifuge, and add the supernatant to the fulvic acid fraction. Both fractions should be freeze-dried, weighed to determine yield, and stored in a desiccator. The residual soil contains the humin fraction and it can also be freeze-dried for subsequent analysis. The amount of total C and 14C in the humic and fulvic acids can be determined by dry combustion or wet oxidation of the organic materials. To determine the amount of total C and 14C in the humin fraction, the residual soil can be analyzed. The specific details for the oxidation procedures are given by Nelson and Sommers (1982). The amount of CO2 produced from the oxidation can be determined by methods also given in chapter 38 and the details for determining 14C02 levels are given in 39-1.3.2. 39-4.2.4 Comments

The humic and fulvic acid fractions will contain inorganic components or ash. Typical ash values are :510% for humic acid and 2:40% for fulvic acid. If the investigator is only interested in the amount of 14C from the original substrate incorporated into the humic or fulvic acid fraction, it is not necessary to determine the percentage ash. However, because of the high ash contents in the extracted materials, the percentage C values will generally be much lower than values reported on a dry, ash-free basis. Procedures for purification of humic and fulvic acids have been detailed by Schnitzer (1982) and Stevenson (1982a). Air or oven drying is not recommended for humic or fulvic acids. The best method for drying humic acid is lyophilization. Because of the large volume of liquid containing the fulvic acid, a flash evaporator is often used to concentrate the fulvic acid prior to freeze drying.

WOLF ET AL.

896

The excessive Cl- levels in the organic matter fractions can cause problems during combustion or wet oxidation of the organic materials. It is necessary to frequently change the halide trapping units in the combustion or oxidation trains. 39-4.3 Extraction of Organic Matter Containing Labeled Nitrogen 39-4.3.1 Introduction Classical methods for the extraction and fractionation of organic N were based upon the assumption that much of the N is proteinaceous in nature, and procedures developed for characterizing various chemical groups in proteins were used. With 15N-Iabeled soils, such methods required certain modifications to accommodate isotopic analyses (Bremner, 1965; Cheng & Kurtz, 1963). In the modified procedure, the soil hydrolysate is neutralized without prior removal of excess acid, and the different forms of N in the neutralized hydrolysate are measured as ammonium that is readily converted to N2 for isotope-ratio analysis. The methods described in the following sections are basically a condensed version of the ones presented by Bremner (1965) and Stevenson (1982b), and these publications should be consulted for further information. The procedure is relatively simple and permits rapid estimation of total N, ammonium N, hexosamine N, amino acid N, and (serine + threonine) N in soil hydrolysates. 39-4.3.2 Acid Hydrolysis of Soils 39-4.3.2.1 Special Apparatus 1. Micro-Kjeldahl digestion unit. 2. Steam distillation apparatus. 3. Distillation flasks: 50- and 100-mL Pyrex Kjeldahl flasks with 19/38 ground glass joints and glass hooks. 4. Microburette, 5 mL, graduated at O.Ol-mL intervals. 39-4.3.2.2 Reagents 1. Sulfuric acid (H2S04), concentrated. 2. Hydrochloric acid (HCI) , approximately 6 M: Add 513 mL of concentrated HCI (specific gravity 1.19 glcm3 ) to about 400 mL of water, cool, and dilute to volume in a 1-L volumetric flask. 3. n-Octyl alcohol. 4. Potassium sulfate-catalyst mixture: Prepare an intimate mixture of 200 g of K2S04, 20 g of CuS04·5H20, and 2 g of Se. Powder the reagents separately before mixing, and grind the mixture in a mortar to powder the cake that forms. 5. Sodium hydroxide (NaOH), approximately 10 M. 6. Sodium hydroxide, approximately 5 M.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

897

7. Sodium hydroxide, approximately 0.5 M. 8. Boric acid indicator solution: Place 80 g of H 3B03 in a 5-L flask marked at a 4-L volume, add about 3800 mL of water, and heat and swirl the flask until the H 3B03 is dissolved. Cool the solution and add 80 mL of mixed indicator solution. The indicator is prepared by dissolving 0.099 g of bromocresol green and 0.066 g of methyl red in 100 mL of ethanol. To the boric acid + indicator solution, add 0.1 M NaOH cautiously until the solution assumes a reddish purple tint (pH about 5.0), and make the solution to 4 L with water. Mix thoroughly before use. 9. Sulfuric acid, 0.0025 M standard. 10. Magnesium oxide (MgO): Heat heavy MgO in a muffle furnace at 600 to 700 °C for 2 h. Cool in a desiccator containing KOH pellets and store in a tightly stoppered bottle. 11. Ninhydrin: Grind 10 g of reagent grade ninhydrin in a mortar and store in a small widemouth bottle. 12. Phosphate-borate buffer, pH 11.2: Place 100 g of sodium phosphate (Na3P04·12H20), 25 g of borax (Na2B407·10H20), and about 900 mL of water in a l-L volumetric flask, and shake the flask until the phosphate and borate are dissolved. Dilute the solution to 1 L and store in a tightly stoppered bottle. 13. Citric acid: Grind 100 g ofreagent grade citric acid (C6H g0 7·H20) in a mortar, and store in a small widemouth bottle. 14. Citrate buffer, pH 2.6: Mix 2.06 g of powdered sodium citrate dihydrate (Na3C6Hs07·2H20) and 19.15 g of powdered citric acid in a mortar, and grind to a fine powder with a pestle. Store in a small widemouth bottle. 15. Periodic acid (HI0 4·2H20) solution, approximately 0.2 M: Dissolve 4.6 g of HI0 4·2H20 in 100 mL of water and store in a glass stoppered bottle. 16. Sodium metaarsenite (NaAs02) solution, approximately 1.0 M: Dissolve 13 g of powdered, reagent grade NaAs0 2 in 100 mL of water and store in a tightly stoppered bottle. 17. Standard (NH4 + + amino sugar + amino acid)-N solution: Dissolve 0.189 g of (NH4hS04' 0.308 g of glucosamine·HCI, and 0.254 g of alanine in water. Dilute the solution to 2 L in a volumetric flask and mix thoroughly. If prepared from pure, dry reagents, this solution contains 20 I-tg of NH4 + -N, 10 I-tg of amino sugar-N, and 20 I-tg of a-amino acid-N/mL. Store the solution for no more than 7 d in a refrigerator at 4°C. 18. Standard (serine + threonine)-N solution: Dissolve 0.150 g of serine and 0.170 g of threonine in water. Dilute the solution to 2 L in a volumetric flask and mix. If prepared from pure, dry reagents, this solution contains 10 I-tg of serine-N and 10 I-tg of threonine-N/mL. Store the solution for no more than 7 d in a refrigerator at 4°C.

898

WOLFETAL.

39-4.3.3 Preparation and Sampling of Soil Hydrolysate Place a sample of finely ground ($100 mesh) soil containing about 10 mg of N in a round-bottom flask fitted with a standard taper (24/40) ground-glass joint. Add two drops of octyl alcohol and 20 mL of 6 M HCI, and swirl the flask until the acid is thoroughly mixed with the soil. Place the flask in an electric heating mantle, connect the flask to a Liebig condenser fitted with a 24/40 ground-glass joint, and heat the soil-acid mixture so that it boils gently under reflux for 12 h. After completion of hydrolysis, wash the reflux condenser with a small quantity of distilled water, allow the flask to cool, and remove the flask from the condenser. Filter the mixture through a Buchner funnel fitted with Whatman no. 50 filter paper, using a suction filtration apparatus that permits collection of the filtrate in a 200 mL tall-form beaker marked to indicate a volume of 60 mL. Wash the residue with 5- to 1O-mL portions of distilled water until the filtrate reaches the 50-mL mark on the beaker. Immerse the lower half of the beaker in crushed ice, and neutralize to pH 6.5 ± 0.1 by cautious addition of NaOH, using a pH meter to follow the course of neutralization. Add the alkali slowly with constant stirring to ensure that the hydrolysate does not become alkaline at any stage of the neutralization process. Use 5 M NaOH to bring the pH to about 5 and complete the neutralization using 0.5 M NaOH. Transfer the neutralized hydrolysate by means of a small funnel to a 100-mL volumetric flask, and dilute to volume with the washings obtained by rinsing the beaker, electrodes and stirrer several times with small quantities of distilled water. Stopper the flask and invert several times to mix the contents. To determine the different forms of N in the hydrolysate, after thorough mixing, usually a 5- to 10-mL sample is pipetted into a 50- or 100-mL distillation flask and the flask is connected to a steam distillation apparatus. It is necessary to use pipettes with wide tips that permit rapid delivery to avoid sampling errors. The form ofN under analysis is determined from the NH3-N liberated by steam distillation for 2 to 4 min. 39-4.3.4 Total Hydrolyzable Nitrogen Place 5 mL of the neutralized hydrolysate in a 50-mL distillation flask, add 0.5 g of K2S04-catalyst mixture and 2 mL of concentrated H 2S04 , and heat the flask cautiously on a micro-Kjeldahl digestion unit until the water is removed and frothing ceases. Increase the heat until the mixture clears, and complete the digestion by boiling gently for 1 h. After digestion, allow the flask to cool, and add about 10 mL of water (slowly and with shaking). Cool the flask under a cold-water tap, and place it in a beaker containing crushed ice. Add 5 mL of H 3B03 indicator solution to a 50-mL Erlenmeyer flask that is marked to indicate a volume of 35 mL, and place the flask under the condenser of the steam distillation apparatus so that the tip of the condenser is about 4 cm above the surface of the H 3B03. Connect the cooled distillation flask to the distillation ap-

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

899

paratus, place 10 mL of 10 M NaOH in the entry funnel, and run the alkali slowly into the distillation flask. When about O.S mL of alkali remains in the funnel, rinse the funnel rapidly with about S mL of water, and allow about 2 mL to run into the distillation flask before sealing the funnel. Commence steam distillation and stop when the distillate reaches the 3S-mL mark on the receiver flask (distillation time about 4 min). Rinse the condenser, and determine the NH4+ -N in the distillate by titration with 0.002S M H 2 S0 4 from a microburette (1 mL = 70 !lg NH4+-N). The color change at the endpoint is from green to a faint, permanent pink.

39-4.3.5 Acid-insoluble Nitrogen This form of N is the difference between total soil N (Bremner & Mulvaney, 1982) and total hydrolyzable N (section 39-4.3.4). It can also be determined directly by acid digestion of the soil residue remaining after hydrolysis (Cheng & Kurtz, 1963).

39-4.3.6 Amino Acid-Nitrogen Place S mL of the hydrolysate (section 39-4.3.3) in a SO-mL distillation flask, add 1 mL of O.S M NaOH, and heat the flask in boiling water until the volume of the sample is reduced to 2 to 3 mL (approximately 20 min). Allow the flask to cool, add SOO mg of citric acid and 100 mg of ninhydrin, and place the flask in a vigorously boiling water bath, so that its bulb is completely immersed in boiling water. After about 1 min, swirl the flask for a few seconds without removing it from the bath, and allow it to remain in the bath for an additional 9 min. Then cool the flask, add 10 mL of phosphate-borate buffer and 1 mL of S M NaOH, and connect the flask to the steam distillation apparatus. Determine the amount of NH3-N liberated by steam distillation as in section 39-4.3.4 (distillation period about 4 min).

39-4.3.7 Ammonia-Nitrogen Place 10 mL of the hydrolysate (section 39-4.3.3) in a So- or 100-mL distillation flask, add 0.07 ± 0.01 g of MgO, and connect the flask to the steam distillation apparatus. Determine the amount of NH3-N liberated by steam distillation as in section 39-4.3.4, but collect the distillate in a SO-mL Erlenmeyer flask that contains S mL of H 3B0 3-indicator solution and marked to indicate a volume of 20 mL. Discontinue distillation when the distillate reaches the 20-mL mark (distillation period about 2 min).

39-4.3.8 (Ammonia

+

Amino Sugar)-Nitrogen

Place 10 mL of the hydrolysate (section 39-4.3.3) in a 100-mL distillation flask, add 10 mL of phosphate-borate buffer, and connect the flask to the distillation apparatus. Determine the amount of NH3-N liberated by steam distillation as described in section 39-4.3.4 (distillation period about 4 min).

WOLFET AL.

900

39-4.3.9 Amino Sugar-Nitrogen This form of N is taken as the difference between the amounts of N recovered in the preceding two sections. 39-4.3.10 (Serine + Threonine)-Nitrogen Proceed as described in section 39-4.3.8, but after removal of (NH3 + amino sugar)-N by steam distillation with phosphate-borate buffer, detach the flask from the distillation apparatus, and rinse the steam inlet tube with 3 to 5 mL of water. Collect the rinse water in the distillation flask, and cool the flask under a cold water tap. Add 2 mL of periodic acid solution, swirl the flask for about 30 s, add 2 mL of sodium arsenite solution, and connect the flask to the distillation apparatus. Determine the amount of NH3-N liberated by steam distillation as described in section 39-4.3.4 (period of distillation about 4 min). 39-4.3.11 Comments One of the main advantages of the described hydrolysis method is that the N in the different fractions is measured as NH4 + -N, and this can be readily converted to N2 for isotope-ratio analysis. Total N in a given sample may be insufficient for mass spectrometer analysis, and duplicate analyses may have to be combined for 15N determinations. If the 15N percentage is relatively high, it may be diluted with a measured amount of unlabeled N to provide sufficient total N for mass spectrometer requirements (Hauck, 1982). The use of 15N-labeled soils provides a means of tracing the movement of added N into the various organic fractions and determination of the rate at which this occurs if a proper time sequence is employed. Allen et al. (1973) present some typical data that can be obtained by this means. The recommended hydrolysis procedure can be modified in several ways, but the more extensive discussion of the procedure by Bremner (1965) and Stevenson (1982b) should be consulted before doing so. It should also be pointed out that the hydrolysis method presented here causes greater decomposition of amino sugars than more conventional methods. The correction factor for hydrolysis losses of amino sugars is about 1.4 (Bremner, 1965). 39-5 CONCLUSIONS During the past 50 yr, the use of isotopes in the study of soil organic matter dynamics has led to a tremendous increase in knowledge of the system that would not have been possible otherwise. Numerous methods for the use of isotopes have been developed over the years with specific objectives in mind and equipment available at the time. This chapter outlines basic methods currently applicable to organic matter studies, recognizing that improvements and modifications are constantly being made.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

901

The References section provides detailed information on techniques that have been developed.

REFERENCES Allen, A.L., F.J. Stevenson, and L. T. Kurtz. 1973. Chemical distribution of residual fertilizer nitrogen in soil as revealed by nitrogen-15 studies. J. Environ. Qual. 2:120-124. Allison, F.E. 1973. Soil organic matter and its role in crop production. Elsevier Scientific Publ. Co., New York. Andersen, A., G. Nielsen, and H. Sorensen. 1961. Growth chamber for labeling plant material uniformly with tadiocarbon. Physiol. Plant. 14:378-383. Anderson, J.P.E. 1982. Soil respiration. p. 831-871. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Andreux, F., C. Cerri, P.B. Vose, and V.A. Vitorello. 1990. Potential of stable isotope, 15N and BC, methods for determining input and turnover in soils. p. 259-275. In A.F. Harrison et al. (ed.) Nutrient cycling in terrestrial ecosystems. Elsevier Applied Sci., New York. Atlas, R.M., and R. Bartha. 1972. Degradation and mineralization of petroleum by two bacteria isolated from coastal waters. Biotechnol. Bioeng. 14:297-308. Balesdent, J., A. Mariotti, and D. Boisgontier. 1990. Effect oftillage on soil organic carbon mineralization estimated from l3C abundance in maize fields. J. Soil Sci. 41:587-596. Balesdent, J., A. Mariotti, and B. Guillet. 1987. Natural J3C abundance as a tracerfor studies of soil organic matter dynamics. Soil BioI. Biochem. 19:25-30. Balesdent, J., G .H. Wagner, and A. Mariotti. 1988. Soil organic matter turnover in long-term field experiments as revealed by carbon-13 natural abundance. Soil Sci. Soc. Am. J. 52:118-124. Bartha, R., and D. Pramer. 1965. Features of a flask and method for measuring the persistence and biological effects of pesticides in soil. Soil Sci. 100:68-70. Boutton, T. W. 1991a. Stable carbon isotope ratios of natural materials: I. Sample preparation and mass spectrometric analysis. p. 155-171. In D.C. Coleman and B. Fry (ed.) Carbon isotope techniques. Academic Press, New York. Boutton, T.W. 1991b. Stable carbon isotope ratios of natural materials: II. Atmospheric, terrestrial, marine, and freshwater environments. p. 173-185. In D.C. Coleman and B. Fry (ed.) Carbon isotope techniques. Academic Press, New York. Boutton, T.W., A.T. Harrison, and B.N. Smith. 1980. Distribution of biomass of species differing in photosynthetic pathway along an altitudinal transect in southeastern Wyoming grassland. Oecologia 45:287-298. Boutton, T.W., W.W. Wong, D.L. Hachey, L.S. Lee, M.P. Cabrera, and P.D. Klein. 1983. Comparison of quartz and Pyrex tubes for combustion of organic samples for stable carbon isotope analysis. Anal. Chem. 55:1832-1833. Bremner, J.M. 1965. Organic forms of nitrogen. p. 1238-1255. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Bremner, J.M., and C.S. Mulvaney. 1982. Nitrogen-total. p. 595-624. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Broadbent, F.E., and T. Nakashima. 1967. Reversion of fertilizer nitrogen in soils. Soil Sci. Soc. Am. Proc. 31:648-652. Broadbent, F.E., and T. Nakashima. 1974. Mineralization of carbon and nitrogen in soil amended with carbon-13 and nitrogen-IS labeled plant material. Soil Sci. Soc. Am. Proc. 38:313-315. Cerri, c.c., B. Volkoff, and F. Andreux. 1991. Nature and behavior of organic matter in soils under natural forest, and after deforestation, burning and cultivation, near Manaus. Forest Ecol. Manage. 38:247-257. Cheng, H.H., and L.T. Kurtz. 1963. Chemical distribution of added nitrogen in soils. Soil Sci. Soc. Am. Proc. 27:312-316. Cheshire, M.V., and B.S. Griffiths. 1989. The influence of earthworms and cranefly larvae on the decomposition of uniformly 14C labeled plant material in soil. J. Soil Sci. 40:117124. Chichester, F.W. 1970. Transformation of fertilizer nitro~en in soil: II. Total and lsN labeled nitrogen of soil organomineral sedimentation fractIOns. Plant Soil 33:437-456.

902

WOLF ET AL.

Chichester, F.W., J.O. Legg, and G. Stanford. 1975. Relative mineralization rates of indigenous and recently incorporated 15N-Iabeled nitrogen. Soil Sci. 120:455-460. Coleman, D.C., and F.T. Corbin. 1991. Introduction and ordinary counting as currently used. p. 3-9. In D.C. Coleman and B. Fry (ed.) Carbon isotope techniques. Academic Press, New York. Corbin, F.T., and B.A. Swisher. 1986. Radioisotope techniques. p. 265-276. In N.D. Camper (ed.) Research methods in weed science. Southern Weed Sci. Soc., Champaign, IL. Craig, H. 1957. Isotopic standards for carbon and oxygen and correction factors for mass spectrometric analysis of carbon dioxide. Geochim. Cosmochim. Acta 12:133-149. Crawford, D.L. 1978. Lignocellulose decomposition by selected Streptomyces strains. Appl. Environ. Microbiol. 35:1041-1045. . Crawford, D.L., and R.L. Crawford. 1976. Microbial degradation of lignocellulose: The lignin component. Appl. Environ. Microbiol. 31:714-717. Crawford, D.L., R.L. Crawford, and A.L. Pometto. 1977. Preparation of s~ecifically labeled l"C-(lignin)- and 14C-(cellulose)-lignocelluloses and their decomposition by the microflora of soil. Appl. Environ. Microbiol. 33:1247-1251. Deines, P. 1970. Mass spectrometer correction factors for the determination of small isotopic composition variations of carbon and oxygen. Int. J. Mass Spectrom. Ion Phys. 4:283295. Des Marais, D., and J.M. Hayes. 1976. Tube cracker for opening glass-sealed ampoules under vacuum. Anal. Chem. 48:1651-1652. Dzurec, R.S., T.W. Boutton, M.M. Caldwell, and B.N. Smith. 1985. Carbon isotope ratios of soil organic matter and their use in assessing community composition changes in Curlew Valley, Utah. Oecologia 66:17-24. Engel, M.H., and R.J. Maynard. 1989. Preparation of organic matter for stable carbon isotope analysis by sealed tube combustion: A cautionary note. Anal. Chem. 61:19961998. Frazer, J.W., and R. Crawford. 1963. Modifications in the simultaneous determination of carbon, hydrogen, and nitrogen. Mikrochim. Acta 3:561-566. Fried, M., and L.A. Dean. 1952. A concept concerning the measurement of available soil nutrients. Soil Sci. 73:263-271. Gauch, H.G. 1972. Inorganic plant nutrition. Dowden, Hutchinson & Ross, Stroudsburg, PA. Goh, K.M. 1991. Bomb carbon. p. 147-151. In D.C. Coleman and B. Fry (ed.) Carbon isotope techniques. AcadeIDlc Press, New York. Gonfiantini, R. 1981. The 6-notation and the mass spectrometric measurement techniques. p. 35-84. In J.R. Gat and R. Gonfiantini (ed.) Stable isotope hydrology. IAEA, Vienna, Austria. Haider, K., and J.P. Martin. 1975. Decomposition of specifically carbon-14 labeled benzoic and cinnamic acid derivatives in soil. Soil Sci. Soc. Am. Proc. 39:657-662. Haider, K., J.P. Martin, and E. Rietz. 1977. Decomposition in soil of 14C-Iabeled coumaryl alcohols; free and linked into dehydropolymer and plant lignins and model humic acids. Soil Sci. Soc. Am. J. 41:556-562. Harris, D., and E.A. Paul. 1991. Techniques for examining the carbon relationships of plant-microbial symbioses. p. 39-52. In D. C. Coleman and B. Fry (ed.) Carbon isotope techniques. Academic Press, New York. Harrison, A.F., D.D. Harkness, and P.J. Bacon. 1990. The use of bomb- 14C for studying organic matter and N and P dynamics in a woodland soil. p. 246-258. In A.F. Harnson et al. (ed.) Nutrient cycling in terrestrial ecosystems. Elsevier Applied Sci., New York. Hauck, R.D. 1982. Nitrogen-isotope ratio analysis. p. 735-779. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Hayes, J.M. 1983. Practice and principles of isotopic measurements in organic geochemistry. p. 5.1-5.31. In W.G. Meinschein (ed.) Organic geochemistry of contemporaneous and ancient sediments. Soc. for Econ. Paleontologists and Mineralogists, Bloomington, IN. Hurst, H.M., and G.H. Wagner. 1969. Decomposition of 14C-Iabeled cell wall and cytoplasmic fractions from hyaline and melanic fungi. Soil Sci. Soc. Am. Proc. 33:707-711. Insam, H., M.M. Ding, and A. Mariotti. 1991. Utilization of a l3C-enriched tracer for carbon flux studies in a tropical Eucalyptus exserta forest. p. 515-519. In Stable isotopes in plant nutrition, soil fertility, and environmental studies. IAEA, Vienna, Austria.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

903

Jackson, M.L. 1969. Soil chemical analysis-Advanced course. 2nd ed. 11th printing. Published by the author, Madison, WI. Jansson, S.L. 1958. Tracer studies on nitrogen transformations in soil. Annu. Roy. Agric. Coll. Sweden 24:101-361. Jansson, S.L., and J. Persson. 1982. Mineralization and immobilization of soil nitrogen. p. 229-252. In F.J. Stevenson (ed.) Nitrogen in agricultural soils. Agron. Monogr. 22. ASA, CSSA, and SSSA, Madison, WI. Jenkinson, D.S. 1960. The production of ryegrass labeled with carbon-14. Plant Soil 13:279290. Jenkinson, D.S. 1971. Studies on the decomposition of 14C-labeled organic matter in soil. Soil Sci. 111:64-70. Jenkinson, D.S., and J.N. Ladd. 1981. Microbial biomass in soil: Measurement and turnover. p. 415-471. In E.A. Paul and J.N. Ladd (ed.) Soil biochemistry. Vol. 5. Marcel Dekker, New York. Jenkinson, D.S., and D.S. Powlson. 1976. The effects of biocidal treatments on metabolism in soil. V: A method for measuring biomass. Soil BioI. Biochem. 8:209-213. Jenkinson, D.S., and J.H. Rayner. 1977. The turnover of soil organic matter in some Rothamsted classical experiments. Soil Sci. 123:298-305. Kassim, G., J.P. Martin, and K. Haider. 1981. Incorporation of a wide variety of organic substrate carbons into soil biomass as estimated by the fumigation procedure. Soil Sci. Soc. Am. J. 45:1106-1112. Kassim, G., D.E. Stott, J.P. Martin, and K. Haider. 1982. Stabilization and incorporation into biomass of phenolic and benzenoid carbons during biodegradation in soil. Soil Sci. Soc. Am. J. 46:305-309. Kearney, P.C., and A. Kontson. 1976. A simple system to simultaneously measure volatilization and metabolism of pesticides from soils. J. Agric. Food Chern. 24:424-426. Keeney, D.R., and D.W. Nelson. 1982. Nitrogen-inorganic forms. p. 643-698. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Kelley, K.R., and F.J. Stevenson. 1985. Characterization and extractability of immobilized 15N from the soil microbial biomass. Soil BioI. Biochem. 17:517-523. Kemper, W.D., and R.C. Rosenau. 1986. Aggregate stability and size distribution. p. 425442. In A. Klute (ed.) Methods of soil analysis. Part 1. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Ladd, J.N., and M.A. Amato. 1986. The fate of nitrogen from legume and fertilizer sources in soils successively cropped with wheat under field conditions. Soil BioI. Biochem. 18:417-425. Ladd, J.N., and 1.K. Martin. 1984. Soil organic matter studies. p. 67-98. In M.F. L'Annunziata and J.O. Legg (ed.) Isotopes and radiation in agricultural sciences. Vol. 1. Academic Press, London. Ladd, J.N., J.M. Oades, and M. Amato. 1981. Distribution and recovery of nitrogen from legume residues decomposing in soils sown to wheat in the field. Soil BioI. Biochem. 13:251-256. L'Annunziata, M.F. 1979. Radiotracers in agricultural chemistry. Academic Press, New York. L'Annunziata, M.F. 1984. The detection and measurement of radionuclides. p. 141-231. In M.F. L'Annunziata and J.O. Legg (ed.) Isotopes and radiation in agricultural sciences. Vol. 1. Academic Press, London. Lavy, T.L. 1975. Effects of soil pH and moisture on the direct radioassay of herbicides in soil. Weed Sci. 23:49-52. Le Feuvre, R.P., and R.J. Jones. 1988. Static combustion of biological samples sealed in glass tubes as a preparation for Ol3C determination. Analyst 113:817-823. Legg, J.O., F.W. Chichester, G. Stanford, and W.H. DeMar. 1971. Incorporation of 15N_ tagged mineral nitrogen into stable forms of soil organic nitrogen. Soil Sci. Soc. Am. Proc. 35:273-276. Levin, I., B. Kromer, D. Wagenback, and K.O. Munnich. 1987. Carbon isotope measurements of atmospheric CO 2 at a coastal station in Antarctica. Tellus 39B:89-95. Loos, M.A., A. Kontson, and P.C. Kearney. 1980. Inexpensive soil flask for 14C-pesticide degradation studies. Soil BioI. Biochem. 12:583-585. Martin, A., A. Mariotti, J. Balesdent, P. Lavelle, and R. Vuattoux. 1990. Estimate of organic matter turnover rate in a savanna soil by \3C natural abundance measurements. Soil BioI. Biochem. 22:517-523.

904

WOLFET AL.

Martin, J.P., and K. Haider. 1979. Biodegradation of 14C-labeled model and cornstalk lignins, phenols, model phenolase humic polymers, and fungal melanins as influenced by a readily available carbon source and soil. Appl. Environ. Microbiol. 38:283-289. Martin, J.P., K. Haider, and D.C. Wolf. 1972. Synthesis of phenols and phenolic polymers by Hendersonula toruloidea in relation to humic acid formation. Soil Sci. Soc. Am. Proc. 36:311-315. Martin, J.P., H. Zunino, P. Peirano, M. Caiozzi, and K. Haider. 1982. Decomposition of 14C-labeled lignins, model humic acid polymers, and fungal melanins in allophanic soils. Soil BioI. Biochem. 14:289-293. Marvel, J.T., B.B. Brightwell, J.M. Malik, M.L. Sutherland, and M.L. Rueppel. 1978. A simple apparatus and quantitative method for determining the persistence of pesticides in soil. J. Agric. Food Chern. 26:1116-1120. Mayaudon, J. 1971. Use of radiorespirometry in soil microbiology and biochemistry. p. 202-256. In A.D. McLaren and J. Skujins (ed.) Soil biochemistry. Vol. 2. Marcel Dekker, New York. Melillo, J.M., J.D .. Aber, A.E. Linkins, A. Ricca, B. Fry, and K.J. Nadelhoffer. 1989. Carbon and nitrogen dynamics along the decay continuum: Plant litter to soil organic matter. p. 53-62. In M. Clarholm and L. Bergstrom (ed.) Ecology of arable land. Kluwer Acad. Publ., Dordrecht, Netherlands. Mook, W.G., and P.M. Grootes. 1973. The measuring procedure and corrections for the high precision mass spectrometric analysis of isotopic abundance ratios, especially referring to carbon, oxygen, and nitrogen. Int. J. Mass Spectrom. Ion Phys. 12:273-298. Mulvaney, R.L. 1993. Mass spectrometry. p. 11-57. In R. Knowles and T.H. Blackburn (ed.) Nitrogen isotope techmques. Academic Press, San Diego. Myrold, D.D., and J.M. Tiedje. 1986. Simultaneous estimation of several nitrogen cycle rates using 15N: Theory and application. Soil BioI. Biochem. 18:559-568. Nadelhoffer, K.J., and B. Fry. 1988. Controls on natural nitrogen-15 and carbon-13 abundances in forest soil organic matter. Soil Sci. Soc. Am. J. 52:1633-1640. Nakas, J.P., and D.A. Klein. 1979. Decomposition of microbial cell components in a semiarid grassland soil. Appl. Environ. Mlcrobiol. 38:454-460. Nelson, D.W., and L.E. Sommers. 1982. Total carbon, organic carbon, and organic matter. p. 539-580. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Oades, I.M., and G.H. Wagner. 1971. Biosynthesis of sugars in soils incubated with I'C_ glucose and 14C-dextran. Soil Sci. Soc. Am. Proc. 35:914-917. O'Leary, M.H. 1988. Carbon isotopes in photosynthesis. BioScience 38:328-336. Osmond, C.B., K. Winter, and H. Ziegler. 1982. Functional significance of different pathways of CO2 fixation in photosynthesis. p. 479-547. In O.L. Lange et al. (ed.) Plant physiological ecology II: Water relations and carbon assimilation. Springer-Verlag, Berlin. Paul, E.A., and F.E. Clark. 1989. Soil microbiology and biochemistry. Academic Press, New York. Paul, E.A., and I.A. van Veen. 1978. The use of tracers to determine the dynamic nature of organic matter. p. 61-102. Trans. 11th Int. Congr. Soil Sci. Vol. 3. Edmonton, Canada. Post, W.M., W.R. Emanuel, P.l. Zinke, and A.G. Stangenberger. 1982. Soil carbon pools and world life zones. Nature (London) 298:156-159. Post, W.M., J. Pastor, P.J. Zinke, and A.G. Stangenberger. 1985. Global patterns of soil nitrogen storage. Nature (London) 317:613-616. Prentice, K.C., and I.Y. Fung. 1990. The sensitivity of terrestrial carbon storage to climate change. Nature (London) 346:48-51. Reyes, V.G., and J.M. Tiedje. 1973. Metabolism of 14C uniformly labeled organic material by woodlice (Isopoda: Oniscoidea) and soil microorganisms. Soil BioI. Biochem. 5: 603-611. Russell, E.W. 1961. Soil conditions and plant growth. John Wiley and Sons, New York. Santrock, J., S. Studley, and J. Hayes. 1985. Isotopic analyses based on the mass spectrum of carbon dioxide. Anal. Chern. 57:1444-1448. Scharpenseel, H.W., and H.U. Neue. 1984. Use of isotopes in studying the dynamics of organic matter in soils. p. 273-310. In Organic matter and rice. Int. Rice Res. Inst., Manila, Philippines. Schlesinger, W.H. 1990. Evidence from chronosequence studies for a low carbon storage potential of soils. Nature (London) 348:232-234.

ISOTOPIC METHODS FOR ORGANIC MATTER DYNAMICS

905

Schnitzer, M. 1982. Organic matter characterization. p. 581-594. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Scott, H.D., and R.E. Phillips. 1972. Diffusion of selected herbicides in soil. Soil Sci. Soc. Am. Proc. 36:714-719. Scully, N.J., W. Chorney, G. Kostal, R. Watanabe, J. Skok, and J.w. Glattfeld. 1956. Biosynthesis in 14C-labeled plants: Their use in agricultural and biological research. Vol. 12. p. 377-385. In Proc. Int. Conf. Peaceful Uses of Atomic Energy, Geneva, Switzerland, 8-20 Aug. 1955. United Nations, New York. Sissons, C.H. 1976. Improved technique for accurate and convenient assay of biological reactions liberating 14C0 2 . Anal. Biochem. 70:454-462. Skjemstad, J.O., R.P. Le Feuvre, and R.E. Prebble. 1990. Turnover of soil organic matter under pasture as determined by BC natural abundance. Aust. J. Soil Res. 28:267-276. Smith, B.N., and S. Epstein. 1971. Two categories of BC/12C ratios for higher plants. Plant Physiol. 47:380-384. Smith, J.H., F.E. Allison, and J.F. Mullins. 1962. Design and operation of a carbon-14 biosynthesis chamber. USDA-ARS Misc. Publ. 911. U.S. Gov. Print. Office, Washington, DC. Sofer, Z. 1980. Preparation of carbon dioxide for stable carbon isotope analysis of petroleum fractions. Anal. Chern. 52: 1389-1391. Stevenson, F.J. 1965. Gross chemical fractionation of organic matter. p. 1409-1421. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Stevenson, F.J. 1982a. Humus chemistry, genesis, composition, reactions. John Wiley and Sons, New York. Stevenson, F.J. 1982b. Nitrogen-organic forms. p. 625-641. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2, 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Stevenson, F.J. 1986. Cycles of soil: Carbon, nitrogen, phosphorus, sulfur, micronutrients. John Wiley and Sons, New York. Stevenson, F.J., and E.T. Elliott. 1989. Methodologies for assessing the quantity and quality of soil organic matter. p. 173-199. In D.C. Coleman et al. (ed.) Dynamics of soil organic matter in trop'ical ecosystems. NifTAL Project, Dep. Agronomy and Soil Science, Univ. of Hawaii, Honolulu. Stewart, B.A., L.K. Porter, and D.D. Johnson. 1963. Immobilization and mineralization of nitrogen in several organic fractions of soil. Soil Sci. Soc. Am. Proc. 27:302-304. Stotzky, G. 1965. Microbial respiration. p. 1550-1572. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Stotzky, G., M.W. Broder, J.D. Doyle, and R.A. Jones. 1993. Selected methods for the detection and assessment of ecological effects resulting from the release of genetically engineered microorganisms to the terrestrial environment. In S.L. Neidleman and A.I. Laskin (ed.) Adv. Appl. Microbiol. 38:1-93. Stout, J.D., K.M. Goh, and T.A. Rafter. 1981. Chemistry and turnover of naturally occurring resistant organic compounds in soil. p. 1-73. In E.A. Paul and J.N. Ladd (ed.) Soil biochemistry. Vol. 5. Marcel Dekker, New York. Swerhone, G.D.W., K.A. Hobson, C. van Kessel, and T.W. Boutton. 1991. An economical method for the preparation of plant and animal tissue for OBC analysis. Commun. Soil Sci. Plant Anal. 22:177-190. Teeri, J.A., and L.G. Stowe. 1976. Climatic patterns and the distribution of C4 grasses in North America. Oecologia 23:1-12. Tiessen, H., and J.W.B. Stewart. 1983. Particle size fractions and their use in studies of soil organic matter: II. Cultivation effects on organic matter composition in size fractions. Soil Sci. Soc. Am. J. 47:509-514. Verma, L., J.P. Martin, and K. Haider. 1975. Decomposition of carbon-14-labeled proteins, peptides: and amino acids; free and complexed with humic polymers. Soil Sci. Soc. Am. Proc. 39.279-284. Vitorello, Y.A., C.c. Cerri, F. Andreux, C. Feller, and R.L. Victoria. 1989. Organic matter and natural carbon-13 distribution in forested and cultivated Oxisols. Soil Sci. Soc. Am. J.53:773-778. Voroney, R.P., J.P. Winter, and E.G. Gregorich. 1991. Microbe/plant/soil interactions. p. 77-99. In D.C. Coleman and B. Fry (ed.) Carbon isotope techniques. Academic Press, New York.

WOLF ET AL. Wagner, G.H. 1975. Microbial growth and carbon turnover. p. 269-305. In E.A. Paul and A.D. McLaren (ed.) Soil biochemistry. Vol. 3. Marcel Dekker, New York. Wagner, G.H., and K.S. Chahal. 1966. Decomposition of carbon-14 labeled atrazine in soil samples from Sanborn Field. Soil Sci. Soc. Am. Proc. 30:752-754. Wagner, G.H., and A.M. Krzywicka. 1975. Decomposition of algal tissues in soil. p. 202-207. In G. Kilbertus et al. (ed.) Biodegradation et humification. Pierron Publ., Sarreguemines, France. Warembourg, F.R., and J. Kummerow. 1991. Photosynthesis/translocation studies in terrestrial ecosystems. p. 11-37. In D.C. Coleman and B. Fry (ed.) Carbon isotope techniques. Academic Press, New York. Wolf, D.C., and J.O. Legg. 1984. Soil microbiolo~. p. 99-139. In M.F. L'Annunziata and J.O. Legg (ed.) Isotopes and radiation in agncultural sciences. Vol. 1. Academic Press, London. Zunino, H., F. Borie, S. Aguilera, J.P. Martin, and K. Haider. 1982. Decomposition of 14C-labeled glucose, plant and microbial products and phenols in volcanic ash-derived soils of Chile. Soil BlOl. Biochem. 14:37-43.

Published 1994

Chapter 40 Practical Considerations in the Use of Nitrogen Tracers in Agricultural and Environmental Research R. D. HAUCK, Tennessee Valley Authority, Muscle Shoals, Alabama

J. J. MEISINGER, USDA-ARS, Beltsville, Maryland R. L. MULVANEY, University of Illinois, Urbana, Illinois

Agricultural research is challenged to develop crop production systems that are productive and economically viable over time, socially and politically acceptable, and conserving of natural resources. Efficient use of N, regardless of its source, is imperative for achieving these ends. Many of the questions centered about efficient N use cannot be answered satisfactorily, if at all, without use of the stable isotopes, 14N and 15N, as tracers. First used in agronomic studies by Norman and Werkman (1945), N tracer methodology has been used almost routinely in recent years to study N transformation processes in plants and animals, air, soils, and waters, and to understand, evaluate, and monitor the effects of different approaches to N management in production agriculture. Stable (nonradioactive) isotopes are used almost exclusively when tracer techniques are needed in agricultural and environmental studies. The extremely short half-lives of the four radionuclides of N, ranging from 0.0125 to 603 s, militate against their use for biological studies. Of these radionuclides, only 13N has been used to a limited extent as a highly sensitive tracer in metabolic studies (mainly of biological N2 fixation) that can be completed within about 2 h. Use of the stable isotopes of masses 14 and 15 as tracers is based on the fact that these isotopes occur naturally in an almost constant ratio. Except for slight variations in the N isotopic composition of natural nitrogenous substances, the ratio of 14Nj15N is about 272:1 (i.e., naturally occurring N contains about 0.366 at. % 15N or about 3660 ppm 15N). The significance of natural variations from these values will be discussed later. Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 907

908

HAUCK ET AL.

Stable N tracers are materials that contain an unnaturally high or low concentration of either 14N or 15N. Until the 1970s, almost all N-tracer experiments were conducted with substances that were enriched with 15N, i.e., had higher than natural concentrations of 15N. Since that time, several agricultural studies have been conducted with 15N-depleted (14N-enriched) materials. Use of either type of tracer material requires that its N-isotopic composition at the time of sampling be significantly different from that of other N in the system under study. Adding 14N_ or 15N-enriched material to a system results in an increase in the total respective 14N or 15N content of the system. When the tracer N mixes with other N having the same form as the tracer, both the tracer and non tracer N lose their isotopic identities (an exception is Nz formed during denitrification [Hauck, 1982]) and the mixture will have a ratio of 14N/15N which is different from that of its components. The change in N-isotope ratio permits the investigator to follow the course of the tracer within the system. The extent of change in isotope ratio is used to calculate the extent to which the tracer has become part of the system. Measuring the total amount of tracer in the system may permit in some studies accurate determination of N loss. Generally, addition of a tracer-labeled material to a system (e.g., soil) enables one to determine the amount of N from the labeled material taken up by plants, the distribution of that N in plant parts or soil physical and chemical fractions, and the relative contributions of the added and resident N to N components moving from soil into air or waters. Most N-tracer studies in agriculture involve some variation of the above technique of isotope-dilution analysis. Several important isotope dilution expressions and their application were discussed by Hauck and Bremner (1976). Although the technique is simple in concept, problems are encountered in correctly interpreting the results obtained. Such problems often result from questionable assumptions made when calculating quantities involved in gross N transformations. Hauck and Bremner (1976) listed three fundamental assumptions central to the use of N isotopes as tracers in biological systems and 12 assumptions specifically relevant to soil N-transformation studies. Mulvaney (1991) discussed the significance of three common assumptions made in N-isotope pool dilution studies: (i) tracer N is distributed uniformly throughout the system or portion of system under study; (ii) the processes under study occur at constant rates; and (iii) N leaving the labeled N pool does not return. Other selected references from the extensive literature in which N tracer data interpretation is discussed include papers by Kirkham and Bartholomew (1954, 1955), Jansson (1958, 1966), Hauck (1973, 1978), Fried (1978), Koike and Hattori (1978), Blackburn (1979), Guiraud (1984), Shen et al. (1984), Jenkinson et al. (1985), Hart et al. (1986), Myrold and Tiedje (1986), Barraclough and Smith (1987), Guiraud et al. (1989), and chapter 42 in this book. Sample variability usually is the greatest single source of methodological error in N-tracer research. Analytical error (i.e., the determination of N-isotope ratio) generally is a negligible fraction of the cumulative error involved in sample collection, processing, and chemical analysis. Refer-

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

909

ence to most of the 15N literature in which procedural errors are discussed can be found in reviews by Hauck and Bremner (1964, 1976), Bremner (1965a), Bremner et al. (1966), Martin and Ross (1968), Fiedler and Proksch (1975), Hauck (1982), Fiedler (1984), and Mulvaney (1993). Until recently, methods for N-isotope-ratio analysis were developed largely by Rittenberg and Sprinson, and most of the analytical procedures currently used in many laboratories are modifications of methods described in their papers (Rittenberg et al., 1939; Rittenberg, 1948; Sprinson & Rittenberg, 1948, 1949). Subsequent articles providing additional details of 15N methodology in agricultural studies include those by Burris and Wilson (1957), San Pietro (1957), Capindale and Tomlin (1957), Junk and Svec (1958), Smith et al. (1963), Cho and Haunold (1966), Martin and Ross (1968), Fiedler and Proksch (1975), Hauck and Bremner (1976), Edwards (1978), Bergersen (1980), Buresh et al. (1982), Chen et al. (1991), Mulvaney (1993), and in the bibliography compiled by Hauck and Bystrom (1970). Since the 1970s, improved instrumentation for sample preparation and N-isotope-ratio analysis has permitted both automation of analytical procedures and the routine determination of isotopic composition of microgram quantities of N. Details of these recent developments are reviewed by Mulvaney (1993). The methods outlined here for N-isotope-ratio analysis of samples obtained from studies using N tracers are similar to but far less comprehensive than those given in the treatments by Bremner (1965a) and Hauck (1982) in the first and second editions of this book. Knowledge of procedural detail as well as new developments is imperative for successful use of N tracers by investigators who perform all phases of their research. However, researchers who make use of analytical services now available may not require detailed knowledge of the analytical procedures used. The level of detail given here may be sufficient to heighten awareness of procedural problems and determine the extent to which analytical precision and accuracy are affected by all phases of the research operation. Accordingly, in this chapter less emphasis is placed on the determination of N-isotope ratio and more on the overall conduct of N-tracer research, including the preparation of tracer materials, calculation of tracer needs, and collection and preparation of sample, depending on kind of study and objectives. 40-1 PREPARING lSN-LABELED MATERIALS 40-1.1 Principles Most N-tracer materials are derived from NH/ or N0 3- enriched in 15N (i.e., containing an unnaturally high concentration of 15N). Materials with an unnaturally low 15N concentration (i.e., 14N-enriched or 15N-depleted) also can be prepared. The commercial preparation of the stable N isotopes resulting in the concentration of either depends on their

910

HAUCK ET AL.

differences in mass and, therefore, their physical properties; 14N and 15N in the form of NH3, NH/ , or N oxides behave differently in exchange columns or distillation columns. Until the 1960s, 15N was concentrated mainly by counter-current exchange of NH3 with NH/, the 15N atoms being extracted from the NH3 vapor and concentrated in the liquid phase. Currently, 15N is concentrated mainly by exchange of NO with HN03 or by cryogenic distillation of NO, the labeled product being collected as HN03 which may be subsequently reduced to NH3. A variety of tracer materials is prepared from the labeled NH3 or N0 3- through chemical synthesis or biosynthesis. The term atom % 15N or at. % 15N commonly is used to denote 15N concentration: nt

at.'lO

15

N

=

no. of 15N atoms x 100 no. of 4N + 15N) atoms

e

[1]

Reference is made in the following discussion to atom % 15N and atom % 15N excess; either term can be used for expressing 15N concentration. Atom % 15N excess refers to the measured atom % concentration of 15N minus the natural background concentration. Users of this term assume a constant value for natural 15N abundance, which is incorrect, or they determine this value for the natural material that is being used as a standard, which may not be the reference material used by others. A recommended approach is to relate all N-isotope ratios to the accepted standard value of 0.3663 at. % 15N for atmospheric N2 (Junk & Svec, 1958) and not use the term atom % 15N excess (Hauck, 1982). The term delta 15N (6 15N) commonly is used when expressing 15N concentration in the range of natural abundance; it is the 15N concentration expressed as parts per thousand differences from the 15N/14N ratio in a standard, usually atmospheric N2. 6 15N

=

e5N/ 14N)x-e5NI 14N)atm x 1000 e SN/ 14N)atm

[2]

where (15N/14N)x and (15N/14N)atm are the N-isotope ratios of the sample and atmospheric N2, respectively. The terms per mil difference and per mil enrichment also have been used. Sometimes 615N values are based on the expression s. 15N

u

=

(at. % lsN)x-(at. % 15N)atm (at. % 15N)atm

0 00

x1

[3]

Values of 6 15N calculated from the expression based on the use of isotope ratios (Eq. [2]) are slightly lower than those based on the use of 15N concentrations (Eq. [3]), but the differences due to method of calculation are well within the limits of analytical error and can be considered negligible.

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

911

The level of 15N (or 14N) enrichment needed in the labeled material is determined by the amount of isotope dilution that occurs during the experiment, and this, in tum, is determined by several factors, including the nature, objectives, and duration of the study. Usually, 15N is purchased in a concentration higher than needed for most studies even when it is in the form to be used. When preparing relatively large amounts of labeled material of low enrichments (e.g., tens to hundreds of grams in the range of I to 5 at. % 15N), highly enriched material is intimately mixed with an amount of the same material of natural N-isotopic composition such that the mixture has the desired N-isotopic composition. This procedure is especially more convenient and less time-consuming when the purchased 15N is in a chemical form other than needed and must be chemically converted. For example, highly enriched NH3 is used to synthesize a small amount of highly enriched urea that is then diluted to the needed isotopic composition and amount with unlabeled urea. To ensure homogeneity, the separate components differing in N-isotopic composition are dissolved together and recrystalized from solution. Depending on experimental objectives, the labeled material can be used as fine crystals, redissolved and used in liquid form, or compressed into particles within a specified size range through granulation or pelleting, followed, if necessary, by crushing and screening. 40-1.2 Determining Nitrogen-iS Concentration Needed The general formula for assaying a compound in a mixture of compounds using isotope dilution calculations is: X2

= [(ClIC2) -

1]·XI·(M2IMI)

[4]

where Xl X2 C2

=

= the weight of the unknown (unlabeled) compound, = the isotopic concentration (expressed as at. % excess) of the com-

the weight of the tracer compound added,

CI

=

pound recovered from the mixture, the isotopic concentration of the original tracer compound.

The term M21MI corrects for the change in molecular weight of the compound as its isotopic composition changes upon dilution. This calculation is unnecessary when amounts are expressed as equivalents, and, where tracer concentrations are low, the correction is negligible. The experimental accuracy of agronomic studies may be affected by the level of 15N enrichment in the applied N source. For example, where N tracer is used to determine plant uptake of applied N or the amount of applied N remaining in soil, standard errors often increase with decrease in 15N concentration in the applied N. This can readily be seen from Eq. [5] for calculating the percent recovery of applied N by crop plants.

HAUCK ET AL.

912

% N recovered = 100 P(c-b) f(a-b)

[5]

where P

f

= total plant N (e.g., in milliequivalents),

= fertilizer N applied (e.g., in milliequivalents), and a, b, and c = the at. % 15N concentrations of the fertilizer, soil, and plants, respectively.

When 15N-enriched materials are used, the value for a always is larger than for c (plants cannot take up more fertilizer N than applied). Though not always possible, when a, b, and c are determined with the same level of precision, the absolute error is constant but the relative error in determining these values increases as the values for a and c approach b, i.e., as (c-b) approaches zero. Mathematical expressions for estimating error in N uptake studies clearly show a marked increase in error as the values for a and c decrease, i.e., as the 15N enrichment of the fertilizer decreases. For 15N-depleted fertilizer, error increases with increase in 15N enrichment, i.e., as the 15N concentration in the fertilizer approaches the level of natural abundance. The significance of this error in relation to other errors can be estimated from Eq. [6] (adapted from Hauck, 1978), which is for propagation of errors for assessing the relative errors in calculating the mass of 15N:

!:1LN LN

1[!:1DM]2 [!:1TN]2 [ !:1C DM TN (c-b)

-y

]2 [

!:1a (a-b)

]2 [ (c-a) (!:1b) ]2 (c-b) (a-b)

[6]

where

LN = plant labeled N (mass 15N/ha), DM = plant dry matter (mass/ha), TN = total N concentration (g N/kg), a, b, c = at. % 15N concentrations in the 15N source, control plants, 15N_ treated plants, respectively, and !:1 = the uncertainty in the respective parameter. This equation shows that the relative uncertainty in the mass of labeled N will be controlled by the component with the largest relative uncertainty (largest coefficient of variation, CV). In the majority of isotope experiments enough labeled N will be added to give an enrichment/depletion of at least 0.2% in the final material of interest; therefore, the terms in Eq. [6] involving (c-b) and (a-b) will be very small. However, these terms can become large in samples with small enrichments, such as from studies tracing N through variation in natural abundance, or studies of labeled N entering a large and slowly reactive pool (e.g., soil organic matter). For most agronomic studies, typical CVs for plant dry matter would be 6 to

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

913

15%, for total N concentration, 1 to 3%, and for 15N enrichment, 0.1 to 0.3% when 15N enrichment is 0.2 at. % excess or higher. A general ranking of relative variations would be: field plots > soil/plant samples > subsamples > total N determinations > 15N determinations (Bartholomew, 1964; Hauck, 1978; Broadbent & Carlton, 1980; Pruden et aI., 1985; Saffigna, 1988). Materials with a low level of 15N enrichment (about 1 at. % or less) and 15N-depleted materials are useful for conducting single-season N uptake studies and tracing the movement of N0 3- derived from the labeled material. In many studies, a low-level tracer can be used for determining the amount of added material remaining in the system and, occasionally with soils of low organic matter content, for determining the amount of N applied one season taken up by plants in the succeeding season. Some typical calculations will illustrate the basis for determining the needed level of 15N enrichment. Assume that soil at the experimental site contains 0.2% N (about 4500 kg of Nlha) , to which is added 15N-enriched fertilizer (1.0000 at. %) at a rate equivalent to 150 kg N/ha. Assume that 25% of the applied N remains in soil after harvest of the first crop. Question: Can the applied N remaining in soil be detected with a satisfactory level of precision? A simplified Eq. [4] is used but in the following calculation the term Xl is the amount of tracer N added, X2 is the amount of (unlabeled) soil N, C2 is the isotopic concentration of N in the soil sample taken after harvest (expressed as at. % excess), and C1 is the isotopic concentration (at. % excess) of the added tracer N. The term M21 M1 that corrects for the change in molecular weight of the tracer N is not used. Assume soil to have a natural 15N abundance of 0.3663 at. % and analytical precision to be ± 0.002 at. % 15N. X2 4500

=

=

[(ClIC2) - l)·Xl

- 0.3663) ] 0 025 [ (1.0000 C2 - 1 . 15 . . 121 C2

=

0.6337

Solving for C2, the at. % 15N excess (15N concentration in excess of natural abundance) of the total soil N is found to be 0.0052, which is barely within the limits of detection. In the above example, use of a fertilizer containing 2.0000 at. % 15N results in an average 15N concentration in total soil N of 0.3795 or 0.0132 at. % excess, which is well within analytical precision and sufficiently different from natural 15N abundance to decrease some of the interpretive errors associated with spatial variability among soil samples. Use of 15N-depleted fertilizer (e.g., containing 0.009 at. % 15N) results in a value for C2 that is only 0.003 at. % 15N different from natural abundance; therefore, this tracer material would not be suitable for studying N remaining in soil under the experimental conditions cited.

914

HAUCK ET AL.

Assume as before that 150 kg of N/ha is added to soil containing 4500 kg of total N/ha, and that 25% of the added N remains immobilized in the organic matter (mostly biomass) after harvest. Assume that 75% of the immobilized N is in the labile (biodegradable) fraction and has an isotopic composition identical to that of the initially added N, and that the remaining 25 % of the immobilized N is in refractory, unavailable forms and need not be considered in the calculations. Further, assume that in the second growing season 10% of the immobilized labeled N in the biomass (labile organic matter) and 2% of the total soil N originally present are mineralized and that plants take up 50% of the N made available from each source (one labeled, the other unlabeled). Question: What is the minimum 15N concentration of the N added the first season such that plants grown during the second season contain 15N at a concentration no lower than 0.4663 at. %? The general isotope dilution formula, Eq. [4], is the basis of Eq. [7] used to answer the above question.

x

=

_(T_'N--,--)-,---(c_-b--,-)

[7]

a

where X is the amount of labeled N in the plant, TN is the total amount of N in the plant (from all sources), a is the at. % excess in the N originally added, and band c are the 15N concentrations expressed as at. % in the original soil and plants, respectively. Labeled N originally added, 150 kglha, 15N concentration, a, is unknown Soil N, 4500 kglha, at. % 15N = 0.3663 Labeled N immobilized, first season, 150· (0.25) = 37.5 kg/ha Labeled N in biomass, 37.5· (0.75) = 28.13 kg/ha Original soil N mineralized, second season, 4500· (0.02) = 90 kg/ha Labeled biomass N mineralized, second season, 28.13· (0.1) = 2.813 kg/ha Total N in plants, second season, (90 + 2.813) . (0.5) = 46.4065 kg/ha Average 15N concentration in plants, 0.4663 at. %. a=

46.4065(0.4663 - 0.3663) 2.813 (0.5) = 3.2994

Minimum 15N concentration in added N required: (3.2994

+ 0.3663) = 3.6657 at.

%

Use of somewhat higher concentrations of 15N would be advisable to compensate for differences in the estimated vs. actual dilution occurring during the study. Expressions other than Eq. [7] can be used to calculate the needed amount of 15N. Many modifications of Eq. [4] for calculating isotope dilution are found in the literature and several are summarized by Hauck and

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

915

Bremner (1976). The assumptions that were made in the above example are based on typical values for immobilization, mineralization, and plant uptake as determined from N-tracer studies. Such assumptions have been found useful in anticipating the extent of isotope dilution to estimate 15N needs, depending on study objectives and experimental circumstances. The above calculations do not consider the almost negligible fraction of N released from refractory soil constituents, return of crop residue to the available N pool, between-season and second-year N inputs, or corrections for the change of atom mass of the labeled N. The general isotope dilution formula can be modified as needed to make crude estimates including such factors. More precise estimates of N mineralized from crop residues differing in e/N ratio can be made using the equations given by Vigil and Kissel (1991). 40-1.3 Fertilizers 40-1.3.1 Enrichment Level and Form Nitrogen tracer studies in agriculture and related fields usually are conducted using labeled materials in the chemical forms available at time of purchase or in forms readily synthesized from the purchased stock. Except for 15N-depleted materials, most often 15N is purchased at an enrichment level higher than needed for a study, followed by dilution to the needed isotopic composition. The relatively high cost of 15N has been a main reason why most agronomic N-tracer studies have been conducted with small grains (high population density) in greenhouse pots or microsize field plots. Field experiments with maize (Zea mays L.) requiring larger plots (e.g., 6 x 10 m) commonly are conducted with lower-cost 15N-depleted materials or materials of low enrichment ( < 1 at. % 15N). Often, the labeled N source is applied as a dilute solution to facilitate uniform application (see 40-2.3.1). However, spurious results can be obtained when the physical form of the fertilizer affects its immediate chemical and biochemical reactions in soil microsites, and these reactions, in turn, significantly affect subsequent transformations, movement, and plant uptake of N (Hauck, 1984). Unpublished studies with 15N-Iabeled soil or fertilizers have shown significant differences in the behavior in soil between fertilizer particles vs. their dilute solution, including rate of nitrification and effect on urease activity, ammonia volatilization, and solubilization followed by mineralization of soil organic matter, as affected by fertilizer type, rate, and disposition in soil. However, these results were obtained in laboratory and greenhouse studies and the practical significance of differences in effects of solids vs. dilute solutions has not been established in the field. Little attention has been paid to the potential importance of micro site reactions as affected by the physical characteristics of the N source, neither in field studies using 15N nor in nontracer studies. Because published data on this subject appear not to be available, one can only speculate on its significance. In all studies under the program described by Hauck and

916

HAUCKET AL.

Kilmer (1975), granulated 15N-enriched or 15N-depleted fertilizers were used to obviate unknown differences in fertilizer behavior resulting from differences in physical form of the labeled and unlabeled fertilizers used. Whenever feasible, we suggest using a physical form of a labeled material that closely matches that of its commercial equivalent used by farmers. 40-1.3.2 Conventional

Conventional fertilizers as used here means water-soluble, fast-acting, usually inorganic materials used in crop production, such as (NH4hS04' NH4N0 3 , (NH 4hHP0 4, and urea. 40-1.3.2.1. Dilution to Needed 15N Concentration. The general isotope dilution formula, Eq. [4], can be used to determine the amounts of unlabeled and labeled materials that are mixed to obtain the desired 15N concentration. Assume that 100 g of urea with a 15N concentration of 2.0000 at. % is to be made from urea containing 10.3663 at. % 15N. Equation [4] can be rewritten as follows: (X1·C1)

+ (X2. C2)

z=-------100

[8]

where Xl and X2 = the weights of labeled and unlabeled N CI and C2 = the 15N concentrations of Xl and X2, respectively, expressed as at. % Z = the at. % 15N concentration of the mixture

For the example given, (Xl + X2) 2.0000

=

=

X1(1O.3663)

100.

+ i~~O -

X1)(0.3663)

Xl = 16.337 (g N as labeled urea) X2 = 83.663 (g N as unlabeled urea)

Correcting for the differences in mass, the corresponding values are 16.326 and 83.674. Such correction usually need not be applied unless an exact 15N concentration is needed. In any case, the exact 15N concentration of the final mixture should be determined through isotope-ratio analysis. 40-1.3.2.2 Product Preparation. The labeled source material and its unlabeled diluent are dissolved together to ensure uniform mixing. Dry mixing is not recommended. Following dissolution (usually in water) the diluted source material is crystallized through evaporation, and then redissolved in, or washed with methyl alcohol, recrystallized, and dried. Use

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

917

of methyl alcohol is recommended because it is usually more chemically pure than ethyl alcohol. The dry, labeled N source can be used as fine crystals, redissolved for sprinkle application, or made into particles. Granulation of materials such as (NH4hS04 requires experience and apparatus not usually found in the research laboratory. However, labeled source material can be compacted into hard cylinders or cakes in a Carver press at about 10 000 psi. The cakes are then crushed and screened to the required particle size. Moistening with water sometimes is required for adequate compaction. 40-1.3.3 Slow-Release Many chemicals have been evaluated as slow-release N fertilizers (e.g., see Hauck, 1985) but the ones that have been studied the most extensively are the ureaforms, isobutylidene diurea, oxamide, and sulfurcoated urea. Only a few of the studies have been made with 15N-Iabeled materials, e.g., with ureaform (Brown & Volk, 1966), oxamide (Westerman et aI., 1972), and oxamide and isobutylidene diurea (Rubio & Hauck, 1986). The synthesis of these materials labeled with 15N is described by Hauck (1994). Preparation of relatively small (kilogram) amounts of S-coated urea for use in field studies is not recommended because the laboratory-scale preparation most probably will result in a product with significantly different dissolution rates than commercial products. Differences between commercially and laboratory synthesized products are less apparent for products such as oxamide whose slow-release characteristics are a function of their molecular structure rather than a physical coating process. 40-1.3.4 Organic Residues 40-1.3.4.1 Plant Tissues. The value of 15N-Iabeled plant tissues as tracers should not be overlooked. Large quantities of grain and stover are produced as valuable by-products of agronomic studies using 15N conducted over a wide range of fertilizer practices and crop management systems. These plant parts can be used in many types of studies (e.g., determining the distribution of applied N in different plant parts and in different N fractions) as affected by N application rate and management. The materials should be available to researchers unable to conduct their own field experiments but who would make measurements other than those made by the original investigator(s) having different research objectives. Or, plant tissues with a sufficient level of 15N enrichment can be used elsewhere from the originating site to study N release and movement from different crop residues. Further details on the use of 15N-Iabeled plant residues for field studies of mineralization/immobilization are given in chapters 39 and 42. 40-1.3.4.2 Animal Wastes. Large amounts of 15N-Iabeled animal manure suitable for field studies of manure-N movement have been produced

918

HAUCKET AL.

as by-products of cattle-feeding experiments (Faust et aI., 1963). Currently, at least three groups of investigators have fed 15N-enriched grain to cattle or poultry specifically to produce 15N-Iabeled animal manures for studies in soil columns or field. How the excreta are handled, processed, and stored depends on study objectives. Important considerations include preventing NH3 loss from NH/ salts and hydrolyzed urea and protein derivatives, accurately determining the N-isotopic composition of different manure components, and separating undigested material such as plant grain and fiber from the manure components of interest. Kirchmann (1985), for example, found that the N transformations of 15N-labeled poultry manure were markedly affected by manure pretreatment. Some investigators have added 15N-labeled urea to urine to study NH3 evolution from simulated urine patches. Although this approach has merit for the purpose intended, it cannot be used to study the reactions and movement of different N forms in manure. Uniform methodologies for producing, handling, and studying 15N-labeled animal wastes have not been established. Suggestions for preparing and characterizing labeled manures can be found in Rauhe and Bornak (1970), Rauhe et aI. (1973), and Kirchmann (1989, 1990). 40-1.3.5 Soil Eight methods have been used to accurately determine the amount of atmospheric N2 fixed by plants (Hauck, 1979). The most common N-tracer techniques for measuring N2 fixation in the field involve isotope dilution. The main problem associated with all isotope dilution approaches for this purpose is that their accuracy depends on how well the nonfixing control plants simulate the N uptake patterns of the N-fixing plants under study (for discussion, see Rennie, 1986; Weaver, 1986; Phillips et aI., 1986; Vose & Victoria, 1986). The use of labeled soil for field measurement of biological N2 fixation probably requires the fewest assumptions that cannot be experimentally validated. Legg and Sloger (1975) labeled soil organic matter with 15N by stimulating rapid turn-over of applied 15N-enriched NH/ and glucose. Using the remineralized amount of initially immobilized 15N as an index of mineralize able soil organic N, they estimated from isotope dilution calculations the amount of total N in soybeans [Glycine max(L.)] that was fixed from the atmosphere. The calculations assumed uniform labeling of the soil organic matter fractions or the fractions that release plant-available N during crop growth. This is not a valid assumption, but the release of 15N from soil organic matter accurately reflects mineralizeable N when the 15N, initially immobilized in the biomass, has equilibrated with other mineralizeable soil organic matter fractions. Such equilibration can be determined by following the 15N through the main chemical fractions (e.g., hydrolyzable distillable, nonhydrolyzable nondistillable, etc.) until no further change in 15N distribution is observed. Three or more years may be required to prepare 15N-Iabeled soil that can be used to accurately study the release of mineralize able N. Once

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

919

prepared and characterized, the labeled soil is a valuable tool that can be used to study, for example, N release as affected by a particular treatment or the relative uptake of labeled soil N and unlabeled applied N, as affected by plant species. 40-2 FIELD STUDY TECHNIQUES 40-2.1 Principles Agricultural scientists study N cycle processes to: (i) improve production, (ii) control N losses to the environment, and (iii) refine and expand understanding of these processes. They have used two fundamentally different approaches to these objectives (i.e., through use of either labeled or nonlabeled N sources). Nonlabeled N studies focus on the total N inputs, total N outputs, and the net change in various soil N pools (organic and inorganic). Studies using labeled N focus on the interaction of the labeled N within the soil-plant system by tracing the course of the 15N atoms throughout the system. Consider the addition of 15N-labeled ammoniacal fertilizer to a soil where 10% of the 15N is exchanged with native soil N from the soil biomass during the cycles of mineralization-immobilization. If only simple isotopic interchange between fertilizer N and biomass N occurs (i.e., a one-for-one substitution of 15N with 14N atoms), the data for total N would not be affected by the substitution because total N available for plant uptake, leaching, or involvement in other processes would not be changed. However, such simple interchange does not occur. As revealed by the tracer technique, a significant sequestering of fertilizer N into a slowly available form occurs with concomitant decreases in 15N involvement in other processes (e.g., plant uptake). Thus, different information is obtained about the soil N cycle through the use of 15N-Iabeled vs. nonlabe led fertilizer. The above differences in labeled and nonlabeled results demonstrate the need to carefully define the research objectives of a study and determine how the use of N tracers will contribute toward meeting these objectives before beginning expensive field-scale tracer experiments. Factors to consider concerning the use of labeled vs. nonlabeled N include the added costs of tracer studies and the availability of trained personnel and analytical facilities, in addition to the research objectives. In selecting either, it is important to understand that the two approaches are not equivalent. Basic problems encountered with use of labeled N in field studies result from (i) nonuniform distributions of N within the field and (ii) natural and experimentally induced variation in composition of subsamples. The variation from nonuniform distribution of N may be spatial (differences in horizontal and vertical distribution), temporal (differences in the rates of change among different N forms over time), and process level (as affected by qualitative and quantitative differences in N cycle processes within the area under study). Spatial variability arises from nonuniform

HAUCKET AL.

920

application of labeled N and the natural heterogeneity in the occurrence and extent of N-cycle processes. For example, soil denitrification has been shown to be lognormally distributed (chapter 2) by several investigators (Folorunso & Rolston, 1984; Duxbury & McConnaughey, 1986; Parkin et aI., 1987). One detailed study isolated a single microsite that accounted for 85% of the denitrifying activity associated with particulate organic matter which was < 0.1 % of the sample weight (Parkin, 1987). Other investigators have reported water flux and hydraulic conductivity to be lognormally distributed (Warrick et aI., 1977; Jones & Wagenet, 1984) and soil N0 3--N content to be log-normal (Tabor et aI., 1985; White et aI., 1987; Cameron & Wild, 1984). Not surprisingly, therefore, the distribution of labeled N is highly heterogeneous, even for uniformly applied N tracer materials (Broadbent & Carlton, 1980; Selles et aI., 1986). Temporal variability is an inherent result of N transformations occurring with different constituents at different rates, times, and places in soil. This happens, for example, when particulate organic matter becomes a denitrification microsite while a nearby area contains an active root taking up 15N. These lognormally distributed N transformations change over time; the particulate organic matter site may become a zone where N is immobilized and the root uptake site may become a decomposing root channel for macropore transport of 15N out of the root zone. As a result of such events, soil samples collected at the end of a study typically contain quite different 15N contents in plant roots, the soil N0 3-N pool, the soil total N pool, and other N components of the soil system. For example, Saffigna (1988) reported 15N concentrations for 100 d-old wheat roots of 6.01 at. % while the 15N concentrations in the soil N0 3- and total N pools were 1.62 and 0.49 at. %, respectively. This nonuniform distribution of labeled N among different components of the system under study permits one to calculate the net extent to which the labeled N has participated in a particular N transformation or movement process. However, large spatial variations in the Nand N-isotopic composition within a particular component resulting from the heterogeneous dynamic features of soil N processes can result in serious sampling errors and errors of data interpretation that can be minimized only through careful sampling. Unquestionably, the need for collecting representative soil and plant samples is greater when conducting field studies with labeled N than with nonlabeled N. 40-2.2 Managing Field Variability Problems associated with field variability can be reduced by changing type and size of plot and borders, and improving plot sampling and sample preparation techniques. No set of "best-experimental practices" can be recommended for all sites because the best practices may be determined largely by the dominant N cycle processes occurring at the site. For example, if N transport through surface run-off and erosion is a major pathway, then using confined microplots which alter infiltration and surface run-off should be avoided.

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

921

Clear definition of research objectives is imperative before experimental techniques are decided upon. Often, objectives may conflict, requiring that objectives be ranked in order of priority. The impact of compromise should be considered (e.g., cost may determine plot size, number of replications, level of 15N enrichment, labor supply, and duration of study, among other considerations). The better the matching of techniques with objectives, the greater will be the accuracy and completeness of the information that can be obtained through use of N tracer methodology. 40-2.3 Application Techniques Which application technique to use will depend on the objectives of the study, physical form (and for solids, the particle size) of the N source, and availability of a suitable application device (e.g., for applying anhydrous or aqua ammonia or animal manure slurries). Other considerations include manner of placement (e.g., incorporated band vs. surface broadcast), time of application, and presence or absence of vegetation. Obviously, no standard application technique can be recommended but the investigator may be guided by the suggestion and comments that follow. 40-2.3.1 Most Common Technique A goal common to most field studies with labeled N is to apply the tracer material in a manner that best represents production practices with the understanding that it is applied as uniformly as possible to minimize variability among replicate samples. Because it is difficult to scale down farm operations to experimental plot dimensions, investigators have paid less attention to duplicating production application techniques than to striving for uniform application, the latter being especially important in studies of residual N or N balance. Most investigators have found that applying labeled fertilizers in solution is the most satisfactory way because solutions are easy to handle under field conditions and can be dispensed in several ways from surface sprays to banded injections. Solutions also allow use of the same liquid without further processing that was prepared when 15N-enriched material was diluted uniformly to the desired isotopic composition. Critical aspects for solution applicators are: (i) accurate calibration of equipment before use (so that appropriate concentrations of labeled solution can be prepared and the correct amounts applied), (ii) preselection of spray nozzles for uniformity, and (iii) frequent monitoring of application rates during field operation. 40-2.3.2 Comments Water-soluble 15N-Iabeled N sources most often are applied in solution by injection or spraying. Jokela and Randall (1987) used a manual refilling syringe to inject a labeled solution 8-cm deep on a 19-cm grid over a 3.5-m2 microplot, which simulated a broadcast incorporated application, requiring

922

HAUCKET AL.

an application time of about 10 min per plot. A microplot point injector for liquid fertilizer placement in high residue systems was described by Benjamin et al. (1988). Broadbent and Carlton (1980) used a positive displacement pump system, which delivered precise volumes of N solutions to an injection shank that was capable of applying 15N to a field plot with a CV of < 1%. Woodcock et al. (1982) gave a detailed description of a peristaltic pump system mounted on a portable frame for 4-m2 wheat plots. Small plot constant-pressure spray equipment, commonly used for herbicide research, can also be used for broadcast surface applications. Applying labeled fertilizer as a dilute solution may result in anomalous results under conditions where the chemistry and biochemistry occurring within the fertilizer-soil microsite soon after application affect the subsequent reactions and movements of the applied tracer N. For example, in acid soils urea-N nitrifies at a faster rate than an equivalent amount of N from (NH4hS04 when applied as solid particles but not as dilute solution; alkaline-hydrolyzing fertilizer particles solubilize more organic matter at the fertilizer-soil microsite than acid-hydrolyzing ones (Hauck, 1982). As indicated earlier (section 40-1.3.1), the practical significance of such differences in the behavior of solid fertilizers as compared with dilute solutions of these chemicals has not been established, but such differences have consistently been observed in laboratory, greenhouse, and unpublished field studies and merit consideration when planning studies using tracer N. Solid N tracer materials have been successfully broadcast on field plots by hand-spreading in most studies with 15N-depleted (NH4hS04 and NH4N0 3 (Hauck & Kilmer, 1975) and slow-release fertilizers (Brown & Yolk, 1966; Westerman et aI., 1972), and under minimal tillage (Meisinger et al., 1985), among others. For successful application, the fertilizer should be in good physical condition [preferably -6+ 16 mesh (1.23-0.58 mm)] particles, and for large plots, the area should be traversed several times during application. Another technique is to subdivide the plot into small areas and apply separately weighed tracer material to each small area (Olson, 1980). Hand broadcast applications offer the advantages of savings in time and money (if plot numbers are small) and the ability to apply 15N under suboptimal conditions or in remote areas where special equipment or apparatus may not be practical. Disadvantages include the extra cost, time, or effort needed to prepare solid materials of the desired N-isotopic composition and physical form, the extra care needed to apply the solid material uniformly, and, sometimes, a greater labor requirement. When the labeled material is in the form of fine crystals, application on windy days presents problems. Labelled N in gaseous forms is not commonly used in field studies because of inherent handling difficulties. However, Sanchez and Blackmer (1987) described an apparatus for applying 15N-Iabeled anhydrous ammonia (NH3) to small field plots (4.6 m2 ) that uses a flexible stainless steel capillary tube to inject 15NH3 from a cylinder held at constant temperature. This apparatus permits field research with 15N-Iabeled NH3 (the commercial N fertilizer used in largest amount in U.S. agriculture) and also allows

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

923

study of volatile additives such as certain nitrification inhibitors. The apparatus gave CVs of 3% or less and a soil environment that was considered by the authors to be representative of that around conventional ammonia applicators (Sanchez & Blackmer, 1987). Vanden Heuvel and Harrold (1990) described an apparatus for dispensing 15NH3 with an applicator knife pulled by an electric winch along a stationary track. The apparatus permits mechanical injection of 15NH3 to field plots without need for a tractor, marks the injection band location, and dispenses NH3 from the same cylinders used by Vanden Heuvel (1988) for preparing the labeled NH3. A point injector for applying 15N-labeled fluid fertilizers such as urea ammonium nitrate is described by Benjamin et al. (1988). 40-2.4 Plot Type and Size Field research with labeled N has been conducted in plots ranging in size from < 1 to 200 m 2 . The appropriate plot size will depend on the study's goals and objectives, duration of the study, and available funding and labor supply. 40-2.4.1 Confined Microplots Confined microplots are small areas of soil (0.1-1 m2) that are isolated from surrounding soil by metal or plastic barriers. The soil is generally kept undisturbed, although the surface plow layer may be mixed. The barriers are pressed into the soil or placed around an intact block of soil exposed by excavation. They are most frequently used for short-term 15N balance work where the principle objective is to study a specific N-cycle pathway or to construct a 15N budget. Simulating field-scale plant growth is usually a secondary objective. Advantages of confined microplots include: (i) precise definition of system boundaries that allows simplified soil sampling (e.g., complete removal of soil); (ii) restriction of lateral dispersion; (iii) prevention of surface run-off loss (important for 15N crop residue studies); and (iv) lowering cost for 15N materials. Disadvantages include: (i) possible alteration of soil drainage characteristics by confining barriers; (ii) restriction of plant root systems (depends on species); and (iii) possible alteration of surface run-off processes. Confined microplots were used frequently before 1970 (see Legg & Meisinger, 1982) with small stature crops such as small grains and forage grasses. Their use subsequently decreased, especially during the 1980s because of the desire to more closely approximate farm-scale crop growth in 15N studies. Although the comparison of confined vs. unconfined microplots is rarely reported, Saffigna (1988) found similar 15N recoveries with wheat in 0.5-m diam. cylinders vs. 1 m2 unconfined microplots. He gave further details on the use of confined vs. unconfined microplots, including installation techniques and materials. 40-2.4.1.1 Suggestions. We suggest that confined microplots be used when the primary goal is to study N pathways (e.g., residue mineralization and other organic N transformations, N transport within the soil or

924

HAUCKETAL.

ammonia loss) and when simulation of field crop growth is not essential. We suggest using small stature crops (when vegetative cover is needed), large microplots with shallow confinements, and soils selected to minimize possible compaction during installation.

40-2.4.2 Large Field Plots Large plots are usually employed in long-term N tracer studies and when large-scale treatments are being studied such as tillage, irrigation, or crop rotation variables. Problems with lateral movement of 15N can be minimized by using large plots ( > 20 m 2) and conventional two-row borders. Researchers choosing this option usually employ 15N-depleted materials to reduce isotope costs (Hauck & Kilmer, 1975; Bigeriego et aI., 1979; Broadbent & Carlton, 1980; Hills et aI., 1983; Kitur et aI., 1984; Meisinger et aI., 1985). Depleted 15N materials are available in large quantities, require no dilution, and are somewhat cheaper than equivalent 15N-enriched materials. They have a tracer value equivalent to a material containing 0.7 at. % 15N. Depleted N sources can easily be traced into crop plants during the year of application. They have also been used to follow N into the soil N0 3- -N pool (Bigeriego et aI., 1979; Broadbent & Carlton, 1980) and the soil organic N pool on soils with moderate to low organic N contents, i.e., soils with < 1.5 g N/kg (Broadbent & Carlton, 1980; Kitur et aI., 1984). Kilmer et aI. (1974) gave an outline of research topics for use of 15N-depleted vs. 15N-enriched fertilizers, with special reference to studies of N fertilizer use and water quality. Most studies with 15N-depleted materials have been conducted with (NH 4)zS04 as the carrier (available in largest amount) although some studies have used the more expensive 15N_ depleted NH 4N0 3 .

40-2.4.2.1 Suggestions. Large Plot Techniques. Use of large field plots and 15N-depleted materials is suggested for studies on soils with low total N contents « 1 g N/kg) and where typical field crop growth is to be attained, as affected by large-scale variables (e.g., tillage) expressed over several years. Large plots would probably not be suitable on soils of high organic N content ( > 2 g N/kg), for intensive studies of N transformations over short time intervals, or with studies where the 15N concentration in the main N pool of interest probably would be diluted to a level < 0.1 at. % different from that of natural 15N abundance (the background level). 40-2.4.3 Unconfined Microsubplots A third plot type is comprised of unconfined, 15N-treated microplots (1-10 m 2 ) placed within larger plots treated in the same manner but with unlabeled N. Either 15N-enriched or 15N-depleted materials are used on the microsubplots to estimate the percent of plant N derived from the added source. The larger surrounding field plot is used to estimate total N removals and dry matter production. Nitrogen transformation rates and

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

925

pathways also can be obtained on subplots treated with materials enriched with 15N (usually in the concentration range of 2-5 at. %). Unconfined microsubplots offer the advantages of lowering isotope costs while maintaining the ability to detect 15N from enriched sources in different soil N pools after considerable dilution. Use of the microsubplot technique is suggested for studies where single-season plant uptake of applied N is a primary goal, for N source, rate, and placement studies, with soils of high total (organic) N content ( > 2 mglkg), and when soil sampling and 15N balance studies are of secondary interest. Labeled subplots have been used to a limited extent in long-term studies of residual N but, usually, the dispersion of 15N during field preparation and planting will limit their usefulness for long-term studies, especially for studies where plot boundaries are difficult to maintain, (e.g., studies involving tillage, irrigation, or rotation variables). 40-2.4.3.1 Microsubplot Size. A common question with unconfined microplots is: what size microplot will produce valid 15N data while avoiding costly investment in labeled N? Two factors influence the microplot area: (i) lateral plant root distribution and (ii) lateral movement of 15N from the microplot via surface/subsurface flow in soil water or through inadvertent removal of crop residue (e.g., by wind). The optimum microplot size is determined, in part, by crop plant to be used, soil type, weather conditions, and 15N placement. Several investigators have studied microplot size requirements for corn, which typically has a lateral root growth of 50 to 80 cm from the crown (Allmaras & Nelson, 1971; Follett et aI., 1974). Olson (1980), from a study using 15N surface-broadcast and then incorporated into a silt loam soil in Kansas, concluded that accurate data on 15N uptake by corn (Zea mays L.) could be obtained by sampling the center row of a three-row plot (71 cm between rows) and leaving 71 cm of unsampled border at each end of the center row. He also suggested that if residual fertilizer N or N balance work was to be studied, the microplot size should be increased. In Minnesota, Jokela and Randall (1987) in a 2-year study found that reliable plant 15N data could be obtained from the center row of a three-row plot (76 cm between rows) if at least 38 cm of border was left at the ends of the center row. This plot size was adequate for both soil types studied, a well-drained silt loam and a poorly drained, tiled clay loam. In Iowa, Sanchez et ai. (1987) found 2 by 2 m microplots adequate for measuring Ist-yr crop recovery of spring-applied, banded N, but significant lateral movement of N that was surface-applied in the fall occurred; this movement was attributed to lateral flow during the winter when soils were saturated. The lateral movement increased in the 2nd and 3rd yr for all treatments, which was attributed to redistribution of labeled plant residues and to tillage. Lateral movement was greatest with nonincorporated treatments and with moldboard plow tillage. In Nigeria on three sandy loam soils cropped to corn, Stumpe et ai. (1989) found that harvesting the two center rows of a four-row plot (75 cm row spacing)

926

HAUCK ET AL.

would provide valid 15N uptake data if at least 50 cm of row bordered the microplot on each end. Further, soil samples taken from the center area were used to measure total 15N balance (average total recoveries of 99% with a CV of 8% were obtained). In England, Powlson et al. (1986) evaluated 12-row microplots spaced 16.7 cm apart (2 m2 plots) cropped to wheat on a silty clay loam. Finding uptake of nonlabeled N by plants in the two outer rows, they advised sampling the center six rows and leaving a 50-cm border on each end of these rows. Little or no contamination was observed in the third border row, but this row was not harvested as a precaution. 40-2.4.3.2 Suggestions. The above results clearly show that unconfined microplots can yield valid 15N plant uptake data for the year of application, but border areas are essential. The size of the surrounding border will depend on crop and soil type, 15N application method, and the tillage system under study. It is best to determine the optimum size for each individual experimental site. However, in the absence of detailed sitespecific information, we would conservatively suggest that about onefourth of the plot dimension should surround the harvested microplot, (i.e., the harvest area for a 2 by 2 m plot would be the center square meter). 40-2.5 Sample Collection and Preparation Collection and preparation of representative soil, plant, and water samples are important parts of any field study using 15N. As indicated previously (40-1.2) in the discussion relating to Eq. [5], the relative uncertainty of data for total 15N uptake decreases as sample accuracy increases because the errors resulting from field heterogeneity are larger than those involving chemical and isotope-ratio analysis. 40-2.5.1 Plants Field studies with 15N have been conducted with a wide range of plants, including cereal, forage, oilseed, fiber, sugar, vegetable, and tree crops. Each crop will have its own unique plant sampling problems resulting from differences in size, differences in harvested product, and relative importance of quality factors. Discussion of specific sampling techniques for all of these crops would exceed the scope of this chapter. Rather, the reader is referred to the individual crop-specific literature for detailed examples of sampling techniques suitable for a given crop. The general principles of plant sampling, drying, and grinding are discussed by Jones and Steyn (1973) and LeClerg et al. (1962). Plant sampling in 15N studies usually involves sampling the total aboveground portion of the crop that is then divided into its constituent plant parts that are relatively homogeneous with respect to N content or which represent meaningful harvestable products. For example, com is commonly divided

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

927

into grain (about 1.5% N) and stover (about 0.8% N) to obtain more homogeneous analytical samples. This grouping also produces data meaningful for corn grain systems, where the stover is recycled, and for corn silage systems. Tree crops may be divided into many groups to isolate the labeled N into the most actively growing tissue and thus avoid a major dilution problem. Obtaining homogeneous samples is especially important where subsequent analytical methods use small (10-50 mg) samples. If it is not possible to divide the plant into homogeneous tissue, then it is important not to subsample the heterogeneous mixture until all of the sample has been dried and thoroughly ground to a small particle size (e.g., < 100 mesh). This minimizes physical segregation of plant parts and the risk of using a biased subsample. The optimum sample size needed to estimate the dry matter production or N content of a field plot will also vary with crop and soil type and growing conditions. Sometimes only 5 to 10% of a plot may be needed to estimate a certain parameter, but in most cases, 10 to 60% of the plot is sampled. Gomez and Gomez (1984) and LeClerg et al. (1962) emphasized the need for estimating the variances among plots in an experiment, between samples within a field plot, and between analytical samples to design an efficient sampling system. In most cases, even with a small percentage of the plot sampled, the greatest variability will be among plots within an experiment (Gomez & Gomez, 1984). This variability can be managed by increasing the number of replicates. A discussion of the statistical aspects of optimizing sample numbers, replication numbers, site selection, and general field plot technique can be found in LeClerg et al. (1962) and in Gomez and Gomez (1984). 40-2.5.1.1 Recommendation. We recommend that plants be sampled by harvesting the total aboveground biomass, plus any harvested root/ tuber products, then subdivided into plant parts of similar N contents (homogeneous analytical sample) and which represent N flow paths (e.g., harvested products vs. recycled residues). The fresh samples should be immediately oven-dried, then weighed, finely ground, mixed, and subsampled. It is suggested that at an early stage of field research using 15N, the variance among field plots, among samples within plots, and among subsamples to be taken for chemical analysis be estimated to design an efficient sampling strategy that considers site-specific field variability, crop variability, and facilities at the experimental site for drying and otherwise handling plant samples. 40-2.5.1.2 Comments. During multi season studies or studies where plants are sampled more than once, special care should be taken to ensure that the sampling or harvesting operations do not cause cross-contamination from plant material inadvertently carried from one plot to another. Such contamination can be particularly serious when harvesting and transporting across the field large amounts of material (e.g., corn stover) with a relatively high 15N enrichment.

928

HAUCKETAL.

40-2.5.2 Soils Sampling soils for representative 15N content presents a difficult challenge when the tracer, as is usual, is very heterogeneously distributed in soil. Several approaches to soil sampling have been taken by investigators using 15N in field studies, to be discussed below. For general discussions of sampling soil, see Cline (1944) and Petersen and Calvin (1986). 40-2.5.2.1 Complete Excavation. Complete soil removal is undoubtedly the best method to obtain quantitative SOil 15N data and is used when total 15N content in a microplot is to be determined with high accuracy. The early work of Carter et al. (1967), in a study where 15N had been added to soil confined in steel cylinders 68 cm in diameter, clearly showed that complete excavation of soil was superior to taking soil cores. The amount of 15N recovered from sets of seven 1.9 cm diam. cores (composited and thoroughly mixed before chemical analysis) ranged from 87 to 135% ofthe N applied (average, 113%); and from representative subsamples of the entire soil excavated from the cylinders, the recovery values ranged from 98 to 103% (average, 100%). However, the complete removal of soil is only practical with confined microplots < 1 m2 because of the large volume of soil that must be processed (a plot 1 m2 , 30 cm deep will yield a volume of soil weighing about 500 kg). Complete excavation is usually done only to a depth of 15 cm (which usually contains most of the labeled N) or, for more complete recovery, to a 45-cm depth. 40-2.5.2.2 Subplot Excavation. For large unconfined microplots (1 m2 or more) the soil is often sampled by excavating a subplot area. Powlson et al. (1986) took two soil cores (4.75 cm diam. and 23 cm deep) from the edges of each 1 m2 microplot with a power driven auger. Deeper cores (23-40 cm and 60-100 cm) were taken from the spaces between microplots. Moraghan et al. (1984), while constructing a 15N budget for plots treated with band-applied fertilizer, reduced sampling errors by sampling 14 blocks of soil, each 20 by 45 by 30 cm deep from within unconfined microplots. In a study of point placement of 15N-Iabeled urea supergranules to flooded rice soil, Mohanty et al. (1989) removed two blocks of soil (40 by 30 cm by 15 cm deep) containing the supergranules. If leaching of labeled N is of interest, intense sampling of lower soil depths is necessary (e.g., Kissel & Smith, 1978; Bigeriego et aI., 1979; Khanif et aI., 1984) because macropore transport may considerably increase the heterogeneous distribution of nitrate moving downward through soil. Priebe and Blackmer (1989) cite evidence for preferential horizontal and vertical movement of solution through macropores of an Iowa soil (mesic Aquic Hapludolls) under a reduced tillage, continuous corn management system. Undisturbed soil cores (10 cm diam., 50 cm long) were taken from the field (a novel method is described). The movement of water and solute through the cores was followed over a 24-h period with

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

929

I80-labeled water and I5N-Iabeled urea. Determinations of labeled water and N compounds (urea, N0 3-, and NH/) in both effluent and soil (5-cm increments) lead Priebe and Blackmer to suggest that preferential flow of water through soil macropores may be an important factor affecting N movement. 40-2.5.2.3 Small-core Sampling. Sampling large plots for I5N is usually done with conventional small diameter soil cores ( < 5 em diam.). Investigators using large plots to which I5N is added usually are interested in tracing the applied N into both the nitrate and total N pools. A problem with taking small cores is that the core samples may not be intercepting in a representative manner labeled N that is heterogeneously distributed in soil. The greater the spatial variability and the fewer the number of cores taken, the greater the risk of error from nonuniform sampling. Following review of several field studies, Meisinger (1984) characterized the spatial variability of soil N0 3- as (i) being a large, small-scale component, that is, 50 to 75% of the total variability is already present within a few square meters (Beckett & Webster, 1971; Cameron et aI., 1971), and (ii) as having a large CV that usually ranges between 30 and 60% with 45% being a typical value. The spatial variability of total N is less than that of N0 3- -N, having CVs ranging between 10 and 25%, typically 15%. By estimating the approximate CV and by assuming that the final bulking of soil cores will produce a sample with a mean that is approximately normally distributed (central limit theorem holds), one can calculate the number of cores needed to estimate the plot mean with a given degree of precision. Gomez and Gomez (1984) give the appropriate formula for this calculation. n

=

(Z)2(CV)2/(d)2

[9]

where n Z

= the number of cores required

standard normal deviate for the alpha level (Z = 1.96 for 0.05 and 1.65 for 0.10) CV = coefficient of variation (as a decimal) d = margin of error in the plot mean (as a decimal) =

Using a CV of 45% for N0 3- -N and a desired precision of ± 10%

(d

= 0.10) in 90% of the plots (alpha = 0.10), one can calculate that 55

cores would be required. Obviously, core sampling for N0 3- will involve substantial labor. Alternatively, the researcher can adjust expectations and calculate, for example that 14 cores per plot would be sufficient to estimate the N0 3- -N content with an accuracy of about 20% on 90% of the plots. For total N (CV = 15%), the same 14 cores per plot should estimate the plot mean to within 8% on 95% of the plots. It is apparent that many core samples must be taken to permit accurate determination of soil I5N content.

930

HAUCK ET AL.

40-2.5.2.4 Soil Sample Preparation. The method of processing soil after sampling should not alter the N pools under study and should ensure that representative subsamples can be taken. Large soil masses resulting from complete plot excavation can be mechanically mixed with a concrete mixer, wet sieved, then subsampled and dried (e.g., Powlson et aI., 1986; White et aI., 1986). Some investigators have added water to the bulk soil to produce a slurry that can be more easily mixed and subsampled (Kissel & Smith, 1978). Using this technique, subsamples should be chemically analyzed immediately to prevent change through microbial action. Whatever the procedure, care must be taken to avoid loss of N or change of N form during sample preparation. Alternatively, the soil can be dried, pulverized, sieved, and mixed for subsampling. The best order of these operations will depend on local facilities, but in all cases thoroughly mixing the sample before subsampling is imperative. Saffigna (1988) reported that the CV for determining 15N-labeled N0 3- -N and total N of greenhouse soils decreased after thorough mixing from 42% to 3% and from 10% to 2 %, respectively. Thorough mixing and fine pulverizing of soil samples is especially important when using analytical instruments with small (e.g., 50 mg) sample requirements. When one considers that a 50-mg sample represents 450 x 106 mg of heterogeneous field soil from a microplot 1 m 2 by 30 cm deep, it is apparent that soil sample collection, mixing, and subs ampling should be given careful attention. 40-2.5.2.5 Soil Sampling Suggestions. The appropriate soil sampling technique will depend on the goals of the study, the 15N application technique, and the size of the plot. For studies that require the most accurate estimates of soil 15N content (e.g., 15N budgets, residual soil 15N) or studies with localized 15N placements (banded or point placements), we suggest either (i) the complete excavation method for plots < 1 m 2 in area, or (ii) the excavation of smaller subplots within larger microplots (1-10 m 2 ), with the excavated area corresponding in size to the distance between plant rows or between localized zones of application (e.g., between adjacent fertilizer bands). For studies that require less accurate soil 15N estimates, have received uniform 15N applications over several seasons, or were conducted on large field plots ( > 20 m 2), we suggest high intensity (10-40 cores/plot) small-core sampling with more intense sampling when 15N03- -N is to be determined and fewer cores taken for determining total 15N. 40-2.5.3 Soil Water No less formidable than collecting representative soil samples is the collection of representative samples of soil water. The leaching of water and solute through soil is a complex, dynamic process that no longer can be described as a simple piston flow process. Water percolation often involves a small but rapidly moving portion of water that penetrates deeply into the soil through large pores and by-passes much of the soil solution (e.g., see Priebe & Blackmer, 1989). Interconnected with this large-pore water is the small-pore water, which is also displaced downward during percolation,

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

931

but at a slower rate. As water and N0 3- percolate through soil, a portion of the solution will move to relatively deep horizons but most of the N0 3usually will remain at shallower depths (Boswell & Anderson, 1964; Thomas & Phillips, 1979). Sampling multipore systems is difficult because the chemical composition of the large-pore water is different from that of the small-pore water. Interpreting soil water data is also difficult because different field samplers collect different portions of the large-pore vs. smallpore water. Soil water samples generally are obtained using a suction device, a free-drainage device, or some type of lysimeter. As with most field methods, there is no ideal method to sample soil water. Only a general overview of methods will be given here; the reader is directed to several recent articles for more detailed discussion (e.g., Rhoades & Oster, 1986; Soileau & Hauck, 1987; Litaor, 1988; Starr et aI., 1991).

40-2.5.3.1 Porous Ceramic Cups. Though frequently used to collect water samples, porous ceramic cups preferentially sample the large-pore sequences immediately surrounding the cup, resulting in sampling error. If the objective is to sample all the water passing a given depth, then several samples must be collected over time; otherwise, sample bias is highly probable (Hansen & Harris, 1975; Alberts et aI., 1977). Porous cups are relatively easy to install but their usefulness is limited by the large errors inherent in spatial variability which use of this sampling approach cannot overcome (Biggar & Nielsen, 1976). Broadbent and Carlton (1980) used ceramic cups in field 15N studies to estimate leaching in conjunction with water flux measurements. Nitrate leaching data derived from porous cups are generally regarded as "point samples" that provide relative comparisons among treatments but quantitative estimates of N0 3- leaching require a simultaneous detailed study of water flux at the site (Biggar & Nielsen, 1976; Rhoades & Oster, 1986). Linden (1977) described the installation and use of porous cup samplers. Rhoades and Oster (1986) recommended reducing some of the sampler variability by preselection for uniform perme abilities and size and by using uniform sampling intervals and suction. 40-2.5.3.2 Trough Extractors. Large ceramic extractors (3 m by 15 cm) have also been placed at the bottom of 20 to 45 cm long troughs that are then placed in soil about 1.5-m deep to collect leachate (Duke & Haise, 1973; Hergert, 1986; Montgomery et aI., 1987). These devices have been placed under disturbed soil with standard trenching techniques and under undisturbed soil blocks with the aid of horizontal tunnels and inflatable bladders that force them up into contact with the soil. These extractors attempt to intercept macropore water with the trough and to sample smallpore water with the evacuated ceramic tube. Montgomery et al. (1987) compared large extractors in filled-in lysimeters with the lysimeter percolate and concluded that the extractors provided a good estimate of deep percolation and NOi flux on a loamy fine sand in North Dakota. However, they also reported that similar extractors covered with disturbed soil were suitable only for estimating N0 3- concentrations (rather than flux), the

932

HAUCK ET AL.

concentration values obtained comparing well with those estimated using ceramic cups. Hergert (1986) reported that the extractors adequately estimated drainage under low leaching conditions (52 mmlyr), but adjustment of the vacuum level was needed under high leaching conditions (165 mmlyr). The vacuum inside the trough should be maintained equal to that in the surrounding soil to avoid too much flow into the trough (excess vacuum) or too little flow (suction too low). Large extractors are difficult to install and require regular care and attention, but they can overcome some of the problems associated with small cup extractors by sampling a large area, collecting macropore events, and sampling small-pore sequences. To our knowledge no 15N studies have been reported with these large extractors. 40-2.5.3.3 Tile Discharge. Sampling tile line discharge is another method of collecting percolate samples from soils with high water tables or restricted drainage. This procedure involves conventional tile line installation, sometimes with provision for isolating the plots with surface/subsurface flow barriers, and installation of automatic collection devices (Zwerman et at., 1972; Gast et at., 1978; Bergstrom, 1987). Hallberg et at. (1986) reviewed the literature on tile drain monitoring, and several research studies have used tile drains to assess the impact of agricultural practices on water quality (Gast et at., 1978; Baker & Johnson, 1981). Tile drain data have been shown to include macropore flow events (Richard & Steenhuis, 1988), but they more typically represent flow with long lag times (months or years depending on the hydrologic cycle) between imposition of surface treatments and tile discharge. The lag times are caused by macropore processes causing slow solute equilibrium within the unsaturated zone and by long travel times for drainage to reach the tile line within the saturated zone. Tile discharge data are best used to measure N0 3- concentrations over a long-term study for large-plot treatments. Over the period 1973 to 1979, Nelson and Randall (1983) and Buzicky et at. (1983) studied movement of 15N-depleted (NH4hS04 in 36 individually drained field plots (6.1 x 9.2 m), each isolated to a depth of 1 m with plastic sheeting. Tile drainage waters flowed to underground collection chambers where they were sampled. Estimating the mass of N0 3- leached is difficult with tile drain data because of uncertainties in delineating exact drainage areas and uncertainties in the quantity of percolation that escapes between tile lines (Hallberg et at., 1986; Bergstrom, 1987). 40-2.5.3.4 Lysimeters. Three general types of lysimeters have been used to collect soil water: disturbed (or filled-in), undisturbed (or monolith), and pan (or ebermayer). The literature on lysimetry is extensive and the reader is referred to classic reviews, such as Khonke et at. (1940) and Allison (1955), and a recent review by Soileau and Hauck (1987) for detailed discussions on lysimeters. Agricultural scientists have used filled-in lysimeters with caution because of the radical disturbance of percolation paths during construction. They are best suited for coarse-textured soils with little or no structural

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

933

development and may be equipped with vacuum extractors or be freedraining. Montgomery et aI. (1987) produced bulk densities similar to those under field conditions by saturating the soil intermittently with water entering from the bottom of the lysimeter. Filled-in lysimeters have been used with 15N sources to study leaching as affected by time of fertilizer application and forage crop species (Jones et aI., 1977) and to evaluate nitrification inhibitors and fertilizer placement (Walters & Malzer, 1990). Of particular interest in the work of Walters and Malzer (1990) is the fact that the values for amount of N leached were threefold greater when calculated from nontracer data obtained on the leachate containing 15N than when isotope dilution calculations were used. The investigators attributed this discrepancy to 15N equilibration in the soil biomass, thereby reducing the amount of labeled N available for leaching. Most investigators have noted increased N0 3- -N losses and higher percolation the first few years after lysimeter filling. Filled lysimeters are best suited to soils where macro-pore flow is thought to be insignificant and to long-term studies that allow ample time for settling and structural development. Undisturbed monolith lysimeters are being used more frequently today to preserve macro-pore processes and more closely approach field conditions. Constructing monolith lysimeters is described in several reports, including the excavation approach (Khonke et aI., 1940; Brown et aI., 1985), pressing cylinders into moist soil with static weight (Tackett et aI., 1965) or hydraulic pressure (Cassel et aI., 1974), and a drilling technique to isolate a core and simultaneously encase it in plastic (Bergstrom, 1987). Undisturbed lysimeters include both free-draining types (Chichester & Smith, 1978; Bergstrom, 1987) and vacuum-assisted types (Cassel et aI., 1974; Brown et aI., 1985). Undisturbed lysimeters offer the advantages of precise definition of system boundaries, inclusion of macropore processes, and the ability to monitor both nitrate concentration and the mass of N leached. They suffer the disadvantages of being expensive, and labor intensive, and are quite variable even for monoliths collected near each other. Nitrogen tracers have been used in monolith lysimeters to study nitrate leaching as affected by soil type and precipitation (Owens, 1960) and no-tillage vs. conventional tillage culture (Chichester & Smith, 1978). We suggest using undisturbed monolith lysimeters when determining the mass of N0 3- -N leached is the primary research objective, macropore transport processes are important, and sufficient labor and time are available. Soil solution samples are also frequently collected from beneath undisturbed blocks of soil by trenching, followed by inserting a horizontal collection device (ebermayer lysimeter). The collector usually is a freedraining glass or metal pan (e.g., Tyler & Thomas, 1977; Shaffer et aI., 1979) that samples gravitational water flowing under saturated conditions. Jordan (1968) modified the free-draining pan by placing glass wool, a fiberglass screen, and moist soil in the pan to collect water under "zero-tension" conditions. Haines et aI. (1982) found that at the 30-cm depth, a low tension ceramic plate collected twice as much percolate as a

934

HAUCK ET AL.

Jordan-type, zero-tension lysimeter, but the N0 3--N concentration from the ceramic plate was only about one-third of that from the Jordan collector. Barbee and Brown (1986) and Shaffer et al. (1979) also compared free-draining vs. suction cup samplers and concluded that porous cups were ineffective in well-structured soil where much of the percolation was associated with rapid leaching through large pores. The ebermayer-type lysimeters offer the advantages of better focus on rapid transport processes, easier installation, and lower labor and maintenance expenses than conventional lysimeters. Disadvantages include a high degree of variability and a lack of system boundaries that make estimation of the mass of N leached nearly impossible. The ebermayer-type lysimeter is useful when macro-pore flow and N0 3- -N concentrations are of primary interest, the soil is well structured, leaching events occur primarily under ponded conditions, and labor and time are limited. 40-2.5.3.5 Suggestions. No single method of sampling soil water can be recommended for all research applications. Several of the techniques outlined above differ in that they sample different proportions of the largepore vs. small-pore soil water and, therefore, give different estimates of the quantity and composition of the soil water. Porous ceramic cups are best suited for qualitative comparisons of N0 3- -N concentrations among treatments in soils where macro-pore transport is minimal. Tile drain samples are suited to long-term studies in soils with high water tables where comparison of N0 3- -N concentrations is of main interest. If sufficient money and labor are available, some type of undisturbed lysimeter is best suited for estimating the mass of N leached and the N0 3--N concentration. Filled lysimeters are less variable and can give useful relative comparisons among treatments for N0 3- -N leaching in nonstructured soils. Collection pans installed beneath undisturbed soil are useful for studying macro-pore processes and N0 3- -N concentrations but give variable results and their use is questionable for accurately estimating the mass of N leached. Soil physicists are still learning about macro-pore flow in soils and its impact on N0 3transport. This knowledge should aid in designing improved field methods for studying N0 3- transport in soil water. 40-2.5.4 Soil Gases

Accurately collecting and determining gaseous forms of N and their exchange between soil, air, and the atmosphere above continues to be an unresolved problem, especially in field studies. The two N cycle processes of greatest interest contributing to gaseous exchange are biological dinitrogen fixation (chapter 43) and denitrification (chapter 44). Many methodologies and techniques have been developed and used with moderate success, depending on experimental objectives. The use of 15N is indispensable for many of these methods. Rather than make a cursory review of this subject, we refer the reader to the review by Mosier and Heinemeyer

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

935

(1985) and the publication, Field Measurement of Dinitrogen Fixation and Denitrification (Hauck & Weaver, 1986), which in turn refer to other articles in this area. 40-3 PREPARING FOR AND MEASURING NITROGEN-ISOTOPE RATIO 40-3.1 Principles

Of the several methods available for determining N-isotopic composition, use of the mass spectrometer continues to be the method of choice because of its accuracy, precision, convenience, and general applicability in studies of N-cycle processes. The only practical alternative is emission spectroscopy, which has the advantage of being technically simpler (e.g., high vacuum is not required) and less costly (emission spectrometers suitable for use in N tracer research are considerably cheaper than mass spectrometers). An attractive feature offered by emission spectroscopy is that this method permits N-isotope ratio measurements to be made on microgram amounts of sample N. However, systems for automatic sample gas preparation and analysis by mass spectrometry have now been developed that permit accurate and acceptable determination of N-isotope ratio in < 10 f,tg of N. Other methods of N-isotope analysis, such as those using infrared spectroscopy, nuclear magnetic resonance, electronparamagnetic resonance, or microwave spectroscopy, have importance in N research but are not used in routine determination of N-isotope ratio. Before the N-isotopic composition of a sample can be determined by mass spectrometry or emission spectroscopy, all N forms in the sample must be quantitatively converted to a gas that is simple in molecular structure and isotopic composition, of low molecular weight, readily prepared from inorganic and organic compounds, preferably unreactive with components of the isotope-analyzing system, and easily pumped from the instrument. Dinitrogen (N 2) best meets these requirements and is used for the isotope-ratio analysis of N in most solids and liquids. The traditional and probably most commonly used procedure for converting combined N to N2 for isotope-ratio analysis involves a three-step procedure: (i) conversion of sample N to NH/-N by means of acid digestion (e.g., Kjeldahl procedure) or by other processes (e.g., hydrolysis); (ii) oxidation of NH/ -N to N2; and (iii) determination of the isotopic composition of the N2. This procedure originally was developed by Rittenberg and colleagues (Rittenberg et aI., 1939; Rittenberg, 1948; Sprinson & Rittenberg, 1948, 1949) for use in biochemical research but has been applied extensively to the agricultural and biological sciences. Following the development of improved instruments and their coupling with emission spectrometers and some automatic mass spectrometer systems, dry combustion techniques for direct conversion of sample N to N2 are gaining favor. Reference to other

HAUCKETAL.

936

techniques for generating N2 for special application can be found in Rittenberg (1948), Bremner (1965a), Kennedy (1965), and in the methodology section of the bibliography by Hauck and Bystrom (1970). Many modifications in analytical methodology have been developed to improve the speed, convenience, and accuracy of N-isotope-ratio analysis. The three-step procedure, especially, remains a complicated and timeconsuming process. Possible errors encountered in both the three-step and direct combustion procedures are discussed by Hauck (1982) and Mulvaney (1993), among others. The intent here is not to give complete procedural details of the many steps leading from sample preparation to determination of N content and isotopic composition. Special procedures, such as for small sample handling and diffusion techniques, are not discussed. For in-depth discussions of preparing for and measuring N-isotope ratio, the reader is referred to reviews by Bremner (1965a), Hauck (1982), and Mulvaney (1993), and to the many references cited in these reviews. 40-3.2 Conversion of Labeled Nitrogen to Ammonium 40-3.2.1 Principles

The method to be chosen for converting labeled N to NH/ depends on the forms of N in the sample under study and whether there is a need to distinguish between them. For total N, a Kjeldahl method usually is used. Steam distillation methods that result in the collection of NH/ -N are used for the determination of NH/-N, N0 2--N, and N03--N. Because only inorganic and total N are determined on most of the samples obtained from agronomic and environmental studies using 15N, procedures will be outlined for only these N forms. 40-3.2.2 Total Nitrogen

Numerous modifications of the Kjeldahl method have been described (see Bremner & Mulvaney, 1982). The basic method is a two-step process: (i) digestion with concentrated H2S04 to convert organic N to NH/-N, and (ii) determination of the amount of NH/ -N in the digest. The speed and completeness of digestion are increased by adding a salt (commonly K2S04 ) to increase the temperature of the digesting solution and a catalyst (Cu, Hg, or Se, alone or in combination) to accelerate oxidation of organic matter. Upon addition of alkali to the digest, followed by steam distillation, NH3 is liberated, trapped, and the NH/ -N content determined by titration with standard acid. Although both macro- and semimicro-methods have been used for converting 15N-Iabeled organic N to NH/-N, semimicro-methods usually are preferable because they provide an appropriate amount of N (about 1 mg) for N-isotope-ratio analysis and are less subject to error resulting from contamination by residues from preceding samples (cross-contamination). The permanganate-reduced Fe modification of the Kjeldahl digestion is recommended here because it can be used

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

937

for determining total N, including N0 2- -N and N0 3- -N, in either plant or soil samples. 40-3.2.2.1 Special Apparatus. Micro (30 or 50 mL) or semimicro (100 mL) digestion flasks are used with an appropriate steam distillation apparatus (for detailed descriptions and discussions, see Bremner & Mulvaney, 1982; Hauck, 1982; Keeney & Nelson, 1982). 40-3.2.2.2 Reagents. Prepare as follows: 1. Salt-catalyst mixture. Intimately mix and grind to a powder 100 g of K2S04, 10 g of CuS04·5H20, and 1 g of Se, using a laboratory jar mill or comparable device. 2. Concentrated sulfuric acid (H2S0 4), 3. Dilute sulfuric acid. To 500 mL of water in a 2-L Pyrex flask, slowly add with continuous stirring 500 mL of cone. H 2S0 4, 4. Permanganate solution. Dissolve 25 g of KMn0 4 in 500 mL of water and store in a dark container. 5. Reduced Fe. Finely divided, to pass a 100-mesh screen. 6. Sodium hydroxide solution (about 10 N). Place 3.2 kg of NaOH into a heavy walled 10-L Pyrex bottle marked to indicate a volume of 8 L, add 4 L of CO2-free water, and swirl contents until the NaOH is dissolved. Stopper the bottle until the solution is cool, then fill to the 8-L mark with CO2-free water. Connect the bottle with a suitable dispensing device that prevents CO2 absorption by the alkali solution. 7. Boric acid-indicator solution. Dissolve 80 g of H 3B0 3 in about 3000 mL of water contained in a 5-L flask marked to indicate a volume of 4 L. Heating and vigorous stirring will speed dissolution. Cool and add 80 mL of mixed indicator solution (99 mg of bromcresol green plus 66 mg methyl red in 100 mL ethyl alcohol). Add 0.1 N NaOH carefully until the solution turns reddish purple (pH about 5.0), make to the 4-L mark with water, and mix. 8. Standard acid. 0.01 N (e.g., 0.00714 N H 2S0 4), 40-3.2.2.3 Procedure. Place soil or plant sample containing about 1 mg of N in a micro- or semimicro-Kjeldahl digestion flask, add 1 mL of KMn04 solution and swirl for about 30 s. Holding the flask at an angle, very slowly pipette 2 mL of dilute H 2S0 4 down the side of the flask, swirling continuously. Cool for about 5 min. Through a dry funnel with stem reaching into the bulb of the digestion flask, add 0.50 ± 0.01 g of reduced Fe, swirl, and let the mixture stand about 15 min until the effervescence ceases. Insert a small Erlenmeyer flask or other loosely fitting glass obstruction in the flask opening and simmer (reflux) gently for 45 min, avoiding significant water loss. Cool, remove glass insert, add two glass beads, 1.1 g of salt-catalyst mixture, and 3 mL of cone. H 2S0 4, Heat gently, then vigorously, turning the flask occasionally. After about 30 min the digest forms a clear greenish yellow solution, the clearing time depending on type of sample. Continue boiling until the entire digestion time is at

938

HAUCK ET AL.

least 1.5 times the clearing time (some soils with refractory N forms may require digestion for as long as 5 h, although < 1 h is usual for plant tissues and 1-2 h for soils). Cool slightly. To the still warm digest, add water drop by drop down the side of the flask until violent reaction subsides, then add an additional 15 to 25 mL of water. If the digest has solidified before any water addition, heat gently until the salt cake disintegrates and redissolves, after which the digest can be diluted. Transfer the diluted digest to an appropriate steam distillation apparatus, add 20 mL of 10 N NaOH, and distill into a 50-mL Erlenmeyer flask containing 5 mL of H 3B0 3-indicator solution. Collect about 35 mL of distillate. Determine the NH/ -N concentration in the distillate by titration with standard acid (the color change at the end point is from green to pink). 40-3.2.2.4 Comments. Significant error in the determination of N-isotope-ratio can result if all forms of N in the sample are not quantitatively converted to NH/ even though failure to include all of the N is within the experimental error of the Kjeldahl procedure (commonly ranging between 1.5 and 4%). For example, the N0 3- -N content of a sample may represent a small fraction ( < 1%) of a sample's total N content, but if its N-isotopic composition is different from that of other total N components, incomplete recovery of N0 3- -N will markedly affect the accuracy by which the isotopic composition of the total N is measured. Modifications of the basic Kjeldahl method are used to obviate such difficulty, for example, use of reduced Fe-permanganate or salicylic acid pretreatment for complete recovery of N02- -N and N0 3- -N or longer digestion times (up to 18 h) for conversion of N in refractory compounds to NH/ -N. Further details and examples of the magnitude of errors associated with incomplete digestion of sample are given by Hauck (1982). To achieve the highest level of precision and accuracy, samples must be ground to pass a sieve of 100 mesh or smaller. This ensures that tissue components of different physical and chemical characteristics are intimately mixed so that aliquot samples are virtually identical. Use of finely ground samples becomes increasingly more important as sample size decreases. Special grinding techniques are required for the preparation of samples to be analyzed for 15N content by some of the automated instrument systems now available. Grinding can be accomplished conveniently by rotating glass bottles containing stainless steel rods of different diameters along with the sample to be ground (Hauck, 1982; Kelley, 1994). Soil, plant tissue, and many seeds can be ground to face-powder consistency with little effort. Cross-contamination from residues in a grinding mill is eliminated because each bottle is a separate grinding chamber. 40-3.2.3 Specific Nitrogen Forms Simple, rapid, and reproducible methods have been developed for determining the amount and isotope ratio of different N forms in soil extracts and hydrolyzates, plant extracts, and waters including exchangeable and nonexchangeable NH/ , N0 2- , N0 3-, a-amino acids, amino sug-

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

939

ars, hydroxyamino acids, and urea. An outline of these procedures is given in Table 40-1, with references to articles that provide more detail. See also Bremner (1965b) for detailed discussion of determining inorganic forms of N. All of the methods described in Table 40-1 involve steam distillation to liberate NH3 and are readily applicable to many kinds of agricultural and environmental studies. Problems occur when the sample concentration of N is low requiring the use of large aliquots or multiple distillations to secure sufficient N for accurate N-isotope-ratio analysis (Hauck, 1982). These problems are less severe today because improved instrumentation permits accurate determination of isotope ratio on smaller amounts ( < 50 flg) of N. 40-3.2.3.1 Ammonium, Nitrite, and Nitrate N. The recommended procedures for chemical analysis of these N forms in preparation for N-isotope-ratio analysis are those described by Keeney and Nelson (1982). They are based on the finding that NH/ -N can be quantitatively liberated from solution containing a small amount of MgO by steam distillation in the presence of alkali-labile organic N compounds and that NH/ -N + N0 2-N + N0 3- -N can be determined accurately by the same method if finely divided Devarda's alloy is added to the solution containing these N forms immediately before MgO addition and steam distillation. Correct use of these procedures in 15N research requires that each N form be completely liberated from solution and recovered in the distillate, that steam distillation rates and times be carefully controlled, and that cross-contamination is negligible. The importance of understanding the several sources of error inherent in the steam distillation procedures cannot be overstated and the reader is urged to review the procedural details and commentaries given in the articles by Hauck (1982), Keeney and Nelson (1982), and Mulvaney (1993), among others. 40-3.3 Direct Conversion of Labeled Nitrogen to Dinitrogen The dry combustion or Dumas technique typically involved heating the sample (550-650 0c) with CuO to convert inorganic and organic N to N2 and N oxides (mainly N20), which are subsequently reduced to N2 with hot Cu. Many modifications have been developed to overcome some of the limitations of this method (e.g., incomplete conversion of N03- to N2) for use in N-isotope-ratio analysis (e.g., see Fiedler and Proksch, 1975). For example, heating to 800 to 900°C in quartz or vycor sealed tubes greatly improves quantitative conversion of all N forms to N2. Limitations of the dry combustion method also have been successfully resolved with the development of the automatic C/N analyzer (ANCA), with which complete conversion to N2 is achieved through flash combustion (= 1700 0c) in the presence of a catalyst (Cr203). The combustion products are swept over Cu at 600°C and then purified chromatographically. Commercial systems are available for direct coupling to a mass spectrometer, permitting automatic determination of both N content and N-isotope ratio. The combustion method merits consideration because it

940

HAUCK ET AL.

Table 40-1. Method for conversion of different forms of soil N to NHi and separation of the NHi for N isotope-ratio analysis. Form ofN In soils containing no NO) or NOi In soils containing NO) In soils containing NO) or NOi

Exchangeable NH4+ NO)

Exchangeable NH4+ and NO) NO) or NOi Exchangeable NH4+ and NO) orNOi Nonexchangeable NH4+

Total NH+ 4

NHi and hexosamine Serine and threonine

Amino acid

Method Total N Kjeldahl digestion of soil and distillation of digest with alkali Kjeldahl digestion and distillation after pretreatment of soil with salicylic acid and thiosulfate to convert NO)-N to amino-N Kjeldahl digestion and distillation after pretreatment of soil with acidified permanganate to oxidize NOi or NO) and with reduced Fe and H 2S04, to reduce NO) to NHi. Inorganic N Steam distillation of soil extract with MgO Steam distillation of soil extract with MgO and Devarda alloy after destruction of NOi with sulfamic acid and removal of NH4+ by steam distillation with MgOt Steam distillation of soil extract with MgO and Devarda alloy after destruction of NOi with sulfamic acidt Steam distillation of soil extract with MgO and Devarda alloy after removal of NH4+ by steam distillation with MgO Steam distillation of soil extract with MgO and Devarda alloy Steam distillation of soil with KOH after removal of exchangeable NHi and labile organic N compounds by KOBr and treatment of residue with HF to decompose minerals Hydrolyzable N Kjeldahl digestion of soil hydrolyzate and steam distillation of digest with alkali Steam distillation of soil hydrolyzate with MgO Steam distillation of soil hydrolyzate with pH 11.2 buffer Steam distillation of soil hydrolyzate with pH 11.2 buffer after steam distillation with same buffer to remove (NHi - hexosamine)-N and treatments with periodate to convert (serine + threonine)-N to NH4+-N and with arsenite to reduce excess periodate Steam distillation of soil hydrolyzate with pH 11.2 buffer after treatments with NaOH to decompose hexosamines and remove NH4+ and with ninhydrin to convert a-amino-N to NH4+-N

t Includes specific chapter and page numbers in monograph. t If NOi is absent, the treatment with sulfamic acid is omitted.

Referencet Bremner & Mulvaney, 1982 Bremner & Mulvaney, 1982 Bremner & Mulvaney, 1982

Keeney & Nelson, 1982 Keeney & Nelson, 1982 Keeney & Nelson, 1982 Keeney & Nelson, 1982 Keeney & Nelson, 1982 Keeney & Nelson, 1982

Stevenson, 1982 Stevenson, 1982 Stevenson, 1982 Stevenson, 1982

Stevenson, 1982

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

941

involves few steps and may be more accurate and precise than the KjeldahlNaOBr method for determining total N and producing N2 for isotope-ratio analysis. For a comparison of Kjeldahl and Dumas-type procedures for use in determining total Nand 15N contents of several organic N sources, see Minigawa et al. (1984). A recent example of the many modifications of the Dumas method is the use of CaO to simplify removal of CO 2 and water during the conversion of combined N to N2 (Kendall & Grim, 1990).

40-3.4 Measuring Nitrogen-Isotope Ratio 40-3.4.1 Mass Spectrometry A mass spectrometer separates molecular ions into a spectrum according to their masses or, more accurately, according to their mass to charge ratio (m/e). The separation for N molecules is achieved by electron bombardment of N2 to form molecular ions (mainly positively charged) of m/e 28,29, and 30, corresponding to 14N2, 14N15N, and 15N20 respectively. The positive ions are drawn from the ionizing chamber into the magnetic sector by a small positive potential on a repeller and a larger negative potential across accelerating electrodes. The ions are accelerated through a magnetic (or magnetic/electrostatic) field where the combination of accelerating voltage and field magnetic strength determine the curvature of each ion path. By adjusting the acceleration voltage, ions of a given m/e can be focused on a target. The intensities of the ion beams generated corresponding to m/e 28,29, and 30 are directly related to the isotopic composition of the N2 being analyzed. Sample N2 is obtained either by direct combustion or the oxidation of NH/-N with alkaline NaOBr or LiOBr. Appropriate measures are taken to ensure that the N2 is free of impurities that could give rise to interfering molecular ions (e.g., CO or NO with m/e of 28 and 30, respectively). Procedural details and discussion of the hypobromite reaction in relation to N-isotope-ratio analysis are given by Hauck (1982) and Mulvaney (1993). Two automated systems of N-isotope-ratio analysis by mass spectroscopy have been developed. The automated Rittenberg analysis system (ARA-MS) involves the same three-step procedure used in conventional N-isotope analysis, the main difference being that the hypobromite oxidation of NH/ -N and the determination of N2 isotopic composition are done automatically. The ARA-MS system is described in detail by Mulvaney (1993). Mass spectrometers also have been coupled to automatic Nand C analyzers (ANCA) that directly combust sample N to N2. Plant, soil, or other solid samples (usually containing 20 to 150 [tg of N) are encased in miniature Sn capsules for loading into an ANCA autosampler, from which individual samples are dropped into a combustion chamber. After combustion, the products (C0 2, N20 NO x , and H 20) are swept into a tube containing hot Cu wire to convert NO x to N2, then through traps to remove CO 2 and H 20. The N2 is further purified by gas chromatography before an aliquot of the effluent is admitted into the mass spectrometer. One of the

HAUCK ET AL.

942

advantages of ANCA-MS is that both N amount and isotopic composition can be obtained during the same sample run. 40-3.4.2 Emission Spectrometry In an emission spectrometer, Nz molecules emit characteristic light in the UV spectrum when excited at low pressure in an electrodeless discharge tube. The light emitted by the excited molecules is resolved by a monochromator and the characteristic light signals are detected by a photomultiplier. The amplified signal peaks are recorded and related directly to the abundance of the three molecular species of N, permitting calculation of N-isotope ratio. Emission spectroscopy is used for N-isotope-ratio analysis mainly in eastern Europe, Japan, and South Africa. As indicated earlier, one advantage of its use is that only a few micrograms of N are needed for 15N analysis but with the advent of ANCA-MS systems that can determine the isotopic composition of microgram quantities of N with a high level of precision, this advantage of emission spectroscopy is less attractive. However, the sample requirement for some nonautomated emission spectrometers can be reduced with special techniques to as low as 0.2 IJ.g which is an important advantage when 15N content is to be determined on small amounts of nitrogenous material separated by biochemical techniques such as thin layer chromatography. For selected references to an abundant literature on this subject, see Hauck and Bremner (1976). A fully automated emission spectrometer system capable of determining N content and isotopic composition of 10 to 20 samples per hour (2-10 IJ.g N per sample) is described by Therion et al. (1986).

40-4 SOURCES OF NITROGEN-1S SUPPLY AND ANALYTICAL SERVICE At least 13 commercial sources of 15N-enriched compounds were available two decades ago (Hauck & Bremner, 1976). Most of these suppliers now are no longer in business or are of questionable reliability as suppliers. Currently, recommended suppliers of 15N for use in agricultural studies are Cambridge Isotope Laboratories, Isotec, Inc., and ICON Services, Inc. In recent years, the U.S. Department of Energy was the main producer and supplier of both 15N-enriched and 15N-depleted compounds. The labeled materials were produced at the Los Alamos Scientific Laboratory in New Mexico and could be purchased from Monsanto Research Corporation's Mound Laboratory (now EG & G Mound Applied Technologies, Miamisburg, OH). The Department of Energy (DOE) has been requested by another U.S. producer of isotopically enriched compounds to withdraw from the production and distribution of such compounds and a formal petition and request was released for public comment. Despite letters of

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

943

support from 15N users, DOE is phasing out its direct involvement as a supplier of 15N. A large number of inorganic and organic compounds, including N gases, labeled with 15N are offered by Cambridge Isotope Laboratories (CIL), 20 Commerce Way, Woburn, MA 0180l. Merck and Co., Inc.lIsotopes, 4545 Oleatha Ave., St. Louis, MO 63116 still supplies a variety of 15N-Iabeled organic compounds but reportedly is divesting its holdings in this market. MSD ISOTOPES, the stable isotope division of Merck Frosst Canada Inc., Montreal, Quebec, has been offered for sale and its status as a supplier of 15N is unclear. Although several laboratories will perform N-isotope-ratio analyses upon special arrangement, two suppliers of 15N-Iabeled compounds also provide analytical service particularly directed toward agricultural research. Labeled materials can be purchased from ICON Services, Inc., 19 Ox Bow Lanes, Summit, NJ 0790l. This supplier through its marketing arm, U.S. Services Inc., makes available automated N analyses provided by Isotope Services, Inc. (ICI). Samples can be submitted in the form of NH/ (e.g., from Kjeldahl digests), from which N2 is generated via an automated Rittenburg procedure, or as dry plant or soil material, from which N2 is produced via automated Dumas combustion for both total N and N-isotope-ratio determinations. Isotec, Inc., a primary producer of isotopically enriched materials, offers a wide range of 15N-enriched compounds plus 15N-depleted NH 4N0 3 and (NH4hS04' but may not continue to offer analytical services using an automated Dumas-type procedure. The company is a subsidiary of Matheson, USA and is located at 3858 Benner Road, Miamisburg, OH 45342. In addition to these commercial ventures, analytical services also are offered by university laboratories. Buyers of 15N-Iabeled compounds should be aware that companies differ in their method of quoting prices. All prices should be compared on the basis of contained 15N rather than on the weight of total material. Some prices quoted may include container, handling, packaging, or shipping charges while others may not. A guaranteed analysis also should be insisted upon. Wise use of 15N begins with a wise purchase of the amount and enrichment level needed to achieve research objectives at minimum acceptable cost. REFERENCES Alberts, E.E., R.E. Burwell, and G.E. Schuman. 1977. Soil nitrate-nitrogen determined by coring and solute extraction techniques. Soil Sci. Soc. Am. J. 41:90-92. Allison, F.E. 1955. The enigma of soil nitrogen balance sheets. Adv. Agron. 7:213-250. Allmaras, R.R., and W.W. Nelson. 1971. Corn (Zea mays L.) root configuration as influenced by some row-interrow variants of tillage and straw mulch management. Soil Sci. Soc. Am. Proc. 35:974-980. Baker, J.L., and H.P. Johnson. 1981. Nitrate-nitrogen in tile drainage as affected by fertilization. J. Environ. Qual. 10:519-522. Barbee, G.C., and K.W. Brown. 1986. Comparison between suction and free-drainage soil solution samplers. Soil Sci. 141:149-154.

944

HAUCKET AL.

Barraclough, D., and M.J. Smith. 1987. The estimation of mineralization, immobilization, and nitrification in nitrogen-15 field experiments using computer simulation. J. Soil Sci. 38:519-530. Bartholomew, W. V. 1964. Guides in extending the use of tracer nitrogen in soils and fertilizer research.!? 81-96. In Soil and fertilizer nitrogen research, a projection into the future. A symposIUm, Publ. no. T 64-4 SF. TVA, Wilson Dam, AL. Beckett, P.H.T., and R. Webster. 1971. Soil variability: A review. Soils Fert. 34:1-15. Benjamin, J.G., R.M. Cruse, A.D. Blaylock, and L.V. Vogl. 1988. A small-plot point injector for liquid fertilizer application. Soil Sci. Soc. Am. J. 52:1194-1195. Bergersen, F.J. 1980. Measurement of nitrogen fixation by direct means. p. 65-110. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen fixation. John Wiley and Sons, Chichester, England. Bergstrom, L. 1987. Nitrate leaching and drainage from annual and perennial crops in tiledrained plots and Iysimeters. J. Environ. Qual. 16:11-18. Bigeriego, M., R.D. Hauck, and R.A. Olson. 1979. Uptake, translocation, and utilization of 15N-depleted fertilizer in irrigated com. Soil Sci. Soc. Am. J. 43:528-533. Biggar, J.w., and D.R. Nielsen. 1976. Spatial variability of the leaching characteristics of a field soil. Water Resour. Res. 12:78-84. Blackburn, T.H. 1979. Method for measuring rates of NHl turnover in anoxic sediments, using a 15N-NH4+ dilution technique. Appl. Environ. Microbiol. 37:760-765. Boswell, F.e., and O.E. Anderson. 1964. Nitrogen movement in undisturbed profiles of fallowed soils. Agron. J. 56:278-281. Bremner, J.M. 1965a. Isotope-ratio analysis of nitrogen in nitrogen-15 tracer investigations. p. 1256-1286. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Bremner, J.M. 1965b. Inorganic forms of nitrogen. p. 1179-1237. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA, Madison, WI. Bremner, J.M., H.H. Cheng, and A.P. Edwards. 1966. Assumptions and errors in nitrogen-15 tracer research. p. 429-442. In Report of the FAOIIAEA Tech. Meet. (Braunschweig, Germany, 1963). Pergamon Press, Elmsford, NY. Bremner, J.M., and C.S. Mulvaney. 1982. Nitrogen-total. p. 595-624. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Broadbent, F.E., and A.B. Carlton. 1980. Methodology for field trials with nitrogen-15depleted nitrogen. J. Environ. Qual. 9:236-242. Brown, K.W., J.C. Thomas, and M.W. Aurelius. 1985. Collecting and testing barrel sized undisturbed soil monoliths. Soil Sci. Soc. Am. J. 49:1067-1069. Brown, M.A., and G.M. Volk. 1966. Evaluation of ureaform fertilizer using nitrogen-15labeled materials in sandy soils. Soil Sci. Soc. Am. Proc. 30:278-281. Buresh, R.J., E.R. Austin, and E.T. Craswell. 1982. Analytical methods in 15N research. Fert. Res. 3:37-62. Burris, R.H., and P.W. Wilson. 1957. Methods for measurement of nitrogen fixation. p. 355-366. In S.P. Colowick and N.D. Kaplan (ed.) Methods in enzymology. Vol. 4. Academic Press, New York. Buzicky, G.C., G.w. Randall, R.D. Hauck, and A.C. Caldwell. 1983. Fertilizer N losses from a tile drained Mollisol as influenced by rate and time of 15-N depleted fertilizer application. p. 213. In Agronomy abstracts. ASA, Madison, WI. Cameron, D.R., M. Nyborg, J.A. Toogood, and D.H. Laverty. 1971. Accuracy of field sampling for soil tests. Can. J. Soil Sci. 51:165-175. Cameron, K.C., and A. Wild. 1984. Potential aquifer pollution leaching following the plowing of temporary grassland. J. Environ. Qual. 13:274-278. Capindale, J.B., and D.H. Tomlin. 1957. Mass-spectrometric assay of elementary nitrogen. Nature (London) 180:701-702. Carter, J.N., O.L. Bennett, and R.W. Pearson. 1967. Recovery of fertilizer nitrogen under field conditions using nitrogen-15. Soil Sci. Soc. Am. Proc. 31:50-56. Cassel, D .K., T.H. Krueger, F. W. Schroer, and E.B. Norum. 1974. Solute movement through disturbed and undisturbed soil cores. Soil Sci. Soc. Am. Proc. 38:36-40. Chen, D., P.M. Chalk, and J.R. Freney. 1991. External-source contamination during extraction-distillation in isotope-ratio analysis of soil inorganic nitrogen. Anal. Chim. Acta 245:49-55.

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

945

Chichester, P.w., and S.J. Smith. 1978. Disposition of 15N-labeled fertilizer nitrate applied during corn culture in field lysimeters. J. Environ. Qual. 7:227-233. Cho, C.M., and E. Haunold. 1966. Some problems encountered in the preparation of nitrogen-15 gas samples and mass spectrometric work. p. 443-445. In Use of isotopes in soil organic matter studies. Report of the FAOIIAEA Tech. Meet. (Braunschweig, Germany. 9-14 Sept. 1963). Pergamon Press, Elmsford, NY. Cline, M.G. 1944. Principles of soil sampling. Soil Sci. 58:275-288. Duke, H.R., and H.R. Haise. 1973. Vacuum extractors to assess deep percolation losses and chemical constituents of soil water. Soil Sci. Soc. Am. Proc. 37:963-964. Duxbury, J.M., and P.K. McConnaughey. 1986. Effect of fertilizer source on denitrification and nitrous oxide emissions in a maize-field. Soil Sci. Soc. Am. J. 50:644-648. Edwards, A.P. 1978. A guide to the use of 14N and 15N in environmental research. Spec. Rep. 78-18. U.S. Army Cold Regions Research and Engineering Lab., Hanover, NH. Faust, H., H. Gurtler, H. Huebner, H. Mielke, W. Rommel, M. Ulbrich, and K. Wetzel. 1963. Report on cooperative research on feeding ammonium bicarbonate to cattle. Wiss. Karl Marx Univ. Leipzig 12:711-718. Fiedler, R. 1984. The measurement of 15N. p. 233-282. In M.P. L'Annunziata and J.O. Legg (ed.) Isotopes and radiation in agricultural sciences. Academic Press, London. Fiedler, R., and G. Proksch. 1975. The determination of nitrogen-15 by emission and mass spectrometry in biochemical analysis: A review. Anal. Chim. Acta 78:1-62. Follett, R.F., R.R. Allmaras, and G.A. Reichman. 1974. Distribution of corn roots in sandy soil with a declining water table. Agron. J. 66:288-292. Folorunso, O.A., and D.E. Rolston. 1984. Spatial variability of field-measured denitrification gas fluxes. Soil Sci. Soc. Am. J. 48:1214-1219. Fried, M. 1978. Critique of field trials with isotopically labeled nitrogen fertilizer. p. 43-62. In D.R. Nielsen and J.G. MacDonald (ed.) Nitrogen in the environment. Vol. 1. Nitrogen behavior in field soil. Academic Press, New York. Gast, R.G., w.w. Nelson, and G.W. Randall. 1978. Nitrate accumulation in soils and loss in tile drainage following nitrogen applications to continuous corn. J. Environ. Qual. 7:258-261. GOfilez, K.A., and A.A. Gomez. 1984. Statistical procedures for agricultural research. 2nd ed. John Wiley and Sons, New York. Guiraud, G. 1984. Contribution du marquage isotopique a l'evaluation des transferts d'azote entre les compartiments organiques et mineraux dans les systemes sol-plantes. These Etat, Universlte P. et M. Curie, Paris. Guiraud, G., C. Marol, and M.C. Thibaud. 1989. Mineralization of nitrogen in the presence of a nitrification inhibitor. Soil BioI. Biochem. 21:29-34. Haines, B.L., 1.B. Waide, and R.L. Todd. 1982. Soil solution nutrient concentrations sampled with tension and zero-tension lysimeters: Report of discrepancies. Soil Sci. Soc. Am. J. 46:658-661. Hallberg, G.R., J.L. Baker, and G.w. Randall. 1986. Utility of tile-lined effluent studies to evaluate the impact of agricultural practices on groundwater. p. 298-325. In Proc. of Agric. Impacts on Groundwater, Omaha, NE. 11-13 Aug. 1986. Natl. Well Water Assoc., Dublin, OH. Hansen, E.A., and A.R. Harris. 1975. Validity of soil-water samples collected with porous ceramic cups. Soil Sci. Soc. Am. Proc. 39:528-536. Hart, P.B.S., J.H. Rayner, and D.S. Jenkinson. 1986. Influence of pool substitution on the interpretation of fertilizer experiments with 15N. J. Soil Sci. 37:389-403. Hauck, R.D. 1973. Nitrogen tracers in nitrogen cycle studies-past use and future needs. J. Environ. Qual. 2:317-327. Hauck, R.D. 1978. Critique of field trials with isotopically labeled nitrogen fertilizer. p. 63-77. In D.R. Nielsen and J.G. MacDonald (ed.) Nitrogen in the environment. Vol. 1. Nitrogen behavior in field soil. Academic Press, New York. Hauck, R.D. 1979. Methods for studying N transformations in paddy soils: Review and comments. p. 73-93. In Nitrogen and rice. Int. Rice Res. Inst., Manila, Philippines. Hauck, R.D. 1982. Nitrogen-isotope-ratio analysis. p. 735-779. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Hauck, R.D. 1984. Significance of nitrogen fertilizer microsite reactions in soil. p. 507-519. In R.D. Hauck et al. (ed.) Nitrogen in crop production. ASA, CSSA, and SSSA, Madison, WI.

946

HAUCK ET AL.

Hauck, RD. 1985. Slow-release and bioinhibitor-amended nitrogen fertilizers. p. 293-322. In D.P. Engelstad (ed.) Fertilizer technology and use. 3rd ed. SSSA, Madison, WI. Hauck, RD. 1994. Synthesis of 15N-labeled isobutylidene diurea, oxamide, and ureaforms for use in agronomic studies. Commun. Soil Sci. Plant. Anal. 25:191-197. Hauck, R.D., and J.M. Bremner. 1964. The methodology ofNl5 research. p. 97-110. In Soil and fertilizer nitrogen research, a projection into the future, A symposium. Publ. no. T64-4 SF. TVA, Wilson Dam, AL. Hauck, RD., and J.M. Bremner. 1976. Use of tracers for soil and fertilizer nitrogen research. Adv. Agron. 28:219-266. Hauck, RD., and M. Bystrom. 1970. 15N - A selected bibliography for agricultural scientists. The Iowa State Univ. Press, Ames. Hauck, RD., and V.J. Kilmer. 1975. Cooperative research between the Tennessee Valley Authority and land-grant universities on nitrogen fertilizer use and water quality. p. 655-660. In E.R Klein and P.D. Klein (ed.) Proc. 2nd Int. Congr. Stable Isotopes, Oak Brook,lL. U.S. Energy Res. Dev. Admin., Agronne, IL. Hauck, R.D., and R.W. Weaver (ed.) 1986. Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18: SSSA, Madison, WI. Hergert, G.W. 1986. Nitrate leaching through sandy soil as affected by sprinkler irrigation management. J. Environ. Qual. 15:272-278. Hills, F.J., F.E. Broadbent, and O.A. Lorenz. 1983. Fertilizer nitrogen utilization by corn, tomato, and sugarbeet. Agron. J. 75:423-426. Jansson, S.L. 1958. Tracer studies on nitrogen transformations in soil with special attention to mineralisation-immobilisation relationships. Kgl. Lantbruks-Hoegsk. Ann. Roy. Agric. Coll., Swed. 24:101-361. Jansson, S.L. 1966. Nitrogen transformation in soil organic matter. p. 283-296. In Report of the FAO/IAEA Tech. Meet., (Braunschweig, Germany. 1963). Pergamon Press, Elmsford, New York. Jenkinson, D.S., R.H. Fox, and J.H. Rayner. 1985. Interactions between fertilizer nitrogen and soil nitrogen-the so-called 'priming' effect. J. Soil Sci. 36:425-444. Jokela, W.E., and G.W. Randall. 1987. A nitrogen-15 microplot design for measuring plant and soil recovery of fertilizer nitrogen applied to corn. Agron. J. 79:322-325. Jones, A.J., and R.J. Wagenet. 1984. In situ estimation of hydraulic conductivity using simplified methods. Water Resour. Res. 20:1620-1626. Jones, J.B., and W.J.A. Steyn. 1973. Sampling, handling, and analyzing plant tissue samples. p. 249-270. In L.M. Walsh and J.D. Beaton (ed.) Soil testing and plant analysis. SSSA, Madison, WI. Jones, M.B., C.c. Delwiche, and W.A. Williams. 1977. Uptake and losses of 15N applied to annual grass and clover in Iysimeters. Agron. J. 69:1019-1023. Jordan, C.F. 1968. A simple, tension-free Iysimeter. Soil Sci. 105:81-86. Junk, G., and H.J. Svec. 1958. The absolute abundance of the nitrogen isotopes in the atmosphere and compressed gas from various sources. Geochim. Cosmochim. Acta 14:234-243. Keeney, D.R., and D.W. Nelson. 1982. Nitrogen-inorganicforms. p. 643-698. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Kelley, K.R. 1994. Conveyor-belt apparatus for fine grinding of soil and plant materials. Soil Sci. Soc. Am. J. 58:144-146. Kendall, C., and E. Grim. 1990. Combustion tube method for measurement of nitrogen isotope ratios using calcium oxide for total removal of carbon dioxide and water. Anal. Chem. 62:526-529. Kennedy, I.R. 1965. Release of nitrogen from amino acids with ninhydrin for 15N analysis. Anal. Biochem. 11:105-110. Khanif, Y.M., O. Van Cleemput, and L. Baert. 1984. Interaction between nitrogen fertilization, rainfall and groundwater pollution in sandy soil (Belgium). Water, Air, Soil Pollut. 22:447-452. Khonke, H., F.R. Dreibelbis, and J.M. Davidson. 1940. A survey and discussion of Iysimeters and a bibliography on their construction and performance. Misc. Pub!. no. 372. USDA, Washington, DC. Kilmer, V.J., R.D. Hauck, and O.P. Engelstad. 1974. Nitrogen isotopes and water quality research. p. 35-43. ~n ~ontribution of irrigation and drainag~ ~o world food supply. Spec. Conf. Proc., BilOXI, MS. 14-16 Aug. 1974. Am. Soc. ClVlI Eng., New York.

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

947

Kirchmann, H. 1985. Losses, plant uptake and utilisation of manure nitrogen during a production cycle. Acta Agric. Scand. (Suppl. 24). Kirchmann, H. 1989. A 3-year balance study with aerobic, anaerobic and fresh 15N-Iabeled poultry manure. p. 113-125. In I.A. Hansen and K. Henniksen (ed.) Nitrogen in organic wastes applied to soils. Academic Press, London. Kirchmann, H. 1990. Nitrogen interactions and crop uptake from fresh and composted 15N_ labeled poultry manure. 1. Soil Sci. 41:379-385. Kirkham, D., and W.v. Bartholomew. 1954. Equations for following nutrient transformations in soil, utilizing tracer data. Soil Sci. Soc. Am. Proc. 18:33-34. Kirkham, D., and W.v. Bartholomew. 1955. Equations for following nutrient transformations in soil, utilizing tracer data: II. Soil Sci. Soc. Am. Proc. 19:189-192. Kissel, D.E., and S.I. Smith. 1978. Fate of fertilizer nitrate applied to coastal bermudagrass on a swelling clay soil. Soil Sci. Soc. Am. 1. 42:77-80. Kitur, B.K., M.S. Smith, R.L. Blevins, and W.w. Frye. 1984. Fate of 15N-depleted ammonium nitrate applied to no-tillage and conventional tillage corn. Agron. 1. 76:240-242. Koike, I., and A. Hattori. 1978. Simultaneous determinations of nitrification and nitrate reduction in coastal sediments by 15N dilution technique. Appl. Environ. Microbiol. 35:853-857. LeClerg, E.L., W.H. Leonard, and A.G. Clark. 1962. Field plot technique. 2nd ed. Burgess Publ. Co., Minneapolis. Legg, 1.0., and 1.1. Meisinger. 1982. Soil nitrogen budgets. p. 503-566. In F.l. Stevenson (ed.) Nitrogen in agricultural soils. Agron. Monogr. 22. ASA, CSSA, and SSSA, Madison, WI. Legg, 1.0., and C. Sloger. 1975. A tracer method for determining symbiotic nitrogen fixation in field studies. p. 661-667. In Proc. 2nd Int. Conf. on Stable Isotopes, Oakbrook, IL. Argonne Nat. Lab., U.S. Dep. of Energy, Washington, DC. Linden, D.R. 1977. Design, installation and use of porous ceramic samplers for monitoring soil-water quality. U.S. Dep. Agric. Tech. Bull. 1562. U.S. Gov. Print. Office, Washington, DC. Litaor, M.I. 1988. Review of soil solution samplers. Water Resour. Res. 24:727-733. Martin, A.E., and P.l. Ross. 1968. Significance of errors in 15N measurements in soil:plant research. Int. Congr. Soil Sci. Trans., 9th (Adelaide) 3:521-529. Meisinger, 1.1.1984. Evaluating plant-available nitrogen in soil-crop systems. p. 391-416. In R.D. Hauck et al. (ed.) Nitrogen in crop production. ASA, CSSA, and SSSA, Madison, WI. Meisinger, 1.1., V.A. Bandel, G. Stanford, and 1.0. Legg. 1985. Nitrogen utilization of corn under minimal tillage and moldboard plow tillage. I. Four-year results using labeled N fertilizer on an Atlantic Coastal Plain soil. Agron. 1. 77:602-611. Minagawa, M., D.A. Winter, and I.R. Kaplan. 1984. Comparison of Kjeldahl and combustion methods for measurement of nitrogen isotope ratios in organic matter. Anal. Chern. 56: 1859-1861. Mohanty, S.K., S.P. Chakravorti, and A. Bhadrachalam. 1989. Nitrogen balance studies in rice using 15N-Iabelled urea and urea supergranules. 1. Agric. Sci., Cambridge 113:119121. Montgomery, B.R., L. Prunty, and I.W. Bauder. 1987. Vacuum trough extractors for measuring drainage and nitrate flux through sandy soils. Soil Sci. Soc. Am. 1. 51:271-276. Moraghan, 1.1., T.l. Rego, R.I. Buresh, P.L.G. Vlek, I.R. Burford, S. Singh, and K.L. Sahrawat. 1984. Labelled nitrogen fertilizer research with urea in the semi-arid tropics. Plant Soil 80:21-33. Mosier, A.R., and O. Heinemeyer. 1985. Current methods used to estimate N20 and N2 emissions from field soils. p. 79-99. In H.L. Golterman (ed.) Denitrification in the nitrogen cycle. Plenum Press, New York. Mulvaney, R.L. 1991. Some recent advances in the use of nitrogen-15 for research on nitrogen transformations in soil. p. 283-296. In Stable isotopes in plant nutrition, soil fertility and environmental studies. FAO/IAEA, Vienna, Austria. Mulvaney, R.L. 1993. Mass spectrometry. p. 11-57. In R. Knowles and T.H. Blackburn (ed.) Nitrogen isotopes techniques. Academic Press, San Diego. Myrold, D.D., and I.M. Tiedje. 1986. Simultaneous estimation of several nitrogen cycle rates using 15N: Theory and application. Soil BioI. Biochem. 18:559-568. Nelson, W.W., and G.W. Randall. 1983. Fate of residual nitrate-N in a tile-drained Mollisol. p. 215. In Agronomy abstracts. ASA, Madison, WI.

948

HAUCKET AL.

Norman, A.G., and C.H. Werkman. 1943. The use of N15 in determining N recovery from plant materials decomposing in soil. J. Am. Soc. Agron. 35:1023-1025. Olson, R.V. 1980. Plot size requirements for measuring residual fertilizer nitrogen and nitrogen uptake by corn. Soil Sci. Soc. Am. J. 44:428-429. Owens, L.D. 1960. Nitro~en movement and transformations in soils as evaluated by a Iysimeter study utilizing Isotopic nitrogen. Soil Sci. Soc. Am. Proc. 24:372-376. Parkin, T.B. 1987. Soil microsites as a source of denitrification variability. Soil Sci. Soc. Am. J. 51:1194-1199. Parkin, T.B., J.L. Starr, and J.J. Meisinger. 1987. Influence of sample size on measurement of soil denitrification. Soil Sci. Soc. Am. J. 51:1492-1501. Peterson, R.G., and L.D. Calvin. 1986. Sampling. p. 33-51. In A. Klute (ed.) Methods ofsoil analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9. ASA, Madison, WI. Phillips, D.A., M.B. Jones, and K.W. Foster. 1986. Advantages of the nitrogen-15 dilution technique for field measurement of symbiotic dinitrogen fixation in legumes. p. 11-21. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18. ASA, CSSA, and SSSA, Madison, WI. Powlson, D.S., G. Pruden, A.E. Johnston, and D.S. Jenkinson. 1986. The nitro~en cycle in the Broadbalk wheat experiment: Recovery and loss of 15N-Iabelled fertihzer applied in ~pring and inputs of nitrogen from the atmosphere. J. Agric. Sci., Cambridge 107.591-609. Priebe, D.L., and A.M. Blackmer. 1989. Preferential movement of oxygen-18-labeled water and nitrogen-15-labeled urea through macropores in a Nicollet soil. J. Environ. Qual. 18:66-72. Pruden, G., D.S. Powlson, and D.S. Jenkinson. 1985. The measurement of 15N in soil and plant material. Fert. Res. 6:205-218. Rauhe, K., and H. Bornak. 1970. Die Wirkung von 15N-markiertem Rinderkot mit verschiedenen Zusatzen im Feldversuch unter besonderer Berucksichtigung der Reproduktion der organischen Substanz im Boden. Albrecht-Thaer-Arch. 14:937-948. Rauhe, K., E. Fichtner, F. Fichtner, E. Knappe, and W. Drauschke. 1973. Quantifizierung der Wirkung organischer und mineralischer Stickstoffdiinger auf Pflanze und Boden unter besonderer Berucksichtigung 15N-markierter tierischer Exkremente. Arch. Acker-, Pflanzenbau Bodenkd. 17:907-916. Rennie, R.J. 1986. Advantages and disadvantages of nitrogen-15 isotope dilution to quantify dinitrogen fixation in field-grown legumes-A critique. p. 43-58. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18. ASA, CSSA, and SSSA, Madison, WI. Rhoades, J.D., and J.D. Oster. 1986. Solute content. p. 985-1006. In A. Klute (ed.) Methods of soil analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Richard, T.L., and T.S. Steenhuis. 1988. Tile drain sampling of preferential flow on a field scale. J. Contam. Hydrol. 3:307-325. Rittenberg, D. 1948. The preparation of gas samples for mass spectrographic analysis. p. 31-42. In D.W. Wilson et al. (ed.) Preparation and measurement of isotopic tracers. J. W. Edwards, Ann Arbor, MI. Rittenberg, D., A.S. Keston, F. Rosebury, and R. Schoenheimer. 1939. Studies in protein metabolism. II. The determination of nitrogen isotopes in organic compounds. J. BioI. Chern. 127:291-299. Rubio, J.L., and R.D. Hauck. 1986. Uptake and use patterns of nitrogen from urea, oxamide, and isobutylidene diurea by rice plants. Plant Soil 94:109-123. Saffigna, P. 1988. N-15 methodology in the field. p. 433-451. In J.R. Wilson (ed.) Advances in nitrogen cycling in agricultural ecosystems. Proc. Symp. at Brisbane, Australia. 11-15 May 1987. CAB Int., Wallingford, UK. Sanchez, C.A., and A.M. Blackmer. 1987. A method for application of nitrogen-15-labeled anhydrous antmonia to small plots. Soil Sci. Soc. Am. J. 51:259-261. Sanchez, C.A., A.M. Blackmer, R. Horton, and D.R. Timmons. 1987. Assessment of errors associated with plot size and lateral movement of nitrogen-15 when studying fertilizer recovery under field conditions. Soil Sci. 144:344-351. San Pietro, A. 1957. The measurement of stable isotores. p. 473-488. In S.P. Colowick and N.O. Kaplan (ed.) Methods in enzymology. Vo . 4. Academic Press, New York.

NITROGEN TRACERS IN AGRICULTURAL RESEARCH

949

Selles, F., R.E. Karamanos, and R.G. Kachanoski. 1986. The spatial variability of nitrogen-IS and its relation to the variability of other soil properties. Soil Sci. Soc. Am. J. 50:105-110. Shaffer, K.A., D.D. Fritton, and D.E. Baker. 1979. Drainage water sampling in a wet, dual-pore soil system. J. Environ. Qual. 8:241-246. Shen, S.M., G. Pruden, and D.S. Jenkinson. 1984. Mineralization and immobilization of nitrogen in fumigated soil and the measurement of microbial biomass nitrogen. Soil BioI. Biochem. 16:437-444. Smith, J.H., J.O. Legg, and J.N. Carter. 1963. Equipment and procedures for N-15 analysis of soil and plant material with the mass spectrometer. Soil Sci. 96:313-318. Soileau, J.M., and R.D. Hauck. 1987. A historical review of U.S. lysimeter research with emphasis on fertilizer percolation losses. p. 208-304. In Yu-S Fox (ed.) Infiltration principles and practices. Water Resources Res. Center, Univ. of Hawaii, Honolulu. Sprinson, D.B., and D. Rittenberg. 1948. Preparation of gas samples for mass spectrometric analysis of isotope abundance. p. 82-93. U.S. Nav. Med. Bull., March-April Suppl. Sprinson, D.B., and D. Rittenberg. 1949. The rate of utilization of ammonia for protein synthesis. J. BioI. Chern. 180:707-714. Starr, J.L., J.J. Meisinger, and T.B. Parkin. 1991. Experience and knowledge gained from vadose zone sampling. p. 279-290. In R.G. Nash and A. Leslie (ed.) Agrichemical residue sampling design and techniques: Soil and groundwater. Am. Chern. Soc. Symp., 23-24 Apr. 1990. Boston, MA. Am. Chern. Soc., Washington, DC. Stevenson, EJ. 1982. Nitrogen-organic forms. p. 625-641. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Stumpe, J.M., P.L.G. Vlek, S.K. Mughogho, and F. Ganry. 1989. Microplot size requirements for measuring balances of fertilizer nitrogen-IS applied to maize. Soil Sci. Soc. Am. J. 53:797-800. Tabor, J.A., A.W. Warrick, D.E. Myers, and D.A. Pennington. 1985. Spatial variability of nitrate in irrigated cotton: II. Soil nitrate and correlated variables. Soil Sci. Soc. Am. J. 49:390-394. Tackett, J.L., E. Burnett, and D.W. Fryrear. 1965. A rapid procedure for securing large, undisturbed soil cores. Soil Sci. Soc. Am. Proc. 29:218-220. Therion, J.J., H.G.C. Human, C. Clase, R.I. Mackie, and A. Kistner. 1986. Automatic nitrogen-IS analyser for use in biological research. Analyst 111:1017-1021. Thomas, G.W., and R.E. Phillips. 1979. Consequences of water movement in macropores. J. Environ. Qual. 8:149-152. Tyler, D.D., and G.W. Thomas. 1977. Lysimeter measurements of nitrate and chloride losses from soil under conventional and no-tillage corn. J. Environ. Qual. 6:63-66. Vanden Heuvel, R.M. 1988. Improved apparatus for preparing nitrogen-IS labeled anhydrous ammonia. Soil Sci. Soc. Am. J. 52:1483-1486. Vanden Heuvel, R.M., and S. Harrold. 1990. Field apparatus for knife injection of nitrogenIS-labeled liquid anhydrous ammonia. Soil Sci. Soc. Am. J. 54:531-534. Vigil, M.E, and D.E. Kissel. 1991. Equations for estimating the amount of nitrogen mineralized from crop residues. Soil Sci. Soc. Am. J. 55:757-761. Vose, P.B., and R.L. Victoria. 1986. Re-examination of the limitations of nitrogen-IS isotope dilution technique for the field measurement of dinitrogen fixation. p. 23-41. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18. ASA, CSSA, and SSSA, Madison, WI. Walters, D.T., and G.L. Malzer. 1990. Nitrogen management and nitrification inhibitor effects on nitrogen-IS urea: II. Nitrogen leaching and balance. Soil Sci. Soc. Am. J. 54: 122-130. Warrick, A.W., G.J. Mullen, and D.R. Nielsen. 1977. Scaling field measured soil hydraulic properties using a similar media concept. Water Resour. Res. 13:355-362. Weaver, R.W. 1986. Measurement of biological dinitrogen fixation in the field. p. 1-10. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18. ASA, CSSA, and SSSA, Madison, WI. Westerman, R.L., L.T. Kurtz, and R.D. Hauck. 1972. Recovery of 15N-labeled fertilizers in field experiments. Soil Sci. Soc. Am. Proc. 36:82-86. White, R.E., R.A. Haigh, and J.H. McDuff. 1987. Frequency distributions and spatially dependent variability of ammonium and nitrate concentrations in soil under grazed and ungrazed grassland. Fert. Res. 11:193-208.

950

HAUCKETAL.

White, P.J., 1. Vallis, and P.G. Saffigna. 1986. The effect of stubble management on the availability of 15N-labeled residual fertilizer nitrogen and crop stubble nitrogen in an irrigated black earth. Aust. J. Exp. Agric. 26:99-106. Woodcock, T.M., G. Pruden, D.S. Powlson, and D.S. Jenkinson. 1982. Apparatus for applying 15N-labelled fertilizer uniformly to field micro-plots. J. Agric. Eng. Res. 27:369-

372.

Zwerman, P.J., T. Greweling, S.D. Klausner, and D.J. Lathwell. 1972. Nitrogen and phosphorus content of water from tile drains at two levels of management and fertilization. Soil Sci. Soc. Am. Proc. 36:134-137.

Published 1994

Chapter 41 Nitrogen Availability Indices L. G. BUNDY, University of Wisconsin, Madison, Wisconsin

J. J. MEISINGER, USDA-ARS, Beltsville, Maryland

Available N in soils is that N present in forms, concentrations, and spatial position that allow utilization by plants growing in the soil. Most plant species can effectively use either NH4 or N0 3 , but a few species use NH4 preferentially. Organic N compounds such as urea and amino acids can also be taken up by plants directly, but the contribution of these organic N compounds to the overall plant N requirement is minimal. Since NH4 is converted to N0 3 in most soils, N0 3 is usually the predominant form of available N in the plant root zone. Available N in soils can originate from many sources including fertilizer N additions and mineralization of organic N from soil organic matter, crop residues, and organic wastes (Keeney, 1982; Meisinger, 1984). Most soils contain 0.08 to 0.4% of N, and 97 to 99% of this occurs as organic N compounds in soil organic matter (Keeney, 1982; Dahnke & Johnson, 1990). If 1 to 3% of this organic N is mineralized annually, 8 to 120 kg of N ha- 1 is released in a plant-available form (Bremner, 1965a). Nitrogen availability indices are N analyses or chemical or biological tests to measure or predict the amounts of available N released from soil under a specific set of conditions.

41-1 CURRENT STATUS OF NITROGEN AVAILABILITY INDICES 41-1.1 Previous Summaries Estimating soil N availability has been a goal of soil scientists since the early 1900s. Summaries of early research (Harmsen & Van Schreven, 1955; Allison, 1965; Bremner, 1965a) reached the general conclusions that: (i) chemical extraction methods were likely to be unsuccessful because they could not imitate the action of soil microorganisms; (ii) biological Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711. USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 951

952

BUNDY & MEISINGER

incubations under standardized conditions were most successful because they used the same microbial agents active under field conditions; (iii) crop response methods were too costly and labor intensive for practical use, but were absolutely essential for calibration of laboratory methods; and (iv) methods based on soil N0 3-N levels were of "very limited value" because the soil N0 3-N pool was too transient to be useful. A 1973 review by Dahnke and Vasey emphasized the need to include some measure of preplant mineral N. However, they emphasized that preplant N is quite variable from year to year depending on factors such as previous crop yield, previous N fertilization, nongrowing season precipitation, and soil-leaching characteristics. Therefore, they recommended annual deep preplant soil sampling to assess N availability in subhumid climates. They also reviewed chemical and biological indexes but reached no consensus regarding a recommended method. In the 1980s, reviews by Stanford (1982) and Keeney (1982) considered residual nitrate, short- and long-term incubations, and chemical extractants. The general conclusions of these reviews were that: (i) long-term biological mineralizations were most suitable but were not practicable, (ii) short-term mineralizations were acceptable but were affected by sample handling and pretreatment (Stanford, 1982) with NH4-N production after 7 d of anaerobic incubation being recommended (Keeney, 1982); and (iii) mild chemical extracts were acceptable for the ranking of soils with NH4-N hydrolyzed by 0.01 M CaCl2 upon overnight (16 h) autoclaving being the recommended method (Keeney, 1982). Keeney and Stanford also endorsed the preplant soil nitrate test in climates without extensive overwinter leaching. This was supported by the fact that most western U.S. states were already using some type of preplant residual N0 3 -N test in 1982. Evaluating plant-available N from a systems perspective was discussed by Meisinger (1984), who emphasized the need to systematically include all available N pools. This review discussed underlying principles, steady-state approaches, crop N requirements, N-efficiency estimates, and methods of estimating residual N0 3 -N and mineralizable N. Meisinger (1984) concluded that improving N-availability assessments should involve: use of both residual N0 3-N and N-mineralization tests, expanded use of local soil properties through soil taxonomic information, and an integration of this information for each specific site by use of computer models and local weather data. 41-1.2 Selective Review of Recent Work

Since the earlier reviews cited in section 41-1.1 were prepared, the need for reliable prediction of N availability in soils has increased due to economic and environmental incentives to use available N more efficiently in crop production and minimize losses of N from cropland to the environment. These incentives have stimulated continued research on N availability indices in several areas including: (i) development of new chemical

NITROGEN AVAILABILITY INDICES

953

indices; (ii) modification and improvement of aerobic incubation procedures to provide more accurate assessments of N availability and facilitate an improved understanding of the characteristics of mineralizable N in soils; and (iii) development and evaluation of field tests for N availability. A selective review of recent work in these areas is provided below. 41-1.2.1 Chemical Methods Development and Comparison with Other Indices Chemical methods of assessing N availability are appealing because they offer a simple and rapid approach to estimation of crop N needs for use in production and a convenient assessment of relative differences among various experimental treatments in research projects. To be successful, chemical procedures must detect innate differences among soils or treatments in the quantities of N likely to mineralize during a specific period (often the crop N uptake period) and must reflect the influences of environmental conditions on the rates and amounts of N mineralized. Given these demanding specifications, none of the many chemical indices proposed have been adopted for routine use in production or research, and Fox and Piekielek (1984) and Meisinger (1984) stated a widely accepted view that any single soil index is unlikely to provide sufficiently accurate predictions of the economic optimum fertilizer N recommendations for corn (Zea mays L.). Recent work with chemical methods includes development and evaluation of new techniques and extensive comparisons of various chemical indices with other methods of assessing N availability including aerobic and anaerobic incubation tests, soil inorganic N measurements, and crop N uptake in field studies. New methods proposed include determination of NH4-N released by heating soil samples with 2 M KCI (Gianello & Bremner, 1986a; 0ien & Selmer-Olsen, 1980), measurement of NH 4-N released by steam distillation of soil samples with pH 11.2 phosphate-borate buffer (Gianello & Bremner, 1988), and UV absorbance of NaHC0 3 soil extracts at 200 nm (Hong et aI., 1990). 41-1.2.2 Undisturbed Soil Core Incubation Nitrogen mineralized during incubation of undisturbed soil cores can provide a more reliable assessment of N availability than standard methods using disturbed soil samples (Rice et aI., 1987; Cabrera & Kissel, 1988a, b). Measurements of N mineralized from disturbed soil samples often overestimate field N availability due to stimulation of mineralization by drying, crushing, and sieving the soil. The relatively large number of undisturbed soil cores needed to assess N mineralization and the process required to obtain these samples (Myers et aI., 1989) probably makes this approach impractical for field-scale predictions of N mineralization (Cabrera & Kissel, 1988a). However, Cabrera and Kissel (1988a) found that estimates of N mineralization in undisturbed soil could be obtained using data from disturbed samples or measured soil characteristics. Use of the undisturbed

954

BUNDY & MEISINGER

core technique is essential for N mineralization studies where variation in the degree of soil disturbance is an integral component of the systems under evaluation such as in tillage studies, pastures, forests, and fallow soils. 41-1.2.3 Field Tests for Nitrogen Availability A major emphasis in recent work on N availability indices (NAI) is the development and evaluation of field tests for N availability. These studies involve measurement of inorganic N (usually N03 ) in soil before planting a test crop or at a speGific time during the crop growing season and establishment of the relationship between the soil inorganic N content and the amounts of N available to a test crop. Once this relationship is known, results from the soil N test are used to provide N recommendations for crops. Preplant soil profile N03 tests have long been recommended as a method of assessing N availability and predicting crop N needs in subhumid areas in the western USA and Canada (Hergert, 1987; Dahnke & Johnson, 1990). Although early work (King & Whitson, 1901, 1902; Buckman, 1910; Call, 1914) and periodic studies during the 1950 to 1970 era (see Keeney, 1982; Stanford, 1982; Meisinger, 1984; Dahnke & Johnson, 1990 for reviews) showed that soil profile N0 3 provided important amounts of available N to crops in at least some growing seasons in humid climates, soil N0 3 tests were not adopted in the higher rainfall areas of the USA. Recent studies of soil profile N03 accumulation and retention and its effects on crop N needs (Sheppard & Bates, 1986; Bundy & Malone, 1988; Roth & Fox, 1990; Liang et aI., 1991) reconfirms earlier findings that substantial amounts of N03 can remain in the medium-to fine-textured soils during the overwinter period and can contribute substantial amounts of available N to subsequent crops. These results suggest that preplant measurement of soil profile N0 3 can provide a useful assessment of the probable contribution of profile N0 3 to crop N needs in areas with humid climates (Bock & Kelley, 1992; Bundy et aI., 1992). Another approach to use of soil N0 3 measurements to assess N availability in the field is the pre-sidedress soil N0 3 test (PSNT) for com proposed by Magdoff et ai. (1984). This procedure involves determination of N0 3-N concentration in the 0- to 30-cm soil layer when com plants are 15 to 30 cm tall, and is discussed in detail in section 41-2.1.2. The delayed sampling time used in the PSNT method essentially provides for an in situ incubation in that soil and climatic factors affecting N availability are allowed to interact during the period prior to soil sampling (Magdoff et al., 1990). Therefore, the PSNT has potential for predicting gains in available N due to net N mineralization from soil organic matter, crop residues, including those from previous legumes, and manures or other organic N sources. Similarly, available N losses through leaching or denitrification occurring before soil sampling should be reflected by the PSNT. This procedure

NITROGEN AVAILABILITY INDICES

955

has been evaluated in several locations in humid regions of the USA and is used to develop N-fertilizer recommendations for com in several states. It should be noted that the PSNT differs from the preplant soil profile N0 3 test in that the PSNT provides an index of N availability that can be used with appropriate calibration data to predict crop N response or the amount of additional N required by the crop. In contrast, preplant profile N0 3 tests provide a direct measurement of available N in some portion of the crop root zone, and this available N is usually directly credited against the crop N needs. 41-2 METHODS 41-2.1 Field Methods 41-2.1.1 Residual Profile Nitrate Tests 41-2.1.1.1 Field Sampling. Field sampling techniques for residual profile nitrate tests are based on studies of spatial variability of soil nitrate in the Western and Great Plains regions of the USA and Canada (Cameron et aI., 1971; Bole & Pittman, 1976; Reuss et aI., 1977). Results from these and other studies indicate that a composite of about 20 cores per field will estimate the mean soil N0 3 content within 15% about 80% of the time (Meisinger, 1984; Dahnke & Johnson, 1990). The sampling intensity needed to achieve this level of accuracy and precision is not greatly affected by field size, but the sampling area should be generally uniform with respect to past management, topography, and soil characteristics. 41-2.1.1.2 Procedure 1. Composite 20 soil cores from 10- to 40-ha areas with similar soils, drainage, and past management. Sample to a minimum depth of 60 cm in 30-cm depth increments and composite samples separately from each 30-cm layer. Follow suggestions from individual states or regions on the value of sampling soil to depths > 60 cm. If deeper samples are recommended, composite soil from these depths in 30-cm increments as indicated above. 2. Thoroughly mix soil from each depth increment in the field, discard nonsoil material, and place a minimum of 0.25 L of field moist soil into an appropriately labeled sample bag. 3. Dry the soil by spreading the sample out in a thin layer (about 1 cm thick) at room temperature with moderate air flow (e.g., household fans) for 24 h. Alternatively, the soil can be dried at low temperatures ( < 40°C) in a forced-air oven. 4. Crush the dried soil to pass a 2-mm screen and thoroughly mix the sample. 41-2.1.1.3 Laboratory Analysis. Analyze the soil for NOrN as described in section 41-2.3.1.1 or 41-2.3.1.2.

956

BUNDY & MEISINGER

41-2.1.1.4 Interpretation of Test Results. Most areas where research has been conducted to evaluate use of a residual profile N03 soil test have established the relationships between test values and crop yields. Calibration information indicating the amounts of added N needed at various test values has also been determined from crop N response experiments (Hergert, 1987; Bock & Kelley, 1992). Although profile N03 test results are usually used to make direct credits against crop N requirements (Keeney, 1982), Hergert (1987) noted that the algorithms or other calibration data used to make N recommendations from N test results may differ substantially from the experimental data. The interpretation of soil profile N03 tests to develop fertilizer recommendations for crops varies among states and regions, and local information should be consulted for specific recommendations. In general, it involves calculation of the total amount of N03-N in the profile depth sampled and direct crediting of this N against the estimated N need of the crop to be grown. 1. Calculate the profile N03-N content in kg of N03-N ha- 1 from the soil N03-N concentration and the assumption that 1 ha (30 cm of soil) weighs 4.48 x 103 Mg. If site-specific values for soil bulk density are available, these should be used in the calculation. 2. Adjust total profile N03-N content for background N03 levels or for expected differences in use efficiency of N03-N found at greater soil depths. 3. Calculate the recommended application rate from the difference between the N requirement of the crop to be grown and the adjusted soil profile N03 -N content. Adjustments for legume or manure N contributions must be made separately from those based on the soil N03 test results. 41-2.1.1.5 Comments. Residual profile N03 tests differ in principle from the PSNT discussed in the following section (41-2.1.2) in that the preplant profile tests measure N03-N remaining in the root zone at or before crop establishment, and this N03 is credited against the N needs of the subsequent crop. In contrast, the PSNT provides an index of N availability measured during the early part of the growing season, which must be related to crop needs through test calibration information. Advantages of the residual N03 tests are that they provide a direct measurement and accounting of available N in the plant root zone that does not depend on predicting the environmentally dependent release of available N from various sources. In addition, the fall or early spring sampling time used for the preplant test is frequently more convenient for users and allows more time for soil analysis and fertilizer applications than the PSNT. The associated disadvantage is that preplant soil N03 tests are not useful for assessing available N contributions from organic N sources such as soil organic matter, previous legume crops, and organic wastes. Other disadvantages of preplant soil N03 tests are the risk of N03 loss from the root zone during the interval between soil sampling and crop N uptake and the increased difficulty and expense of obtaining the soil profile samples.

NITROGEN AVAILABILITY INDICES

957

41-2.1.2 Pre-Sidedress Nitrate Test This NAI is designated as the PSNT because soil nitrate is sampled a few weeks before the normal time for sidedress N applications to corn, and to clearly distinguish it from pre plant nitrate tests. The PSNT also differs from preplant tests because only the surface 30 cm is sampled, while preplant tests sample to 60 cm or more. The literature also refers to the PSNT as the late spring nitrate test, the Vermont or Iowa test, and the June or mid-June nitrate test. Magdoff et ai. (1984) first suggested the corn PSNT test. They reported a good relation (R2 = 0.74) between corn silage yields and soil N0 3 -N concentrations in the surface 30 cm when the corn plants were 15 to 30 cm tall. A review and evaluation of the PSNT test in humid regions was recently provided by Bock and Kelley (1992). 41-2.1.2.1 Principles. The PSNT is based on a timely sampling of the spring accumulation of soil N0 3 -N under natural field conditions, just before the warm-season corn crop begins its period of rapid N uptake (Magdoff, 1991; Meisinger et aI., 1992b). The PSNT assumes that most of the corn fertilizer N will be applied as a sidedressing and that only small quantities of N ( < 50 kg of N ha- 1) were applied at planting, either as starter fertilizer or uniformly applied broadcast N. The N0 3 -N content of a typical agricultural soil represents the net balance between nitrate production processes vs. nitrate loss processes. In a humid-temperate climate, a typical silt loam surface soil will have nitrate contents that are: lowest in winter (due to leaching), rise in the spring and early summer (due to commencement of mineralization), decrease during summer (due to crop uptake), and increase again in the fall (Harmsen & Van Schreven, 1955; Stevenson, 1986). Figure 41-1 summarizes recent data from Sarrantonio and Scott (1988) that exemplifies this pattern (solid symbols) for central New York conditions. Surface soil NOTN concentration after winter leaching (or excess irrigation) would commonly be 5 to 10 mg of NOrN kg- 1 but this may increase two- to sixfold in the late spring, depending on recent additions of manure or crop residues (Magdoff et aI., 1984; Fox et aI., 1989; Meisinger et aI., 1992a). The PSNT has succeeded in humid regions because leaching is not an efficient nitrate removal process in fine- and medium-textured soils. That is, nitrate is not as mobile and subject to complete leaching losses as once thought, due to preferential water flow. Furthermore, preferential flow is enhanced by reduced tillage systems (especially no-tillage; e. g., Tyler & Thomas, 1977; Thomas et aI., 1989) that have greatly expanded in the past 10 yr. Preferential flow is characterized by rapid and deep percolation through large-pore sequences that can by-pass spring mineralized N produced in small-pore sequences. The PSNT has also succeeded because it has been applied to a warmseason crop (corn), which allows ample time for the spring mineralization period before the crop begins its period of rapid N uptake (see dashed symbols of Fig. 41-1). However, the close juxtaposition of the spring N0 3 -N maximum and the corn N uptake period can also cause logistical

BUNDY & MEISINGER

958 SOil & CORN N vs. TIME • =Soil N, Check • =Soil N, H. Vetch

(1J

s::. "Z Cl ~

t:.. =Corn N, Check o =Corn N, H. Vetch

PSNT

90

n

0- - _0--

E C)

0

60

(")

z

.=

P

/

/

/

o

/

/

/

"

--

__ 0

120

90

III

z

Cl ~

.,;L!.---l:;.---

_-t:..

CI)

60

,:,(, (1J

0.::J

30

30 Z

...

c:

~

o 0

(1J

s::. "-

o

May

Oct

o

o

TIME

Fig. 41-1. Soil mineral N contents (solid symbols) and corn N uptake (open symbols) throughout the growing season in a central New York silt loam soil previously receiving no fertilizer N (triangUlar symbols) or after a hairy vetch winter cover crop (circular symbols) (Sarrantonio & Scott, 1988).

problems. Only about 2 to 3 wk are available to measure the soil N0 3-N and to apply the appropriate side dress fertilizer N. The above principles explain why the com PSNT has been successful in humid-temperate climates, but they also contain the basis for limitations of this test. For example, the PSNT would be less useful in: (i) unusual spring weather conditions that affect mineralization, such as cool or dry springs; (ii) on highly leachable nonstructured soils, such as loamy sands; (iii) with cool-season crops that rapidly assimilate spring mineralized N, such as winter wheat; and (iv) in circumstances where soil N transformations perturb the N0 3-N pool, such as short-term denitrification events or short-term immobilization events. 41-2.1.2.2 Interpretation. The PSNT must be interpreted in relation to field crop response to determine its usefulness as a NAI. Field evaluations have been reported from 34 locations in Vermont (Magdoff et al., 1984), nine experiments over 2 yr in Iowa (Blackmer et al., 1989), 47 treatment-year combinations in Maryland (Meisinger et al., 1992a), and 272 site-years of combined data throughout the Northeast (Magdoff et al., 1990). These field studies have convincingly shown that the PSNT test can. successfully identify N-sufficient sites. There is a remarkable consensus that PSNT N0 3 -N concentrations of 20 to 25 mg of N0 3-N kg-lor more are associated with N sufficiency for com. Using the PSNT as a quantitative index of fertilizer N needs has met with varied success. For example, Pennsylvania, Iowa, and Vermont use PSNT values below the critical level as a direct input into fertilizer N

NITROGEN AVAILABILITY INDICES

959

recommendations (Beegle et al., 1989; Jokela, 1989; Blackmer et aI., 1991). Others have concluded that variability in the PSNT vs. relative yield relation limits its usefulness as a quantitative index (Fox et aI., 1989; Meisinger et aI., 1992a). We recommend that the quantitative adoption of the PSNT should be appraised by each individual state, because this issue involves a critical examination of: (i) current field calibration data, (ii) fertilizer N recommendation philosophies, and (iii) alternative fertilizer N-recommendation systems. Each of these elements has unique solutions that are specific to the soils, climate, and cropping systems of each state. 41-2.1.2.3 Method. The PSNT basically involves collecting a representative soil sample at a specific time and determining the N0 3-N concentration. Modifications of the preplant soil-nitrate sampling procedure have been made to accommodate the unique aspects of PSNT sampling. We have recommended an automated laboratory N0 3-N method suited to fast analysis of many samples because the PSNT requires rapid sample tum-around. A manual N0 3-N method is also described.

41-2.1.2.3.1 Field Sampling The PSNT requires collection of a representative soil sample from the surface 30 cm when the com is 15 to 30 cm tall. This is not a small task because soil N0 3-N variability is large, CVs routinely range between 30 and 80%, with 45% being a common value (Meisinger, 1984). The PSNT assumes that most of the N will be applied as a sidedressing with only modest N rates applied at planting ( < 50 kg of N ha- 1) as banded starter fertilizer or as uniform broadcast N application (e.g., herbicides with VAN).

41-2.1.2.3.1.1 Procedure 1. Use the same basic soil sampling protocol as section 41-2.1.1.2, except sample only the surface 30 cm. 2. Pay special attention to identify areas that received manure, sludges, or other organic amendments within the past 3 yr and sample them separately. Carefully avoid areas with preplant fertilizer N (starter bands) by sampling between rows. 3. Dry, crush, and mix soil sample as per section 41-2.1.1.2. 41-2.1.2.3.1.2 Laboratory Analysis. Analyze the soil for N0 3-N as described in section 41-2.3.1.1 or 41-2.3.1.2. 41-2.1.2.3.1.3 Comments. The field sampling problem for N0 3-N has been summarized by Meisinger (1984), who concluded that large numbers of cores would be needed to accurately estimate N0 3-N means. Common soil sampling instructions call for 10 to 20 cores from an area selected to contain similar soils, past management, etc. With a CV of 45%, this sampling intensity would likely estimate the N0 3-N mean to ± 20% on 9 out of 10 fields. Doubling the number of cores would improve the estimate to about ± 14% on 9 out of 10 fields. There is a need for research studies to design efficient sampling plans for the PSNT.

BUNDY & MEISINGER

960

Location of the samples in relation to preplant N application zones is important, due to the high N concentrations near fertilizer bands. The best way to avoid problems is to avoid the fertilizer bands, which can be accomplished with N applied as starter near the row. At this time, we do not recommend PSNT sampling if anhydrous ammonia has been applied preplant because the location of these injection zones is not known. However, Iowa researchers (Davis & Blackmer, 1990) may devise a suitable sampling scheme in the future. The time of sample collection is critical. Sampling when the corn is 15 to 30 cm tall allows for an increase in soil N0 3-N due to spring mineralization, plus time for analysis and sidedress N applications, before rapid corn N uptake causes soil N0 3-N levels to dramatically decline (see Fig. 41-1). If sampled too early or too late, the soil N0 3-N levels will probably be biased low. The short period available for PSNT sampling places a premium on soil sampling and limits the number of sites an individual can sample. Experience in the northeastern states indicates that one consultant can service about 1000 ha (about 2500 acres) using conventional manual sampling techniques. The 30-cm PSNT sample depth represents a compromise between sampling the entire root zone vs. conventional plow layer sampling. Rootzone sampling would greatly increase labor requirements. The 30-cm depth has been shown to produce less variability in critical N0 3 -N concentrations than shallow 0- to 15-cm samples (Blackmer et aI., 1989). Sampling the entire 30-cm depth is important because many soils will contain high NOrN levels in the surface layers due to recent mineralization of surface residues or surface broadcast applications of fertilizer N. The objectives of soil sample drying and processing are to stop biological activity as soon as feasible and to homogenize the sample. Microbial activity can be stopped by rapid air drying, by low-temperature oven drying, or freezing; but freezing is usually only a temporary preservation measure. After drying, it is important to mix the sample well because only 10 g of soil will be analyzed out of 2000 g collected. Saffigna (1988) reported a reduction in the within sample NOrN CV from 25 to 5% due to simple soil mixing. One should also remember that the lO-g analytical sample ultimately represents nearly 1010 g of soil if a 2.5-ha area were sampled to 30 cm. 41-2.2 Laboratory Methods Laboratory methods of assessing N availability include biological and chemical techniques (see section 41-1.1). The biological methods usually involve incubation of soil under conditions that promote N mineralization from organic sources, and measurement of the inorganic N produced. In this chapter, both short-term (ammonium production under water-logged conditions) and long-term (aerobic incubation with periodic leaching) incubation methods will be recommended. The potential use of several promising chemical methods will also be discussed.

NITROGEN AVAILABILITY INDICES

961

41-2.2.1 Biological Methods 41-2.2.1.1 Theory and Potential Application of Methods. As noted in the introduction (41-1.1), biological N availability indices are based on the assumption that the same biological processes that cause release of plantavailable N in the field are also responsible for production of inorganic N in the laboratory procedures. The results from biological N availability indices must be viewed as relative indications of soil N availability. 41-2.2.1.2 Ammonium Production During Waterlogged Incubation

41-2.2.1.2.1 Principles In this method, soil N availability is estimated from NH 4-N produced during a 7-d waterlogged incubation. The procedure was initially proposed by Waring and Bremner (1964), and this technique was the biological index recommended by Keeney (1982). While limitations of biological indices stated earlier also apply to this method, it has several advantages that make it attractive where a rapid biological index is needed to provide a relative assessment of N availability. These advantages include its simplicity and ease of adaptation to laboratory routine, a short incubation time (7 d), little or no influence of sample pretreatment on test results, elimination of concerns related to optimum water content and water loss during incubation, and minimal apparatus and reagent requirements (Keeney, 1982). Although previous reviews have cited numerous reports of satisfactory relationships between results from this method and other indices (Keeney, 1982; Stanford, 1982; Meisinger, 1984), several studies have found poor correlations between NH 4-N production under waterlogged conditions and field measurements of N availability (Fox & Piekielek, 1984; McCracken et aI., 1989; Hong et aI., 1990). Boone (1990) suggests that the apparent differences between N availability measured via anaerobic incubation and field data are not contradictions but instead reflect differences in N transformations measured by the two methods. Specifically, field measurements represent the net effect of N mineralization and N immobilization under aerobic conditions, while the waterlogged incubation likely measures N mineralization from aerobic soil organisms killed by the anaerobic test conditions.

41-2.2.1.2.2 Method 41-2.2.1.2.2.1 Special Apparatus. Automated analytical equipment for continuous flow analysis of NH4-N (see section 41-2.3.2.1) or perform NH4-N analyses using a manual method (section 41-2.3.2.2). If NH4-N analyses are to be performed by steam distillation, use the procedures described by Keeney (1982). 41-2.2.1.2.2.2 Reagents. Potassium chloride (KCI) solution, approximately 4 M: Dissolve 3 kg of KCl in 10 L of water.

962

BUNDY & MEISINGER

41-2.2.1.2.2.3 Procedure. Place 12.5 ± 0.1 mL of water in a 16 by 150-mm test tube and add 5 g of soil. Stopper the tube and place it in an incubator at 40°C. After 7 d, add 12.5 ± 0.1 mL of 4 M KCI to the tube and shake the contents for 1 h on a mechanical shaker. Filter the contents of the tube through acid-washed Whatman no. 42 filter paper. Determine the NH4-N concentration in the filtrate using the automated method described in section 41-2.3.2. Determine the amount of NH4-N in the soil before incubation by the same procedure used for the incubated samples and calculate mineralizable N from the difference between the two analyses. 41-2.2.1.2.2.4 Comments. For routine use of the method, soil samples can be measured on a volume basis rather than a weight basis (Keeney, 1982). Determination of initial NH4-N in the soil can often be omitted without greatly affecting the results, since most agricultural mineral soils contain limited amounts of exchangeable NH4-N. Initial NH4-N should be determined in any correlation or calibration studies with the method. Ammonium-N determinations on filtered extracts using the automated technique recommended may produce lower mineralizable N values than direct steam distillation of the incubated samples with MgO (Sahrawat & Ponnamperuma, 1978; Keeney, 1982).

41-2.2.1.3 Inorganic Nitrogen Production During Long-Term Aerobic Incubation 41-2.2.1.3.1 Principles Use of cumulative inorganic (NH4 + N0 3 )-N production in long-term incubations to estimate potentially mineralizable N in soils was initially proposed by Stanford and Smith (1972). They suggested that the rate of N mineralization is proportional to the amount of potentially mineralizable N in the soil, that the pattern of N mineralization follows first-order kinetics, and that an N mineralization potential value (No) can be estimated from 10g(No - Nt) = log No - ktl2.303. Stanford (1982) concluded that the No values are a definable soil characteristic that may be useful for estimating the N-supplying capacities of soils under specific environmental conditions. Work by Stanford et al. (1973) showed that the rate constant k has a 010 of 2 within the temperature range of 5 to 35°C, and Stanford and Epstein (1974) found that soil water content near field capacity was directly correlated with N mineralization. The possibility of making adjustments for temperature and soil water content stimulated research interest in estimating N mineralization in the field based on N mineralization potentials (Smith et aI., 1977). The technique involves measurement of inorganic N produced during aerobic incubation of soil or soil amended with sand or vermiculite under near-optimum conditions of temperature, moisture, and aeration for up to 30 wk. Inorganic N is usually removed by periodic leaching of soil samples incubated in a combination filtration-incubation container. The total incu-

NITROGEN AVAILABILITY INDICES

963

bation time required varies with the objective and conditions of individual experiments, but the incubation should be continued until an adequate description of the relationship between cumulative N mineralization and time of incubation is obtained (Stanford, 1982). Although the initial work of Stanford and Smith (1972) involved a 30-wk incubation, subsequent studies (Stanford et aI., 1974) indicate that adequate estimates of potentially mineralizable N can often be obtained with 8- to lO-wk incubation periods. Initially, Stanford and Smith (1972) suggested that N mineralized during the first 7 to 14 d of incubation should be disregarded because of the strong and variable influence of sample pretreatment on the amounts of N mineralization during this period. However, subsequent work indicates that the pattern of N mineralization during the first few weeks of incubation may be critical for understanding the N availability status of soils. In general, N released during the first few weeks of incubation likely reflects the readily mineralizable or active fraction of soil organic N, while that released later probably originates from a much larger but more stable soil organic matter pool. The concepts of two separate soil organic N pools, each with its own decomposition rate, are reflected in much of the recent work using long-term incubation methods to assess soil N availability and develop appropriate models to describe the N mineralization process (Molina et aI., 1983; Lindemann & Cardenas, 1984; Beauchamp et aI., 1986; Deans et aI., 1986; Bonde & Rosswall, 1987; Bonde & Lindberg, 1988; Ellert & Bettany, 1988; Boyle & Paul, 1989). In addition, several studies have used simultaneous measurements of Nand C mineralization to describe the observed patterns of N mineralization during long-term incubations (Molina et aI., 1983; Robertson et aI., 1988; Houot et al., 1989). Because of the substantial time and apparatus requirements for this method, it is usually used only when long-term N mineralization information is essential. For example, long-term incubation data provide a standardized method of assessing the potential long-term N supplying capacities of soils (Stanford, 1982). In addition to possible use for estimating N mineralization in field soils, the long-term incubation technique is useful for evaluating the effects of past management practices or experimental treatments on soil N-supplying capability. Long-term incubations are also frequently used in work to model soil N mineralization and characterize various components of the labile N pool in soils.

41-2.2.1.3.2 Method 41-2.2.1.3.2.1 Special Apparatus 1. Filter units, polystyrene, lS0-mL (Falcon 7111, Becton Dickinson Co., Lincoln Park, NJ)1, fitted with cellulose acetate filter 1 Mention of product names or companies are for the benefit of the reader and do not imply endorsement or preferential treatment by ASA, the Univ. of Wisconsin-Madison, or the USDA, to the exclusion of other similar products, which may also be satisfactory.

BUNDY & MEISINGER

964

2.

3. 4. 5. 6.

7. 8.

membranes having a 0.22 J.tm pore size and a bubble point of 373 kPa. The pressed fiberglass prefilter media supplied with this unit are also needed. Fiberglass prefilters: a. 47-mm diam., 0.5 J.tffi pore size (Micron Separations, no. GIS, Westboro, MA). b. 47-mm diam., 1.3-mm thick, with acrylic binders for additional wet strength (no. 66078, Gelman Sciences Inc., Ann Arbor, MI). Fiberglass screen, I-mm mesh size cut to fit inside the 65-mm diam. filter units (Gallagher Tent and Awning Co., Madison, WI). Vacuum pump and manifold (with stopcocks for control of vacuum to filter units) capable of applying a suction of 80 kPa to each filter unit. Vacuum regulator and manometer to control and monitor vacuum level. Drip irrigation system (Bonde & Rosswall, 1987) for addition of leaching solution to filter units. Apparatus consists of 150-mL funnels connected with Tygon tubing to the center port of the filter unit's detachable lid. The tubing is fitted with a screw clamp to control flow rate from the funnel. Filter flasks, Erlenmeyer, 250-mL, heavy walls, with sidearms. Analytical equipment for determination of inorganic N in leachates (see section 41-2.3).

41-2.2.1.3.2.2 Reagents 1. Calcium chloride (CaCI2) solution, approximately 0.01 M. Dissolve 15 g of calcium chloride dihydrate (CaCI2·2H20) in 10 L of water. 2. Nutrient solution, minus N (Stanford & Smith, 1972), approximately 0.002 M CaS04·2H20; 0.002 M MgS04; 0.005 M Ca(H2P04h·H20; and 0.0025 M K2S04. Dissolve 3.4 g of CaS04·2H20, 2.4 g of MgS04, 12.6 g of Ca(H2P04h·H20, and 4.4 g of K2S04 in 10 L of water. 3. Silica sand, acid washed, 0.42 to 0.59 mm particle diameters. 41-2.2.1.3.2.3 Procedure. Assemble the filter units by removing the threaded funnel from the filter unit base and positioning the unit's cellulose acetate membrane and O-ring gasket in the base of the filter unit. Place the Micron Separations GIS fiberglass prefilter on top of the membrane and O-ring gasket, and replace the funnel assembly. This prefilter prevents soil particles from clogging the membrane pores and avoids microbial growth on the membrane surface (Beauchamp et al., 1986). Place the pressed fiberglass filter provided with the filter unit on. top of the Micron Separations prefilter to protect the prefilter and membrane from clogging. Add deionized water to the filter unit to thoroughly moisten the prefilters and membrane before adding the soil. Uniformly mix field-moist samples of the soil to be evaluated (30 g oven-dried weight) with 30 g of acid-washed silica sand and transfer the

NITROGEN AVAILABILITY INDICES

965

mixture to the filter unit. Determine the water content in a subsample of the field-moist soil by oven drying (105°C) before preparing the soil-sand mixture. Place two layers of the 1-mm mesh fiberglass screen on top of the soil-sand mixture, and place the Gelman no. 66078 fiberglass prefilter on the fiberglass screen. The screen and prefilter are permanently positioned in the filter unit to minimize soil dispersion during leaching. Attach the lid of the filter unit to the funnel, place the cotton-plugged fitting provided with the unit on the off-center port in the unit lid to allow air exchange during incubation. Seal the vacuum port in the filter unit base with the rubber cap provided. Place the filter unit containing the soil-sand sample on a 250-mL Erlenmeyer filtering flask with its sidearm connected to a vacuum manifold adjusted to deliver a suction of 80 kPa. Attach the tube from the drip irrigation system to the center port in the filter unit lid. Add 100 mL of 0.01 M CaCl2 solution to the irrigation system funnel, adjust the screw clamp on the inlet tube to a flow rate of approximately 150 mL h -1, and apply suction to the filter unit. After the 0.01 M CaCl2 solution has been added to the sample in the filter unit, place 25 mL of the minus-N nutrient solution into the irrigation system funnel and continue to collect the leachate. When leaching is complete (approximately 1 h), continue suction on the leaching unit for 0.5 h to provide a uniform moisture content in the soil-sand mixture for the subsequent incubation period. Disconnect the filter unit from the vacuum and irrigation systems, seal the center port in the unit lid with parafilm, and place the filter units in an incubator maintained at 35°C and containing an open pan of water for humidity control. Measure the volume of leachate ( ± 1 mL) collected from each sample to allow calculation of N mineralized on a soil basis. Analyze an aliquot of the leachate for inorganic N as described in sections 41-2.3. The procedure described in the preceding paragraph is performed immediately prior to the initial incubation and is repeated after each incubation interval. The frequency of leaching and the total duration of the incubation period depend on the objectives of the individual investigation. General information on the rates and amounts of soil N mineralization can often be obtained with four to six leachings over an 8- to 12-wk incubation period. Detailed studies may require 12 to 14 leachings and incubation times up to 40 wk.

41-2.2.1.3.2.4 Comments. The procedure described follows the basic concepts of long-term incubation and periodic leaching proposed by Stanford and Smith (1972), and employs improvements developed in more recent long-term N mineralization studies. Filter units similar to those described above have been employed in numerous long-term incubation studies (MacKay & Carefoot, 1981; Bonde & Rosswall, 1987; Bonde et aI., 1988; Marion & Black, 1988; Robertson et aI., 1988; Nadelhoffer, 1990). These membrane filter units provide the advantages of standardized conditions and more uniform control of soil moisture content during incubation. A filter membrane with a bubble point exceeding the suction applied

BUNDY & MEISINGER

for leaching provides accurate control of soil water content during incubation (MacKay & Carefoot, 1981). Increased variability in N mineralization measurements has been attributed to less precise control of soil moisture where filter membranes were not used in the incubation devices (Nadelhoffer, 1990). Although MacKay and Carefoot (1981) initially suggested overnight equilibration at an appropriate suction to achieve reproducible N mineralization values (CV = 3.9%), much shorter equilibration periods appear to be adequate. Using the O.S-h equilibration suggested above in the author's laboratory, average CV values of < S.O% are routinely obtained for 4O-wk cumulative N mineralization values in experiments involving 20 treatments and four replications. These results suggest that extensive laboratory replication of incubated treatments may not be required for many studies. An alternative technique for controlling soil moisture contents in N mineralization studies by adjusting soil-to-sand mixing ratios has been described by Lueking and Schepers (1986). Simultaneous measurements of CO 2 evolution are often of interest in long-term incubation N mineralization studies to obtain C mineralization information (Paustian & Schnurer, 1987; Gale & Gilmour, 1988; Robertson et al., 1988; Boyle & Paul, 1989; Houot et al., 1989; Nadelhoffer, 1990). The units described (Falcon 7111) can be accommodated inside a 1.9-L wide-mouth terephthalate jar (No. YB-06043-7S, Cole-Parmer Instrument Co., Chicago, IL) along with a SO-mL Erlenmeyer flask containing 20 mL of O.S M NaOH to absorb CO2 and a 70-mL vial of water to minimize soil water loss during the incubation. When the incubated soil samples are removed from the jar for periodic leaching, the alkali traps are replaced, and the sorbed CO2 is determined using an automated colorimetric method adapted from Chaussod et al. (1986). Enclosing the filter units inside jars during incubation also provides very precise control of sample water content relative to units incubated without enclosure. In the author's laboratory, enclosed samples lost only about 2 to 3 mL of water during 4 wk of incubation based on the average volume of leachate retained at the next leaching. Use of field-moist soil samples is recommended to avoid the enhanced N mineralization frequently observed during the first week of incubation with air-dried soils (Stanford & Smith, 1972; Beauchamp et al., 1986; Cabrera & Kissel, 1988c). Beauchamp et al. (1986) reported that, due to the initial flush of mineralized N during the first 7 d of incubation, the total amount of N mineralized in 42 d from air-dried soil was greater than from frozen or field-moist soils. The flush of N released when air-dried soils are incubated may originate from soil microbes killed by the air-drying pretreatment (Richter et al., 1982). Beauchamp et al. (1986) concluded that this N release is an experimental artifact and should not be considered part of the true soil mineralization potential. Use of air-dried samples may produce satisfactory results where a relative comparison of soil N availability is desired. A concern with incubation procedures involving periodic leaching to remove mineralized N is that some soluble organic N may be removed

NITROGEN AVAILABILITY INDICES

967

during the leaching process (Smith et aI., 1980; Beauchamp et aI., 1986; Smith, 1987). The amounts of organic N found in leachates has varied from 37 to 70% of the total N leached (Smith et aI., 1980) to to to 20% of the total N leached (Beauchamp et aI., 1986). Beauchamp et ai. (1986) found no differences in soluble organic N among soils or between air-dried or field moist sample pretreatments. Working with air-dried soils, Smith (1987) found that most of the soluble organic N was removed in the initial leaching, and that organic N represented 5% or less of the inorganic N produced during an 84-d incubation. Smith (1987) also found that the soluble organic N in leachates was not highly susceptible to mineralization. Although several authors have noted the need to include soluble organic N in estimates of total N mineralization, it appears that adequate estimates of soil N availability can be obtained from inorganic N production alone (Smith, 1987). 41-2.2.2 Chemical Extraction Methods In view of the many chemical methods available and the low correlation of many of these techniques with field-measured N availability, no single chemical method is recommended here. The selection and use of these techniques should be determined from the objectives of the work in which the methods are used. If the goal is to obtain a rapid, relative indication of N availability among soils differing in past management, several recent techniques including digestion with 2 M KCI at tOO °C (Gianello & Bremner, 1986a), steam distillation with pH 11.2 phosphate-borate buffer (Gianello & Bremner, 1988), and UV absorbance at 200 nm of NaHC0 3 soil extracts (Hong et aI., 1990), are likely to produce satisfactory results. These methods have advantages over previously proposed chemical indices in that they are convenient, rapid, and well correlated with N availability determined by incubation methods (Gianello & Bremner, 1986b) or field measurements (Hong et aI., 1990). If the objective is to predict soil N availability for determination of N recommendations in crop production, methods other than chemical indices should be considered because chemical indices are usually not well correlated with field-measured N availability (Hong et aI., 1990). A possible exception is the procedure evaluated by Hong et ai. (1990) involving determination of UV absorbance at 200 nm of NaHC0 3 soil extracts. This measurement reflects both N0 3 (Norman et aI., 1985) and organic matter (Fox & Piekielek, 1978) in the soil extract, but the method has not been widely evaluated. In addition, Fox and Piekielek (1984) found that N mineralized in laboratory incubation methods was not well correlated with field measurements of soil N availability. This suggests that chemical indices well correlated with aerobic or anaerobic incubation methods may not provide good estimates of N availability in the field. Hong et ai. (1990) showed that the best relationships between field-measured N availability and the indices evaluated in their work occurred with either direct or indirect measurements of soil N0 3 at corn planting or at the pre-side dress

BUNDY & MEISINGER

968

growth stage. This finding and the wide-spread interest in development and use of soil N0 3 tests to predict crop N requirements (Magdoff et aI., 1984; Hergert, 1987; Blackmer et aI., 1989; Fox et aI., 1989; Dahnke & Johnson, 1990; Magdoff et aI., 1990; Magdoff, 1991) suggests that appropriately timed soil N0 3 measurements will likely provide the best assessment of field N availability. 41-2.3 Methods for Inorganic Nitrogen 41-2.3.1 Nitrate Method Nitrate is extracted from a representative soil sample by shaking 10 g of soil with 100 mL of 2 M KCI for 1 h, followed by filtration. The basic analytical steps are: N0 3 reduction to N0 2 by passage through a copperized-cadmium column, and determination of N0 2 with a modified GriessIllosvay procedure by diazotizing with sulfanilamide and coupling with N-(l-naphthyl)-ethylenediamine to form a purple-colored dye, which is measured by absorbance at 520 nm. The method is described in detail by Bremner (1965b) and Keeney and Nelson (1982); it has excellent sensitivity and few interferences with filtered extracts. The general working range is 0.01 to 5 mg of N0 3-N L -1 of soil extract (Henriksen & Selmer-Olsen, 1970; Keeney & Nelson, 1982) and a multirange manifold has been described to extend the range to 100 mg of N0 3-N L -1 (Jackson et aI., 1975). Both an automated and a manual method of this procedure are described below. Alternative N0 3 -N analysis methods may also be acceptable and are briefly discussed in the comments section below. 41-2.3.1.1 Laboratory Analysis, Automated. The following automated Cu-Cd reduction procedure has been adapted for soil analysis from EPA method no. 353.2 (USEPA, 1983) and the American Public Health Association Standard Method no. 4500-N03-F (APHA, 1989), which were designed for N0 3- -N analysis of water and wastewater using a Technicon auto analyzer system (Technicon Instrument Corp., 1977b). The procedure has been successfully applied to soil extracts as reported by Henriksen and Selmer-Olsen (1970), Ananth and Moraghan (1987), Gelderman and Fixen (1988), and Tel and Heseltine (1990). 41-2.3.1.1.1 Special Apparatus

Automated analytical equipment for continuous flow instrumentation consisting of: a sampler, a manifold, and N0 3 -N analytical cartridge, a proportioning pump, a flow cell colorimeter capable of measuring absorbance at 520 nm (although 510-540 nm may be used), and a recorder or other data output storage device. The detailed configuration of the apparatus depends on manufacturer's specifications for specific instruments. Consult the manufacturer's operating instructions for specifications on component operation, flow rates, tube sizes, and analysis rates.

NITROGEN AVAILABILITY INDICES

969

41-2.3.1.1.2 Reagents

1. Potassium chloride, 2 M: Dissolve 1490 g of KCI in 8 L of distilled water and dilute to 10 L. 2. Copper sulfate solution, 0.08 M: Dissolve 20 g of CuS04·5H20 in water and dilute to 1 L. 3. Dilute HCI, 6 N: Dilute 50 mL of conc. HCI to 100 mL. 4. Granulated Cd: 40 to 60 mesh. 5. Ammonium chloride-EDTA solution: Dissolve 85 g of NH4Cl and 0.1 g of disodium ethylenediamine tetraacetate in distilled water; adjust pH to 8.5 with conc. NH40H; dilute to 1 L, and add 0.5 mL of the wetting agent polyoxyethylene 23 lauryl ether (Brij-35). Store in refrigerator to minimize bacterial growth. 6. Color reagent: To approximately 800 mL of distilled water, add, while stirring, 100 mL conc. phosphoric acid, 40 g of sulfanilamide, and 2 g of N-1-naphthylethylenediamine dihydrochloride. Stir until dissolved and dilute to 1 L. Store in brown bottle in refrigerator when not in use. This solution is stable for several months. 7. Stock nitrate solution: Dissolve 7.218 g of pure dry KN0 3 in distilled water and dilute to 1 L. This solution contains 100 mg of N0 3- N L -1. Store in refrigerator. 8. Working nitrate solution: Using the stock nitrate standard solution (reagent no. 7 above), prepare working nitrate standards by pipetting 0, 1, 5, 10, 20, 30, and 50 mL into 1-L volumetric flasks and diluting to volume with soil extracting solution (2 M KCI, reagent no. 1 above). These working standards contain 0,0.1,0.5, 1,2,3, and 5 mg of NOrN mL -1. Store in refrigerator. 9. Stock nitrite solution: Dissolve 0.493 g of pure dry NaN0 2 in distilled water and dilute to 100 mL. This solution contains 100 mg of N0 2-N mL -1. Store in refrigerator at 4°C. 10. Using stock nitrite solution (reagent no. 10 above), dilute 1.0 mL to 500 mL with 2 M KCI (reagent no. 1 above). This solution contains 2.0 mg of N0 2-N mL -1. This solution is not stableprepare as needed. 41-2.3.1.1.3 Procedure

1. Prepare Cd column: Place about 10 g of granulated Cd in a beaker and wash with about 50 mL 6 N HCI for 1 min, followed by a thorough rinse with distilled water (Cd should be silver). Add about 100 mL of 0.08 M CUS04 solution and swirl for about 5 min or until blue color fades, decant solution and repeat with fresh CUS04 until a brown colloidal precipitate of Cu forms. Gently rinse with distilled water to remove all precipitated Cu, repeat distilled water rinse about 10 times until all blue and light-gray color disappear from the wash (Cd should be black).

BUNDY & MEISINGER

970

2. Set up continuous flow analysis system for (N0 3 + N0 2 )-N as per manufacturer's instructions. 3. Extract soil by placing 10 g of dry soil in a suitable 250-mL vessel, add 100 mL of 2 M KCI, stopper, and shake vigorously. Continue shaking for 1 h on a mechanical shaker or with frequent manual shaking, filter the soil-KCl suspension through a Whatman no. 42 filter to obtain a clear extract, discard the first portion of the filtrate but collect the remainder for analysis. 4. If the soil extract pH is below 5 or above 9, adjust the pH to between 5 and 9 with either HCI or KOH. 5. Allow continuous flow analysis system to warm up according to manufacturer's instructions, allow all reagents and soil extracts to reach room temperature, obtain a stable baseline output with all reagents, supplying 2 M KCl extracting solution in place of unknowns. A new reduction column may have to be conditioned with a nitrate working standard prior to use; see manufacturer's instructions. 6. Place appropriate N0 3-N working standards in sampler (use at least 5), which will bracket the expected N0 3-N concentration in the soil extracts. Place unknown soil extracts in the sampler with periodic insertion of a mid-range N0 3-N standard for quality control. 7. Switch sample line to sampler and start analysis with due attention to output device gain to be sure all working standards are within range. 8. Calculate results by preparing an appropriate standard curve derived from plotting peak heights (or other output signal) vs. N0 3 -N concentration. Compute the final soil N0 3 -N concentrations (mg NOrN kg- 1 soil) by converting soil extract peak heights to N0 3 -N concentrations via the standard curve and multiplying by 10 (for the 10:1 dilution from soil extraction). For more precise work, soil hygroscopic moisture may be taken into account (usually 1-4%, depending on clay content) by drying a separate soil sample at 105°C. 41-2.3.1.1.4 Comments The precision of this method has been reported to be ± 0.06 mg of N0 3-N L -1 (Henriksen & Selmer-Olsen, 1970), with CVs being 1 to 2% for typical soil extract N0 3-N concentrations (Skjemstad & Reeve, 1978; APHA, 1989). This method determines (N03 + N0 2 )-N in the soil extract. Separation of N0 3 from NOz can be accomplished by conducting the same analysis without the Cu-Cd column, which will quantify the N0 2-N directly. However, soils generally contain insignificant levels of N02 , so the determination and interpretation of (N0 3 + N02 )-N data are equivalent to N0 3-N for most applications.

NITROGEN AVAILABILITY INDICES

971

The efficiency of the Cu-Cd column can be determined by passing the 2 mg of N0 3 -N L -1 standard through the column and comparing it to the 2 mg of N0 2-N L -1 standard, which was not passed through the column. If < 100% efficiencies are detected, the column is still likely to be usable because the quantity of N0 2-N produced will still be proportional to the N0 3-N content. After several hundred samples, the column efficiency will reach a point where Cd metal should be re-copperized; however, this procedure is only effective a few times before a new Cd must be used. The potential problem of N0 2-N reduction within the column can be evaluated by passing the N0 2-N standard through the column and comparing it to the original N0 2-N standard. Nitrite reduction can be controlled by increasing the flow rate. The NH4Cl reagent has EDTA included to prevent interferences from high concentrations of Fe, Zn, and other metals. Most soil extracts, however, will not contain high enough concentrations of these metals to cause interference. Indeed, Henriksen and Selmer-Olsen (1970) found no interference from 1% solutions of several heavy metal salts. The EDTA has been included as a precaution to ensure quantitative N0 3-N analysis even in unusual situations (e.g., soils treated with sludges containing heavy metals), but several investigators have reported it to be unnecessary for routine soil analysis (Henriksen & Selmer-Olsen, 1970; Dorich & Nelson, 1984). The 2 M KCI extract is recommended because it is the most common soil extractant for inorganic N and has proven satisfactory for a wide range of soils. Several other extracts have also been successfully employed such as: 1 M KCI (Bremner, 1965b), 0.5 M K2S04 (Magdoff et aI., 1984),0.025 M AI2(S04h (Roth & Fox, 1990), and various Ca salts (Bremner, 1965b; Sims & Jackson, 1971). If a different extractant is selected, it should be evaluated with standard N0 3 -N additions to representative soils to ensure quantitative extraction and freedom from interferences. Prepare the working standards with the same solution used for soil extraction. The method described is suitable for N0 3 -N analysis in the 0.01 M CaCl2 leachates obtained from aerobic incubation experiments (41-2.2.1.3). The 1-h shaking time recommended can be reduced to 15 min if only N0 3 is of interest. However, 1-h shaking time must be used if exchangeable NH4-N is to be determined. Where simultaneous extraction of large numbers of samples is desired, 2.5 g of soil and 25 mL of 2 M KCI can be shaken for 1 h in stoppered 25 by 200 mm test tubes. If the tubes are stored vertically in a refrigerator (5°C) overnight, a sample of the extract can be decanted for analysis without filtration for most soils. This modification has performed satisfactorily in the authors' laboratory for both N0 3-N and NH4- N analyses. Turbid, colloidal, or highly colored extracts can lead to an accumulation of suspended material on the reduction column. This potential problem is minimized by the 10: 1 dilution of the extraction step and filtration. Filtration should use a low nitrate filtering media. Two recent reports (Sparrow & Masiak, 1987; Scharf & Alley, 1988) have observed significant

BUNDY & MEISINGER

972

NOrN contamination from some filtering media, which can be controlled by using quantitative grade filter paper (Whatman no. 42 or equivalent) or glass fiber filters, by pre-washing the filters, or by using centrifugation instead of filtration. The installation of a dialyzer can also control problems from colloidal material (Henriksen & Selmer-Olsen, 1970), although this reduces sensitivity. Smith and Scott (1991) have also reviewed the background and general use of continuous-flow and discrete analysis systems in soil science. Several other N0 3 -N methods suited to automation have been applied to soil extracts, such as those using hydrazine to reduce N0 3 -N to N0 2-N, followed by the Griess-Illosvay reaction (Best, 1976; Markus et aI., 1985). These methods can also provide satisfactory soil N0 3 -N analyses, but take precautions to minimize interferences; for example, Mg can interfere with hydrazine reduction (Ananth & Moraghan, 1987). For any N0 3-N procedure, a careful quality control program should be followed that includes: (i) periodic comparative N0 3-N analyses by independent procedures (instrumental vs. "wet chemistry" procedures), (ii) periodic N0 3 -N recovery checks from known NOrN additions to soils, and (iii) regular "blind analysis" of standard soils containing known levels of N0 3 -N. 41-2.3.1.2 Laboratory Analysis, Manual. For a simplified N0 3 -N analysis, we recommend the manual Cu-Cd reduction method described in detail by Keeney and Nelson (1982) and further investigated by Dorich and Nelson (1984). The manual method is designed for use with small numbers of samples, with traditional laboratory equipment, and with uncomplicated reagents. The method is reliable, sensitive, and has few interferences. 41-2.3.1.2.1 Special Apparatus 1. Spectrometer: With 1-cm light path capable of measuring absorbance at 540 nm, although 510 to 540 nm could be used. 2. Reducing columns: 1 by 30 cm Pyrex tubes with fritted glass plates and stopcock at lower end and an upper reservoir capable of holding 75 mL. Connect a glass tube (4 mm i.d.) implanted in a twohole no. 00 stopper to the column stopcock. A 4-mm (i.d.) glass tube in the other hole of the stopper is connected to a flow regulator and vacuum source (see Keeney and Nelson, 1982, for details). 41-2.3.1.2.2 Reagents 1. Reagents 1 through 3 are the same as section 41-2.3.1.1.2. 4. Cadmium metal: Approximately 1-mm diam. granules. 5. Ammonium chloride, 4 M: Dissolve 202 g of NH4 CI in 800 mL deionized water and dilute to 1 L. 6. Ammonium chloride, 0.1 M: Dilute 50 mL of concentrated NH4 CI solution (reagent no. 5 above) to 2 L with deionized water.

NITROGEN AVAILABILITY INDICES

973

7. Diazotizing reagent: Dissolve 0.5 g of sulfanilamide in 100 mL of 2.4 M HCI. Store in refrigerator at 4 °C. 8. Coupling reagent: Dissolve 0.3 g of N-(l-naphthyl)-ethylenediamine dihydrochloride in 100 mL of 0.12 M HCI. Store this solution in a brown bottle in a refrigerator at 4 0c. 9. Standard N03-N solutions and standard N0 2-N solution are the same as reagents no. 7 through 10 of section 41-2.3.1.1.2. 41-2.3.1.2.3 Procedure 1. Prepare Cu-Cd as described in procedure 1 of section 41-2.3.1.1.3, treating enough Cd for several columns. 2. Fill Pyrex reducing column with the dilute NH4C1 solution and pour in the Cu-Cd particles to a depth of 20 cm. Ensure that all air bubbles are removed from the column, drain off excess NH4C1, and wash thoroughly with dilute NH4 CI solution (about 10 pore volurnes), using a slow flow rate of about 8 mL min -1. Always ensure that the Cu-Cd metal is covered with at least 1 cm of dilute NH4C1 solution. 3. Add 1 mL of 4 M NH4 CI solution to top reservoir and lower liquid to top of column. Add 75 mL of 0.1 M NH4 CI solution to reservoir, attach a 100-mL volumetric flask to the stopper on the outlet, and pass the NH4 CI through the column at a flow rate of 110 mL min- 1 by manipulating the stopcock and vacuum flow regulator. 4. Drain off excess NH4 CI from column until solution is just above the Cu-Cd metal. Add 1 mL of 4 M NH4 CI solution to top of column, then pipette an aliquot of 2 M KCI soil extract containing < 20 lAg of N0 3-N (usually 2-5 mL) onto the top of the column. Attach a 100-mL volumetric flask to the stopper, allow the extract to enter the Cu-Cd column by draining some NH4 C1 from the bottom, rinse the inside of the column with 2 mL of 0.1 M NH4CI, then add 75 mL of 0.1 M NH4 CI to the top reservoir and begin passage through the column at a flow rate of 110 mL min-I. 5. Collect the entire eluant in the 100-mL volumetric flask, containing 2 mL of diazotizing reagent, and mix. After 5 min, add 2 mL of coupling reagent, mix, bring to volume with 0.1 M NH4 C1, and let stand for 20 min. 6. Measure absorbance at 540 nm with instrument zeroed against a reagent blank. 7. Prepare a N0 3-N standard curve by passing 4 mL of each NOTN working standard (reagent no. 8 of section 41-2.3.l.l.2) through the column and developing the color as described above. A 4-mL aliquot of each N0 3-N working standard should contain 0,0.4,2, 4, 8, 12, and 20 lAg of N0 3-N and should produce a linear plot of absorbance vs. N0 3-N. 8. Calculate the soil N0 3-N concentration (mg N0 3-N kg- 1 of soil) by converting the soil extract absorbance reading to N0 3-N via the

BUNDY & MEISINGER

974

standard curve and making appropriate adjustments for the volume of the soil extract analyzed and the soil:2 M KCI extraction ratio used (usually a 1:10 ratio). 41-2.3.1.2.4 Comments

The comments for the automated Cu-Cd reduction method (section 41-2.3.1.1.4) should be reviewed since most are applicable to the manual method as well. The reagents for the manual method are less complex than the automated method and are in keeping with the objective of recommending a simple N0 3-N method. The elimination of the EDTA and pH adjustment of the NH4 CI solution should not cause metal interference problems for most soil extracts (Henriksen & Selmer-Olsen, 1970; Dorich & Nelson, 1984). However, if heavy metals are a concern, then a buffered (pH 8.5) NH4 CI solution containing EDTA is recommended as prescribed by Gales and Booth (1975). Careful control of the eluant flow rate is important because slow rates can cause further reduction of N02-N and low recoveries, while high flow rates can produce poor conversion of N0 3-N to N02- N, Dorich and Nelson (1984) studied effluent flow rates and column lengths and concluded that a 110 mL min- 1 flow rate with a 20-cm column gave rapid, precise, and accurate results with 2 M KCI soil extracts. The size of the Cd granules can vary from 0.25 to 1 mm, but if granules < 1 mm are used, the column flow rates may have to be increased to avoid further reduction of N0 2-N within the column. Particular care should be given to the preparation and use of the Cu-Cd columns. Difficulties are most likely to arise from unsatisfactory copperization of the Cd, from incomplete washing off of excess Cu from the treated Cd, or from air entering the column. If the Cu-Cd metal is inadvertently exposed to air, the column should be repacked (procedure 2 above) to completely exclude air bubbles. Further readings on the Cd reduction method for N0 3-N and the Griess-Illosvay N0 2-N reactions can be found in Bremner (1965b) and Keeney and Nelson (1982). Other manual N0 3-N methods for small numbers of samples include steam distillation (Bremner, 1965b), micro-diffusion (Bremner, 1965b), nitrosation of salicylic acid (Cataldo et aI., 1975; Vendrell & Zupancic, 1990), nitrate specific electrode (Keeney & Nelson, 1982; Gelderman & Fixen, 1988), chromotropic acid (Sims & Jackson, 1971), and direct UV absorption (Cawse, 1967). The steam distillation method has been discussed in detail by Bremner (1965b) and Keeney and Nelson (1982) and is usually the standard to which other methods are compared. Steam distillation is an excellent method and we recommend it where its requirements for special apparatus and relatively long analysis times are not serious problems.

NITROGEN AVAILABILITY INDICES

975

41-2.3.2 Ammonium Method 41-2.3.2.1 Laboratory Analysis, Automated. Ammonium in soils or soil extracts is determined by reacting NH4 with phenol and hypochlorite in alkaline solution to form an intense blue color that is measured by absorbance at 630 nm. The colorimetric procedure is widely known as the Berthelot reaction or the indophenol blue method. It has been used to determine NH4-N in a wide range of applications. Agricultural uses of the technique include determination of NH4-N in Kjeldahl digests (Mann, 1963) and determination of NH4 in soils and soil extracts using manual (Kempers, 1974; Keeney & Nelson, 1982; Dorich & Nelson, 1983) and automated techniques (Rice et ai., 1984; Markus et ai., 1985; Gentry & Willis, 1988; Tel and Heseltine, 1990). The method has excellent sensitivity and precision and is suited to determination of NH4 in most solutions used to extract inorganic N from soils. Potential interferences by di- and trivalent cations can be minimized by addition of EDTA (Dorich & Nelson, 1983) or potassium sodium tartrate (Technicon Instrument Corp., 1977a) during the analysis. The working concentration range for NH4-N in sample solutions is 0.2 to 10 mg of N L -1. This sample concentration range can be expanded at least lO-fold through use of a sample dilution loop or automatic dilution during the continuous flow analysis. The procedure described here was developed for analysis of water and wastewater using a Technicon auto analyzer (Technicon Instruments Corp., 1977a; Kopp & McKee, 1978). This procedure has performed well for analysis of 2 M KCI soil extracts and leachates from long-term soil incubation experiments in the author's laboratory. This method has been adapted for automated NH4 analyses by other workers (Rice et ai., 1984; Markus et ai., 1985; Gentry & Willis, 1988). 41-2.3.2.1.1 Special Apparatus

Automated analytical equipment for continuous flow analysis of NH4-N consisting of: a sampler, a manifold and NH4-N analytical cartridge, a proportioning pump, a flow cell colorimeter equipped to measure absorbance at 630 nm, and a recorder or other output storage device. The detailed configuration of the apparatus depends on manufacturer's specifications for specific instruments. Consult the manufacturer's operating instructions for specifications on component operation, flow rates, tube sizes, and analysis rates. 41-2.3.2.1.2 Reagents 1. Potassium chloride (KCl), 2 M: Dissolve 1490 g of KCI in 8 L of distilled water and dilute to 10 L.

BUNDY & MEISINGER

976

2. Alkaline phenol solution: Dissolve 200 g of NaOH in about 500 mL of distilled water. Cool the solution during preparation by placing the vessel in circulating cold water. In a fume hood, slowly add 243 g of phenol to the NaOH solution with continuous stirring and continue cooling the vessel with cold water. Add 0.5 mL of Brij-35 wetting agent and dilute the cooled solution to 1 L with distilled water. Store in a refrigerator when not in use. 3. Sodium hypochlorite solution: Commercially available household bleach containing at least 5.25% (wtlvol) available CI is satisfactory. 4. Potassium sodium tartarate solution: Dissolve 150 g of KNaC4H 40 6 ·4HzO in about 800 mL of distilled water, add 0.5 mL of Brij-35, and dilute to 1 L. Store in a refrigerator. 5. Stock ammonium standard solution: Dissolve 0.3821 g of dry ammonium chloride (NH4CI) in a l-L volumetric flask, and dilute to volume with distilled water. This solution contains 100 mg of NH4-N L -1. Store in a refrigerator. 6. Working ammonium standard solutions: Using the stock ammonium standard solution (no. 5 above), prepare working standard solutions by pipetting 0, 2, 4, 6, 8, and 10 mL into l00-mL volumetric flasks and diluting the solutions to volume with the soil extracting solution (2 M KCI, reagent no. 1 above, or other extractant used for the samples to be analyzed). These standard solutions contain 0, 2, 4, 6, 8, and 10 mg of NH4-N L -1. Store in a refrigerator. 41-2.3.2.1.3 Procedure

1. Set up automated continuous flow analysis system for NH4-N according to manufacturer's instructions. 2. Prepare soil extracts according to procedures described for N03-N analysis (section 41-2.3.1.1.3). If exchangeable NH4-N is to be determined, the extraction must be performed by shaking soil with 2 M KCI (1:10 soil: solution ratio) for 1 h. 3. Allow the automated analysis system to warm up according to manufacturer's instructions, and allow refrigerated reagents and sample solutions to reach room temperature before use. Obtain a stable baseline output from the analysis system using all reagents (described previously) and extracting solution in place of soil extract samples. 4. Analyze working standard solutions containing 0 to 10 mg of NH4-N L -1 (reagent no. 6, section 41-2.3.2.1.2), and adjust gain to obtain appropriate output readings. The output response should be linear within the range of NH4-N concentrations used. 5. Place soil extracts to be analyzed into the instrument sampler, include a mid-range NH4-N working standard after each 8 to 10 samples for quality control, and start the analysis. Some micropro-

NITROGEN AVAILABILITY INDICES

1)77

cessor-controlled analytical systems provide for automatic standard analysis and recalibration if required. 6. Determine the sample NH4-N concentration from a comparison of sample output peak heights with the standard curve obtained from working standard peak heights and NH4-N concentrations. Calculate the soil NH4-N concentration from the soil extract concentration and the dilution factor used in the soil extraction procedure (10:1 in the 2 M KCI extraction described above). Microprocessorcontrolled instruments usually perform these calculations using system software. 41-2.3.2.1.4 Comments The general use of continuous flow analysis systems for inorganic N determinations in soils has been reviewed by Smith and Scott (1991). The precision and accuracy of the NH4 method described compares favorably with steam distillation and other manual methods (Rice et aI., 1984; Markus et aI., 1985; Adamsen et aI., 1985). Typical CV values for automated methods range from 2 to 5% (Adamsen et aI., 1985). Use of salicylate as a substitute for phenol in the procedure has been proposed for both manual (Nelson, 1983; Kempers & Zweers, 1986) and automated (Adamsen et aI., 1985; Gentry & Willis, 1988) versions of the indophenol ammonium method. Procedures using salicylate have equal or improved sensitivity relative to procedures using phenol and avoid the volatility and safety concerns associated with the phenol reagent. Improved sensitivity of the procedure through use of nitroprusside [Na2Fe(CN)sNO·5H20] to increase the intensity of color development was first proposed by Lubochinsky and Zalta (1954) and has been incorporated into many current procedures for NH4 using the indophenol procedure (Dorich & Nelson, 1983; Adamsen et al., 1985; Kempers & Zweers, 1986; Gentry & Willis, 1988; Tel & Heseltine, 1990). A 10-fold increase in sensitivity of the method is usually provided by addition of nitroprusside (Keeney & Nelson, 1982). 41-2.3.2.2 Laboratory Analysis, Manual. Where NH4 analyses are needed for a relatively small number of samples or where automated analytical equipment is not available, the steam distillation method described by Bremner (1965b) and Keeney and Nelson (1982) and the indophenol blue colorimetric procedure recommended by Keeney and Nelson (1982) provide excellent alternatives to the automated techniques described above. The manual method described here is virtually identical to the Keeney and Nelson (1982) colorimetric procedure.

41-2.3.2.2.1 Special Apparatus 1. Variable wavelength spectrometer, equipped with a l-cm light path and capable of absorbance measurements at 636 nm.

BUNDY & MEISINGER

978

41-2.3.2.2.2 Reagents

1. Potassium chloride (KCI) , 2 M: Prepare as described in section 41-2.3.2.1.2. 2. Stock ammonium standard solution: Prepare as described in section 41-2.3.2.1.2. Immediately before use, dilute 4 mL of the stock NH4 solution to 200 mL. The resulting working solution contains 2 mg of NH4-N L -1. 3. Phenol-nitroprusside reagent: Dissolve 7 g of phenol and 34 mg of sodium nitroprusside [disodium pentacyanonitrosylferrate, Na2Fe(CNhNO·2H20] in 80 mL of distilled water, and dilute to 100 mL. Mix well, and store in a dark-colored bottle in a refrigerator. 4. Buffered hypochlorite reagent: Dissolve 1.480 g of sodium hydroxide (NaOH) in 70 mL of distilled water, add 4.98 g of sodium monohydrogen phosphate (Na2HP04) and 20 mL of sodium hypochlorite (NaOCI) solution (5-5.25% NaOCI). Use less or more hypochlorite solution if the NaOCI concentration is higher or lower than that indicated. Check the pH to ensure a value between 11.4 and 12.2. Add a small amount of additional NaOH if required to raise the pH. Dilute to a final volume of 100 mL. 5. Ethylenediaminetetraacetic acid (EDTA) reagent: Dissolve 6 g of ethylenediaminetetraacetic acid disodium salt (EDTA disodium) in 80 mL of distilled water, adjust to pH 7, mix well, and dilute to a final volume of 100 mL. 41-2.3.2.2.3 Procedure

1. Prepare filtered soil extract as described in section 41-3.3.1.1.3. 2. Pipette an aliquot (not more than 5 mL) of the filtered 2 M KCI extract containing between 0.5 and 12 I-lg of NH4-N into a 25-mL volumetric flask. Aliquots of ::53 mL normally contain sufficient NH4-N for quantitation. Add 1 mL of the EDTA reagent, and mix the contents of the flask. Then allow the flask contents to stand for 1 min. Add 2 mL of the phenol-nitroprusside reagent, followed by 4 mL of the buffered hypoc1orite reagent, and immediately dilute the flask to volume with distilled water and mix well. Place the flask in a water bath maintained at 40°C, and allow it to remain 30 min. Remove the flask from the bath, cool to room temperature (about 10 min), and determine the absorbance of the colored complex at a wavelength of 636 nm against a reagent blank solution. 3. Determine the NH4-N concentration of the sample by reference to a calibration curve plotted from the results obtained with a 25-mL standard sample containing 0,2,4,6, 8, 10, and 12 I-lg of NH4-N. To prepare this curve, add an appropriate amount of 2 M KCI solution (same volume as that used for aliquots of soil extract) to a series of 25-mL volumetric flasks. Then add 0, 1,2,3,4,5, and 6 mL of the 2 mg of NH4-N L -1 solution to the series of flasks, and

NITROGEN AVAILABILITY INDICES

979

measure the intensity of blue color developed with these standards by the procedure described for analysis of the extract.

41-2.3.2.2.4 Comments Several factors affect the successful use of the indophenol blue method for measurement of NH4 in soil extracts. Divalent cations such as Ca2+ and Mg2+ in KCl extracts of soil interfere with the described method unless EDTA is added. The maximum amounts of these cations that can be tolerated when EDTA is included is about 4 mg of Mg2 + or 6 mg of Ca2+ per 25 mL of final diluted volume (Dorich & Nelson, 1983). If a precipitate develops in samples after the addition of the buffered hypochlorite reagent, a smaller sized aliquot of soil extract should be used to reduce the amounts of divalent cations in the system, or 2 mL of the EDTA reagent should be added to provide additional capacity for complexing divalent cations. For most soils, interference from divalent cations has not been observed if 5 mL or less of KCI extract is analyzed, but the method of Kempers (1974) can be used if precipitation of divalent cations is a major problem. A solution pH of 11.4 to 12.0 during color development is necessary for accurate and sensitive NH4 measurements. Although the NaOHhypochlorite reagent is buffered at the optimum pH for color development, basic or acidic extracts may influence the results obtained. All basic or acidic extracts should be neutralized before NH4 determination to ensure satisfactory color development. The phenol-nitroprusside reagent must be added before the buffered hypochlorite reagent to obtain proper color development. The phenol-nitroprusside reagent is stable in the dark, but weekly preparation is recommended for optimum sensitivity in NH4 analysis. A sufficient NaOCl concentration in the buffered hypochlorite reagent is critical for accurate NH4 measurements. The hypochlorite reagent should be prepared immediately before use to obtain optimum results, because the NaOCI concentration in this reagent decreases on standing. Commercial bleach is a satisfactory source of hypochlorite, but technicalgrade NaOCI solution from laboratory supply companies may give more reproducible results. Color development at 40°C for 30 min, gives the most sensitive and reproducible determinations of NH4 in KCI extracts of soils. Alternatively the color may be developed at room temperature (about 25°C) if the time is extended to 1 h. However, higher molar absorptivity values are obtained when color is developed at 40°C. The color produced at 25 or 40 ° is stable for at least 7 h (Dorich & Nelson, 1983).

REFERENCES Adamsen, F.l., D.S. Bigelow, and G.R. Scott. 1985. Automated methods for ammonium, nitrate, and nitrite in 2 M KCI-phenylmercuric acetate extracts of soil. Commun. Soil Sci. Plant Anal. 16:883-898.

980

BUNDY & MEISINGER

Allison, F.E. 1965. Estimating the ability of soils to supply nitrogen. Agric. Chem. 11:46-48, 139. American Public Health Association. 1989. Nitrogen (nitrate). p. 4-137 to 4-139. In L.S. Clescern et al. (ed.) Standard methods for the examination of water and wastewater. 17th ed. Am. Public Health Assoc., Washington, DC. Ananth, S., and J.T. Moraghan. 1987. The effect of calcium and magnesium on soil nitrate determination by automated segmented-flow methods. Soil Sci. Soc. Am. J. 51:664667. Beauchamp, E.G., W.D. Reynolds, D. Brasche-Villeneuve, and K. Kirby. 1986. Nitrogen mineralization kinetics with different soil pretreatments and cropping histories. Soil Sci. Soc. Am. J. 50:1478-1483. Beegle, D., G. Roth, and R. Fox. 1989. Nitrogen soil test for com in Pennsylvania. Pennsylvania State Univ., University Park, Agron. Facts no. 17 (revised). Best, E.K. 1976. An automated method for determining nitrate-N in soil extracts. Queensl. J. Agric. Anim. Sci. 33:161-166. Blackmer, A.M., T.F. Morris, D.R. Keeney, and R.D. Voss. 1991. Estimating nitrogen needs for com by soil testing. Iowa State Univ. Publ. Pm 1381. Blackmer, A.M., D. Pottker, M.E. Cerrato, and J. Webb. 1989. Correlations between soil nitrate concentrations in late spring and com yields in Iowa. J. Prod. Agric. 2:103-109. Bock, B.R., and K.R. Kelley. 1992. Predictin~ N fertilizer needs for com in humid regions. TVA Bull. Y-226. Natl. Fert. and EnViron. Res. Ctr., Tennessee Valley Authority, Muscle Shoals, AL. Bole, J.B., and U.J. Pittman. 1976. Sampling southern Alberta soils for N and P soil testing. Can. J. Soil Sci. 56:531-535. Bonde, T.A., and T. Lindberg. 1988. Nitrogen mineralization kinetics in soil during long-term aerobic laboratory incubations: A case study. J. Environ. Qual. 17:414-417. Bonde, T.A., and T. Rosswall. 1987. Seasonal variation of potentially mineralizable nitrogen in four cropping systems. Soil Sci. Soc. Am. J. 51:1508-1514. Bonde, T.A., J. Schnurer, and T. Rosswall. 1988. Microbial biomass as a fraction of rotentially mineralizable nitrogen in soils from long-term field experiments. Soil Bio . Biochem.20:447-452. Boone, R.D. 1990. Soil organic matter as a potential net nitrogen sink in a fertilized cornfield, South Deerfield, Massachusetts, USA. Plant Soil 128:191-198. Boyle, M., and E.A. Paul. 1989. Carbon and nitrogen mineralization kinetics in soil previously amended with sewage sludge. Soil Sci. Soc. Am. J. 53:99-103. Bremner, J.M. 1965a. Nitro~en availability indexes. p. 1324-1345. In C.A. Black et al. (ed.) Methods of soil analYSIS. Part 2. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Bremner, J.M. 1965b. Inorganic forms of nitrogen. p. 1179-1237. In C.A. Black et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Buckman, H.O. 1910. Moisture and nitrate relations in dry-land agriculture. J. Am. Soc. Agron. 2:121-138. Bundy, L.G., and E.S. Malone. 1988. Effect of residual profile nitrate on com response to applied nitrogen. Soil Sci. Soc. Am. J. 52:1377-1383. Bundy, L.G., M.A. Schmitt, and G.W. Randall. 1992. Predicting N fertilizer needs for com in humid regions: Advances in the Upper Midwest. p. 73-89. In B.R. Bock and K.R. Kelley (ed.) Predicting N fertilizer needs for com in humid re~ons. TVA Bull. Y-226. Natl. Fert. and Environ. Res. Ctr., Tennessee Valley Authonty, Muscle Shoals, AL. Cabrera, M.L., and D.E. Kissel. 1988a. Evaluation of a method to predict nitrogen mineralized from soil organic matter under field conditions. Soil Sci. Soc. Am. J. 52:10271031. Cabrera, M.L., and D.E. Kissel. 1988b. Length of incubation time affects the parameter values of the double exponential model of nitrogen mineralization. Soil Sci. Soc. Am. J. 52:1186-1187. Cabrera, M.L., and D.E. Kissel. 1988c. Potentially mineralizable nitrogen in disturbed and undisturbed soil samples. Soil Sci. Soc. Am. J. 52:1010-1015. Call, L.E. 1914. The effect of different methods of preparing a seed bed for winter wheat upon yield, soil moisture, and nitrates. J. Am. Soc. Agron. 6:249-259. Cameron, D.R., M. Nyborg, J.A. Toogood, and D.H. Laverty. 1971. Accuracy of field sampling for soil tests. Can. J. Soil Sci. 51:165-175.

NITROGEN AVAILABILITY INDICES

981

Cataldo, D.A., M. Haroon, L.E. Schrader, and V.L. Young. 1975. A rapid colorimetric determination of nitrate in plant tissue by nitration of salicylic acid. Commun. Soil Sci. Plant Anal. 6:71-80. Cawse, P.A. 1967. The determination of nitrate in soil solutions by ultraviolet spectrophotometry. Analyst (London) 92:311-315. Chaussod, R., B. Nicolardot, and G. Catroux. 1986. Mesure en routine de la biomasse microbienne des sols par la methode de fumigation au chloroforme. Sci. Sol 25:201-211. Dahnke, W.C., and G.Y. Johnson. 1990. Testing soils for available nitrogen. p. 127-139. In R.L. Westerman (ed.) Soil testing and plant analysis. 3rd ed. SSSA, Madison, WI. Dahnke, W.C., and E.H. Vasey. 1973. Testing soils for nitrogen. p. 97-114. In L.M. Walsh and J.D. Beaton (ed.) Soil testing and plant analysis. ASA, CSSA, and SSSA, Madison, WI. Davis, J.G., and A.M. Blackmer. 1990. Sampling soils for late-spring nitrate in fields fertilized with anhydrous ammonia. p. 266. In Agronomy abstracts. ASA, Madison, WI. Deans, J.R., J.A.E. Molina, and C.E. Clapp. 1986. Models for predicting potentially mineralizable nitrogen and decomposition rate constants. Soil Sci. Soc. Am. J. 50:323-326. Dorich, R.A., and D.W. Nelson. 1983. Direct colorimetric measurement of ammonium in potassium chloride extracts of soil. Soil Sci. Soc. Am. J. 47:833-836. Dorich, R.A., and D. W. Nelson. 1984. Evaluation of manual cadmium reduction methods for determination of nitrate in potassium chloride extracts of soils. Soil Sci. Soc. Am. J. 48:72-75. Ellert, B.H., and J.R. Bettany. 1988. Comparison of kinetic models for describing net sulfur and nitrogen mineralization. Soil Sci. Soc. Am. J. 52:1692-1702. Fox, R.H., and w.P. Piekielek. 1978. Field testing of several nitrogen availability indexes. Soil Sci. Soc. Am. J. 42:747-750. Fox, R.H., and W.P. Piekielek. 1984. Relationships among anaerobically mineralized nitrogen, chemical indexes, and nitrogen availability to corn. Soil Sci. Soc. Am. J. 48:10871090. Fox, R.H., G.w. Roth, K.V. Iversen, and W.P. Piekielek. 1989. Soil and tissue nitrate tests compared for predicting soil nitrogen availability to corn. Agron. J. 81:971-974. Gale, P.M., and J.T. Gilmour. 1988. Net mineralization of carbon and nitrogen under aerobic and anaerobic conditions. Soil Sci. Soc. Am. J. 52:1006-1010. Gales, M.E., and R.L. Booth. 1975. A copper-cadmium column for manually determining nitrate. In News of environmental research in Cincinnati. 28 Feb. 1975. USEPA, Cincinnati. Gelderman, R.H., and P.E. Fixen. 1988. Recommended nitrate-N test. p. 10-12. In W.C. Dahnke (ed.) Recommended chemical soil test procedures for the North Central region. North Dakota Agric. Exp. Stn., Fargo, North Central Regional Pub!. 221. Gentry, C.E., and R.B. Willis. 1988. Improved method for automated determination of ammonium in soil extracts. Commun. Soil Sci. Plant Anal. 19:721-737. Gianello, C., and J.M. Bremner. 1986a. A simple chemical method of assessing potentially available organic nitrogen in soil. Commun. Soil Sci. Plant Anal. 17:195-214. Gianello, C., and J.M. Bremner. 1986b. Comp'arison of chemical methods of assessing potentially available organic nitrogen in SOlI. Commun. Soil Sci. Plant Anal. 17:215236. Gianello, c., and J.M. Bremner. 1988. A rapid steam distillation method of assessing potentially available organic nitrogen in soil. Commun. Soil Sci. Plant Anal. 19:15511568. Harmsen, G.W., and D.A. Van Schreven. 1955. Mineralization of organic nitrogen in soil. Adv. Agron. 7:299-398. Henriksen, A., and A.R. Selmer-Olsen. 1970. Automatic methods for determining nitrate and nitrite in water and soil extracts. Analyst 95:514-518. Hergert, G.W. 1987. Status of residual nitrate-nitrogen soil tests in the United States. p. 73-88. In J.R. Brown (ed.) Soil testing: Sampling, correlation, calibration, and interpretation. ASA Spec. Publ. 21. ASA, CSSA, and SSSA, Madison, WI. Hong, S.D., R.H. Fox, and W.P. Piekie1ek. 1990. Field evaluation of several chemical indexes of soil nitrogen availability. Plant Soil 123:83-88. Houot, S., J.A.E. Molina, R. Chaussod, and C.E. Clapp. 1989. Simulation by NCSOIL of net mineralization in soils from the Deherain and 36 Parcelles fields at Grignon. Soil Sci. Soc. Am. J. 53:451-455.

982

BUNDY & MEISINGER

Jackson, w.L., C.E. Frost, and D.M. Hildreth. 1975. Versatile multi-range analytical manifold for automated analysis of nitrate-nitrogen. Soil Sci. Soc. Am. Proc. 39:592-593. Jokela, W. 1989. The Vermont nitrogen soil test for corn. Coop. Ext. Serv., Univ. of Vermont Publ. no. FS 133, Burlington. Keeney, D.R. 1982. Nitrogen-availability indexes. p. 711-733. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9 ASA and SSSA, Madison, WI. Keeney, D.R., and D.W. Nelson. 1982. Nitrogen inorganic forms. p. 643-698. In AL. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, WI. Kempers, AJ. 1974. Determination of submicrogram quantities of ammonium and nitrates in soil with phenol, sodium nitroprusside and hypochlorite. Geoderma 12:201-206. Kempers, A.J., and A. Zweers. 1986. Ammonium determination in soil extracts by the salicylate method. Commun. Soil Sci. Plant Anal. 17:715-723. King, F.H., and A.R. Whitson. 1901. Development and distribution of nitrates and other soluble salts in cultivated soils. Wisconsin Agric. Exp. Stn. Bull. no. 85. King, F.H., and A.R. Whitson. 1902. Development and distribution of nitrates in cultivated soils. (Second paper.) Wisconsin Agric. Exp. Stn. Bull. no. 93. Kopp, J.F., and G.D. McKee. 1978. Methods for chemical analysis of water and wastes. Nitrogen ammonia-Method 350.1. USEPA Environ. Monitoring and Support Lab., Cincinnati. Liang, B.C., M. Remillard, and A.F. MacKenzie. 1991. Influence of fertilizer, irrigation, and non-growing season precipitation on soil nitrate-nitrogen under corn. J. Environ. Qual. 20:123-128. Lindemann, W.C., and M. Cardenas. 1984. Nitrogen mineralization potential and nitrogen transformations of sludge-amended soil. Soil Sci. Soc. Am. J. 48:1072-1077. Lubochinsky, B., and J.P. Zalta. 1954. Colorimetric microdetermination of ammoniacal nitrogen. Bull. Soc. Chim. BioI. 46:1363. Lueking, M.A., and J.S. Schepers. 1986. Achieving desired moisture conditions in potentially mineralizable nitrogen incubation studies. Soil Sci. Soc. Am. J. 50:1370-1373. MacKay, D.C., and J.M. Carefoot. 1981. Control of water content in laboratory determination of mineralizable nitrogen in soils. Soil Sci. Soc. Am. J. 45:444-446. Magdoff, F. 1991. Understanding the Magdoff pre-sidedress nitrate test for corn. J. Prod. Agric. 4:297-305. Magdoff, F.R., W.E. Jokela, R.H. Fox, and G.F. Griffin. 1990. A soil test for nitrogen availability in the northeastern United States. Commun. Soil Sci. Plant Anal. 21:11031115. Magdoff, F.R., D. Ross, and J. Amadon. 1984. A soil test for nitrogen availability to corn. Soil Sci. Soc. Am. J. 48:1301-1304. Mann, L.T., Jr. 1963. Spectrophotometric determination of nitrogen in total micro-Kjeldahl digests. Anal. Chern. 35:2179-2182. Marion, G.M., and C.H. Black. 1988. Potentially available nitrogen and phosphorus along a chaparral fire cycle chronosequence. Soil Sci. Soc. Am. J. 52:1155-1162. Markus, K.K., J.P. Mckinnon, and A.F. Buccafuri. 1985. Automated analysis of nitrite, nitrate, and ammonium nitrogen in soils. Soil Sci. Soc. Am. J. 49:1208-1215. McCracken, D.V., S.J. Corak, M.S. Smith, W.W. Frye, and R.L. Blevins. 1989. Residual effects of nitrogen fertilization and winter cover cropping on nitrogen availability. Soil Sci. Soc. Am. J. 53:1459-1464. Meisinger, J.J. 1984. Evaluating plant-available nitrogen in soil-crop systems. p. 391-416. Evaluating plant-available mtrogen in soil-crop systems. p. 391-416. In R.D. Hauck et al. (ed.) Nitrogen in crop production. ASA, Madison, WI. Meisinger, J.J., V.A. Bandel, J.S. Angle, B.E. O'Keefe, and C.M. Reynolds. 1992a. Presidedress soil nitrate test evaluation in Maryland. Soil Sci. Soc. Am. J. 56:1527-1532. Meisinger, J.J., F.R. Magdoff, and J.S. Schepers. 1992b. Predicting N fertilizer needs for corn in humid regions: Underlying principles. p. 7-27. In B.R. Bock and K.R Kelley (ed.) Predicting N fertilizer needs for corn in humid regions. TVA Bull. Y-226. Natl. Fert. and Environ. Res. Ctr., Tennessee Valley Authority, Muscle Shoals, AL. Molina, J.AE., C.E. Clap, M.J. Schaffer, F.W. Chichester, and W.E. Larson. 1983. NCSOIL, A model of nitrogen and carbon transformations in soil: Description, calibration, and behavior. Soil Sci. Soc. Am. J. 47:85-91.

NITROGEN AVAILABILITY INDICES

983

Myers, R.G., C.W. Swallow, and D.E. Kissel. 1989. A method to secure, leach, and incubate undisturbed soil cores. Soil Sci. Soc. Am. J. 53:467-471. Nadelhoffer, K.J. 1990. Microlysimeter for measuring nitrogen mineralization and microbial respiration in aerobic soil incubations. Soil Sci. Soc. Am. J. 54:411-415. Nelson, D.W. 1983. Determination of ammonium in KCI extracts of soils by salicylate method. Commun. Soil Sci. Plant Anal. 14(11):1051-1062. Norman, R.J., J.C. Edberg, and J.W. Stucki. 1985. Determination of nitra~e in soil extracts by dual-wavelength ultraviolet spectrophotometry. Soil Sci. Soc. Am. J. 49:1182-1185. 0ien, A., and AR. Selmer-Olsen. 1980. A laboratory method for evaluation of available nitrogen in soil. Acta Agric. Scand. 30:149-156. Paustian, K., and J. Schnurer. 1987. Fungal growth response to carbon and nitrogen limitation: Application of a model to laboratory and field data. Soil Biol. Biochem. 19:621629. Reuss, J.O., P.N. Soltanpour, and A.E. Ludwick. 1977. Sampling distribution of nitrates in irrigated fields. Agron. J. 69:588-592. Rice, C.W., J.H. Grove, and M.S. Smith. 1987. Estimating soil net nitrogen mineralization as affected by tillage and soil drainage due to topographic position. Can. J. Soil Sci. 67:513-520. Rice, C.W., M.S. Smith, and J.M. Crutchfield. 1984. Inorganic N analysis of soil extracts by automated and distillation procedures. Commun. Soil Sci. Plant Anal. 15:663-672. Richter, J., A Nuske, W. Habenicht, and J. Bauer. 1982. Optimized N-mineralization parameters of loess soils from incubation experiments. Plant Soil 68:379-388. Robertson, K., J. Schnurer, M. Clarholm, T.A. Bonde, and T. Rosswall. 1988. Microbial biomass in relation to C and N mineralization during laboratory incubations. Soil BioI. Biochem. 20:281-286. Roth, G.W., and R.H. Fox. 1990. Soil nitrate accumulations following nitrogen-fertilized corn in Pennsylvania. J. Environ. qual. 19:243-248. Saffigna, P. 1988. N-15 methodology in the field. p. 433-451. In J.R. Wilson (ed.) Advances in nitrogen cycling in agricultural ecosystems. Proc. Symp. at Brisbane, Australia. 11-15 May 1987. CAB Int., Wallingford, UK. Sahrawat, K.L., and F.N. Ponnamperuma. 1978. Measurement of exchangeable NH4 in tropical rice soils. Soil Sci. Soc. Am. J. 42:282-283. Sarrantonio, M., and T.W. Scott. 1988. Tillage effects on availability of nitrogen to corn following a winter green manure crop. Soil Sci. Soc. Am. J. 52:1661-1668. Scharf, P., and M.M. Alley. 1988. Centrifugation: A solution to the problem posed by ammonium and nitrate contamination of filters in soil extract analysis. Soil Sci. Soc. Am. J. 52:1508-1510. Sheppard, S.C., and T.E. Bates. 1986. Changes in nitrate concentration over winter in three southern Ontario soil profiles. Can. J. Soil Sci. 66:537-541. Sims, J.R., and G.D. Jackson. 1971. Rapid analysis of soil nitrate with chromotropic acid. Soil Sci. Soc. Am. Proc. 35:603-606. Skjemstad, J.O., and R. Reeve. 1978. The automatic determination of ammonia, nitrate plus nitrite, and phosphate in water in the presence of added mercury (III) chloride. J. Environ. Qual. 7:137-141. Smith, J.L., R.R. Schnabel, B.L. McNeal, and G.S. Campbell. 1980. Potential errors in the first-order model for estimating soil nitrogen mineralization potentials. Soil Sci. Soc. Am. J. 44:996-1000. Smith, K.A., and A Scott. 1991. Continuous-flow, flow-injection, and discrete analysis. p. 183-227. In K.A. Smith (ed.) Soil analysis: Modern instrumental techniques. 2nd ed. Marcel Dekker, New York. Smith, S.J. 1987. Soluble organic nitrogen losses associated with recovery of mineralized nitrogen. Soil Sci. Soc. Am. J. 51:1191-1194. Smith, S.J., L.B. Young, and G.E. Miller. 1977. Evaluation of soil nitrogen mineralization potential under modified field conditions. Soil Sci. Soc. Am. J. 41:74-76. Sparrow, S.D., and D.T. Masiak. 1987. Errors in the analyses for ammonium and nitrate caused by contamination from filter papers. Soil Sci. Soc. Am. J. 51:107-110. Stanford, G. 1982. Assessment of soil nitrogen availability. p. 651-688. In F.J. Stevenson (ed.) Nitrogen in agricultural soils. Agron. Monogr. 22. ASA and SSSA, Madison, WI. Stanford, G., J.N. Carter, and S.J. Smith. 1974. Estimates of potentially mineralizable soil nitrogen based on short-term incubations. Soil Sci. Soc. Am. Proc. 38:99-102.

984

BUNDY & MEISINGER

Stanford, G., and E. Epstein. 1974. Nitrogen mineralization-water relations in soils. Soil Sci. Soc. Am. Proc. 38:103-107. Stanford, G., M.H. Frere, and D.H. Schwaninger. 1973. Temperature coefficient of soil nitrogen mineralization. Soil Sci. 115:321-323. Stanford, G., and S.J. Smith. 1972. Nitrogen mineralization potentials of soils. Soil Sci. Soc. Am. Proc. 38:99-102. Stevenson, F.J. 1986. Cycles of soil. John Wiley and Sons, New York. Technicon Instrument Corp. 1977a. Ammonia in water and wastewater. Industrial Method no. 98-70W/A. Tarrytown, NY. Technicon Instrument Corp. 1977b. Nitrate and nitrite in soil extracts. Industrial Method no. 487-77A. Tarrytown, NY. Tel, D.A., and C. Heseltine. 1990. The analysis of KCl soil extracts for nitrate, nitrite and ammonium using a TRAACS 8000 analyzer. Commun. Soil Sci. Plant Anal. 21:16811688. Thomas, G.W., M.S. Smith, and R.E. Phillips. 1989. Impact of soil management practices on nitrogen leaching. p. 247-276. In R.E. Follett (ed.) Nitrogen management and ground water protection. Elsevier Sci. Publ. Co., New York. Tyler, D.D., and G. W. Thomas. 1977. Lysimeter measurements of nitrate and chloride losses from soil under conventional and no-tillage corn. J. Environ. Qual. 6:63-66. Vendrell, P.F., and J. Zupancic. 1990. Determination of soil nitrate by transnitration of salicylic acid. Commun. Soil Sci. Plant Anal. 21:1705-1713. U.S. Environmental Protection Agency (USEPA). 1983. Nitrogen, nitrate-nitrite. p. 353.2-1 to 352.2-7. In Methods of chemIcal analysis of water and wastes. EPA-600 4-79-020. 1983 ed. ESEPA, Cincinnati. Waring, S.A., and J.M. Bremner. 1964. Ammonium production in soil under waterlogged conditions as an index of nitrogen availability. Nature (London) 201:951-952.

Published 1994

Chapter 43 Dinitrogen Fixation R. W. WEAVER, Texas A&M University, College Station, Texas 77843 SETH K. A. DANSO, International Atomic Energy Agency, Vienna, Austria

The quantity of dinitrogen (N2 ) fixed by diverse groups of microorganisms is of great importance to the N cycle of the biosphere. It has been estimated that approximately 83% of the N fixed annually originates from biological N2 fixation and by contrast 14% is from manufacture of fertilizers (Burns & Hardy, 1975). Well-nodulated plants may fix in excess of 200 kg of N ha- I ye l and estimates for bacteria associated with grass roots generally are in the range of a few kg ha- I . Because of the extensive nature of grasslands, however, the total quantity of N2 contributed by grasslands to the soil is equivalent to that of legumes (Burns & Hardy, 1975). The primary limiting factor to N2 fixation by microorganisms in nature is availability of energy to drive the N2-fixation process. In soil systems, the major energy source is from plant photosynthates made available to microorganisms in nodule structures or, in the case of nonsymbiotic N2 fixation, from root exudates. Without plants, N2 fixation is minimal and of little agronomic importance. The purpose of this chapter is to describe methods that may be used to measure or estimate the quantity of N2 fixed in systems that include plants. Measurement of N2 fixation by pure cultures of microorganisms is described in chapter 11 by Knowles and Barraquio in this book. Methods that will be described include acetylene reduction, total N difference, isotope dilution, and I5N2 incorporation. 43-1 ACETYLENE REDUCTION 43-1.1 Principles Discovery that nitrogenase is able to reduce acetylene (CzH2 ) to ethylene (C2H 4 ) resulted in the development of a highly sensitive method having great utility for laboratory measurements and some application for Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

1019

1020

WEAVER & DANSO

field measurement of N2 fixation. Burris (1974, 1975) provides an excellent overview of the method's development. The method has been used extensively for the past 20 yr to estimate nitrogenase activity that has sometimes been extrapolated to amounts of N2 fixed. A theoretical conversion factor of 3 mol of ~H4 produced to 1 mol of N2 reduced was first suggested (Hardy et aI., 1968). During N2 fixation, some electrons are consumed in production of H2 but H2 production is restricted in the presence of C2H 2. Thus, a theoretical conversion factor of 4 mol of C2H 4 produced to 1 mol of N2 reduced is more commonly used today (Boddey, 1987). Actually, conversion factors range between 1.5 to 25 and must be determined experimentally for each system (Hardy et aI., 1973). Use of ~H2 reduction (AR) to quantitatively measure rates of N2 fixation has fallen into disfavor for both legume (Witty & Minchin, 1988) and grass systems (Giller, 1987). The reasons are multifold. The assay is a point in time measurement; therefore, it is not an appropriate method for measuring seasonal rates of N2 fixation because the rate may vary with time. Problems also occur in using the method for point in time measurements. In the case of legume nodule systems, exposure of nodules to C2H 2 may reduce O 2 permeability of the nodule resulting in reduced nitrogenase activity (Sheehy et al., 1983). Because acetylene inhibits loss of H2 by nitrogenase, N2 fixation would be overestimated unless the particular organism has an efficient uptake hydrogenase (Arp, 1992). The principal utility of the method was that a nodulated root could be removed from the plant and assayed in a closed system. However, removal of the plant top and disturbance of the root system are detrimental to AR activity of many legumes because of perturbation of nodule oxygen barriers (Minchin et a1., 1986; Herdina & Silsbury, 1990). Under field conditions there is always the risk of losing many nodules during excavation of the root. In situ methods for legumes have been proposed (Lindstrom, 1984), but generally are not used because of labor requirements. In grass systems, the primary problem is the vulnerability of the N2fixation system to changes in the gaseous environment that invariably occur during collection of soil or root samples. Additionally, the long incubation periods needed for detection of AR activity also allow time for changes in O 2 content, microbial populations, and production of ~H4 that occur independently of AR (Nohrstedt, 1983). Methods for estimating the magnitude ofthis problem are described by Nohrstedt (1983) and Witty (1979). 43-1.2 Acetylene Reduction for Nodulated Root Systems 43-1.2.1 Principles Procedures are described separately for nodulated and non-nodulated root systems because requirements for the two systems are considerably different. For nodulated plants, measurable AR activity occurs for a single effective nodule almost immediately on exposure to ~H2' The closed system that is described is useful for determining if nodules are active in N2

DINITROGEN FIXATION

1021

Fig. 43-1. Apparatus for generating C2 H 2 from CaC2 . The top reservoir may be supported by a ring stand.

fixation, but it should not be considered quantitative in measuring nitrogenase activity or extrapolating to a quantity of N2 fixed. To quantitatively measure nitrogenase activity, it is necessary to use a flow through system that will not be described in this chapter because of limited practical application (Sheehy, 1991; Gerbaud, 1990; McNeill et aI., 1989; Weisz & Sinclair, 1988; Mederski & Streeter, 1977). Even with flow through systems it is necessary to experimentally determine a conversion factor if AR activity is to be converted to amount of N2 fixed. 43-1.2.2 Materials 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Calcium carbide (CaC2), available from sporting goods stores. Ethylene for preparation of standards. Apparatus for generating C2H 2 (Fig. 43-1). Wide mouth canning jars (500 mL) with lids. Rubber septa, 5.5 mm diam. Plastic syringes, 50 mL. Plastic syringes,S mL. Plastic syringes, 1 mL. Vacutainers, 5 mL. Nodulated roots. Gas chromatograph equipped with H2 flame ionization detector. Porapak Q column, 1.5 m long and 32 mm diam. Nitrogen carrier gas.

1022

WEAVER & DANSO

43-1.2.3 Procedure Calculate the quantity of ~H2 needed to provide an atmosphere of 10% ~H2' on a volume basis, in the vessel used for incubation of the sample. Generate the ~H2 from Ca~ by adding it to the water in the apparatus (Fig. 43-1) described by Tann and Skujins (1985). The apparatus allows collection of the generated ~H2 in a reservoir created by the displacement of water. The apparatus may be constructed from convenient size flasks depending on the amount of ~H2 to be generated. The flask on the top should have a larger capacity than the flask on the bottom so that all of the water displaced may be contained. Fill the lower flask with water and assemble the apparatus. Generate ~H2 by dropping large crystals of CaC2 into the lower flask and connect the upper flask. For each gram of Ca~ added approximately 130 mL of C2H 2 is generated. It is generally better to generate ~H2 from CaC2 than using tank C2H 2, because it is not as contaminated with other gases. If large quantities of C2H 2 are needed, it may be most practical to use tank ~H2' Purity of the ~H2 can influence the AR activity (Tough & Crush, 1979; Hyman & Arp, 1987). Acetylene may be partially purified by passing the gas through concentrated sulfuric acid and water and by freezing out some contaminants (Hyman & Arp, 1987). Purity of the C 2H 2 will not likely be a problem for most qualitative applications, but a determination should be made experimentally by comparison with purified C2H 2. The vessel used for incubation should be large enough to contain the sample and provide an adequate volume of air such that O 2 will not become depleted during the incubation. Generally, a large and well-nodulated root or several smaller roots may be contained in a 500-mL canning jar and incubated for 1 h without depleting O 2 enough to interfere with the rate of AR. If nodules are removed from the root, the nodules should not occupy more than 10% of the volume of the incubation vessel to avoid O 2 depletion because of their high respiration. For extended incubations, the O 2 concentration should be monitored and supplemented as needed. After the incubation, remove a gas sample for analysis. Usually, it is convenient to remove approximately 5 mL of gas with a needle and syringe and inject it into a 3-mL vacutainer for storage under pressure. Storage under pressure is desirable so that samples removed in a syringe will be under pressure and contamination by air minimized. The vacutainer should be prepared for receiving the gas sample. First open the container so that gases that have been released from the rubber septum during storage under vacuum will be released. Some of these gases tend to interfere with gas chromatographic analyses. Replace the septum and re-evacuate the container using a vacuum pump or a 50-mL syringe and hypodermic needle. Do not assume that the containers are fully evacuated as received from the manufacturer. It is a good practice to analyze the gases within a few days, because longer-term storage may result in absorption of some ~H4 into the rubber septum.

DINITROGEN FIXATION

1023

Calibrate the gas chromatograph for the range of C2H 4 concentrations expected. One-milliliter capacity plastic syringe may be used for injections. Generally, an operating temperature of 50°C is adequate for the column and a flow rate of 50 mL min- 1 for a N2 carrier gas allows for good sensitivity and separation of gases. Retention times for C2H 4 and C2H 2 are approximately 30 and 60 s, respectively. 43-1.3 Acetylene Reduction for Grass Systems 43-1.3.1 Principles

A major advantage of the AR method is the sensitivity of analytical methods for the detection of C2H 4 . Rates of AR that equate to < 0.5 kg of N2 fixed ha- 1 in 100 d can be measured when 24-h incubations are used. Thus, sensitivity of the method for detecting agronomically significant amounts of N2 fixation is not limiting provided an incubation time of 1 d can be tolerated. Closed systems must be used to allow enough C2H 4 to accumulate, however, in providing a closed system, the material being assayed is disturbed to some extent. The technique used for exposure of plant root systems to C2H 2 depends on the type of plant involved and the degree of root disturbance that can be tolerated. Disturbing the root system is particularly critical because it alters the gaseous environment that strongly influences the rate of AR activity (Zuberer & Alexander, 1986) and for incubation times of hours, alterations in microbial populations may occur. Also, for such long incubation times, removal of plant tops may have a negative effect by reducing availability of plant photosynthates to roots. Leaving the plant top on unfortunately is impractical for many field investigations. The technique we have chosen to describe relies on the use of cores and worked quite well for field surveys of AR by pasture grasses and should work equally well for other plant types (Weaver et aI., 1980). The results of using the technique cannot be considered quantitative for in situ rates because of disturbing the system and poor correlation with measurements based on incorporation of 15N2 (Morris et aI., 1985). A comprehensive review by Boddey (1987) has been published that covers in detail problems with measurement of AR activity of non-nodulating N2 fixing bacteria in association with grasses. 43-1.3.2 Materials

1. 2. 3. 4. 5. 6. 7. 8.

Calcium carbide (see section 43-1.2.3). Gas impermeable plastic bags (saran). Device for taking soil cores. Self-adhering insulation tape. Duct tape. White no. 000 rubber stoppers. Large screw clamp as used for truck radiator hose. Tin cans at least 8-cm diam.

1024

WEAVER & DANSO

43-1.3.3 Procedure

The technique depends on being able to take intact soil cores and leaving them intact. Tin cans serve as the coring device. New cans are used, and may be purchased from a canning company with one end open. Remove the closed end of the can with a can opener. The reinforced edge (rim remains after removing lid) of the can is used in forcing the sharp end of the can into the soil. Cans of 8 cm in diameter by 11 cm in height are convenient. Insert the can into the soil using a specially-constructed core driver. The core driver is a steel plate 2-cm thick that is grooved to a l-cm depth to fit the edge of the can having the rim. Weld a 1.5-cm thick steel rod to the center of the steel plate for a cylindrical sliding weight of 3 kg that can be used to drive the can into the soil. It will be necessary to cut the tops from the plants being sampled. The apparatus for taking the cores helps in pushing the cans into the soil without collapsing the cans. Use of a hammer or a block of wood does not allow for uniform pressure on the can and the can is more likely to collapse. The can is not strong enough to be inserted into dry soil or soil containing stones. Anyway, our experience has been that dry soil does not support measurable rates of AR activity. After inserting the can, remove it from the soil with the aid of a shovel. The soil core is placed inside a 17 by 28 cm plastic bag constructed from saran or mylar and laminated with polyethylene so that the bag is heat sealable. The saran or mylar makes the bag relatively impermeable to gases (Burris, 1974). Prepare one end of the bag for removal of gas samples. It is convenient to use approximately 2-mm thick discs cut from white no. 000 rubber stoppers. A disc is attached to the bag with a piece of gray duct tape that is at least three times the width of the disc. The disc is centered on the sticky side of the duct tape then the tape is sealed to the bag being careful not to have wrinkles that could allow gas to escape around the edges of the tape. Place the soil core into the bag, compress the bag to remove most of the air, and heat seal. Separate the upper and lower portions of the bag by tightly wrapping self-adhering insulation tape horizontally around the middle of the bag and core. Place a radiator hose clamp over the insulation tape and tighten to complete the seal. Separating the core into the two compartments facilitates mixing of gases through the soil core at the end of the incubation. The volume of air space in the containers must be estimated so that enough C2H 2 can be added to achieve a 10% atmosphere. The volume of a soil filled core with bag in place may be estimated by taking an extra core sample and injecting a known amount of C2H 2 into the assembly and determining the degree of dilution by gas chromatography. The calculated value will be used to determine the amount of CzH2 to add to each assembly to achieve an approximate concentration of 10%. Since this concentration of CzH2 is saturating for nitrogenase, the actual concentration in the container is not critical. Add the calculated quantity of C2H 2 to each container. The gas is only added to one end of the assembly and allowed to move into the core by diffusion so as to minimize disturbance of the

DINITROGEN FIXATION

1025

gaseous atmosphere within the core. Incubate the assemblies under the desired temperature conditions for 12 to 24 h. At the end ofthe incubation, mix the gaseous contents by alternately squeezing each end. The exact volume of each container and a representative sample is needed to compute how much CzH 4 was produced. Remove a sample for gas chromatography analyses and store the sample in a vacutainer as indicated under section 43-1.2.3. An accurate gas volume for each container is determined by the degree of CzHz dilution.

43-2 NITROGEN DIFFERENCE 43-2.1 Introduction Nitrogen balance experiments were originally used to determine the role of bacteria and plants in Nz fixation. It may be of interest to read the historical review by Bergersen (1980b) of the development of evidence demonstrating Nz fixation. Nitrogen increases can be readily demonstrated for inoculated plants grown on media lacking mineral N. Demonstration of Nz fixation in soil systems is much more complex because the plant now has two sources of N; N from the soil and Nz fixation. The contribution of N from the soil is estimated by growing plants that do not have the benefit of Nz fixation and assuming the Nz-fixing plant obtains a similar amount of N from the soil. The amount of N in the plant not benefiting from Nz fixation is subtracted from the plant benefiting from N2 fixation to determine the quantity of N2 fixed. The method is not sensitive enough for determining the quantity of N2 fixed over time intervals of less than a few days. Therefore, it is of little use as a point in time measurement for N2 fixation. A description of techniques to measure N2 fixation under controlled conditions of the laboratory and in the field follow. The reader may also refer to descriptions of methods by Vincent (1970), Brockwell (1980), Somasegaran and Hoben (1985), and Wynne et al. (1987).

43-2.2 Nitrogen-Difference Method for Controlled Environment 43-2.2.1 Principles Plants are grown under conditions that allow for control of mineral N inputs and prevention of contamination of Nz-fixing microorganisms. It is not practical or necessary to grow the plants axenically. The technique described is a variation of that described by Leonard (1943). Basically, plants are grown in pots provided water and nutrients by subirrigation. Subirrigation prevents N2-fixing microorganisms from being washed into the pot during irrigation. Seeds are inoculated with the microorganism of interest and grown in a plant support medium deficient in mineral N. Plant nutrients other than N are provided by a nutrient solution. Uninoculated

WEAVER & DANSO

1026

control plants are needed to demonstrate that contamination with nodulating microorganisms did not occur and to evaluate the combined contribution of mineral N from the plant growth medium and that from the seed. Sometimes, it is also useful to have control plants receiving mineral N to demonstrate that lack of mineral N was the main factor limiting plant growth and to compare with the symbiotic system's ability to provide N. Dinitrogen fixed is the difference in total N content between inoculated and uninoculated plants. 43-2.2.2 Materials 1. High quality legume seeds.

2. 3. 4. 5. 6. 7.

Culture of micro symbiont. Plastic pots (1 L) with lids. Absorbent cotton (rolls). Coarse vermiculite. Sodium hypochlorite (household bleach) or ethanol. Chemicals for nutrient solution (see Table 12-1 in this book).

43-2.2.3 Procedure Seed and materials used to culture plants must be free of nodulating organisms. Unless contaminated after receipt, plastic pots, absorbent cotton, vermiculite, and chemicals are free of nodulating microorganisms. It is not necessary to autoclave plant nutrient solutions made with distilled water. Used or contaminated materials, however, will need to be autoclaved. Containers may be reused by washing with a soapy solution and rinsing in a solution of household bleach (5% NaOCl) diluted 1:4 with water on a volume basis. Rinse thoroughly with water to remove the bleach. Two 1-L plastic containers are used for each growth unit (Fig. 43-2). A lid is placed on one container and the other container sits on top of it. The lower container serves as a reservoir for water. The upper container is for the N-deficient plant growth medium. Cut a hole approximately 2 cm in diameter in the center of the lid and the bottom of the top container. Cut a second hole of approximately 6-mm diam. in the lid close to edge of the lid. Make a cotton wick by cutting a 20-cm long strip of absorbent cotton that is wide enough to snugly fit through the holes in the center of the container and lid. The wick will serve to draw nutrient solution into the rooting medium and should extend from the bottom of the nutrient solution up to approximately 6 cm into the pot containing the plants. Set the upper container on the lid of the nutrient reservoir. Snap the lid on the lower container so that it fits snugly to prevent cross-contamination of rhizobia from other pots. Fit a piece of Tygon tubing into the smaller hole of the lid for replenishment of nutrients. Place a cap over the tubing to reduce the likelihood of contamination. After assembling the two contain-

DINITROGEN FIXATION

1027

Fig. 43-2. Cutaway of modified Leonard jar used in growing legumes for evaluation of N z fixation.

ers, fill the top container with potting material. Vermiculite, perlite, or moderately coarse sand perform well, because they provide adequate capillaries for supply water while remaining well aerated. Hold the cotton wick upright while adding the potting materials so that the wick extends into the potting material. Fill the container to within 2 cm of the top. Prepare the N-free nutrient solution according to the composition provided in Table 12-1 in this book. Wacek and AIm (1978) provide a description of an alternate container. Moisten the potting mixture by pouring nutrient solution onto its surface. Fill the bottom reservoir to within 2 cm of the top. After this initial wetting, water and nutrients are supplied by capillary rise through the cotton wick. To minimize experimental error, plant with high quality seed that has been thoroughly washed and treated (see chapter 12 by Weaver and Graham) so that they are free of any nodulating organisms. Plant seed carefully so they are all at equal depth and equally spaced from other seeds. Plant four to six seeds in each pot for large-seeded legumes and 20 to 30 for small-seeded legumes. After planting, inoculate the seeds by dribbling a suspension of inoculum onto the seed. Use the plant nutrient solution to dilute the inoculum to the desired concentration. Cover the seed and place a 1-cm layer of 5 to 10 mm size gravel on the top of the pot to reduce evaporation and reduce the possibility of air-borne contaminants reaching the rooting zone. After seedlings emerge, thin to one or two seedlings per pot for large-seeded legumes and 10 to 15 per pot for small-seeded plants.

WEAVER & DANSO

1028

Each pot should contain the same number of plants. When thinning, remove both unusually large and small plants. It is best to remove the seedlings within 7 d of emergence by pulling them from the pot and not leaving the root in the pot to contribute a source of N. Care must be exercised not to contaminate other pots because the media clinging to the roots have high populations of nodulating organisms. Grow the plants under conditions that provide temperature and light regimes suitable for good plant growth and nodulation. Grow the plants for at least 1 mo and preferably 6 wk or longer to allow time for nodulation and accumulation of symbiotically fixed N2. At harvest, determine plant dry weight, total plant N, and perhaps total numbers of nodules and nodule mass depending on the experimental objectives. See chapter 12 for comments on nodule storage.

43-2.3 Nitrogen Difference Method-Field 43-2.3.1 Principles Quantitative measurement of N2 fixation of plants grown in the field is challenging because there is no absolute method. Basically, three approaches may be taken: constructing a complete N balance for the system, subtracting the quantity of N accumulated in a plant that does not fix N2 from the plant that fixes N2 (N difference), and isotope dilution. The first method will not be described because it is impractical for most situations (Weaver, 1986) and the third method is described in section 43-6. The primary assumption for the N-difference method is that the N2fixing plant accumulates the same amount of mineral N from the soil as the non-fixing plant. Since roots are not normally harvested the expectation is limited to plant shoots. If plant shoots are to contain the same amount of mineral N from the soil at least two conditions must hold. The two plant types must explore the same soil volume and they must be actively growing during the same time period. It is impractical to attempt to measure small contributions from N2 fixation by this method considering soil variability and experimental error. Probably, N2 fixation amounts of at least 20 kg ha- 1 would be required to be statistically significant (Weaver, 1986). Generally, this method would only be useful for symbiotic N2 fixation. Therefore, the technique described will be for symbiotic systems.

43-2.3.2 Materials 1. 2. 3. 4. 5.

Dinitrogen-fixing plant. Non-N2-fixing control plant. Inoculants (see chapter 12). Field site. Planting equipment.

DINITROGEN FIXATION

1029

43-2.3.3 Procedure For maximum opportunity to differentiate between inoculants, select a field site that is as homogeneous as possible to reduce experimental error and is low in plant available N. If most of the N in the plant comes from mineral N, it will not be possible to distinguish between the quantity of N fixed by different treatments. To reduce the chance for contamination between plots, the site should not be prone to flooding or erosion. It may be advisable to grow a cereal crop on the site to reduce the amount of mineral N present or to incorporate some material with a wide C/N ratio to immobilize the mineral N. It is highly unlikely that differentiation between N2 -fixing ability of strains can be accomplished on a site that contains a resident population of the symbiont. It is advisable to measure the population size of the resident microorganisms capable of nodulating the plant well ahead of planting (see chapter 12). Uninoculated control treatments should be included for each N2 -fixing cultivar used in addition to a non-fixing control plant. The uninoculated control plant may be used to determine the ability of the resident microorganisms to fix N2 and if any inoculation response is achieved. Generally, it is not used as the non-fixing control because it is unlikely that nodules will not form somewhere on the root system by the end of the season. The non-fixing control plant should be well adapted to the soil and climate. For grain legumes a non-modulating isoline may be a good choice. Often a non-nodulating isoline of a particular cultivar will not be available but the non-nodulating plant may still be a good control plant if it has a similar maturity date to the nodulating plant. Choose non-fixing plants that have similar growth characteristics to the fixing plant. For instance, in the case of soybean (Glycine max L.), use a non-fixing plant that is planted in a row and is not unduly tall (shading could be a problem) relative to the soybean or has an unusually large root system. Plot size, planting pattern, and experimental design depend on the particular experiment and field conditions. Generally, single row plots are not used because of potential border affects and increased likelihood of contamination between rows. If the soil is not physically moved between rows by cultivation, erosion or water, it is unlikely that contamination will be a problem. Generally, three or four row plots are used for grain legumes with only the middle row or rows harvested. For accurate seed yield measurements and to reduce experimental error, it is advisable to harvest 3 to 5 m of row. For measurement of N in total shoot biomass it is generally not feasible to harvest more than 1 m of row and it should be done before many of the leaves drop at physiological maturity. In the case of forage or pasture plants, the size of the plots should be approximately 4 m2 . Plots will have to be larger on erodable soils or where water may run across plots to reduce contamination between plots. A cereal or forage grass may serve as the non-fixing control plant. Often the plots will be clipped more than once during the growing season. Thus, the

1030

WEAVER & DANSO

non-fixing plant must be able to regrow following clipping. The centrall m 2 area of the plot should be clipped for measurements. For actual evaluation of biological N2 fixation, the total N contained in plant biomass should be measured. Generally, it is not feasible to measure the N contained in the root system, resulting in underestimation of N2 fixation. Sometimes only seed yield or dry matter production is of interest but it is not possible to infer actual amounts of N fixed in the whole plant from any of these measurements alone. Often, it is useful to make some observations on nodulation. Differences between treatments often occur during the first 3 to 4 wk of plant development. It is relatively easier to dig the root system at this time to observe nodulation than later in the season. Plants from the border rows may be sampled for nodule counts. The coefficient of variation (CV) often is quite large for measurements of nodule numbers and mass.

43-2.3.4 Comments The selection of the non-fixing plant largely determines the accuracy of the estimation. Unfortunately, there is no certain way of selecting the correct control plant or even knowing that the correct plant was selected after the experiment. Consequently, the results of the measurements cannot be considered absolute and some argue that relative differences may not even be correct. The largest potential for error is when approximately 50% of the N in the legume comes from N2 fixation and the least error occurs when < 20% or more than 80% comes from fixation. Some authors argue that more than one non-fixing plant should be used and the results averaged. The true value would still not be known, but presumably the determined value would be closer to the true value. Generally, this approach does not merit the extra labor and space required.

43-3 NITROGEN-IS ISOTOPE TECHNIQUES 43-3.1 Introduction The basis for using isotopes as tracers or measuring biological reactions is that different isotopes of an element possess the same chemical properties even though they differ in atomic mass. Thus, one isotope may be substituted for another in a reaction and the product(s) analyzed for the substituted atom. Isotope identification is made possible through differences in radioactivity or atomic weight (see chapter 39 by Legg et al. in this book). Two isotopes of N, 14N and 15N are the most widely used in agricultural research because they are stable and therefore useful for long-term studies. Their use does not involve the potential risks and health hazards commonly associated with radioisotopes. The major drawbacks, high cost of 15N-Iabeled compounds, along with high equipment and maintenance costs, are no longer as serious as they used to be. By judicious use, the cost

1031

DINITROGEN FIXATION

of 15N-Iabeled fertilizers required for N2 fixation studies can be less than $100 for a greenhouse experiment and, unless several treatments are being studied, approximately $1000 for a field study. The commercial analysis of samples presently ranges between $5 to $10, depending on the number of samples. Two 15N-Iabeling techniques are available for measuring N2 fixation: soil labeling with 15N and of more limited use, incubation in an atmosphere of 15N2 gas. The advantages with the 15N soil-labeling method include the fact that N2 fixation can be measured directly in the field, and integrated estimates of N2 fixed for various time periods can be assessed. The labeling of soil with 15N is currently the most popular and perhaps the most reliable method for measuring N2 fixation in field crops and has been reviewed extensively (Chalk, 1985; Danso, 1988; Hardarson et aI., 1987; Hauck & Weaver, 1986). The 15N soil-labeling method is further classified into the A-value, isotope dilution and 15N natural abundance methods. Respectively, these methods involve: (i) the addition of a higher amount of 15N_ labeled fertilizer to the reference than to the fixing plant, (ii) same amount of a 15N-enriched fertilizer is added to reference and fixing plants, and (iii) no labeled fertilizer is applied. Because the isotope dilution method is less controversial, simpler and more popular than the A-value method, we shall not discuss the A-value method in this chapter. Vose and Victoria (1986) have discussed it. 43-3.2 Isotope Dilution 43-3.2.1 Principle The N in biological systems is composed predominantly of two stable isotopes, 15N and 14N, which constitute about 0.3663 and 99.6337%, respectively, of the N in the atmosphere and natural compounds (Rennie et aI., 1978). The nearly constant 15N/14N ratio in nature makes it possible to use materials with artificially altered 15N/14N ratios to trace biological pathways and quantify N products in biological systems by examining for 15N/14N isotope ratio changes. A material containing more than 0.3663% 15N is referred to as 15N enriched or labeled, and the term atom % 15N excess represents the difference between the atom % 15N in the enriched material and natural abundance (0.3663%). A material containing < 0.3663 atom % 15N is referred to as 15N-depleted. Because 15N-depleted fertilizers are less popular for N 2-fixation studies than 15N-enriched materials, we will only describe the use of 15N-enriched materials. Isotope dilution of 15N occurs when the higher than natural 15N abundance in an enriched material becomes diluted by the lower 15N/14N ratio of a natural substance; the extent of 15N dilution will depend upon the relative 15N enrichments and amounts. Where one of the two 15N compounds (x) being mixed is enriched (a 1 atom % 15N excess) and the other (y) is of 15N natural abundance (zero atom % 15N excess), the final 15N dilution or enrichment (a) can be expressed mathematically as:

WEAVER & DANSO

1032

a

= [x

(a 1)] I (x

+ y)

[1]

To measure N2 fixation by the isotope dilution method, a 15N-enriched fertilizer is added to soil to increase the 15N/14N ratio of available soil N and thus create a difference between the 15N/14N ratios of soil Nand atmospheric N2. In principle, the 15N/14N ratio in a plant that totally depends on soil N reflects the integrated 15N/14N ratio of the available soil N for the period of growth. This is not the case with a N2-fixing plant that, in addition to soil N, assimilates N2; the higher 15N/14N ratio absorbed from the 15N-Iabeled soil will be diluted by the lower 15N/14N of the fixed N2. The relative difference in the 15NI 14N ratio in a reference non-N 2-fixing plant and that of the fixing plant grown under identical conditions is then an indication of the capacity to fix N2, and the following equation (McAuliffe et aI., 1958; Fried & Middelboe, 1977) is used to calculate the proportion (% Ndfa) and amount (Ndfa) of N2 fixed in the fixing crop:

,0 Ndfa = ( 1 -

Of.

Ndfa

=

(1 -

atom % 15N excess in fixing Plant) 15' atom % N excess m reference plant

X

100

atom % 15N excess in fixing Plant) Total N in . x . atom % 15N excess m reference plant fixmg plant

[2]

[3]

Plants absorb variable amounts of N at different stages of their growth, and the 15N/14N ratio in soil generally does not remain constant with time. Thus, the 15N/14N ratio of a soil extract at anyone time or an arithmetic mean from extracts at different times is not a reliable indicator of the integrated 15N/14N ratio of plant absorbed N from a 15N-Iabeled soil. Instead, a reference plant that totally depends on soil N is used. The accuracy of the estimates of N fixed will depend on how the value of 15N/14N assessed by the reference plant accurately reflects the 15N/14N ratio of soil-derived N in the fixing plant. It is important to ensure that the reference plant does not fix N2, and this can be checked by examining for nodulation (if it is a nodulating species) or C2H 4 production from C2H 2 (see sections 43-1 and 43-2). The reference plant should absorb its N from the same N pool as the fixing plant, which may be assessed by examining for similarity in rooting pattern. The fixing and reference plants must also grow and obtain their N in a similar pattern with time, a requirement that can be verified by harvesting the different plants at various growth stages and comparing their growth and N-uptake patterns. In addition, the time of 15N application, planting and harvesting should be the same for fixing and reference plants. More detailed criteria for selecting reference plants have been provided by Fried et al. (1983), Danso et ai. (1986), and Peoples et ai. (1989). Many different species have served as reference plants. These include non-nodulating legume isolines (currently available for grain legumes only), uninoculated legumes (provided the soil is devoid of effective

DINITROGEN FIXATION

1033

homologous rhizobia and cross contamination can be avoided) and several non-legume species (e.g., grasses and many cereal crops). One of the greatest strengths of the 15N methodology has often been overlooked! It can be used to reliably rank plants for their % N derived from N2 fixation on the basis of their relative 15N enrichments (Danso et al., 1986). The lower the atom % 15N excess in a cultivar or species, the greater is the contribution from N2 fixation to the total N in the plant. A potential source of error, the reference plant is eliminated with this approach. It is important to realize, however, that this ranking approach is applicable only for plants grown under similar soil and 15N-enrichment conditions and cannot be used to compare N 2-fixing plants harvested at different times. Regarding the form of 15N-Iabeled fertilizer to apply, there is as yet no universal recommendation or a standard 15N-Iabeled fertilizer (Chalk, 1985). Several different 15N fertilizer forms are being used. The only crucial requirement is, that the relative availability of soil N and the applied fertilizer N should be similar for both reference and fixing plants. The traditional agricultural fertilizers, K 15N03 , (15NH4)zS04 and C0(1sNH2)z have been used. The choice of the 15N fertilizer used is frequently dictated by availability and differences in purchase price. The use of 15N labeled slowrelease N fertilizers is becoming increasingly popular. The incorporation of slow N-release fertilizer formulations results in more stable 15N/14N ratios in soil with time than with the traditional mineral fertilizers (Witty, 1983; Witty & Ritz, 1984). A stable 15N/14N ratio enhances the suitability of many reference plants and thus improves the accuracy of N 2-fixation estimates. Most slow-release formulations labeled with 15N are not commercially available, however, thus restricting widespread usage (Danso, 1988). The alternative is to use 15N immobilized within organic forms, such as by incorporating the residues of plants previously grown on 15N-fertilized soil (Witty & Ritz, 1984; chapters 39 by Legg et al. and 40 by Hauck et al. in this book).

43-3.2.2 Materials 1. Seeds of N 2-fixing plant. 2. Seeds of non-N 2-fixing reference plant. 3. Inoculant containing N2 fixers (see chapter 12 by Weaver and Graham in this book). 4. Nitrogen-labeled fertilizer, 5 to 10 atom % 15N excess. 5. Watering can or knapsack sprayer. 6. Tap water for preparing solutions.

43-3.2.3 Procedure The field site selected should be as homogeneous as possible, otherwise significant treatment differences cannot be observed easily. The N level in the soil may be low or high, depending on the objective of the

1034

WEAVER & DANSO

study. The sensitivity of the isotope-dilution method, however, is greater at high than at low levels of N z fixation (Danso & Kumarasinghe, 1990), and the depressive effect of Non N z fixation should be considered. Because the precision of 15N-derived data is generally higher than yield-dependent data (Hardarson et aI., 1984), isotope plots do not need to be as large as yield plots for the same precision. Isotope plot sizes ranging from 1 to 5 mZ , consisting of four to five rows for grain crops, and for forage and pasture crops, 1 to 2 mZ plots are satisfactory. A randomized split plot design is often convenient. The advantage with this design is that it allows contrasting treatments such as different N rates and rhizobial inoculation treatments to be isolated within separate blocks, thus minimizing effects due to erosion, and cross contamination. For trees, individual trees have frequently served as single plots. This has been dictated by cost and labor considerations as a result of the massive sizes of trees. Many multi-purpose trees are, however, small enough (especially when they are young) to allow using a few trees per plot. To quantify N z fixation, a reference plant should be used. When genotypes with different maturity periods are being compared, a reference plant to match each maturity group is necessary or the 15N/14N ratio of mineral N must be constant. To avoid large errors due to soil variability, it is advisable to plant reference plants in close proximity to fixing plants, possibly on adjacent plots. When, how, and how much of the 15N label to apply to soil are influenced by the type of 15N-Iabeling material used and the sensitivity of the instrument for analyzing 15N. If a conventional mineral fertilizer is used, it can be applied to soil just prior to planting or immediately after thinning seedlings. In addition to allowing thinning to be done before 15N application, post-planting 15N application allows for 15N application to be stopped or delayed if seeding emergence is poor or if replanting becomes necessary. Replanting should be done on all plots; otherwise serious errors will result due to the imposed difference in times of initial 15N uptake among plants in the different treatments. Post-planting 15N application requires greater care during application to prevent damaging plants, or to avoid serious errors from mixing of soil and rhizobia under different treatments and must be accomplished during the seedling stage. For ease of application and to ensure a fairly homogeneous distribution of the normally small amounts of 15N-Iabeled fertilizer used, it is advisable to dissolve the appropriate amount of the 15N-Iabeled fertilizer in water and spray the fertilizer solution onto plots (see chapter 39). It may even be advisable to prepare stock solutions from which the required aliquots are taken. The spraying may be done with a small knapsack sprayer, a normal garden watering can, or plastic squeeze bottles with perforated lids. A trial run is needed to decide on the most appropriate watering rate that will moisten the soil but not puddle it (usually between 100 to 500 mL m- z depending on soil type) is required. It is important to consider the type of instrument available when deciding how much 15N to apply to soil (see chapter 39). High precision dual

DINITROGEN FIXATION

1035

inlet mass spectrometers that switch back and forth from measuring a reference gas and the sample in question can measure 15N abundance with a precision of 0.00004 atom %. Thus, an enrichment of 0.0001 would be adequate to indicate 15N uptake. Crasswell and Eskew (1991) have recently compared several different types of commercially available instruments for precision of 15N analysis. With a single inlet mass spectrometer, an enrichment of 0.004 atom % 15N would be necessary and a similar 15N enrichment would be needed with an automated N analyzer coupled to a mass spectrometer. With an emission spectrometer, an enrichment of 0.01 atom % would be needed. Approximately 1 kg of 15N ha- 1 (10 kg of N ha- 1 of 10 atom % 15N excess or 20 kg of N ha -1 of 5 atom % 15N excess) is adequate for detection in most single-year isotope dilution experiments using most analytical systems. Only in exceptional cases (e.g., where N2 fixation was exceedingly high or if the emission spectrometer used is of very low sensitivity) would 1.5 to 2 kg of 15N ha- 1 (equivalent to 0.15-0.2g m- 2) have to be applied. For grain legumes and seasonal crops, a one-time 15N application is generally satisfactory. In the case of perennials, such as pastures and N 2fixing trees, it is recommended to split the required 15N fertilizer in equal amounts and apply at quarterly, half yearly or yearly intervals (Danso et aI., 1988). Where applicable, the 15N additions can be made to coincide with harvests (Vallis et aI., 1967). In case 15N-Iabeled plant residues are used for soil labeling, these should preferably be chopped into small pieces (2-5 cm pieces) and rototilled into soil (chapters 38 and 39). Sometimes the labeling can be achieved in situ, by rototilling plants growing on 15N_ labeled soil back into the soil (Fried et aI., 1983). In all cases, a greener plant residue with a C/N ratio closer to that of soil than using a matured crop is preferable. The high C/N ratios in fully matured crops, especially in cereals, could immobilize soil N and result in unusually high levels of N2 fixation (Papastylianou & Danso, 1991). In addition to being planted at the same time, it is necessary to harvest both fixing and reference plants at the same time. This means that for crops of different maturity groupings, matching reference plants are needed for each. Because of problems associated with quantitative recovery of roots especially in field-grown plants, most estimates of N2 fixation are on aboveground plant parts. It is thus assumed that total N or N2 fixed in roots constitutes a small fraction of the total N in whole plants. This assumption may not be true for all plants. Some trees have been shown to contain more N belowground than above (Sanginga et aI., 1990), and the N partitioned to belowground parts of some pastures can be substantial (Heichel et al., 1981). Thus, wherever possible, roots should also be harvested. For greenhouse-grown plants and some field experiments, the freshly harvested plant sample is small enough to be handled in a single bulk for drying and grinding. With bulky samples, it is not easy to get homogeneous subsampIes when entire plants are involved (chapter 40). It is advisable to split the freshly harvested plants into different plant parts for subsampling. The different parts are then dried separately in the oven at 70°C for 48 h,

WEAVER & DANSO

1036

and ground through a l00-mesh size sieve. One approach is to analyze separately each plant part for Nand 15N/14N isotope ratios, and use the following equation (not an arithmetic mean) to calculate the overall or weighted atom % 15N excess to compute N fixed for the whole plant (Eq. [4]): Weighted atom % 15N excess ::;: AE(A)

X

TN (A) + AE(B) x TN (B) + AE(C) x TN (C) etc. TN (A + B + C)

[4]

Where AE is atom % 15N excess, TN is total N, and A,B, and C represent three harvested plant parts. This approach results in many samples being analyzed. An easier and reliable approach involves drying and grinding the individual plant parts separately as described above, but subsequently mixing to get a composite sample for Nand 15N/14N analysis. From the dry weights of the individual parts before grinding, the relative dry weight ratios to use for preparing the composite mixture for Nand 15N/14N ratio analysis can be calculated. It is easier to get more homogeneous mixtures from dried, ground material than from fresh material. 43-3.3 Nitrogen-1S Natural Abundance Method 43-3.3.1 Principles The 15N natural abundance method operates on the same principle as the isotope dilution method (43-3.2.1). The only difference is that with the 15N natural abundance method, no 15N-enriched material is added to soil. In this case, the higher 15N/14N ratio in soil than atmospheric N2 is the consequence of natural N-cycle processes. During N turnover reactions in soil, 14N is preferentially lost into the atmosphere compared to 15N, creating a natural 15N enrichment of soil N relative to atmospheric N2 (Shearer & Kohl, 1986). It is important to note that the differences between the 15N/14N ratios of soil N and atmospheric N2 are very small, being several orders of magnitude lower than where 15N-enriched materials have been added. These small differences are simplified by multiplying by a thousand and expressed in b units (Amarger et aI., 1979). Thus b15N::;: (R sample - R standard) x 1000 , R standard

[5]

15N

where R

= 14N + 15N

[6]

and R standard::;: R air ::;: approximately 0.0003663, i.e., 1 b 15N unit ::;: 0.00037 atom % b 15N excess, compared to a value of 2730 b units contained in 1 atom % 15N excess. The b 15N units for most soils are between -2 and +15.

DINITROGEN FIXATION

1037

The major drawback with the lsN natural abundance method is that of low sensitivity. To be able to accurately measure these small differences in lsN, the analytical procedures have to be precise (Peoples et aI., 1989). Care must be taken at each step of sample preparation to assure complete recovery of N. Also, the measurement requires a dual inlet, dual collector mass spectrometer equipped with a mode for switching between the gas sample and reference gas. The method is of limited use on soils with extremely low olsN. Such soils are commonly found under natural vegetation (Hansen & Pate, 1987). Here, the variability in 015N may be large enough to obscure any differences between olsN of soil N and atmospheric N 2, and the lsN natural abundance method should not be used. Another potential problem with the 15N natural abundance method is that isotopic discrimination occurring between lsN and 14N within the plant can cause significant errors in estimates of N2 fixation. This is not a serious problem with the isotope dilution method because the isotopic discrimination effect is obscured by the high lsN enrichment used. A procedure devised to correct for isotopic discrimination at the lsN natural abundance level involves analyzing for the lsN/14N ratio in a control fixing plant grown in a N-free medium (see section 43-3.3.3). The 15N natural abundance method, despite the above-stated problems, has provided several useful estimates of N2 fixation. The method has particular merit for studying N2 fixation in systems that should not be disturbed through the application of 15N fertilizer (Danso et aI., 1992). Where the olsN values allow, the method can provide useful information on N2 fixation in forest trees, and, in addition to some quantitative measurements, provide useful evidence of the potential of some species with unknown N2-fixing ability (Shearer & Kohl, 1986). The method's added value in such exploratory studies is that reference plants may not be needed; differences in the b 15 N values of putative N2 can be used to rank them. The use of the 15N natural abundance method does not require the purchase of 15N fertilizers, and the possible suppressive effect of added lsN-labeled fertilizer on N2 fixation is eliminated. 43-3.3.2 Materials

1. 2. 3. 4. 5. 6. 7.

Seeds of N 2-fixing plant. Seeds of non-N2-fixing reference plant. Inoculant containing N2-fixing microorganisms. Complete nutrient solution minus N (see Table 43-1). Plastic pots (1 L size or bigger). Sieved (2 mm diam.) soil collected from 0-15 cm soil layer. Hydroponic system.

43-3.3.3 Procedure Because of the small differences in lsN abundance being measured, the need to select a homogeneous site is crucial. The lsN natural abundance method will work only in soils in which the variation in olsN is

WEAVER & DANSO

1038

smaller than differences in N2 fixed among treatments. For establishing the potential ability of species to fix N2, measurements of the 6 15N in leaves or aboveground samples are sufficient (Shearer et al., 1983). A lower 6 15N compared to putative non-N2 fixers growing in the same locality suggests N2 fixation. For measuring N2 fixation by the 15N natural abundance method, two types of reference plants are needed. One type is for assessing the 15N/14N ratio of soil N. The reference crop selection in this case is similar to that described for the isotope-dilution technique (see section 43-3.2.1), except in established natural stands, where one is compelled to rely on non-N2fixing plants occurring naturally within the locality. To reduce uncertainty of appropriateness of the reference plant and thus the probability of serious errors, it is advisable to use the mean of more than one of the naturally occurring non-Nz-fixing plant species rather than use only one of them as a reference. Another reference is needed for measuring the discrimination between 14N and 15N that occurs during N2 fixation (Amarger et al., 1979; Shearer & Kohl, 1986). This is achieved by planting the inoculated N2fixing plant hydroponically or in washed sand watered regularly with Nfree medium (Amarger et al., 1979) to ensure that the plant is completely dependent on N2 fixation. The analysis of Nand 15N/14N ratios after harvest should be done either on individual plant parts or on composite plant samples (see section 43-3.2.3), but the estimates should reflect N2 fixed in the whole plant (see section 43-3.2.3). A slightly modified isotope-dilution equation is used to calculate N2 fixation as follows: % Ndfa

= (x - y)/(x - f) x 100

[7]

where x y f

= 615N of non-fixing reference plant = 6 15N in the fixing plant grown in soil = 6 15N in the fixing plant grown hydroponically. 43-4 USE OF DINITROGEN·15 GAS 43-4.1 For Measuring Dinitrogen Fixation

43-4.1.1 Principles The detection of 15N in tissues of biological systems exposed to 15N2 gas is the only direct, unequivocal method for demonstrating that N2 fixation occurred. Uptake of 15N from 15N2 gas also allows the fate of fixed or absorbed N to be followed in various intermediates and products within plants. To measure uptake of 15N2 gas, it is necessary to enclose the bio-

DINITROGEN FIXATION

1039

logical system in a gas-tight container with a sufficient enrichment of 15N and for long enough to be able to detect 15N enrichment within the biological system. A major disadvantage of the 15N2 gas exposure method is that it usually involves short-term assays, and higher plants, in particular, require incubation conditions quite different from those of normal field-grown crops. Thus, the major uses of 15N2 are to demonstrate N2 fixation in systems where activity is expected based on other indirect methods, to determine the compounds into which N is incorporated and to follow N transport in plant-microbe systems. Another use has been to determine calibration constants for C2H 2 reduction assays. There is no standard incubation chamber, and several types have been used depending on the N2-fixing system and the purpose of the study. A principal requirement is that conditions within the chamber should match as closely as possible the in situ conditions. For small biological systems such as bacterial cultures or enzymes, a serum bottle or test tube with a serum stopper is adequate (Spiff & Odu, 1972). For intact plants, a desiccator or a perspex chamber (Ross et aI., 1964) may be constructed. To avoid rapid changes in the composition of the atmosphere during assay of plants, Knowles (1980) has suggested a ratio of gas phase volume (mL) to sample fresh weight (g) of at least 300. Too large containers also are undesirable; more 15N2 is required, making the study unduly expensive. In some cases, when only the root needs to be exposed to 15N2, the pot may serve as an incubation chamber after sealing around the stem and the 15N2 may be injected directly into the rhizosphere (Douglas & Weaver, 1986). The 15N2 gas may be purchased already prepared from several suppliers (e.g., Merck Sharp and Dohme, Montreal, Canada and Monsanto Research Corp., Miamisburg, OH). It is also possible to generate the 15N2 in the laboratory. Ammonium sulphate enriched with 15N is placed in an evacuated bottle, and LiOBr is injected to generate 15N2 which is collected in a connected evacuated flask. The 15N2 gas is then transferred to a flask containing an acidified solution to remove any traces of NH3 before use. See Bergersen (1980a) for a detailed description of an apparatus for preparing 15N2 from a 15N4 salt. However, unless many studies are planned, it is probably easier and cheaper to purchase the 15N gas. The 15N enrichments in purchased gases have generally varied from around 40 to 90 atom % 15N excess. The required enrichment is dictated by the 15N measuring equipment, volume of the exposure chamber, the N2-fixing activity of the system, duration of exposure, and other available sources of N. In most systems, a final 15N enrichment in the gas phase of 10 to 50 atom % 15N excess should be adequate. It is necessary to accurately know the 15N enrichment of the gas phase within the incubating chamber to be able to calculate the proportion and amount of N fixed. Thus, the gas within the chamber must be sampled at the beginning and preferably also at the end (using the sampling ports) of the incubation for precise 15N determination. The equation for calculating N fixed is:

1040

WEAVER & DANSO

Ndfa

=

Total N . fix' [8] Atom % 15N excess in tissues of fixing system 15 . . . x m mg Atom % N m gas sample wlthm chamber system (g)

The length of time necessary to detect 15N enrichment depends on the N2-fixing activity of the system, the 15N enrichment of the gaseous atmosphere, and the sensitivity of the analytical technique used to determine 15N abundance. With active biological systems, such as the Azolla-Anabaena symbiosis, exposure times of a few minutes with an atmosphere enriched to 10 atom % 15N excess are adequate. On the other end of the scale, for associative symbioses between grasses and heterotrophic bacteria with low levels of N2 fixation, exposure times of several days with 50 atom % 15N excess or more may be needed (Eskew et aI., 1981). The simplest N2-fixing systems including soil samples and bacterial cultures require in addition to the 15N2 gas, an oxygen-scavenging system. For the lower plants (e.g., blue-green algae), only a light source may be necessary. Higher plants normally have to be exposed to 15N2 for prolonged periods to obtain valid estimates of N2 fixation. Also, the composition of gases changes rapidly due to consumption of CO2 for photosynthesis and the liberation of O 2, Chambers for higher plants, therefore, required supplementary set up with adequate controls for light, CO2, O 2, gas circulation, water, and temperature control. Such an apparatus has been described by Witty and Day (1978) for laboratory investigations, but a simpler apparatus was used by Montange et al. (1981). Ideally, the N2 partial pressure should be in the range 0.8 to 1.0 and certainly ~0.4 to saturate N2-fixing sites (Knowles, 1980). The CO2 and O2 concentrations should be similar to those found in nature, that is, 0.03 and 20%, respectively. Instead of sampling for CO2 and O 2 and supplementing through sampling ports, solenoid valves that allow for the automatic analyses and addition of CO2 and O 2 (when needed) in the circulating gas may be installed. 43-4.1.2 Materials 1. Wide mouth laboratory bottle, approximately 40-mm diam. and 60-mL capacity. 2. Serum stopper seal to fit bottle. 3. Plastic syringe fitted with valve, 50 mL. 4. Nitrogen-free nutrient solution (see Table 12-1). 5. Azolla plant. 6. Dinitrogen gas. 7. Mass or emission spectrometer. 43-4.1.3 Procedure Azolla has been selected because its high rate of growth and N2 fixation make it possible to detect 15N uptake after short exposure periods of

1041

DINITROGEN FIXATION

< 1 h. Within this period rapid changes in the gaseous composition and growth conditions are unlikely, and relatively simple incubation chambers and conditions can thus be used. A laboratory bottle is only one of the many containers that can be used. Place 1 g of a growing culture of Azolla together with 9 mL of nutrient solution into a 60-mL bottle and cap tightly with a serum stopper. Pierce the stopper and withdraw 20 mL of air from the chamber with a 50-ml syringe. Refill with 20 mL of 15N2 purchased from a commercial source or generated in the laboratory (see section 43-4.1.1). It is also possible to pre-evacuate the container, flush a few times with He and refill with a pre-mixed air having the required N2 pressure and 15N enrichment. To estimate the required 15N enrichment, use the isotope-dilution equation (see Eq. [1], section 43-3.2). The "y" in the equation is the total N within Azolla at incubation, "x" is the increment in total N during the period of incubation, "a1" is the required 15N enrichment in the chamber and "a" the 15N enrichment in Azolla after harvest. For a safe level of detection by both mass and emission spectrometry, aim for 0.1 atom % 15N excess in Azolla. Assuming growth is under optimal conditions, Azolla can double its biomass within 3 to 5 d (avg. 4 d). Since there are 96 h in 4 d, the increase in Azolla biomass and N content within a 1-h incubation period would be about 1% of the initial N. Thus: [0.01 (a1)] 1(0.01 + 1) and a1 =

=

0.1

(0.1 x 1.01) 0.01 = 10.1 atom % 15N excess

Because the chamber was not evacuated, a higher 15N enrichment will be required to compensate for dilution from 14N pre-existing in the chamber. Again, the isotope-dilution equation is used to obtain the required 15N enrichment. The air space available in the chamber is approximately 50 mL, 20 of which is occupied by 15N2. that is, [20 (al)] 1(20 + 30) = 10.1 a 1 = 10.1 x 50/20 = 25.2% Therefore, the 20 mL of N2 to be injected should contain a 15N enrichment of approximately 25 atom % 15N excess to give a final 15N enrichment of about 10 atom % 15N excess in the chamber. It is necessary to know accurately the 15N enrichment in the chamber. For this, a sample gas has to be analyzed at the beginning and at the end of incubation. Remove the Azolla plant from the container after 1 h incubation. Dry, grind, and analyze the tissues for 28N, 29N, and 30N composition. It is essential that all three molecular species be determined since distribution of the three ionic species in the head space is not in equilibrium (Knowles, 1980).

1042

WEAVER & DANSO

43-4.2 Calibration of Acetylene-Reduction Method 43-4.2.1 Principles In ecological and agricultural studies where the goal is to measure the input of N, it is necessary to convert AR values obtained to estimates of N fixed, usually expressed in kg of N ha- 1 over a given period. Based on theoretical comparison of electron flow, Hardy et al. (1968) suggested a conversion factor of 3:1 for C2HzlN2 (see section 43-1.1). There are, however, several reasons why the simple theoretical ratio is inadequate and should not be used. Acetylene is approximately 60 times more soluble in water than N2; thus, the enzyme is much more readily saturated with CzH2 than with N2. Nitrogenase also catalyzes the reduction of protons to H2 gas when N2 is the substrate. On the other hand, in the presence of 0.1 atmosphere CzH2' the reduction of protons is almost completely inhibited, and all of the electron flow is diverted to reduction of CzH2. Determined conversion ratios have thus varied widely (see section 43-1.1), and the use of theoretical conversion factor can result in serious errors (Rennie et al., 1978). The most reliable or preferred method for calibrating the AR method is the incorporation of 15N from 15N2 under similar conditions. In this exercise, a CzH2/N 2 conversion ratio will be determined for the AzollaAnabaena symbiosis.

43-4.2.2 Materials 1. 2. 3. 4. 5.

Materials listed in section 43-4.1.2. Source of purified CzH2 (see sections 43-1.2 and 43-1.2.3). Ethylene for preparation of standards. Vacutainers, 5 mL. Gas chromatograph fitted with appropriate column (see section 43-1.2.2). 6. Nitrogen carrier gas. 7. Emission or mass spectrometer.

43-4.2.3 Procedure Incubate the Azolla in an atmosphere containing 10% CzH2 as described in section 43-1.2.3. Analyze for CzH2 and C2~ on a gas chromatograph and harvest after 1 h. Calculate JUIloles CzH4 produced per gram of dry weight Azolla. For the 15N2 incubation, use the same procedures described in sections 43-4.1.2 and 43-4.1.3. The following equations are used to calculate the amount and I-tmoles of N2 fixed: I-tg N2 fixed

=

atom % 15N excess in Azolla atom % 15N excess in gas phase x total N in Azolla (mg)

[9]

DINITROGEN FIXATION

1043

f.tg N2 fixedl28

= f.tIlloles N2

[10]

The f.tmoles N2 fixed is then compared with f.tmoles c;H4 produced to arrive at the actual conversion ratio.

REFERENCES Amarger, N., A. Mariotti, F. Mariotti, J.e. Durr, C. Bourguignon, and B. Lagacherie. 1979. Estimate of symbiotically fixed nitrogen in field grown soybeans using variations in 15N natural abundance. Plant Soil 52:269-280. Arp, D.J. 1992. Hydrogen cycling in symbiotic bacteria. p. 432-460. In G. Stacey et al. (ed.) Biological nitrogen fixation. Chapman and Hall, New York. Bergersen, F.J. 1980a. Measurement of nitrogen fixation by direct means. p. 65-110. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen fixation. John Wiley and Sons, New York. Bergersen, F.J. 1980b. Methods, accidents, and design. p. 3-10. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen fixation. John Wiley and Sons, New York. Boddey, R.M. 1987. Methods for quantification of nitrogen fixation associated with Gramineae. CRC Crit. Rev. Plant Sci. 6:209-266. Brockwell, J. 1980. Experiments with crop and pasture legumes-Principles and practice. p. 417-488. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen fixation. John Wiley and Sons, New York. Burns, R.C., and R.W.F. Hardy. 1975. Nitrogen fixation in bacteria and higher plants. Springer-Verlag, New York. Burris, R.H. 1974. Methodology. p. 10-33. In A. Quispel (ed.) The biology of nitrogen fixation. American Elsevier Publ. Co., New York. Burris, R.H. 1975. The acetylene reduction technique. p. 249-258B. In D.P. Stewart (ed.) Nitrogen fixation by free-living micro-organisms. International Biological Programme. Vol. 6. Cambridge Univ. Press, New York. Chalk, P.M. 1985. Estimation of N2 fixation by isotope dilution: An appraisal of techniques involving 15N enrichment and their application. Soil BioI. Biochem. 17:389-410. Crasswell, E.T., and D.L. Eskew. 1991. Nitrogen and nitrogen-15 analysis using automated mass and emission spectrometers. Soil Sci. Soc. Am. J. 55:750-756. Danso, S.K.A. 1988. The use of 15N enriched fertilizers for estimating nitrogen fixation in grain and pasture legumes. p. 345-358. In D.P. Beck and L.A. Materon (ed.) Nitrogen fixation by legumes in medIterranean agriculture. Martinus Nijhoff Publ., Dordrecht, Netherlands. Danso, S.K.A., G.D. Bowen, and N. Sanginga. 1992. Biological nitrogen fixation in trees in agro-ecosystems. Plant Soil 141:177-196. Danso, S.K.A., G. Hardarson, and F. Zapata. 1986. Assessment of dinitrogen fixation potentials of forage legumes with 15N techniques. p. 26-57. In I. Haque et al. (ed.) Proceedings workshop on potentials of forage legumes in farming systems of subSaharan Africa. International Livestock Centre for Africa (ILCA), Addis Ababa, Ethiopia. Danso, S.K.A., G. Hardarson, and F. Zapata. 1988. Dinitrogen fixation estimates in alfalfaryegrass swards using different nitrogen-15 labeling methods. Crop. Sci. 28:106-110. Danso, S.K.A., and K.S. Kumarasinghe. 1990. Assessment of potential sources of error in nitrogen fixation measurements by the nitrogen-15 isotope dilution technique. Plant Soil 125:87-93. Douglas, L.A., and R. W. Weaver. 1986. Partitioning of nitrogen-15-labeled biologically fixed nitrogen and nitrogen-15-labeled nitrate in cowpea during pod development. Agron. J. 78:499-502. Eskew, D.L., A.R.J. Eaglesham, and A.A. App. 1981. Heterotrophic 15N2 fixation and distribution of newly fixed nitrogen in a rice-flooded soil system. Plant Physiol. 68: 48-52. Fried, M., S.K.A. Danso, and F. Zapata. 1983. The methodology of measurement of N2 fixation by non-legumes as inferred from field experiments with legumes. Can. J. Microbiol. 29:1053-1062. Fried, M., and V. Middelboe. 1977. Measurement of amount of nitrogen fixed by a legume crop. Plant Soil 47:713-715.

1044

WEAVER & DANSO

Gerbaud, A. 1990. Effect of acetylene on root respiration and acetylene reducing activity in nodulated soya bean. Plant Physiol. 93:1226-1229. Giller, K.E. 1987. Use and abuse of the acetylene reduction assay for measurement of "associative" nitrogen fixation. Soil BioI. Biochem. 19:783-784. Hansen, A.P., and J.S. Pate. 1987. Evaluation of the 15N natural abundance method and xylem analysis for assessing N2 fixation of understorey legumes in Jarrah (Eucalyptus marginata Donn ex Sm.) forest in S.W. Australia. J. Exp. Bot. 38:1446-1458. Hardarson, G., S.K.A. Danso, and F. Zapata. 1987. Biological nitrogen fixation in field crops. p. 165-192. In B.R. Christie (ed.) Handbook of plant science in agriculture. CRC Press, Boca Raton, FL. Hardarson, G., F. Zapata, and S.K.A. Danso. 1984. Field evaluation of symbiotic nitrogen fixation by rhizobial strains using 15N methodology. Plant Soil 82:369-375. Hardy, R:W.F., R.C. Burns, and RD. Holsten. 1973. Applications ofthe acetylene-ethylene assay for measurement of nitrogen fixation. Soil BioI. Biochem. 5:47-81. Hardy, R.W.F., R.D. Holsten, E.K. Jackson, and R.C. Burns. 1968. The acetylene-ethylene assay for N2-fixation: Laboratory and field evaluation. Plant Physiol. 43:1185-1207. Hauck, R.D., and RW. Weaver. 1986. Field measurement of dinitrogen fixation and denitrification. Spec. Publ. 18. SSSA, Madison, WI. Heichel, G.H., D.K. Barnes, and C.P. Vance. 1981. Nitrogen fixation of alfalfa in the seeding year. Crop Sci. 21:330-335. Herdina, and J .H. Silsbury. 1990. Estimating nitrogenase activity of faba bean (Vicia faba L.) by acetylene reduction (AR) assay. Aust. J. Plant Physiol. 17:489-502. Hyman, R., and D.J. Arp. 1987. Quantification and removal of some contaminating gases from acetylene used to study gas utilizing enzymes and microorganisms. Appl. Environ. Microbiol. 53:298-303. Knowles, R. 1980. Nitrogen fixation in natural plant communities and soils. p. 557-582. In F.J. Bergersen (ed.) Methods for evaluating biological nitrogen fixation. John Wiley and Sons, New York. Leonard, L.T. 1943. A simple assembly for use in the testing of cultures of rhizobia. J. Bacteriol. 45:523-525. Lindstrom, K. 1984. Analysis of factors affecting in situ nitrogenase (C2H 2 ) activity of Galega orientalis, Trifolium pratense, and Medicago sativa in temperate conditions. Plant Soil 79:329-341. McAuliffe, C., D.S. Chamblee, H. Uribe-Arango, and W.W. Woodhouse, Jr. 1958. Influence of inorganic nitrogen on nitrogen fixation by legumes as revealed by 15N. Agron. J. 50:334-337. McNeill, A.M., J.E. Sheehy, and D.S.H. Drennen. 1989. The development and use of a flow-through apparatus for measuring nitrogenase activity and photosynthesis in field crops. J. Exp. Bot. 40:187-194. Mederski, H.J., and J.G. Streeter. 1977. Continuous, automated acetylene reduction assays using intact plants. Plant Physiol. 59:1076-1081. Minchin, F.R., J.E. Sheehy, 1)nd J.F. Witty. 1986. Further errors in the acetylene reduction assay: Effects of plant disturbance. J. Exp. Bot. 37:1581-1591. Montange, D., F.R. Warembourg, and R. Bardin. 1981. Utilisation du 15N2 pour estimer la fixation d'azote et sa repartition chez les legumineuses. Plant Soil 63:131-139. Morris, D.R, D.A. Zuberer, and R.W. Weaver. 1985. Nitrogen fixation by intact grass-soil cores using 15N2 and acetylene reduction. Soil BioI. Biochem. 17:87-91. Nohrstedt, Hans-Organ. 1983. Natural formation of ethylene in forest soils and methods to correct results given by the acetylene-reduction assay. Soil BioI. Biochem. 15:281-286. Papastylianou, I., and S.K.A. Danso. 1991. Nitrogen fixation and transfer in vetch and vetch-oats mixtures. Soil BioI. Biochem. 23:447-452. Peoples, M.B., A.W. Faizah, B. Rerkasem, and D.F. Herridge. 1989. Methods for evaluating nitrogen fixation by nodulated legumes in the field. Monogr. 11. Australian Centre for Int. Agric. Res., Canberra. Rennie, RJ., D.A. Rennie, and M. Fried. 1978. Concepts of 15N usage in dinitrogen fixation studies. p. 107-133. In Isotopes in biological dinitrogen fixation. Proceedings of an Advisory Group Meeting, Vienna. STIIPUB/478. IAEA, Vienna. Ross, P.J., A.E. Martin, and E.F. Henzell. 1964. A gas-tight growth chamber for investigating gaseous nitrogen changes in the soil:plant:atmosphere. Nature (London) 204: 444-447.

DINITROGEN FIXATION

1045

Sanginga, N., F. Zapata, S.K.A. Danso, and G.D. Bowen. 1990. Effect of successive cuttings on uptake and partitioning of 15N among plant parts of Leucaena leucocephala. BioI. Fertil. Soils 9:37-42. Shearer, G., and D.H. Kohl. 1986. Nz-fixation in field settings: Estimations based on natural 15N abundance. Aust. J. Plant Physiol. 13:699-756. Shearer, G., D.H. Kohl, R.A. Virginia, B.A. Bryan, J.L. Skeeters, E.T. Nilsen, M.R. Sharifi, and P.W. Rundel. 1983. Estimates of Nz-fixation from variation in the natural abundance of 15N in Sonoran Desert Ecosystems. Oecologia 56:365-373. Sheehy, J.E. 1991. Theory of a crop enclosure system for measuring nitrogen fixation, photosynthesis, respiratlOn, and biological processes in the soil. Ann. Bot. 67:123-130. Sheehy, J.E., F.R. Minchin, and J.F. Witty. 1983. Biological control of the resistance to oxygen flux in nodules. Ann. Bot. 52:565-571. Somasegaran, P., and H.J. Hoben. 1985. Methods in legume-Rhizobium technology. NIFTAL Project, Univ. of Hawaii, P.O. Box 0, Paia, Maui, HI. Spiff, E.D., and e.T.1. Odu. 1972. An assessment of non-symbiotic nitrogen fixation in some Nigerian soils by the acetylene reduction technique. Soil BioI. Biochem. 4:71-77. Tann, C.C., and J. Skujins. 1985. Soil nitrogenase assay by 14C-acetylene reduction: Comparison with the carbon monoxide inhibition method. Soil BioI. Biochem. 17:109-112. Tough, H.J., and J.R. Crush. 1979. Effect of grade of acetylene on ethylene production by white clover (Trifolium repens L.) during acetylene reduction assays of nitrogen fixation. N. Z. J. Agric. Res. 22:581-583. Vallis, I., K.P. Haydock, R.J. Ross, and E.F. Henzell. 1967. Isotopic studies on the uptake of nitrogen by pasture plants. III. The uptake of small additions of 15N-Iabeled fertilizer by Rhodes grass and Townsville lucerne. Aust. J. Agric. Res. 18:865-877. Vincent, J.M. 1970. A manual for the practical study of root-nodule bacteria. Int. BioI. Programme Handb. 15. Blackwell Scientific Publ., Oxford. Vose, P.B., and R.L. Victoria. 1986. Re-examination of the limitations of nitrogen-15 isotope dilution technique for the field measurement of dinitrogen fixation. p. 23-41. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. Soil Sci. Soc. Am. Spec. Publ. 18. SSSA, Madison, WI. Wacek, T.J., and D. Aim. 1978. Easy-to-make "Leonard Jar." Crop. Sci. 18:514-515. Weaver, R.w. 1986. Measurement of biological dinitrogen fixation in the field. p. 1-10. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. Soil Sci. Soc. Am. Spec. Publ. 18. SSSA, Madison, WI. Weaver, R.W., S.F. Wright, M.W. Varanka, O.E. Smith, and E.C. Holt. 1980. Dinitrogen fixation (CzHz) by established forage grasses in Texas. Agron. J. 72:965-968. Weisz, P.R., and T.R. Sinclair. 1988. Soybean nodule gas permeability, nitrogen fixation and diurnal cycles in soil temperature. Plant Soil 109:227-234. Witty, J .F., and J .M. Day. 1978. Use of 15Nz in evaluating asymbiotic Nz fixation. p. 135-150. In Isotopes in biological dinitrogen fixation. Proceedings of an Advisory Group Meeting, Vienna. STlIPUB/478. IAEA, Vienna, Austria. Witty, J.F. 1979. Overestimate of Nz-fixation in the rhizosphere by the acetylene reduction method. p. 137-144. In J.L. Harley and R.S. Russell (ed.) The soil-root interphase. Academic Press, London. Witty, J.F. 1983. Estimating Nz-fixation in the field using 15N-labeled fertilizer: Some problems and solutions. Soil BioI. Biochem. 15:631-639. Witty, J.F., and K. Ritz. 1984. Slow-release 15N fertilizer formulations to measure Nz-fixation by isotope dilution. Soil BioI. Biochem. 16:657-661. Witty, J.F., and F.R. Minchin. 1988. Measurement of nitrogen fixation by the acetylene reduction assay; myths and mysteries. p. 331-334. In D.P. Beck and L.A. Materon (ed.) Nitrogen fixation by legumes in mediterranean agriculture. Martinus Nijhoff Publ., Dordrecht, the Netherlands. Wynne, J.e., F.A. Bliss, and J.C. Rosas. 1987. Principles and practice of field designs to evaluate symbiotic nitrogen fixation. p. 371-389. In G.H. Elkan (ed.) Symbiotic nitrogen fixation technology. Marcel Dekker, New York. Zuberer, D.A., and D.B. Alexander. 1986. Effects of oxygen partial pressure and combined nitrogen on Nz-fixation (CzHz) associated with Zea mays and other gramineous species. Plant Soil 90:47-58.

Published 1994

Chapter 44 Measuring Denitrification in the Field A. R. MOSIER, USDAIARS, Fort Collins, Colorado

LEIF KLEMEDTSSON, IVL, Goteborg, Sweden

Denitrification is the stepwise process by which N0 3- is reduced to N2:

The process can both be microbial and chemical, but the microbial process dominates in most soils (Broadbent & Clark, 1965). Microbial denitrification is an alternative respiration process by bacteria in the soil (Focht & Verstraete, 1977) that can use N0 3- under 02-limiting conditions. These bacteria prefer to use O 2 as their electron acceptor, if it is available (see chapter 14 by Tiedje in this book). To measure denitrification in the field, the large temporal and spatial variation of the process must be considered (Folorunso & Rolston, 1984). This requires an understanding of the factors controlling the process. Soil atmosphere O 2 concentration is the main factor controlling denitrification in soils that are not water-logged (Tiedje, 1988). Oxygen status of the soil depends on the O 2 diffusion rate and the consumption rate of the gas in the soil. The diffusion rate depends upon temperature, soil structure, and water content. As the O 2 diffusion rate in water is about 10 000 times slower than in air, soil water content is a primary controlling factor. Oxygen consumption rate depends on the easily degradable organic C content of the soil and soil water and temperature, which affects soil microbial activity (Bremner & Shaw, 1958). The 02 concentration in the bulk soil as well as in soil aggregates will have a large spatial and temporal variation, which will contribute to the variation of denitrification (Parkin, 1987). Regulation of denitrification is furthermore complicated by the dependence of microorganisms on organic substrate, alternative N electron acceptors, for which they have to compete with other microorganisms and plants. Additionally, in soils not receiving nitrate fertilizer, denitrifiers depend on the net mineralization and the nitrification activity in the soil. Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

1047

MoSIER & KLEMEDTSSON

1048

All of these can be looked upon as regulative factors that interact in a synergistic manor with the soil oxygen status creating "hot spots" where much of the denitrification activity is concentrated. These hot spots are within soil aggregates, or on decomposing organic material in the soil (Parkin, 1987; Seech & Beauchamp, 1988). Because denitrification is predominantly confined to active sites in arable soils, it has the largest spatial and temporal variability of any of the N cycle processes (Tiedje et aI., 1989). Denitrification variation imposes a large problem in accurately measuring aerial and yearly emissions. Therefore, special consideration of temporal and spatial variation is needed when measuring denitrification or N20-emissions in the field. Before undertaking denitrification measurements, we suggest reading Hauck and Weaver (1986), Klemedtsson et aI. (1990), Mosier & Heinemeyer (1985), Parkin & Robinson (chapter 2 in this book), and Tiedje et aI. (1989) as reference materials on the measurement and variation of denitrification. Because of this variability and the problems in measuring N2 production against normal atmospheric background, quantifying denitrification is difficult and no "standard, absolute" methods exist. There is also no consensus as to a "best" method for quantifying denitrification in the field.

44-1 METHODS

This is not a comprehensive review of denitrification techniques but rather a description of techniques that we think are best suited for field measurements within the constraints of current technology. We suggest two different methods for measuring denitrification in the field, acetylene inhibition and use of 15N isotopes, using either in situ flux or intact core measurements. 44-1.1 The Acetylene Inhibition Method

In the early 1970s, Federova et aI. (1973) discovered that acetylene inhibited the reduction of N20 to N2 in the denitrification process. This discovery formed the basis for the development of the acetylene inhibition method (Balderston et aI., 1976; Yoshinari & Knowles, 1976).

Previously it was difficult to measure N2 production during denitrification, because of the high background concentration of the gas in the atmosphere (78%). By inhibiting the last step in the denitrification process with acetylene, low denitrification rates can be measured in ambient atmosphere by measuring N20 emissions, given the low natural background concentration ofN20 of about 310 ppb (Duxbury, 1986; Klemedtsson et aI., 1990; Mosier & Heinemeyer, 1985; Ryden et aI., 1979; Tiedje et aI., 1989).

MEASURING DENITRIFICATION

1049

44-1.2 The Nitrogen-15 Method An alternative to the acetylene block method is using the stable isotope lsN. This concept was developed almost 40 yr ago but is now much more practical since the cost of isotopes has decreased and isotope ratio mass spectrometers are now more stable, more readily available, more "user friendly," and less expensive than in past decades. Using lsN-labeled fertilizer to quantify N gas production directly from denitrification permits direct measurement of N2 and N20 from the fertilizer applied (Rolston et aI., 1976, 1978, 1982) and from the total mineral N in the soil (Hauck & Bouldin, 1961). The method is useful and relatively sensitive since the isotopic composition of N2 in the atmosphere is 99.26% mass 28,0.73% mass 29, and 0.0013% mass 30, small changes in mass 30 within a confined atmosphere are readily measured. Direct measurement techniques using lsN, involve applying highly enriched (20-80%) lsN fertilizer to a designated plot of soil and measuring N gas production following denitrification by quantifying the increase in lsN-labeled gases within a closed chamber. Using the 15N method for studying denitrification is constrained by the same assumptions and sources of error that apply to other uses of 15N. These considerations are discussed in detail by Bremner (1965), Hauck and Bremner (1976), Bremner and Hauck (1982), Buresh et al. (1982), Hauck (1982) and will not be detailed herein. 44-2 EXPERIMENTAL PROTOCOLS Both acetylene inhibition and 15N can be used with many of the possible techniques for measuring denitrification. Either method may be used with static cores or closed chambers or both used simultaneously to provide a cross check. We suggest the following techniques because a minimum of field equipment is required to make measurements. 44-2.1 Static Core Protocol In the early 1980s, different soil core techniques were developed, using the acetylene inhibition method to measure denitrification under field conditions. Two basically different soil core systems were developed. In the first, intact soil cores are incubated statically with acetylene inside a container (Aulakh et aI., 1982; Robertson & Tiedje, 1987; Svensson et aI., 1984; Ryden et aI., 1987). In the second, confined intact soil cores are incubated such that soil air and acetylene are recirculated through the macropores of the soil (Parkin et aI., 1984). Positive and negative aspects of both core systems have been detailed in a review by Tiedje et al. (1989). They conclude that both approaches estimate the denitrification process equally well if the inherent limitations of the techniques are considered. We recommend using the static core system in the field as it is less complicated and thus accommodates greater replication.

1050

MOSIER & KLEMEDTSSON

Several different types of static core systems have been used to measure denitrification rates. We suggest using encased cores: (i) to protect them from destruction during the incubation; (ii) to use the same approach on soils with different structure stability, and (iii) for ease of use in the field. According to Tiedje et ai. (1989), the main problem with static cores is limited diffusion of acetylene into the soil core and the NzO out of the core. Since encasement of the core presents a barrier to gas diffusion, the encasement should have holes or slits to facilitate gas diffusion. Core sampling devices should be constructed so that one can determine if soil compaction has occurred during sampling. If compaction exceeds 5% the core should be discarded. If compaction is a persistent problem then it may be reduced by decreasing the length and increasing the diameter of the soil core. Soil cores should be incubated at the in situ temperature in the field or laboratory. We recommend this because using a 010 quotient to calculate changes in the denitrification rates with increasing temperature can result in underestimation. An increase in temperature increases the activity of the denitrifiers, and also promotes anaerobiosis by affecting soil respiration. If soil cores are transported to the laboratory for incubation, they should be transported on ice (Tiedje et aI., 1989). It is important that the soil cores are transported without vibrations that might change soil structure. Denitrification rates in field soils do not always follow a normal distribution in time and space (see chapter 2 by Parkin and Robinson in this book). As a general rule, if denitrification rates have a coefficient of variation (CV) above 100%, then one can assume that the distribution is skewed and that it may be lognormally distributed, at least during some periods over the year. Under these conditions one should use at least 20 replicates for each treatment to be investigated. Analysis of data can be handled as described in Parkin et ai. (1988, 1990) or in Svensson et ai. (1991). If the denitrification rates have a CV that is < 100%, then one can use normal statistical analysis and < 10 replicates. 44-2.1.1 Acetylene Sources First, acetylene is a combustible gas that must be handled with care. Second, acetylene from commercial tanks contains relatively large concentrations of acetone (Hyman & Arp, 1987). Unfortunately, acetone is a good substrate for soil heterotrophic organisms and must be removed when using the acetylene in denitrification experiments. One way to remove acetone from commercial grade acetylene is to pass the gas through gas dispersion tubes in two consecutive concentrated sulfuric acid traps followed by another trap containing distilled water (Mosier et aI., 1985). The acetylene stream exiting the water trap should be analyzed for acetone to ensure that no acetone is being added to the plot. High purity acetylene can also be produced from calcium carbide (CaCz). The acetylene formed from CaCz can be assumed to be clean even though small amounts of Hz, CH4 , CzH 4 , and PH3 are produced (Hyman

MEASURING DENITRIFICATION

1051

& Arp, 1987). These contaminants do not appear to alter soil denitrification (Tiedje et al., 1989). 44-2.1.2 Soil Core Collection Minimally disturbed, intact, soil cores (0-10 cm) are removed from the appropriate plots using a I-m long, drop hammer soil sampler. The diameter of the soil core should be 3 to 8 cm. Collection is facilitated by a sampling device that contains an inner liner made from PVC or stainless steel pipe that is slightly larger than the inside diameter of the sampler tip. The liner is perforated with numerous 2-mm diam. holes made by drilling the pipe to provide ready equilibration of the air within the soil and atmosphere. The liner also serves as the core encasement during incubation. Each soil core can be quickly removed from the corer and placed in an incubation chamber. Each core is placed in a pvc or glass container that is a centimeter or less larger diameter than the outside diameter of the core liner. The headspace above the soil should be about 100 cm3. The top of the incubation cylinder is a rubber stopper drilled to fit a gas tight gas chromatographic septum through which a needle can be inserted for sampling the gas inside the incubation cylinder (Burton & Beauchamp, 1984). Intact soil cores from 10 to 20, 20 to 30 etc. can be collected in the same manner if information concerning denitrification activity deeper in the soil profile is desired. Acetylene (free of contaminants) is added to the gas, normal air unless an oxygen-free atmosphere is desired, headspace above or around the soil core, to a final concentration of 5 to 10% voVvol (5-10 kPa). The acetylene in the gas headspace is mixed with the gas in the larger macro pores by alternatively reducing and increasing the pressure by pumping with a large gas tight syringe (30-50 mL volume). The over pressure due to the added acetylene is vented with a needle. Each cylinder is incubated for up to 24 h, in a hole of slightly larger dimensions, in the ground adjacent to the study area. Headspace atmosphere in the core container is sampled after 3, 12, or 24 h, after mixing the macropore gas with the head space gas with a large syringe. In soils where the nitrate supply may be quickly depleted, short time sampling intervals of 2 or 3 h are suggested. Since acetylene also blocks nitrification, soils may become nitrate deficient and underestimate denitrification. The gas samples could either be analyzed directly or stored in gas tight vials (Aulakh et al., 1991; Ryden et al., 1987). 44-2.1.3 Analysis Air samples are analyzed for N 20 using a gas chromatograph equipped with a 63Ni electron capture detector (ECD), which is operated at high temperature (300-400 0c) using a system like that described by Mosier and Mack (1980). Although detector response and linearity vary from one manufacturer to another, most systems that we know of provide satisfactory capability for N 20. Valco Instruments pneumatically or

1052

MosmR & KLEMEDTSSON

electrically operated lO-port sampling and four-port switching valves are incorporated into the GC system. The sampling and switching valves may be controlled by either the GC microprocessor or an external data acquisition-control unit. Detector, sampling valve, switching, valve and column oven temperatures are 300 to 400, 100, 175, and 70°C, respectively (detector temperature may vary according to manufacturer recommendations). The flow rate of carrier, backflush, and detector purge gases (95% argon + 5% methane) is 18 cm3/min. Gas samples are introduced into a 1 to 5 cm3 gas sampling loop (size depends upon the sensitivity of the ECD being used) through an inlet system (Mosier & Mack, 1980). Both CO2 and water vapor are removed from the gas samples. The two absorbent traps are prepared by packing 10-mm-diam. Millipore syringe filter holders with Ascarite and Mg(CI0 4)2' and connected to the GC inlet by vacuum-tight luer fittings. The traps are used to reduce potential variability in N20 analysis caused by different CO2 and water vapor concentrations between samples. The traps also ensure that water vapor does not accumulate in the sample loop and eliminates any effect of high CO2 concentrations on N20 analysis. To reduce analysis time and eliminate interference from water vapor (when a dryer is not used) and fluorocarbons in the sample gas mixtures, we use a 1 m by 2 mm i.d. nickel precolumn in combination with a 3 m by 2 mm i.d. analytical column. Both columns, packed with Porapak Q (801 100 mesh) are attached directly to the 10-port sampling valve (Mosier & Mack, 1980) as are the sample loop and carrier gas sources. With the valve in the starting position, the precolumn is backflushed with carrier gas while the sample loop is first evacuated and then refilled to local atmospheric pressure ± 0.5 mm Hg with air from a sample syringe. Sample loop pressure is measured precisely with a Setra Systems Inc. 304B electronic manometer. When the sampling valve is rotated, the contents of the sample loop are swept through the precolumn into the analytical column that is vented to the atmosphere through the four-port switching valve located at the exit of the column. The sampling valve is time programmed to return to position A after N20 clears the precolumn (about 2.1 min). This allows the sample loop to be evacuated and refilled and the precolumn to be backflushed while the sample N20 continues to be separated from other atmospheric constituents on the analytical column. After O2 is eluted from the analytical column (about 3 min), the time programmed four-port switching valve diverts the analytical column flow into the detector. Without the switching valve the large amount of O 2 in the gas sample may overload the detector to give a broad, tailing peak for O2 and reduced analytical precision for N20. Oxygen also reduces the life of some EC detectors. Ten to 12 samples per hour can be run with this system, depending upon the retention characteristics of each column, at a routine precision of < 3 ppb (vol/vol) for ambient concentrations of N20. The column system is also amenable to automation through an additional inlet valving system or direct sampling from gas collection systems (Mosier & Heinemeyer, 1985).

1053

MEASURING DENITRIFICATION

Table 44-1. Bunsen absorption coefficients for N2 0 in water at different temperatures (Tiedje, 1982). Temperature °C

at

5

10

15

20

25

30

35

1.06

0.88

0.74

0.63

0.54

0.47

0.41

t a is the milliliter of N2 0 at 0 °C and 760 mm of Hg (STP) that is absorbed by 1 mL of water.

When N20 concentrations are below 100 ppm the above ECD system is used. When using the acetylene inhibition method, one may need to bypass the ECD with the acetylene in the sample, after it exits the chromatographic column. Acetylene will alter both the sensitivity and stability of some ECDs. When N2 0 concentration is > 100 ppm a thermal conductivity (Ryden et al., 1978) or ultrasonic (Bremner & Blackmer, 1982) may be used rather than an ECD because of the nonlinear nature of the ECD. The same inlet and chromatography system as described above may be used or the system described by Ryden et al. (1978) is an alternative. According to Ryden et al. (1978), N20 was separated from other gases in the sample by injection through a 0.46-mL gas sampling loop onto a column of Porapak Q (550 by 0.16 cm, i.d.) heated to 50°C. The thermal conductivity detector used a filament current of 220 rnA and helium carrier gas at 30 cm3/min. 44-2.1.4 Calculations

The flux (Q) of N20 per hour and core is calculated as: [1]

where Mi is the total N20 produced in the soil core during the first gas sampling interval (Ti)' generally 3 h after adding acetylene, and Ms is the total N2 0 produced during the second gas sampling interval (Ts)' 12 or 24 h after initiation. The Ms and Mi are calculated according to Tiedje (1982) using the Bunsen coefficient (a), examples are listed in Table 44-1, from the equation:

[2] where Cs is the N20 concentration in the total gas phase, measured by gas chromatography (ng N20-N/mL); Vi is volume of liquid phase; Vg = total gas volume, which is the sum of the air-filled pore space (Vap) in the soil core (Vap = Vp-V i , where Vp is total pore space), the headspace above the core (Vh) and the free volume between the soil core and the incubation container (Vf ). The total pore space (air and water filled) Vp is calculated from the soil bulk density, (Bd)' soil particle density (Pd = 2.65 glcm3 ) and the volume

1054

MosmR & KLEMEDTSSON

of the soil core (Vc> after soil water content has been determined by drying the soil at 105°C over night:

[3] We recommend that N 20 emission from the soil core be expressed as ng N 2 0-N d- 1 g-l dry weight of soil using the equation: F = (Q*24)/w

[4]

where W is the dry weight of soil in each core. 44-2.2 Protocol for Closed Chamber Field Gas Flux Measurements Using Acetylene or Nitrogen-IS Gas fluxes measured with chambers estimate the soil-atmosphere exchange of gases and are not necessarily a measure of the instantaneous denitrification rate (Jury et aI., 1982). With either the acetylene or the 15N technique, the area of soil from which gas flux is to be measured must be defined. We suggest driving a cylinder (10-60 cm length and 15-30 cm in diameter) into the soil to establish each site. The length and diameter or shape must be suited to the field management with respect to zonal tillage, irrigation, type of crop, traffic patterns, and crop row spacing and orientation. Use of the acetylene method for long-term measurements requires establishment of multiple sites to permit rotation to new sites every 1 to 7 d, to prevent microbial adaptation to acetylene. Prolonged exposure of the soil to acetylene may lead to accelerated acetylene utilization or incomplete blockage of N 2 0 reduction to N 2 . 44-2.2.1 Acetylene Acetylene may be applied by either the radial diffusion of acetylene gas directly (Ryden & Dawson, 1982) or addition of 3 to 6 g of CaC2 (Aulakh et aI., 1991) to each of four 20-cm deep 2-cm diam. holes around each measurement site. Acetylene must be applied before each measurement when using cylinder gas or every 48 h when using CaC2 • Two to 3 h after applying the acetylene, a closed chamber, (Hutchinson & Mosier, 1981) is fitted and sealed over the established flux site. Use of open-base cylinders to which chamber covers can be fitted at the time of sampling prevents disturbance of soil and roots that could alter denitrification (Mosier et aI., 1991). A gas sample is immediately withdrawn by syringe from the chamber, and samples are withdrawn at IS-minute intervals for 1 h. 44-2.2.1.1 Analyses. Gas samples should be returned to the laboratory and analyzed by gas chromatography as described in section 44-3.1.3. There may be occasion to analyze samples for N2 and other fixed gases. The most direct and simple method for analyzing gas samples for oxygen

MEASURING DENITRIFICATION

1055

and N is using a thermal conductivity or ultrasonic detector following a 2 m by 0.16 cm, i.d. stainless steel or nickel column packed with 50 to 80 mesh molecular sieve 5A. The gas sample may either be injected through a conventional GC inlet or sampling loop of 0.5 to 2 mL. The gas sampling valve and sampling loop concept provide more precise analyses than direct injection by syringe. If > 1% precision is not required for the experiment then syringe injection may be used to reduce GC system cost. To analyze atmospheric gases including O 2, N2, and N20 along with CO2, CH4 , and H2 a split column GC configuration (Beard & Guenzi, 1976) using parallel columns of Porapak Q and molecular sieve 5A, or dual Porapak Q columns using differential temperatures (Bremner & Blackmer, 1982) may be used.

44-2.2.1.2 Calculations. Nitrous oxide flux can be computed from the concentration change with time using the following equation (Hutchinson & Mosier, 1981), to correct for the reduction in soil N20 concentration gradient with time as the gas accumulates inside the chamber:

[5] where V is the volume of the chamber, A is the area of soil covered, Co is the initial N20 concentration in the chamber and C 1 and C2 are the N20 concentrations after time tl and t2 where t2 = 2 t 1. To accurately quantify daily N2 and N20 emissions when flux rates are changing frequently it is necessary to make flux measurements at least every 6 h within each 24 h period (Aulakh et aI., 1991) but when flux rates are small and vary with the diel variation in soil temperature, one late morning measurement should characterize the daily flux rate (Mosier, 1989).

44-2.2.2 Nitrogen·IS·Modified Hauck Technique The direct N gas measurement technique using 15N was first proposed by Hauck et aI. (1958). Hauck and Bouldin (1961) showed calculations for 15N enrichment of NO J- undergoing denitrification and N2 evolution but it was not until two decades later that the applicability of the method became evident. Siegel et aI. (1982) revitalized the technique and modified the calculations. Mulvaney (1984) adapted the calculation to triple-collector mass spectrometers (MS), and Mulvaney and Boast (1986) refined the calculation equations to permit using lower 15N abundance N sources. Mulvaney and Kurtz (1982) showed that N20 evolution from soil could be measured with this technique. The method has been used in a variety of agricultural systems ranging from upland crops to flooded rice (Oryza sativa L.) (Mosier et aI., 1986a; Mulvaney & Vanden Heuvel, 1988; Banerjee et aI., 1990; Mosier et aI., 1989; Lindau et aI., 1990; Buresh & DeDatta, 1990; Bronson & Mosier, 1991; Heinemeyer et aI., 1988; Mohanty & Mosier, 1990; Aulakh et aI., 1991).

1056

MOSIER & KLEMEDTSSON

The "Hauck Technique" involves applying highly 15N-enriched fertilizer to soil ( > 20 atom % 15N) and using a chamber cover to isolate the atmosphere above the 15N-fertilized soil for a designated time, to permit determining the rate of change of 15N atoms (N gases) in the chamber atmosphere over time. The calculations (see section 44-2.2.2.3) use the fact that denitrification N-gases, principally N2 under most conditions, evolving from the soil into the chamber head-space, do not randomly mix isotopically with the N-gases in the chamber (78% N2 containing 0.366 atom % atom 15N). Using this nonrandom 15N distribution, the technique permits calculation of not only the amount of N-gas evolved from the added 15N-enriched fertilizer but also that from natural abundance soil N constituents as well. The 15N mole fraction of the N0 3 in the soil that serves as the N-gas source is calculated directly from N2 mass spectral data (29/28 and 30/28 + 29 ratios when using a dual collector MS and 29/28 and 30/28 ratios when using a triple collector MS) (Mulvaney, 1984). It is, therefore, not necessary to disturb the soil in the plot during the course of the experiment to determine the soil 15N03- content. The soil 15N03- when denitrification occurred must be known to calculate the total amount of N2 evolved from the site at any given time. The technique is reasonably sensitive as 5 g ofN ha- 1 d- 1 can be detected when the soil N0 3- 15N content is above 50 atom %. The technique, as described here, requires several assumptions. The first is that the total amount of 28N2 inside the gas collection chamber does not change during the sample collection period. Siegel et al. (1982) noted that providing the chamber 28N2 is 100 times or more greater than the amount of gas produced, the calculations are correct. The equations of Mulvaney and Boast (1986) do not rely upon this assumption. The second, and major assumption is that the 15N label of the N0 3- in the soil is uniform. We assume that N0 3- formed from the 15N-Iabeled fertilizer added to the soil mixes uniformly with the unlabeled N0 3- already in the soil or with unlabeled N0 3- formed from organic N mineralization. Nonuniform mixing of the 15N label may cause underestimation of the total N denitrified. It seems unlikely that complete equilibration of fertilizer Nand soil N ever occurs, but studies by Mulvaney (1988) and Mosier and Schimel (1993) indicate that accurate N-gas production estimates can be made when the soil N03- pool is not uniformly labeled. The depth that cylinders are driven into the soil is important when using 15N techniques. It is prudent to conduct N-balance measurements along with direct N-gas flux measurements when using 15N. The root zone of the vegetation should be enclosed within the cylinder so that plants outside the treated area cannot withdraw labeled N from it. However, if the whole root zone of the plant is not enclosed within the cylinder, the area around the treated site must be sampled (Mosier et aI., 1986a,b). Since rice plants provide a conduit for gaseous transport of N2 and N20, plants must be included inside rice field microplots (Mosier et aI., 1990). After establishing microplots, N fertilizer containing 50 to 80 atom % 15N is added to the soil. Addition is made by either injecting the appro-

MEASURING DENITRIFICATION

1057

priate amount of solution a few centimeters below the soil surface in a grid across the microplot (Schimel & Firestone, 1989); removing the top 10 cm of microplot soil and mixing it thoroughly with the 15N fertilizer (Mosier et aI., 1986a); or pipetting the fertilizer solution uniformly over the surface of the microplot; or applying 15N directly into floodwater for rice (Lindau et aI.,1990). In upland crops, significant denitrification will not occur until the field is irrigated or sufficient rainfall is received to limit O 2 diffusion in the soil. At this time daily monitoring is required. If N20 emissions comprise > 5% of the total N2 + N 20, then flux measurement periods of no more than 2 h can be used. If N 20 makes up < 5% of the total, as in a rice field or heavy textured soil, then overnight cover placement is appropriate (Mosier, 1989). 44-2.2.2.1 Gas Collection. Gas collection chambers (Hutchinson & Mosier, 1981) are placed over the microplots every day at midnight, 0600, 1200, and 1800 h each day for 2 h. Forty cubic centimeter gas samples are taken immediately after covering, and again after 1 and 2 h with 60 cm3 polypropylene syringes fitted with a one-way, gas tight, plastic stopcock. Chambers are removed after 2 h. This procedure is followed for 3 to 4 d following each water event until no fertilizer-derived mineral N remains in the soil. In flooded rice, chambers are placed at 1800 h, samples taken at 0,4, and 14 h later, and chambers removed after the 14-h sampling. 44-2.2.2.2 Analysis. One or 2 mL of gas collected from a soil chamber atmosphere is delivered to the MS by a one-way stopcock fitted syringe. To analyze the gas sample on the MS it is first necessary to construct an inlet system on the instrument so that oxygen, CO2, and water vapor are removed from the gas sample. An inlet is simple and can be constructed for about $500. This inlet (Fig. 44-1) consists of a series of stainless steel components; a 2 to 5 mL sample loop, an oxygen scrubber, and a cold trap, each connected by a gas-tight valve to permit isolating each portion of the inlet. The inlet is arranged to permit introducing a gas sample directly from a syringe fitted with a two-way luer fitted stopcock into the sample loop. The loop can be immersed in a cryogen (liquid N or a dry ice cooled solution) to remove water vapor, CO2 or N20 depending upon the desired analysis. For our purpose we want to look at total N-gas evolution from the soil so will not use a cryogenic trap at this stage. From the sample loop the gas passes into the oxygen scrubber. This scrubber is made by packing a 0.5 m long by 4 mm i.d. (~ in. o.d.) stainless steel tube, with 10 to 15 g of Cu-coated silica O 2 trapping material (Chemical Research Suppliers, Inc., 900 Westwood Ave., Addison, IL 60101) and plugging the ends of the tube with quartz wool. This scrubber can be operated at room temperature but efficiency is much greater if an elevated temperature is used. At a temperature of 250 to 300 °C, the trap effectively removes O 2 and reduces N20 and NO to N2. After several hundred samples, the trap is readily regenerated by passing H2 gas through the trap while heating at 250 to 300 °C (follow manufacturer's regeneration instructions as H2 can be explosive).

1058

MOSIER & KLEMEDTSSON

TO MASS SPEC

Fig. 44-1. Diagram of a simple spectrometer gas sampling inlet, (V) indicates vacuum-tight valve, sample loop is 2 to 5 cm3 volume to be used with or without a cold trap, reduction column is heated to 250 to 300°C.

About 30 s after introducing the sample into the O 2 scrubber the sample is passed through the high-efficiency liquid N trap (a coil of tubing not just a V-tube) to remove CO2 and H 20. The purified sample N2 is then passed into the MS inlet. Both sample and reference gases are introduced into the MS through the gas inlet system so that they are directly comparable. If a dual inlet MS is available, a time zero field gas sample is introduced into the MS through the sample inlet into the reference side to serve as the reference gas. If a single inlet instrument is used then samples and reference gases must be run sequentially. 44-2.2.2.3 Calculations. Rather than go through equation derivation, we will refer you to Siegel et al. (1982), Mulvaney (1984), and Mulvaney and Boast (1986). Table 44-2 summarizes the basic equations used by Mosier et al. (1986a,b, 1989) to calculate the total N gas flux from the soil. Remember that this is total N2 + N20 + NO, not just the N2 derived from the added fertilizer. The equations from Siegel et al. (1982), Mulvaney (1984), and Mulvaney and Boast (1986) were derived to quantify N2 or N20 production. If the N in N2 and N20 evolved during the same gas collection period are greatly different when the calculations are not theoretically valid for measuring N2 + N oxides. Mosier et al. (1986a,b) and Bronson et al. (1992), in field studies, did not find significant differences in the isotopic composition of N2 and N20 measured during the same gas sampling time when separate measurements were made on the N2 and N20 produced. Careful cross checks should be made with each experiment to ensure that large differ-

MEASURING DENITRIFICATION

1059

Table 44-2. Equations used to calculate N gas emissions from mass spectrometric isotope ratio measurements of gas samples. r = (29N2f2 8N2) sample - (29N2f2 8N 2) reference r' = (3°N 2f2 8N2 + 29N2) sample - ON 2 f2 8 N 2 + 29N2) reference 3. Sample = air sample from collection chamber at some time, t, after installing chamber. 4. Reference = air sample from field, i.e., normal air sample, taken from the chamber immediately after installation, 29N)28N 2 and 30N 2f2 8 N 2 + 29N 2 are ion current ratios determined by the mass spectrometer. 5. 15XN = mole fraction of 15N in the soil N0 3- pool 1.

2.

e

~

~

6. d

= 2.015 (~ r'/~ r)/(1 + = the fraction of total N gas = ~ r'1(15XN)z

(2.015

(~ r'/~

r»

in the gas collection chamber attributable to denitrification.

7. Total denitrification N gas evolved from the soil into the collection chamber = total N2 in the chamber volume x d. 8. N2 Flux = ~ CIA t where A = soil surface area covered by chamber t = time that the chamber covered the soil ~ C = the change in concentration of 30N 2 and 29N 2 in the chamber during time t

Table 44-3. Mass spectrometer data to use in the following calculations. Isotope ratios Sample gas 4 h Plot 1

Reference gas 0 h Plot 1

29/28

30/28 + 29

29/28

30/28 + 29

Rep 1 Rep 2 Rep 3

0.00735 0.00735 0.00735

0.0000751 0.0000741 0.0000749

0.00731 0.00729 0.00730

0.0000331 0.0000327 0.0000328

Mean

0.00735

0.0000751

0.00730

0.0000329

r = 0.00735 - 0.00730 = 0.00005 r' = 0.0000751 - 0.0000329 = 0.0000418 15XN =0.651 d =0.00009863

ences, > 10% in isotopic composition of the N in N2 and N20, do not appear. The following is an example calculation using data from gas samples collected during a 1987 rice denitrification experiment in Griffith, N .S. W., Australia, and analyzed on a VG-622 mass spectrometer (a single inlet, dual collector instrument) at the CSIRO Division of Plant Industry in Canberra. The MS was fitted with the gas sampling inlet system described above. Samples from Griffith were collected 0 and 4 h after placing chambers over rice plots that had been fertilized with 79 atom % 15N-urea in the floodwater at pannicle initiation (see Mosier et aI., 1989). The gas samples were injected into Vacutainers and taken to Canberra for analyses. The MS data are shown in Table 44-3 and calculations follow:

1060

MOSIER & KLEMEDTSSON

To calculate the amount of N2 inside the chamber (7775 mL): assume that air is 78% N2 ; correct volume to standard temperature and pressure (STP); 273 K and 760 mm of Hg; assume that the temperature was 20°C (293 K) and the atmospheric pressure was 760 mm of Hg. With these assumptions calculate a total of 5652 mL of N2 inside the chamber. The air temperature should be measured during each sampling time and used in the calculations. The atmospheric pressure for the sampling location should also be used, that is, about 640 mm of Hg in Fort Collins, CO and 760 mm of Hg in Baton Rouge, LA. To calculate the mass of this N2 , we know that 1 mol of a gas at STP occupies 22 400 mL; then 5652 mL of gas is 0.252 mol, 7.065 g. To calculate the amount of N gas evolved from the soil into the gas chamber: N2 evolved = N2 in chamber x d = 7.065 g x 0.00009863 = 697 Ilg N plot- 1 4 h- 1 N2 flux = 697 Ilg of Nt (707 cm2 x 4 h) = 0.247 Ilg N cm- 2 h- 1 • On a hectare basis, then 24.7 g of N h- 1 evolved from the soil during this 4-h sampling period that began 52-h after urea fertilization. 44-2.3 Measuring Dinitrogen Emissions from Applied Nitrogen-IS-Labeled Fertilizer If the objective of a study is to determine the amount of denitrification only from the applied N fertilizer, the "Hauck" calculations do not need to be used. The analytical technique, however, remains the same as that described above. The studies of Rolston et al. (1976, 1978, 1982) show the utility of this technique where they fertilized 1-m2 plots with 40 to 50 atom % 15N KN0 3 and imposed a variety of irrigation and organic C amendments to the plots. By enclosing the air space above the plots for 1 to 4 h each day and measuring the increase in concentration within the chamber headspace of N 20 and 15N2, they were able to show the effects of the various treatments on N loss by denitrification. The isotopic composition of N in gas samples was determined for samples pulled through Ascarite, magnesium perchlorate, and a commercial O2 scrubber by direct injection into the mass spectrometer. With the knowledge of the volume of the chamber headspace, the concentration of N2 in the gas (measured by gas chromatography), the time that the plot was covered, and the atom % 15N of the N in the chamber gas, the flux of N2 from the fertilizer was calculated. The isotopic composition of the N2 evolved from the fertilizer is just as described above for the Hauck Technique, the N2 produced from the added nitrate does not mix isotopically with the N2 in the air. It is, therefore, necessary to calculate atom % 15N by the equation:

atom % 15N = [29 + 2(30)]t [2(28) + 2(29) + 2(30)]

[6]

MEASURING DENITRIFICATION

1061

This requires measuring the 29/28 isotopic ratios as well as the 30/28 ratios or 30/28 + 29 ratios of the N2 and has a sensitivity of about 100 g of N ha- 1 d- 1 .

44-3 PROBLEMS WITH GAS SAMPLING AND STORAGE CONTAINERS Affordable, leak-free, noncontaminating gas sampling and storage devices are a continual concern. We generally recommend using Monoject polypropylene syringes fitted with one-way nylon stopcocks as gas-sampling devices when samples are to be analyzed within a few hours or injected into storage containers. These syringes are uniform in quality from batch to batch and are relatively inexpensive. Gases do, however, diffuse through the walls of the polypropylene syringes. Therefore, they cannot be used as storage devices. They also cannot be used to transport samples within areas where the atmospheric pressure changes. Nylon syringes leak less through their walls but sti11leak when changing atmosphere between the sampling and analytical sites (an elevation change of about 1200 m). Vacutainers have been widely used as gas transport and storage devices because they are relatively inexpensive and convenient to use. Additional problems, to those discussed above, exist. First, the tubes are not completely evacuated as they come from the manufacturer, thus contain a significant amount of gas before the sample is added. The pressure in the tubes is quite constant, however. By measuring existing pressure in a large set of each lot member of evacuated vials, before and after injecting 12 mL of air, usually 75 to 85% of the contents of the Vacutainers come from the injected sample. This initial dilution does not affect the 15XN calculations, see Table 44-3 for definition of terms, but does reduce the calculated d value by 15 to 25%. This same dilution applies to N20 as well. A second problem with the Vacutainers is that the N20 content of the tubes is above that of normal air. When the tubes are filled with 12 mL of normal atmospheric air, the tube air contains> 800 ppb voVvol N20 rather than 310 ppb normally found in air. This is not a problem when measuring total N gas on the MS, but a correction must be made when quantifying N20 emissions from the soil. The excess N20 is readily correctable by collecting time zero gas samples from the field plots at each sampling time and analyzing them to determine the tube N20 content of time zero air. These two problems are reduced by washing new Vacutainers with soap and water, drying them, then reevacuating and resealing them with silicone caulking. Recently, we began using glass serum bottles fitted with gray butyl rubber stoppers (Wheaton) as gas transport and storage containers. We evacuate the bottles and reseal them with silicone caulking. We have found no uptake or production of N20 from the butyl rubber stoppers.

1062

MOSIER & KLEMEDTSSON

44-4 GAS DIFFUSION PROBLEMS Although the above techniques have been used in a variety of agricultural situations, caution is advised in their use and interpretation because of the effect of soil texture and water content on gas diffusion (Letey et aI., 1980). In flooded soils and probably in wet clayey soils, the movement of the gases produced in the soil to the atmosphere above is controlled by water. Since a gas diffuses about 10 000 times more slowly in water than in air, the time required for a gas to move from its production site in the soil to the atmosphere may depend upon soil water content. In the case of flooded rice, gases may be entrapped in the soil and reach the soil surface only slowly unless the gas production zones are in close proximity to plant roots, (Lindau et aI., 1988). Rice plants do facilitate the movement of gases from the root zone to the atmosphere (Cicerone & Shetter, 1981) and have been shown to transport N2 and N20 from flooded soil to the atmosphere (Mohanty & Mosier, 1990; Mosier et aI., 1990). Buresh & DeDatta (1990) have also shown, in a rice field, that 15N2 was present in the soil of 15N-fertilized plots. Although not a quantitative measure, this does show that quantification of denitrification by measuring N-gas flux may not always be complete and some measure of N entrapped in the soil needs to be made. Mosier et ai. (1989) used a multiple phase equilibration (McAullife, 1971) technique to estimate N gases in a rice field soil porewater and dissolved in the floodwater.

REFERENCES Aulakh, M.S., J.W. Doran, and A.R. Mosier. 1991. Field evaluation of four methods for measuring denitrification. Soil Sci. Soc. Am. J. 55:1332-1338. Aulakh, M.S., D .A. Rennie, and E.A. Paul. 1982. Gaseous nitrogen losses from cropped and summer-fallow soils. Can. J. Soil Sci. 62:187-195. Balderston, W.L., B. Sherr, and W.J. Payne. 1976. Blockage by acetylene of nitrous oxide reduction in Pseudomonal perfectomarinus. Appl. Environ. Microbiol. 31:504-508. Banerjee, N.K., A.R. Mosier, K.S. Uppal, and N.N. Goswami. 1990. Use of encapsulated calcium carbide to reduce denitrification losses from urea-fertilized flooded rice. In Proc. Int. Denitrification Workshop, Giessen, Germany. March 1989. Mitt. Dtsch. Bodenforsch. Ges. 60:245-248. Beard, W.E., and W.D. Guenzi. 1976. Separation of soil atmospheric gases by gas chromatography with parallel columns. Soil Sci. Soc. Am. J. 40:319-321. Bremner, J .M. 1965. Isotope-ratio analysis of nitrogen in nitro~en-15 tracer investigations. p. 1256-1286. In C.A. Black (ed.) Methods of soil analYSIS. Agron. Monogr. 9. Part 2. ASA, Madison, WI. Bremner, J.M., and A.M. Blackmer. 1982. Composition of soil atmospheres. p. 873-895. In A.L. Page et al. (ed.) Methods of soil analysis. Agron. Monogr. 9. Part 2. 2nd ed. ASA and SSSA, Madison, WI. Bremner, J.M., and R.D. Hauck. 1982. Advances in methodology for research on nitrogen transformations. p. 479-484. In FJ. Stevenson (ed.) Nitrogen in agricultural soils. Agron. Monogr. 22. ASA, Madison, WI. Bremner,J.M.! and K. Shaw. 1958. Denitrification in soil. I. Methods of investigation. J. Agnc. SCI. 51:22-39. Broadbent, F.E, and FE. Clark. 1965. Denitrification. p. 344-359. In W.v. Bartholomew and FE. Clark (ed.) Agron. Monogr. 10. ASA, Madison, WI.

MEASURING DENITRIFICATION

1063

Bronson, K.F., and A.R. Mosier. 1991. Effect of encapsulated calcium carbide on dinitrogen, nitrous oxide, methane, and carbon dioxide emissions in flooded rice. BioI. Fertil. Soils 11:116-120. Bronson, K.F., A R. Mosier, and S.R. Bishnoi. 1992. Nitrous oxide emissions in irrigated com as affected by nitrification inhibitors. Soil Sci. Soc. Am. J. 56:161-165. Buresh, R.J., E.R. Austin, and E.T. Craswell. 1982. Analytical methods in ISN research. Fert. Res. 3:37-62. Buresh, R.J., and S.K. DeDatta. 1990. Denitrification losses from puddled rice soils in the tropics. BioI. Fertil. Soils 9: 1-13. Burton, A., and E.G. Beauchamp. 1984. Field techniques using the acetylene blockage of nitrous oxide reduction to measure denitrification. Can. J. Soil Sci. 64:555-562. Cicerone, R.J., and J.D. Shetter. 1981. Sources of atmospheric methane: Measurements in rice paddies and a discussion. J. Geophys. Res. 86:7203-7209. Duxbury, J.M. 1986. Advantages of the acetylene method of measuring denitrification. p. 73-91. In R.D. Hauck and R.W. Weaver (ed.) Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18. SSSA, Madison, WI. Federova, R.I., E.1. Melekhina, and N.1. Ilyuchina. 1973. Evaluation of the method of "gas metabolism" for detecting extra terrestrial life. Identification of nitrogen-fixing organisms. Izv. Akad. Nauk SSSR. Ser. BioI. 6:791. Focht, D.D., and M. Verstraete. 1977. Biochemical ecology of nitrification and denitrification. p. 135-214. In M. Alexander (ed.) Advances in microbial ecology. Vol. 1. Plenum Press, New York. Folorunso, O.A, and D.E. Rolston. 1984. Sratial variability of field measured denitrification gas fluxes. Soil Sci. Soc. Am. J. 48:12 4-1219. Hauck, R.D. 1982. Nitrogen-isotope-ratio analysis. p. 735-779. In AL. Page et al. (ed.) Methods of soil analysis. Part 2. Agron. Monogr. 9. 2nd ed. ASA and SSSA, Madison, WI. Hauck, R.D., and R.W. Weaver. 1986. Field measurement of dinitrogen fixation and denitrification. SSSA Spec. Publ. 18. SSSA, Madison, WI. Hauck, R.D., and D.R. Bouldin. 1961. Distribution of isotopic nitrogen in nitrogen gas during denitrification. Nature (London) 191:871-872. Hauck, R.D., and J.M. Bremner. 1976. Use of tracers for soil and fertilizer nitrogen research. Adv. Agron. 28:219-266. Hauck, R.D., S.W. Melsted, and P.E. Yankwich. 1958. Use of N-isotope distribution in . nitrogen gas in the study of denitrification. Soil Sci. 86:287-291. Heinemeyer, 0., K. Haider, and A.R. Mosier. 1988. Phytotron studies to compare nitrogen losses from com-planted soil by the 15-N balance or direct dinitrogen and nitrous oxide measurements. BioI. Fert. Soils 6:73-77. Hutchinson, G.L., and A.R. Mosier. 1981. Improved soil cover method for field measurement of nitrous oxide flux. Soil Sci. Soc. Am. J. 45:311-316. Hyman, M.R., and D.J. Arp. 1987. Quantification and removal of some contaminating ~ases from acetylene used to study gas-utilizing enzymes and microorganisms. Appl. EnVIron. Microbiol. 53:298-303. Jury, W.A, J. Letey, and T. Collins. 1982. Analysis of chamber methods used for measuring nitrous oxide production in the field. Soil Sci. Soc. Am. J. 46:250-256. Klemedtsson, L.K., G. Hansson, and A.R. Mosier. 1990. The use of acetylene for the quantification of N2 and N20 production from biological processes in soil. p. 167-180. In J. Sorensen and N.P. Revsbeck (ed.) Denitrification in soil and sediment. Plenum Press, New York. Letey, J., W.A. Jury, A Hadas, and N. Valoras. 1980. Gas diffusion as a factor in laboratory incubation studies on denitrification. J. Environ. Qual. 9:223-227. Lindau, C.W., R.D. DeLaune, W.H. Patrick, Jr., and P.K. Bollich. 1990. Fertilizer effects on dinin:ogen, nitrous oxide, and methane emissions from lowland rice. Soil Sci. Soc. Am. J. 54.1789-1794. Lindau, C.W., W.H. Patrick, Jr., R.D. DeLaune, K.R. Reddy, and P.K. Bollich. 1988. Entrapment of nitrogen-15 dinitrogen during soil denitrification. Soil Sci. Soc. Am. J. 52:538-540. McAullife, C. 1971. GC determination of solutes by multiple phase equilibration. Chem. Techno!. (Jan.) 0:46-50. Mohanty, S.K., and A.R. Mosier. 1990. Nitrification-denitrification in flooded rice soils. In 14th Int. Congr. Soil Sci. 4:326-331.

MOSIER & KLEMEDTSSON

1064

Mosier, A.R. 1989. Chamber and isotope techniques. p. 175-187. In M.O. Andreae and D.S. Schimel (ed.) Exchange of trace gases between terrestrial ecosystems and the atmosphere. John Wiley and Sons, Chichester. Mosier, A.R., and D.S. Schimel. 1993. Nitrification and denitrification. p. 181-208. In R. Knowles and T.H. Blackburn (ed.) Nitrogen isotope techniques. Academic Press, San Diego. Mosier, A., D. Schimel, D. Valentine, K. Bronson, and W. Parton. 1991. Methane and nitrous oxide fluxes in native, fertilized and cultivated grasslands. Nature (London) 350:330-332. Mosier, A.R., S.K. Mohanty, A. Bhadrachalam, and S.P. Chakravorti. 1990. Evolution of dinitrogen and nitrous oxide from the soil to the atmosphere through rice plants. BioI. Fertil. Soils 9:31-36. Mosier, A.R., S.L. Chapman, and J.R. Freney. 1989. Determination of dinitrogen emission and retention in floodwater and porewater of a lowland rice field fertilized with lsN_ urea. Fert. Res. 19:127-136. Mosier, A.R., W.D. Guenzi, and E.E. Schweizer. 1986a. Determination of dinitrogen and nitrous oxide from irrigated crops in north-eastern Colorado. Soil Sci. Soc. Am. J. 50:831-833. Mosier, A.R., W.D. Guenzi, and E.E. Schweizer. 1986b. Field denitrification estimation by nitrogen-15 and acetylene inhibition techniques. Soil Sci. Soc. Am. J. 50:831-833. Mosier, A.R., and O. Heinemeyer. 1985. Current methods used to estimate N20 and N2 emissions from field soils. 79-99. In H.1. Golterman (ed.) Denitrification in the nitrogen cycle. Plenum Pub. Corp., New York. Mosier, A.R., and L.K. Mack. 1980. Gas chromatographic system for precise, rapid analysis of N 20. Soil Sci. Soc. Am. J. 44:1121-1123. Mosier, A.R., F.M. Melhuish, and W.S. Meyer. 1985. Direct measurement of denitrification using acetylene blockage and infrared gas analysis in a root zone lysimeter. p. 101-115. In W.A. Muirhead and E. Humphreys (ed.) Root zone limitations to crop production on clay soils. Aust. Soc. Soil SCI., Riverina Branch, Griffith, Australia. Mulvaney, R.L. 1988. Evaluation of nitrogen-15 tracer techniques for direct measurement of denitrification in soil. III. Laboratory studies. Soil Sci. Soc. Am. J. 52:1327-1332. Mulvaney, R.L. 1984. Determination of ISN-iabeled dinitrogen and nitrous oxide with triplecollector mass spectrometers. Soil Sci. Soc. Am. J. 48:690-692. Mulvaney, R.L., and C.W. Boast. 1986. Equations for determination of nitrogen-15 labeled dirutrogen and nitrous oxide by mass spectrometry. Soil Sci. Soc. Am. J. 50:360-363. Mulvaney, R.L., and L.T. Kurtz. 1982. A new method for determination of lsN-labeled nitrous oxide. Soil Sci. Soc. Am. J. 46:1178-1184. Mulvaney, R.L., and RM. vanden Heuvei. 1988. Evaluation of nitrogen-15 tracer techniques for direct measurement of denitrification in soil: IV. Field studies. Soil Sci. Soc. Am. J. 52: 1332-1337. Parkin, T.B. 1987. Soil microsites as a source of denitrification variability. Soil Sci. Soc. Am. Proc. 26:238-242. Parkin, T.B., S.T. Chester, and J.A. Robinson. 1990. Calculating confidence intervals for the mean of lognormally distributed variable. Soil Sci. Soc. Am. J. 54:321-326. Parkin, T.B., J.J. Meisinger, S.T. Chester, J.L. Starr, and J.A. Robinson. 1988. Evaluation of statistical estimation methods for lognormally distributed variables. Soil Sci. Soc. Am. J. 52:323-329. Parkin, T.B., H.F. Kaspar, A.J. Sexstone, and J.M. Tiedje. 1984. A gas-flow soil core method to measure field denitrification rates. Soil BioI. Biochem. 16:323-330. Robertson, G.P., and J.M. Tiedje. 1987. Nitrous oxide sources in aerobic soils. Nitrification, denitrification and other biological processes. Soil BioI. Biochem. 19:187-193. Rolston, D.E., A.M. Sharpley, D.W. Toy, and F.E. Broadbent. 1982. Field measurement of denitrification: III. Rates during irrigation cycles. Soil Sci. Soc. Am. J. 46:289-296. Rolston, D.E., D.L. Hoffman, and D.W. Toy. 1978. Field measurement of denitrification: I. Flux of N2 and N20. Soil Sci. Soc. Am. J. 42:863-869. Rolston, D.E., M. Fried, and D.A. Goldhamer. 1976. Denitrification measured directly from nitrogen and nitrous oxide gas fluxes. Soil Sci. Soc. Am. J. 40:256-266. Ryden, J.C., J.H. Skinner, and D.J. Nixon. 1987. Soil core incubation system for the field measurement of denitrification using acetylene-inhibition. Soil BioI. Biochem. 19:753757.

y.

MEASURING DENITRIFICATION

1065

Ryden, J.C., and K.P. Dawson. 1982. Evaluation of the acetylene-inhibition technique for the measurement of denitrification in grassland soils. J. Sci. Food Agric. 33:1197-1206. Ryden, J.C., L.J. Lund, and D.D. Focht. 1978. Direct in-field measurement of nitrous oxide flux from soils. Soil Sci. Soc. Am. J. 42:731-737. Ryden, J.C., L.J. Lund, J. Letey, and D.D. Focht. 1979. Direct measurement of denitrification loss from soils. II. Development and application of field methods. Soil Sci. Soc. Am. J. 43:110-118. Schimel, J.P., and M.K. Firestone. 1989. Nitrogen incorporation and flow through a coniferous forest soil profile. Soil Sci. Soc. Am. J. 53:779-784. Seech, A.G., and E.G. Beauchamp. 1988. Denitrification in soil aggregates of different sizes. Soil Sci. Soc. Am. J. 52:1616-1621. Siegel, R.S., R.D. Hauck, and L.T. Kurtz. 1982. Determination of 3ON2 and application to measurement of N2 evolution during denitrification. Soil Sci. Soc. Am. J. 46:68-74. Svensson, B.H., L. Klemedtsson, S. Simkins, K. Paustian, and T. Rosswall. 1991. Soil denitrification in three cropping systems characterized by differences in nitrogen and carbon supply. I. Rate-distribution frequencies, comparison between systems and seasonal N-Iosses. Plant Soil 138: 257-271. Svensson, B.H., L. Klemedtsson, and T. Rosswall. 1985. Preliminary field denitrification studies of nitrate-fertilized and nitrogen fixing crops. p. 157-169. In H.L. Golterman (ed.) Denitrification and the nitrogen cycle. NAlO Conf. Ser. I. Ecology Vol. 9. Plenium Press, London. Tiedje, J.M. 1988. Ecology of denitrification and dissimilatory nitrate reduction to ammonium. p. 179-243. In A.J .B. Zehnder (ed.) Biology of anaerobic micro-organisms. John Wiley and Sons, New York. Tiedje, J.M. 1982. Denitrification. p. 1011-1024. In A.L. Page et al. (ed.) Methods of soil analysis. Agron. Monogr. 9. Part 2. 2nd ed. ASA and SSSA, Madison, WI. Tiedje, J.M., S. Simkins, and P.M. Groffman. 1989. Perspectives on measurement of denitrification in the field including recommended protocols for acetylene based methods. p. 217-240. In M. Clarholm and L. Bergstrom (ed.) Ecology of arable land. Kluwer Academic Pub!. Dordrecht, Holland. Yoshinari, T., and R. Knowles. 1976. Acetylene inhibition and nitrous oxide reduction by denitrifying bacteria. Biochem. Biophys. Res. Commun. 69:705-710.

Published 1994

Chapter 45 Sulfur Oxidation and Reduction in Soils M. A. TABATABAI, Iowa State University, Ames, Iowa

Soil analysis from various parts of the world indicates that S occurs in soils mainly in organic combinations, and that most cultivated soils do not contain sufficient organic matter to supply the crops need of S (Tabatabai & Bremner, 1972a; Neptune et aI., 1975; Tabatabai, 1984). In addition, the release rate of inorganic sulfate from soil organic matter is too slow to meet the crops demand of this element (Tabatabai & Bremner, 1972b; Tabatabai & AI-Khafaji, 1980). Other factors that contribute to the S need are higher yields, more intensive land use by double cropping, use of high analysis fertilizer, increasing use of irrigation practices, declining use of S as a fungicide and insecticide, and decreasing S02 emissions into the atmosphere because of recent environmental regulations. To meet the S requirements of crop plants, the alternative is to amend the soils with S-containing fertilizer (Walker, 1964; Hagstrom, 1984). A wide variety of S-containing fertilizers in the form of solid and liquid are available. Among these, elemental S (SO) offers the best solution to the problem of cost and grade because of its purity (about 100% S). However, problems of dustiness, unpleasantness, and fire hazard have limited its use as a fertilizer (Beaton & Fox, 1971). To avoid the disadvantages associated with the finely divided So, a product containing 90% So and 10% of bentonite clay and certain proprietary additives has been developed (Sulphur Institute, 1977). Before it can be used by crops, however, So has to be oxidized to sulfate. Elemental S is oxidized in soils by chemical and biochemical processes, and several factors affect these processes. Several of those factors (e.g., soil moisture and temperature, particle size, and soil texture) have been studied by several workers (for review of literature, see Weir, 1975; Konopka et aI., 1984, Wainwright, 1984). Unlike studies on forms of S and the factors affecting oxidation of So in soils, not much information is available on the reduction of sulfate in soils (Konopka et aI., 1984). Therefore, most of the methods available for estimation of sulfate reduction are those used in studies of transformations of S in sediments (J0rgensen, 1978a,b; Fossing & J0rgensen, 1989). Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5.

1067

1068

TABATABAI

Although these methods are not widely used for estimation of sulfate reduction in soils, they will be described in this chapter.

45-1 SULFUR OXIDATION Elemental S is oxidized in soils by biotic processes and to a much lesser degree by abiotic processes. As Wainwright (1984) indicated, a wide spectrum of microorganisms is capable of oxidizing So, including members of the genus Thiobacillus, several heterotrophs, the photosynthetic S bacteria, and the colorless, filamentous S bacteria. Of those, only the thiobacilli and heterotrophs have been shown to play an important role in S oxidation in most agricultural soils. Early work by Guittonneau and Keilling (1932) indicated that polythionates (oxidizable S) are produced in SO-amended soils, but because of analytical problems, no specific intermediate S compounds could be determined. In review of the literature on oxidation of So by the genus Thiobacillus, Vishniac and Santer (1957) suggested that S201-, S40l-, and SOl- are formed as intermediate products. The first two of these compounds have been isolated from the reaction involving bacteria (London & Rittenberg, 1964; Starkey, 1956). Using 35S-labeled thiosulfate, Trudinger (1959) showed that S40l- is the first intermediate in oxidation of thiosulfate to sulfate by a Thiobacillus sp. Gleen and Quastel (1953) have reported similar results for thiosulfate oxidation on perfusion through soils. Formation of such compounds during oxidation of So in soils deserves investigation because work by Audus and Quastel (1947) showed that S20l- inhibits germination and subsequent root growth of several plants and that there is a considerable degree of selectivity of this toxic action of S20l- on growth of plant roots. For example, at a concentration of 250 mg/L S as S201- (on solution basis) root growth of garden pea (Pisum sativum L.) was inhibited by 50%. Studies by Nor and Tabatabai (1977) showed that the amount of S203-' S206-' and S04-S produced during incubation (30°C) of SOamended soils under aerobic conditions varied with time and soil used. They showed that S201- was produced within the first few days of incubation and that S40l- accumulated in some soils. The rate of S oxidation increased with increasing incubation temperatures (5, 15, and 30°C) and with increasing the rate of S application (50, 100, and 200 Ilg of S/g soil). For 100 Ilg of S/g soil, the rates of oxidation of So in 10 Iowa surface soils ranged from 39 to 75 Ilg of S/g soil after incubation at 30°C for 70 d; rates were more rapid in alkaline soils than in acid soils. There was little change in pH of soils even when the S application rate was increased from 50 to 200 Ilg/g soil. The rate of S oxidation was lower in air-dried soils than in field-moist soils (Nor & Tabatabai, 1977). The rate of So oxidation is strongly affected by soil moisture, temperature., and pH; particle size; and soil type (Janzen & Bettany, 1987a,b; Nor & Tabatabai, 1977). Therefore, it is difficult to extrapolate the So oxidation

SULFUR OXIDATION IN SOILS

1069

rate obtained from laboratory experiments to soils under field conditions. Work by Janzen and Bettany, (1987a) showed that the effect of So particle size on So oxidation rate can be minimized by using particle size in the range of 0.106 to 0.150 mm. The effect of soil moisture on So oxidation varies among soil types with the water potential optima ranging from -0.08 to -0.27 MPa, depending on the soil type (Nevell & Wainwright, 1987; Janzen & Bettany, 1987b; Deng & Dick, 1990). The optimum temperature for So oxidation under aerobic conditions has been reported to be at 30 °C (Waksman & Joffe, 1922; Li & Caldwell, 1966; Nor & Tabatabai, 1977). Deng and Dick (1990) reported optimum temperature for So oxidation of 25 to 30 °C. In most studies of So oxidation in soil, the rate is usually determined by measuring the amount of SO]- produced during a specific period after application of So to soil. Therefore, the results are expressed as the proportion of So oxidized during a given time. Janzen and Bettany (1987a) have identified two limitations related to the manner in which the oxidation rate is expressed. They argued that (i) oxidation rate are specific to the particle size of So used. Therefore, rates determined by using one particle size are not applicable to particles of a different size. Furthermore, So oxidation rates reported by different investigators using various sizes of So particles cannot be compared, and (ii) current methods of defining So oxidation rates are biased by confounding effects of diminishing So during oxidation. Because So oxidation is exclusively a function of surface area of the particle used, the amount of SO]- produced during incubation from So present, not of the total mass. In pure culture studies with thiobacilli, Laishley et al. (1983) reported that oxidative activity did not penetrate into the core of So particles, perhaps because of steric hindrance. This has been demonstrated in several studies by demonstrating linear relationships between the amount of SO]- produced and the total surface area of the applied So, irrespective of the amount applied (Fox et aI., 1964; Janzen et aI., 1982; Koehler & Roberts, 1983; Laishley et aI., 1983). One problem encountered in expressing the So oxidation rate as a function of the initial surface area is, however, the change in the surface area during incubation. As Janzen and Bettany (1987a) stated, the expression of So oxidation rates on the basis of instantaneous rather than initial surface area allows independence of both time and particle size. The decline in surface area over time can be mathematically determined if it is assumed that the So particles are relatively uniform spheres. Based on these assumptions, Janzen and Bettany (1987a) derived the following equations to calculate the proportion of So oxidized and the rate constant:

rn / rno = 1 - [1 - 2 kt / gDoP

[1 ]

where m / rno is the proportion of So oxidized as a function of a rate constant (k), time (t), density (g), and initial particle diameter (Do). Rearrangement of Eq. [1] gives:

1070

TABATABAI 1

k = {1- [1- mlmo]~}·gDo/2t

[2]

The rate constant calculated from Eq. [2] has the desired units (g S cm- 2 d- 1) and is independent of both particle diameter and time. 45-1.1 Principles Estimation of So oxidation rate in soils involves determination of SO]- produced when soil is incubated with So under specific conditions. The variables that should be considered include particle size of So, and time and temperature of incubation. These variables must be selected so that sufficient SO]- is produced for accurate determination, S01- produced from organic S mineralization is minimized, acidification of the soil sample is prevented, and So is applied accurately (Janzen & Bettany, 1987a). Any particle size of So can be used for this purpose. In general, So oxidation rate increases with decreasing particle size. If very fine (e.g., < 150 mesh, 106 !AID) So is used, it can be diluted with the same size glass beads to facilitate weighing (Nor & Tabatabai, 1977). More recent work showed that a relatively coarse (0.199 mm) So can be used successfully for estimation of So oxidation rate in soils (Janzen & Bettany, 1987a; Deng & Dick, 1990). According to Janzen and Bettany (1987a), this relatively coarse material has several advantages over other materials they considered: (i) it could be readily produced within narrow limits, (ii) its diameter could be accurately confirmed by microscopic analysis, and (iii) because of its lower specific surface area, coarse So could theoretically be applied at high levels without adversely affecting oxidation rate through secondary effects, such as acidification. As a result, application of required So to a small amount of soil sample is accurate and convenient. 45-1.2 Methods (Nor & Tabatabai, 1977; Janzen & Bettany, 1987a; Deng & Dick, 1990) 45-1.2.1 Special Apparatus

1. French square bottles (8 oz, 250 mL). 2. Stoppers (no. 7). 45-1.2.2 Reagents

1. Elemental S: Sublimed So (J.T. Baker Chemical Co., Phillipsburg,

NJ). 2. Elemental So powder, 0.119 mm: Grind prilled So and collect the particles that passed through a 0.150-mm sieve, but were retained in a 0.106-mm sieve. Wet-sieve in ethanol to ensure complete removal of fine particles adhering to the desired particles. Rinse the ethanol-washed So several times with deionized water and dry it overnight at 65 DC.

SULFUR OXIDATION IN SOILS

1071

3. Glass beads: can be obtained from Cataphote, Inc. (P.O. Box 2369, Jackson, MS). 4. Calcium phosphate monohydrate solution [Ca(H2P0 4 h·H20], 500 mg PIL. Dissolve 2.02 g of Ca(H2P0 4 h·H20 in about 700 mL of deionized water, and make to volume of 1 L with deionized water. 45-1.2.3 Procedure Mix a 20-g sample of soil ( < 2 mm) with So in a 250-mL (8 oz) French square bottle to give the So concentration desired. For convenience, 20 mg So, 0.199 mm or fine So mixed with a similar size glass beads can be used. If air-dried soil is used, add 5 mL of deionized water to bring the moisture content to 30 to 50% of water-holding capacity. Stopper the bottle and incubate it for the time and temperature desired. It is recommended to incubate for 6 d at room temperature (23°C) or at 30°C. Remove the stopper and aerate the bottle every 2 d. After incubation, add 100 mL of 500 mgIL of Pas Ca(HP0 4 )z, stopper the bottle and shake it on an endto-end shaker for 1 h. Filter the soil-solution mixture through a Whatman No. 42 filter paper under suction. For complete removal of any particulate, filter the soil extract through a GA-8 membrane filter (Gelman Instrument Co., Ann Arbor, MI). Analyze this filtrate for S20?- and S40l- by the colorimetric methods described by Nor and Tabatabai (1975, 1976) or total inorganic S (S20?-, S40l-, and SOi-) by the methylene blue method (Tabatabai, 1982), or for SOi- by ion chromatographic method (Dick & Tabatabai, 1979; Tabatabai, 1992). The amount of SOi- produced can be calculated from the amount of S20?- + S40l- + SOi- obtained by the methylene blue method by subtracting the values of S20?- and S40lobtained by the colorimetric methods.

45-2 SULFATE REDUCTION The inorganic S fraction in soils may occur as sulfate and compounds of lower oxidation state such as sulfide, polysulfide, sulfite, thiosulfate, and So. In well-drained, well-aerated soils, most of the inorganic S normally occurs as sulfate, and the amounts of reduced S compounds are generally < 1% (Freney, 1961). Under anaerobic conditions, particularly in tidal swamps and poorly drained or waterlogged soils, the main form of inorganic S in soils is sulfide and often So (Brummer et aI., 1971a,b; Harmsen, 1954; Hart, 1959). Reduced inorganic S compounds produced in or added to soils can be oxidized under aerobic and anaerobic conditions. The main reduced S compounds added to soils are sulfide (S2-), such as metal sulfides, elemental sulfur (SO), thiosulfate (S20?-), tetrathionate (S20i-), and sulfite (SO?-). Oxidation of these compounds in soils can be chemical (abiotic) or microbiological (biotic) or both. The chemical oxidation of aqueous sulfide by O 2 is relatively slow, with reported half-lives ranging from about one to several hours (Zehnder & Zinder, 1980). The studies by

1072

TABATABAI

Krebs (1929), however, showed that the reaction is catalyzed by traces of transition metal ions. Also, various mixtures of SO, polysulfides, S20l-, S40g-, SOl-, and SOl- have been reported as products at near neutral pH values, indicating that a complex set of reactions is involved (Zehnder & Zinder, 1980). Accurate determination of the reduced forms of inorganic S in soils is difficult, partly because of the ease with which they can be oxidized on exposure to air, but mainly because of the limitations of current analytical methods. No procedure has been entirely satisfactory for determination of sulfide in soils. Gilboa-Garber (1971) proposed a method for direct spectrophotometric determination of inorganic sulfide in biological materials. This method involves colorimetric determination of sulfide as methylene blue after extraction with Zn acetate. A modification of this method was successfully used by Howarth et ai. (1983) for determination of sulfide in porewaters of core samples of a salt marsh. Smittenberg et aI., (1951) proposed a method for determination of sulfide, sulfite, thiosulfate, polysulfide, So, and organically bound S in soils. It involves digestion of a soil sample with HCI for determination of the monosulfidic S compounds and digestion with Sn and HCI for determination of the total oxidizable S. In both methods the H 2S released is determined colorimetrically as methylene blue. None of these methods, however, gives accurate results; digestion of soil with HCI does not release the S from acid-insoluble metal sulfides, causing underestimation of the sulfide present, and digestion with Sn and HCI releases S from organic S, causing overestimation of the total oxidizable S (Melville et aI., 1971). The nature of this S fraction, however, has not been identified. Much of the reducible S fraction in soils is extractable with 0.5 M NaOH and is distributed between the fulvic acid and humic acid fractions. Therefore, digestion of soil with Sn and HCI may prove adequate where the amounts of oxidizable S are relatively large, but this method is unsatisfactory for use with soils containing small amounts of reduced inorganic S compounds, especially with surface soils containing appreciable amounts of organic S. Studies of pyrite formation and the measurement of sulfate reduction in salt marsh sediments by Howarth and Merkel (1984) showed that the reduction with Cr(H) of pyrite and So to H 2S is more specific and sensitive than oxidation of these forms of S to sulfate. The Cr(H) reduction method has been used in studies of inorganic S compounds in modern sediments and shales and of formation of 35S-labeled So and pyrite in coastal marine sediments during short-term 35S01- reduction measurements (Howarth & J! 90% of the S from FeS, So, and FeS2' Acetone extraction followed by Cr reduction of the filtrate was specific for So. Hydriodic acid reduction recovered > 90% of the S from FeS, SOj-, and p-nitrophenyl sulfate. The Zn-HCI procedure partially recovered S from SOj-, So, and FeS2' None of the above procedures reduced L-methionine. Analysis of both moist and oven-dried peat by Wieder et al. (1985) showed that ovendrying of peat samples increased the ester-sulfate Sand SOj- fractions and decreased the estimated C-bonded S, which was calculated from the difference between the total S and HI-reducible S. A polarographic method for determination of the reduced S species in marine porewaters has been proposed by Luther et al. (1985). They have demonstrated that, with polarographic techniques, it is possible to measure thiosulfate, sulfide, bisulfide, and polysulfide ions with a mercury electrode. They considered the polysulphide ions, S;-, to be composed of one S atom and in the -2 oxidation state, S(2-), and the remaining (x-I) S atoms in the zero-valent oxidation state S(O). The number of S atoms in each is measurable by this technique. Tetrathionate and other polythionates can be measured by this technique, but Luther et al. (1985) could not detect these reduced S species in the porewaters that they tested. By using this differential pulse polarographic technique, Luther et al. showed that salt marsh and subtidal porewater profiles contain significant concentrations of thiosulfate, bisulfide, and polysulfide. This technique has been successfully used in demonstrating the seasonal cycling of S and Fe in porewaters of a Delaware salt marsh (Luther & Church, 1988). The methods developed for estimating sulfate reduction are mainly used in studies of coastal marine sediments (Fossing & J~rgensen, 1989; J~rgensen, 1978a). Even though these methods are not commonly used in studies of sulfate reduction in soils, they will be described here for potential use in studies involving sulfate reduction in flooded soils. 45-2.1 Principles Estimation of sulfate reduction involves addition of SO]- into a homogenized or undisturbed sediment core or soil sample and determination of the reduced inorganic S produced by reduction with Cr2 + in an acid solution and determination of the H 2S released as methylene blue and by counting the radioactivity of the H 2S released when 35S0j- is used. This Cr-reducible S (single-step method) comprises H 2S, So, FeS, and FeS2' Alternately, sulfate reduction can be measured by a two-step method in which the acid volatile sulfide (H2S + FeS) and Cr-reducible S (SO + FeS2) are sequentially distilled from the sample. The fraction of 35S0]- reduced during incubation is calculated from the sum of 35S in the acid volatile sulfide and Cr-reducible S. The single-step distillation is simpler and faster than the consecutive distillation of acid volatile Sand Cr-reducible S. Comparison of the two methods by Fossing and J~rgensen (1989) showed that the single-step method resulted in higher (4-50%) sulfate reduction

1074

TABATABAI

rates than those obtained from the sum of 35S in acid volatile Sand Crreducible S. The difference was largest when the sediment had been dried after acid volatile S but before Cr-reducible S distillation. Relative to the 35S in the H 2S released by the acid volatile S distillation alone, the 35S of the H 2S released from the total reduced inorganic S (Cr-reducible S) distillation resulted in 8 to 87% higher reduction rates. Studies of sulfate reduction rates in a range of marine sediments and salt marshes have shown that 10 to 15% of the reduced radiolable 35S01- was recovered from Cr-reducible S (for review, see Fossing & Jjjrgensen, 1989), but the percentage varies widely, depending on the type of sediment and on the S chemistry. According to Fossing and Jjjrgensen (1989), the mechanism(s) involved in 35S incorporation into So and FeS2 during short-term incubation is not clear, but 35S labeled Cr-reducible S has to be taken into account in measurements of sulfate reduction rates. Because the single-step method is more accurate, convenient, and more rapid than the two-step method, the former method will be described here. 45-2.2 Methods (Fossing & Jjjrgensen, 1989) 45-2.2.1 Special Apparatus 1. Modified Johnson-Nishita apparatus (Tabatabai, 1982), the apparatus described by Canfield et al. (1986), or that described by Zhabina and Volkov (1978). 45-2.2.2 Reagents 1. Zn(OAch, 5% in 1% acetic acid. 2. Nitrogen gas and ferric ammonium sulfate and p-aminodimethylanaline sulfate solutions: Prepare as described by Tabatabai (1982). 3. Concentrated HCI. 4. Ethanol (95%). 5. Chromic chloride hexahydrate (CrCl3·6H20): Prepare the highly reactive Cr2 + solution from the more stable Cr3+ by percolating 1 M CrCI3·6H20 in 0.5 M HCI through a Jones reactor with amalgamated Zn granules: 2 Cr3+ +

Zn~2

Cr2+ + Zn2+

An efficient reduction is varified by a color change from dark green [Cr(IH)] to bright blue [Cr(H)]. For construction of the Jones reactor, see Kolthoff and Sandell, 1963, p. 569. The Jones reductor can be constructed from a glass-column (40 cm long, 1.5 cm i.d.) with an integral sinter at the bottom and stopcocks in both ends (Fossing & Jjjrgensen, 1989). Wash the granular Zn (0.3-1.5 mm grain size) three times with 1 M HCI and twice with deionized water.

SULFUR OXIDATION IN SOILS

1075

Amalgamate the Zn for a few minutes with a saturated solution of HgCl2 (ca. 0.25 M), transfer it to the glass column, and wash it with three column volumes of 0.5 M HCl. Remove the reduced Cr solution by suction into a large polyethylene syringe in which it can be stored under reduced conditions (bright blue) for several weeks. Otherwise, the Cr(II) solution should be freshly prepared every 2 to 3 d and stored in a ground-glass stoppered bottle (Canfield et aI., 1986). An alternate and faster method of producing large volumes of Cr(II) solution is recommended by Fossing and J~rgensen (1989). In this method, simply fill a glass bottle with 1 M HCl-rinsed "mossy zinc" (Aldrich Chemicals, Milwaukee, WI) and then fill the bottle with the Cr(III) solution (1 M CrCI3 ·6H20 in 0.5 M HCI) under continuous flow of N2. The mossy zinc does not need to be amalgamated. Chromium(III) is reduced to Cr(II) within 10 min and is kept under N2 until it is drawn into syringes through an outlet at the bottom of the bottle. After use, the mossy Zn can be regenerated by washing with 1 M HCl. 45-2.2.3 Procedure Sulfate reduction can be measured in undisturbed sediment core (3 cm in diameter) by the core injection technique described by J~rgensen and Fenchel (1974) and J~rgensen (1978a). In this method, a volume of 2 ILL carrier-free 35S0i- (70 kBq) is injected at a specific depth into replicate cores from each site or station. The sediment core is incubated for 18 to 24 h at room temperature (23°C), then it is cut into segments and transferred to 20 mLof20% (wt/vol) of Zn acetate [Zn(OAchl and frozen to terminate the reaction and fix the sulfides. The reduced S is then distilled as H 2S from the sediment into Zn( OAc h traps as described below and the radioactivity of SOi- and of the precipitated ZnS is determined. Sulfate concentration in porewater of sediments or soil samples can be determined turbidimetrically in replicated parallel core samples as described by Tabatabai (1974). Segments from each depth interval are pooled from the replicated cores and homogenized. The homogenized sediment is centrifuged and 35S0i- radioactivity is measured in a subsample of the supernatant. The sediment pellet is washed twice with a salt solution (e.g., 0.1 M NaCl) to remove 35S0i-. The washed sediment is homogenized and 1 to 2 g is transferred into a distillation flask (modified Johnson-Nishita apparatus, or the apparatus described by Canfield et aI., 1986; or that described by Zhabina and Volkov, 1978) and mixed with 5 mL of distilled water and 5 mL of methanol. The distillation flask is degassed for 20 min with N2, 16 mL of 1 M CrCl2 in 0.5 M HCI and 8 mL of 12 M HCI are added, and the sediment slurry is gently boiled for 40 min. During this distillation, the total reduced inorganic S is dissolved and distilled into Zn(OAch sequential traps containing 10 mL 5% Zn(OAch, buffered with 0.1 % acetate and with a drop of antifoam. More than 98% of the distilled H 2S is recovered as ZnS in the first trap. After distillation, the two traps are pooled, 5 mL is subsampled,

TABATABAI

1076

mixed with 5 mL of scintillation fluid (Dynagel, Baker Chemical), and 35S is counted. The concentration of the Cr-reducible S is determined spectrophotometrically at 670 nm in an aliquot of the trapping solution after adding 2 mL of ferric ammonium sulfate solution, 10 mL of p-aminodimethylanaline solution, and adjusting. the volume to 100 mL with deionized water. The sulfate reduction rate is calculated according to the following equation: SRR

= (SOl-) a (A

x 24 x 1.06 nmol sol- cm-3 d-1 a) h

+

where a is the total radioactivity of ZnS, A is the total radioactivity of SOjafter incubation, h is the incubation time in hours, (SOj-) is sulfate concentration in nmol per cm3 sediment, and 1.06 is a correction factor for the expected isotope fractionation (Jliirgensen & Fenchel, 1974; Fossing & Jliirgensen, 1989).

REFERENCES Audus, L.J., and J.H. Quastel. 1947. Selective toxic action of thiosulfate on plants. Nature (London) 60:264-265. Beaton, J.D., and R.L. Fox. 1971. Production, marketing, and use of sulfur products. p. 335-379. In R.A. Olson (ed.) Fertilizer technology and use. SSSA, Madison, WI. Briimmer, G., H.S. Grunwaldt, and D. Schroeder. 1971a. Contributions to the genesis and classification of Marsh soils. II. On the sulphur metabolism of muds and salt marshes. Z. Pflanzenernaehr. Dueng. Bodenkd. 128:208-220. Briimmer, G., H.S. Grunwaldt, and D. Schroeder. 1971b. Contributions to the genesis and classification of Marsh soils: III. Contents, oxidation status, and mechanisms of bounding of sulphur in polder soils. Pflanzenernaehr. Dueng. Bodenkd. 129:92-108. Canfield, D.E., R. Raiswell, J.T. Westrich, C.M. Reaves, and R.A. Berner. 1986. The use of chromium reduction in the analysis of reduced inorganic sulfur in sediments and shales. Chern. Geol. 54:149-155. Deng, S., and R.P. Dick. 1990. Sulfur oxidation and rhodanese activity in soils. Soil Sci. 150:552-560. Dick, W.A., and M.A. Tabatabai. 1979. Ion chromatographic determination of sulfate and nitrate in soils. Soil Sci. Soc. Am. J. 43:899-904. Fossing, H., and B.B. J!Ilrgensen. 1989. Measurement of bacterial sulfate reduction in sediments: Evaluation of a single-step chromium reduction method. Biogeochemistry 8:205-222. Fox, R.L., H.M. Atesalp, D.H. Kampbell, and H.P. Rhoades. 1964. Factors influencing the availability of sulfur fertilizers to alfalfa and corn. Soil Sci. Soc. Am. Proc. 28:406-408. Freney, J.R. 1961. Some observations on the nature of organic sulphur compounds in soils. Aust. J. Agric. Res. 21:424-432. Gilboa-Garber, N. 1971. Direct spectrophotometric determination of inorganic sulfide in biological materials and in other complex mixtures. Anal. Biochem. 43:129-133. Gleen, H., and J.H. Quastel. 1953. Sulfur metabolism in soils. Appl. Microbiol. 1:70-77. Guittonneau, G., and J. Keilling. 1932. L' evolution et al solubilisation du soufre elementaire dans la terra arable. Ann. Agron. 2:690-725. Hagstrom, G.R. 1984. Fertilizer sources of sulfur and their use. p. 567-581. In M.A. Tabatabai (ed.) Sulfur in agriculture. ASA, Madison, WI. Harmsen, G. W. 1954. Observations on the formation and oxidation of pyrite in the soil. Plant Soil 5:324-348.

SULFUR OXIDATION IN SOILS

1077

Hart, M.G.R. 1959. Sulphur oxidation in tidal mangrove soils of Sierra Leone. Plant Soil 11:215-236. Howarth, RW., A. Giblin, J. Gate, B.J. Peterson, and G.W. Luther III. 1983. Reduced sulfur compounds in the pore waters of a New England salt marsh. Environ. Biogeochem. 35:135-152. Howarth, RW., and B.B. Jl1Irgensen. 1985. Formation of 35S-labeled elemental sulfur and pyrite in coastal marine ~ediments (Limfjorden a~d Kysing Fj?rd, Denmark), during short-term 35S0}- reductIOn measurements. Geochlm. Cosmochim. Acta 48:1807-1818. Howarth, R.W., and S. Merkel. 1984.~te formation and the measurement of sulfate reduction in salt marsh sediments. Llmnol. Oceonogr. 9:598-608. Janzen, H.H., and J.R. Bettany. 1987a. Measurement of sulfur oxidation in soils. Soil Sci. 143:444-452. Janzen, H.H., and J.R. Bettany. 1987b. The effect of temperature and water potential on sulfur oxidation in soils. Soil Sci. 144:81-89. Janzen, H.H., J.R Bettany, and J.W.B. Stewart. 1982. Sulfur oxidation and fertilizer sources. p. 229-240. In Proc. Alberta Soil Science Workshop, Edmonton, Alta. Jl1Irgensen, B.B. 1978a. A comparison of methods for the quantification of bacterial sulfate reduction in coastal marine sediments: I. Measurement with radiotracer techniques. Geomicrobiol. J. 1:11-27. Jl1Irgensen, B.B. 1978b. A comparison of methods for the quantification of bacterial sulfate reduction in coastal marine sediments: II. Estimation from chemical and bacteriological field data. Geomicrobiol. J. 1:49-64. Jl1Irgensen, B.B., and T. Fenchel. 1974. The sulfur cycle of a marine model system. Mar. BioI. 24:189-201. Koehler, F.E., and S. Roberts. 1983. An evaluation of different forms of sulfur fertilizers. p. 833-842. In A.1. More (ed.) Proc. Int. Sulphur '82 Conf., Vol. 2, London. Nov. 1982. The British Sulphur Corporation Limited. Kolthoff, I.M., and E.B. Sandell. 1963. Textbook of quantitative inorganic analysis. 3rd ed. Macmillan, New York. Konopka, A.E., R.H. Miller, and L.E. Sommers. 1984 Microbiology of the sulfur cycle. p. 23-55. In M.A. Tabatabai (ed.) Sulfur in agriculture. ASA, Madison, WI. Krebs, H.A. 1929. Uber der Wirkung der schwermetalle auf die Autoxydation der Alkalisulfide und des Schwefewasserstoffs. Biochim. Z. 204:344-346. Laishley, E.J., R.D. Bryant, B.W. Kobryn, and J.B. Hyne. 1983. The effect of particle size and molecular composition of elemental sulphur on ease of microbiological oxidation. Alberta Sulphur Res. Ltd. Q. Bull. 20:33-50. Li, P., and A.L. Caldwell. 1966. The oxidation of elemental sulfur in soil. Soil Sci. Soc. Am. Proc. 30:370-372. London, J., and S.C. Rittenberg. 1964. Path of sulfur in sulfide and thiosulfate oxidation by thiobacilli. Proc. Natl. Acad. Sci. USA 52:1183-1190. Luther, III, G.W., and T.M. Church. 1988. Seasonal cycling of sulfur and iron in porewaters of a Delaware salt marsh. Mar. Chern. 23:295-309. Luther, III, G.W., E.A. Giblin, and R. Varsolona. 1985. Polarographic analysis of sulfur species in marine porewaters. Limnol. Oceanogr. 30:727-736. Melville, G.E., J.R. Freney, and C.H. Williams. 1971. Reduction of organic sulfur compounds in soil with tin and hydrochloric acid. Soil Sci. 112:245-248. Neptune, A.M.L., M.A. Tabatabai, and J.J. Hanway. 1975. Sulfur fractions and carbonnitrogen-phosphorus-sulfur relationships in some Brazilian and Iowa soils. Soil Sci. Soc. Am. Proc. 39:51-55. Nevell, W., and M. Wainwright. 1987. Influence of soil moisture on sulphur oxidation iIi brown earth soils exposed to atmospheric pollution. BioI. Fert. Soils 5:209-214. Nor, Y.M., and M.A. Tabatabai. 1975. Colorimetric determination of microgram quantities of thiosulfate and tetrathionate. Anal. Lett. 8:537-547. Nor Y.M., and M.A. Tabatabai. 1976. Extraction and colorimetric determination of thiosulfate and tetrathionate in soils. Soil Sci. 122: 171-178. Nor, Y.M., and M.A. Tabatabai. 1977. Oxidation of elemental sulfur in soils. Soil Sci. Soc. Am. J. 41:736-741. Smittenberg, J., G.W. Harmsen, A. Quispel, and D. Otzen. 1951. Rapid methods for determining different types of sulphur compounds in soil. Plant Soil 3:353-360. Starkey, R.L. 1956. Transformation of sulfur by microorganisms. Ind. Eng. Chern. 48:14291437.

1078

TABATABAI

Sulphur Institute. 1977. Agri-Sul starts production in Canada. Sulphur Inst. J. 12:2-3. Tabatabai, M.A. 1974. Determination of sulphate in water samples. Sulphur Inst. J. 10:1113. Tabatabai, M.A. 1984. Importance of sulphur in crop production. Biogeochemistry 1:45-62. Tabatabai, M.A. 1982. Sulfur. p. 501-538. In A.L. Page et al. (ed.) Methods of soil analysis. Part 2. 2nd ed. ASA and SSSA, Madison, WI. Tabatabai, M.A. 1992. Methods of measurement of sulphur in soils, plant materials, and water. p. 307-344. In R.W. Howarth et al. (ed.) Sulphur cycling on the continents: Wetlands, terrestrial ecosystems and associated water bodies. SCOPE 48. John Wiley and Sons, New York. Tabatabai, M.A., and A.A. AI-Khafaji. 1980. Comparison of nitrogen and sulfur mineralization in soils. Soil Sci. Soc. Am. J. 44:1000-1006. Tabatabai, M.A., and J.M. Bremner. 1972a. Forms of sulfur and carbon, nitrogen and sulfur relationships in Iowa soils. Soil Sci. 114:380-386. Tabatabai, M.A., and J.M. Bremner. 1972b. Distribution of total and available sulfur in selected soils and soil profiles. Agron. J. 64:40-44. Trudinger P.A. 1959. The initial products of thiosulfate oxidation by Thiobacillus X. Biochern. Biophys. Acta 31:270-272. Vishniac, W., and M. Santer. 1957. The thiobacilli. Bacteriol. Rev. 21:195-213. Wainwright, M. 1984. Sulfur oxidation in soils. Adv. Agron. 37:349-396. Waksman, S.A., and J.S. Joffe. 1922. Oxidation of sulfur in the soil. J. Bacteriol. 7:231-256. Walker, T.W. 1964. The use of sulfur as fertilizers. Agrochimica 9:1-14. Weir, R.G. 1975. The oxidation of elemental sulphur and sulphides in soil. p. 40-49. In K.D. McLachlan (ed.) Sulphur in Australasian agriculture. Sydney Univ. Press, Sydney, Australia. Wieder, R.K., G.E. Lang, and v.A. Granus. 1985. An evaluation of wet chemical methods for quantifying sulfur fractions in freshwater wetland peat. Limnol. Oceanogr. 30:11091115. Zehnder, A.J.B., and S.H. Zinder. 1980. The sulphur cycle. p. 105-145. In O. Hutzinger (ed.) The natural environment and the biogeochemical cycles. Springer-Verlag, Berlin. Zhabina, N.N., and 1.1. Volkov. 1978. A method for determination of various sulfur compounds in sea sediments and rocks. p. 735-746. In W.E. Krumbeih (ed.) Environmental biogeochemistry and geomicrobiology. Vol. 3. Ann Arbor Sci., Ann Arbor, MI.

Published 1994

Chapter 46 Iron and Manganese Oxidation and Reduction WILLIAM C. GHIORSE, Cornell University, Ithaca, New York

Microbial oxidation and reduction of Fe and Mn are of wide-ranging importance to soil scientists (Alexander, 1977; Paul & Clark, 1989). Indeed, knowledge of the distribution, abundance, identity, and activity of Fe- and Mn-transforming microbes in soils and sediments can greatly enhance studies on such diverse agricultural and environmental problems as Fe and Mn availability to plants, metal accumulation, toxicity and mobility of metals and pesticides, and clogging in wells and wetland drainage systems. Knowledge of the biology of Fe- and Mn-transforming microorganisms may allow for future applications in which the metal mobilization and immobilization activities of these microorganisms are exploited for economic and environmental benefit (Ehrlich & Brierley, 1990). Except for the morphologically recognizable "iron bacteria," relatively little is known of the occurrence of Fe-Mn-transforming organisms in nature. Even less is known of their function in natural systems or the factors controlling their in situ activities. On the other hand, several model organisms have been isolated and characterized taxonomically (e.g., Thiobacillus ferrooxidans, Leptothrix discophora, Shewanella putrefaciens, and Geobacter metallireducens (Lovley et aI., 1993)). In some cases, the biochemical mechanisms underlying their Fe- and Mn-transforming abilities have been investigated. (For reviews, see Ghiorse 1984, 1988; Ehrlich, 1987, 1990; Lovley, 1987, 1991; Nealson et aI., 1988, 1989; Myers & Nealson, 1990; Ehrlich et aI., 1991; Nealson & Myers, 1992). A persistent problem has been the difficulty of distinguishing abiotic from biologically mediated (biotic) transformations, especially in environments like soil where microbial activity may alter the redox chemistry of the microenvironment, causing Fe and Mn redox changes to occur by direct or indirect mechanism (Ehrlich, 1990). These problems also apply to microbial growth media which, in some instances, may be altered by growthinduced changes in pH or Eh or metabolic products that cause chemical oxidation or reduction of Fe and Mn. These possibilities are taken into Copyright © 1994 Soil Science Society of America, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties-SSSA Book Series, no. 5. 1079

1080

GIDORSE

account when media are selected for enumeration, enrichment, and isolation of Fe- and Mn-transforming microorganisms. Bacteria and fungi have been implicated in both oxidation and reduction processes in soil and sediment primarily by applying cultural methods for the detection and enumeration of microorganisms that can cause Fe or Mn oxidation or reduction in selected media. Measurement of the disappearance or production of soluble forms of the metals have been used to indicate activity in soil. Inhibition of the activity by heat, metabolic inhibitors, antibiotics, and other biostatic and biocidal agents is used to establish that microbial activity is involved in the processes. The presence of Fe-Mn-transforming organisms in samples can be taken as evidence for their potential activity in the environment. Relatively few studies have addressed the problem of measuring in situ rates of microbial activity in soil because well-tested methods and standard protocols are not available. The current literature contains ample evidence establishing the potential of soil and aquatic microorganisms to oxidize and reduce Fe and Mn, but definitive experiments showing the relative contributions of abiotic and biologically mediated reactions are lacking. It should be noted that this criticism can be made of almost any microbial process in most natural environments. Development of the required methods and protocols to overcome this problem awaits new methods for estimating activity in samples from natural environments. This may require combinations of conventional methods such as those described in this book with new in situ rate measurement techniques, as well as microscopic molecular probe techniques for distinguishing the genetic potential and expression of specific genes in individual cells. Recent advances in microbial ecology, cell biology and molecular probe technology hold much promise that such methods soon will be available. A future challenge will be to apply these powerful techniques in natural systems such as the soil and sediment that are the focus of this chapter. The approach in this chapter is to present well-established principles and methods for analysis of organisms in soil, sediments, and drainage systems responsible for biological Fe and Mn transformations. Microscopic observations, especially combined phase contrast and epifiuorescence microscopy are recommended as the first step in examining any sample that may contain active microorganisms; however, extensive description of the morphological features of iron bacteria are beyond the scope of the chapter (for recent descriptions see Hanert, 1981a,b; Mulder & Deinema, 1981; Ghiorse, 1984a; Hackett & Lehr, 1986; Jones, 1986; Ghiorse & Ehrlich, 1992). Enumeration and isolation methods form the core of the chapter. In some cases, the organisms are easily counted and isolated directly from colonies on enumeration plates. Others are readily obtained in enrichment cultures, but do not readily form colonies on solid media. Most probable number (MPN) enumeration methods are recommended for the latter.

OXIDATION AND REDUCTION OF FE AND MN

1081

46-1 IRON-DEPOSITING AND MANGANESE-OXIDIZING HETEROTROPHS 46-1.1 Enrichment and Isolation 46-1.1.1 Principles Heterotrophic Fe-depositing and Mn-oxidizing bacteria and fungi occur widely in soil and aquatic sediments. Most are strict aerobes, some are microaerophiles, and a few are facultative anaerobes (Ghiorse, 1984a). Their enrichment and isolation are facilitated by the wide range of possible media for heterotrophs, but excess organic nutrients may hinder detection of Fe- and Mn-depositing activity. Care must be exercised in selecting media with regard to the concentration of organic C, phosphate, and Fe and Mn sources. Possible abiotic reactions of Fe and Mn with media components should be considered. Low organic nutrient and low phosphate media usually produce the best results. Iron and Mn sources must provide enough soluble Fe or Mn to reveal activity without hindering growth. Low solubility sources or low concentrations (mg/Kg) of Fe and Mn generally are most successful. For enrichment and isolation media, consideration of the composition of the soil or sediment is important as well as the chemical composition of the medium. The use of a natural material extract medium, possibly supplemented with a small amount of yeast extract or peptone is recommended. The strategy is to stimulate growth of the heterotrophs while limiting abiotic reactions. Microorganisms that possess the ability to deposit (accumulate) Fe(III) oxides or oxidize Mn(II) should be encouraged to grow, while controlling the growth of other bacteria. Low incubation temperature may help to achieve the latter goal. Oxygen is required for the formation of the metal oxides, but microaerophilic conditions may favor growth of the bacteria involved. Carbon dioxide has been reported to stimulate the processes in soil enrichments under microaerophilic conditions (Uren & Leeper, 1978; Hanert, 1981b; Mulder & Deinema, 1981). Thus, if possible, it is advantageous to use low-oxygen enrichment strategies that encourage the growth of microaerophiles. A variety of complex media have been tested for enrichment of Fedepositing and Mn-oxidizing bacteria from soil and sediments (Bromfield & Skerman, 1950; Chapnick et aI., 1982; Cullimore & McCann, 1977; Hanert, 1981b; Ghiorse, 1984a; Ehrlich, 1990). These media range from natural water or soil slurries to complex mixtures containing high concentrations of organic nutrients. Generally, pH is adjusted to the neutral range (6-8), but more acid media have been used when appropriate (Bromfield, 1978). Media in the circumneutral pH range will not reveal enzymatic Fe(II) oxidation because Fe(II) autoxidizes at pH> 3. The autoxidation

GHIORSE

1082

occurs more rapidly in water at neutral pH under aerobic conditions. Thus, at neutral pH, the propensity of a culture to accumulate colloidal Fe (III) oxide is revealed, rather than the ability to oxidize Fe(II). Contrastingly, enzymatic Mn oxidation is detected in circumneutral pH media; though at a pH above 8, Mn oxidation will occur abiotically. Salts of Fe(lI) such as FeCI2 , or FeS04 are used to produce Fe (III) oxide, which forms spontaneously in Fe (II)-containing neutral pH media exposed to 02' Reduced Fe sources include metallic Fe as iron powder or iron wire, FeS, FeCI2 , FeS04' and various Fe(III)-organic complexes; sources of Mn(lI) include MnC03 , MnCI2 , MnS04' and organic Mn(lI) complexes (Ghiorse & Hirsch, 1978). Organic nutrients are provided with adequate C, N, P, trace elements, and growth factors to support heterotrophic growth in the presence of Mn(II) or Fe (III) oxide. lron- and Mn-oxidizing fungi often will grow on the complex media designed for heterotrophic bacteria. To detect the maximum number of Fe- and Mn-depositing bacteria, it is advisable to use both the natural material based medium and the complex medium listed below. Detection is based largely on color of colonies. The Fe(llI) oxides give a yellowish to reddish brown hue to colonies of Fe-accumulating bacteria, depending on the concentration and mineral type of oxide. The Mn(lIl, IV) oxides produce a yellowish brown to dark brown color in colonies. Confirmation of the presence of Fe (III) or Mn(lIl, IV) oxides is done with spot test reagents; Prussian blue for Fe (III) oxides, a dye reaction (e.g., leucocrystal violet) or chemical test (production of O 2 from H 2 0 2 ) for Mn (III, IV) oxides. 46-1.1.2 Materials 1. Diluted Fe- or Mn-containing extract medium. 2. Diluted Fe- or Mn-containing PYG medium. 3. Spot test reagents for Fe (III) and Mn(III, IV) oxides. Prussian Blue reagents for Fe(III); 1 N of HCI and 1 % potassium ferrocyanide. Add one drop of acid followed by a drop of ferrocyanide solution. Blue color indicates Fe(III). Leuco crystal violent reagent (Kessick et aI., 1972) for Mn(lIl, IV) oxides. Dilute 2 mL of a 0.1% leuco crystal violet l in 0.1 N of perchloric acid with 5 mL 6 M acetic acid-acetate buffer (final pH = 4.0). Add a drop of test material. Violet color indicates Mn(lIl, IV) oxide. The dye solution oxidizes slowly in the presence of O 2 or upon exposure to UV light. Store spot test reagents in dark at 4 °C. 4. 0.01% aqueous acridine orange solution.

1 Leuco crystal violet and other leuco bases of common triphenyl methane dyes such as leuco malachite green are readily oxidized by Mn(III, IV) oxides. The leuco bases are available from organic chemical supply companies.

OXIDATION AND REDUCTION OF FE AND MN

1083

46-1.1.3 Procedures

Natural material based enrichment media can be prepared from extract of soil, or sediment. For enrichments from soil or sediment, prepare an extract from a slurry of material mixed 1:1 with distilled water (e.g., 50 gl50 mL). The slurry is filtered (Whatman no. 1) and the clarified filtrate is filter sterilized (0.2 !tm) and used to prepare 50 mL of Fe- or Mn-containing enrichment medium in 250-mL Erlenmeyer flasks. Add 0.1 g of FeC03 , 0.25 to 0.5 g ofFeo wire or Feo powder or 0.05 g ofMn S04·H20 or MnCI 2, or 0.1 g of MnC0 3 to the enrichment flasks. Addition of 0.1 giL yeast extract is optional. It will provide small amounts of nutrients and, stimulate more rapid growth (Hanert, 1981b). The enrichment cultures are inoculated with a small amount (0.1-1 g) of soil or sediment. Cultures should be incubated in cotton-plugged flasks, stationary in the dark. Shaking may inhibit slow-growing microaerophiles. Incubation temperature should be within ± 5 °C of the mean daily temperature of the environment under investigation. Lower temperature incubation with no added nutrients promotes slower growth of all bacteria in the enrichment culture. Many Feand Mn-depositing bacteria (e.g., prosthecate bacteria) prefer low nutrient concentrations and, therefore, grow more slowly than other heterotrophs that can rapidly exploit available nutrients. Long-term (weeks), low-temperature incubation under low-nutrient conditions may produce different results than short-term ambient temperature incubation. Therefore, it is advisable to set up enrichment cultures under various nutrient and temperature regimes. Microaerobic or oxygen gradient enrichments can be established by adding 0.1 % molten agar (Difco Noble or Difco Purified Agar) or agarose to the medium. If purified agar is not available, standard grade agar can be used if it is washed several times with distilled water to remove soluble organics. Washing is monitored as a decline in intensity of the straw-yellow color of the wash water. The medium is dispensed into 15 x 150 mm screw capped test tubes to fill them completely with a small (max. 5 mm) air space at the top. Proportional amounts of solid Fe or Mn sources are added to each tube to maintain the concentrations indicated above. Most of the O 2 can be stripped from the medium by bubbling with 0.2 !tm-filtered N2 before inoculation. Inoculate with a small amount of soil or sediment. Screw caps are left loose to allow air to enter the tube at the top. Growth of microaerophilic bacteria will occur in the oxygen gradient and form bands of growth and iron or manganese oxide beneath the surface of the tube. Numerous enrichment media for Fe- and Mn-depositing heterotrophs are described in the literature (Ghiorse & Hirsch, 1978; Hanert, 1981b; Cullimore & McCann, 1977). The most successful of these contain a very low concentration of complex organic nutrients (0.25 giL or less of yeast extract, peptone, or beef extract, or mixtures of these components) and mineral salts. The concentration of phosphate salts should be kept low because excess phosphate may interfere with Fe and Mn

1084

GHIORSE

oxidation. Addition of an inorganic N source generally is not required if peptone or beef extract is used. If these complex sources of N are not used, then (NH4hS04 and KN0 3 should be added. A useful general purpose medium for growth of soil and sediment consists of 0.25 giL each of peptone, yeast extract, and glucose (PYG) plus 0.5 gIL of MgS0 4·7H20 and 0.01 gIL of CaCI2 • A vitamin mixture can be added if desired (Staley, 1968). A 10- or 100-fold dilution of this medium with an Fe or Mn source added as indicated above provides a balanced low-nutrient enrichment medium in which many Fe- and Mn-depositing bacteria can grow. Enrichment cultures should be checked periodically (at least once a day at the beginning) for signs of growth (turbidity) as well as for Fe or Mn deposition. When O 2 is low, Fe may oxidize slowly at intermediate levels, or near its source at the bottom of the flask or tube. When O2 is high, gel-like deposits of FeOOH may form quickly. Iron-depositing bacteria usually form rusty colored films or colonies of rust-colored spots on the walls of glass containers. Manganese-oxidizing bacteria similarly form brownish films or small spots. Suspected colonies and films should be examined microscopically for the presence of bacteria and by the spot tests for Fe(IH) or Mn(H) oxide. To test for the presence of bacteria in the Fe-Mn oxide deposits, collect three colonies or fragments of film from the vessel wall with a pasteur pipette. Place one on a piece of filter paper and test with a drop of leuco crystal violet spot test reagent. Mn(HI, IV) oxides turn blue. The second colony is tested with Prussian Blue reagents for Fe(IH) oxide. Mount the third on a glass microscope slide in a drop of 0.1 giL of acridine orange under a coverslip. View by phase contrast and epifluorescence microscopy at a magnification of 500 to 1000 x . The phase contrast image will reveal characteristic shapes and structures of Fe- and Mn-depositing bacteria but, the diagnosis of bacteria in the aggregates is confirmed by epifluorescence. The acridine orange-DNA and -RNA complexes in the bacterial cells will shine through the Fe and Mn oxides with a bright orange, yellow, or apple-green fluorescence (see color plates in Hanert, 1981b). This direct method of detection is recommended for screening enrichment cultures. Once diagnosed, the bacterial growth is streaked on the natural material extract medium or the lO-fold diluted PYG medium solidified with at least 1.5% agar. Higher agar concentrations can be used to suppress rapidly spreading bacteria in the sample. Isolation may be facilitated by a second passage in the enrichment medium followed by streaking on agar plates. Select colonies containing Fe or Mn oxide using the same spot tests described above. Repeat streaking and colony selection until you are satisfied that an axenic (uniform morphology) culture is obtained. Bear in mind that amorphous Fe and Mn oxides formed in the cultures may be mistaken for bacteria in the microscope; also, the oxides may concentrate nutrients that attract bacteria. Satellite bacteria are commonly associated with Fe and Mn deposits in enrichment cultures. The satellite bacteria are difficult to remove by streaking on agar; therefore, it may be advantageous

OXIDATION AND REDUCTION OF FE AND MN

1085

to purify the culture by streaking on media without an added Fe or Mn source. Most Fe- or Mn-depositing heterotrophs obtained in such enrichments do not depend on Fe or Mn deposition for growth. Many Mndepositing bacterial lose this property if they are transferred frequently on artificial media. Iron-depositing bacteria generally do not lose this property. Fungal and actinomycete growth will be recognized by colony morphology and microscopic examination. Fungal hyphae are much thicker (5-10 !-tm) than actinomycete hyphae (0.5-1.0 !-tm). Both can become heavily encrusted with Fe or Mn oxides in older cultures (Bromfield & Skerman, 1950), but Fe or Mn deposits may form at some distance from the hyphae in young cultures (Bromfield, 1978; Emerson et aI., 1989). 46-1.2 Dilution Spread Plate Counts 46-1.2.1 Principles The number of Fe- and Mn-depositing microorganisms in soil and sediment can be estimated by a viable cell plate count. This is achieved by serially diluting the sample in a suitable diluent and plating samples by the spread plate method. As a rule of thumb, the viable count of aerobic heterotrophic bacteria in soil or sediment will be 10% or less of the total count, and the number of viable Fe- and Mn-depositing will be a small fraction of the viable count, frequently < 10%. In principle, Fe depositors should outnumber Mn depositors, because Fe deposition can be a nonspecific Fe-oxide accumulation process. Manganese-oxidizing activity appears to be a more specific, enzyme-linked process. Thus, it is less commonly observed among heterotrophic bacteria than iron deposition (Ghiorse, 1984a). The dilutions to be plated should be selected for each sample by estimating the number of aerobic heterotrophs in the sample. This can be determined from a total direct count (see chapter 8 by Zuberer in this book) or by a preliminary aerobic heterotrophic plate count on lO-fold diluted PYG agar (46-2.1.3). The number of Fe- or Mn-depositing bacteria should be in a range of 0.1 to 10% of the aerobic heterotroph plate count or 0.01 to 1.0% of the total direct count. Three or four of the lO-fold dilution tubes should be plated in triplicate for statistical accuracy. 46-1.2.2 Materials 1. Diluted extract Fe or Mn agar. 2. Diluted Fe or Mn PYG agar. 3. Ninety-milliliter sterile dilution buffer consisting of a 1:800 dilution of 34 g of KH 2POiL of distilled water, adjusted to pH 7.2 with 1 N of NaOH (Seeley & VanDemark, 1981). 4. One hundred-milliliter milk dilution bottles. 5. Pipettes, 1 and 10 mL. 6. Bent glass rod spreader. 7. Rotary spread plate table.

GmORSE

1086

8. Toothpicks. 9. Spot test reagents for Fe and Mn (46-1.1.2). 46-1.2.3 Procedure Prepare diluted extract Fe or Mn agar following procedures described in 46-1.1.3, but adding 20 gIL of agar. Insoluble powders (Feo, FeC03 MnC0 3) can be dispersed evenly in the molten agar and poured into plates. Iron wire is not recommended for plate counts. Follow the usual procedures for spread plate counts of soil or sediment. Aseptically add 10 g of soil or sediment to 90 mL of sterile dilution buffer in a 125-mL milk dilution bottle. Serially dilute the sample in 3 or 4, 10-fold dilutions to arrive at the proper range as estimated above. Dispense 1.0 or 0.1 mL to each plate. Evenly spread the suspension on the plate using a bent glass rod and rotary table. Allow excess liquid to absorb into the agar before inverting. Bind the edge of the plate with parafilm to prevent drying. Incubate at 20°C or appropriate temperature ± 5 °C above mean daily temperature of the environment from which the sample was collected. Check daily at first for growth and appearance of yellow or brown colonies that test positive for Fe or Mn oxide by spot tests (46-1.1.1). Use a toothpick to remove a small piece of the colony. Immerse it in spot test reagents. An initial count can be done at 5 to 7 d, but some colonies will accumulate Fe and Mn oxides more slowly. Allow at least 21 d before final counts are done. Be careful when counting colonies that have grown out from brown or black Fe-Mncontaining particles. These may test positive simply because the original material contains (Fe(III) or Mn(III, IV) oxide. 46-2 IRON-OXIDIZING AUTOTROPHS 46-2.1 Acidopblles (Enrichment, Isolation, and Enumeration) 46-2.1.1 Principles Iron-oxidizing autotrophic bacteria use Fe2 + as an energy source and CO2 as a C source. They are divided arbitrarily into two groups based on the pH range for growth. The acidophiles grow at pH values < 5, optimally at pH 2 to 4; neutrophiles (see section 46-2.2) grow at pH above 5, and optimally at pH 6.5 to 7.5. The aqueous chemistry of Fe governs availability of the energy source, Fe2 +. This species becomes unstable in the presence of O 2 as the pH increases above 3. The rate of abiological Fe2 + oxidation in water or soil solution is slow at pH 2 to 3 or less, but at pH above 5, when O 2 is abundant, the rate increases. Thus, aerobic bacteria depending on Fe2 + as an energy source are either acidophilic in acidic, high O 2 environments or microaerophilic in neutral pH environments. Acidophilic bacteria occur widely in mining regions where coal and mineral deposits contribute sulfide minerals (e.g., pyrite) to the soil and sediments. These minerals are oxidized by the bacteria-producing sulfuric acid and

OXIDATION AND REDUCTION OF FE AND MN

1087

metal ions (Fe 2+) in solution. The Fe2+ is oxidized by acidophilic Feoxidizing bacteria. Thiobacillus ferrooxidans is the best-known acidophilic S- and Feoxidizing bacterium in soi1. This organism is a mesophile (Topt = 30-35 0c) (Alexander, 1977; Kuenen & Tuovinen, 1981; Ehrlich, 1990; Norris, 1990). Several other mesophilic Fe-oxidizing autotrophic acidophiles such as Leptospirillum ferrooxidans (Norris, 1990) have been described. In addition, several thermophilic Fe and S oxidizers are known (Kuenen & Tuovinen, 1981; Ehrlich, 1990; Norris, 1990). Thiobacillus ferrooxidans uses both reduced S and Fe as sources of energy; L. ferrooxidans uses only Fe. Both organisms may occur in the same enrichment cultures when Fe2+ is provided as the sole energy source and CO2 is the C source. Isolation of Fe-oxidizing autotrophs is best achieved by streaking the enrichment culture on a solidified medium. The use of highly purified agar or agarose (see Kuenen & Tuovinen, 1981; Mishra et a1., 1983; Holmes & Yates, 1990) is necessary because organic compounds tend to inhibit growth of the autotrophs or promote the growth of acidophilic heterotrophs. Once isolated colonies are obtained, the culture must be checked thoroughly for the presence of satellite heterotrophs that frequently are found associated with these organisms (Ehrlich, 1990). No standard enumeration procedure has been published for acidophilic Fe-oxidizing autotrophs in soil per se; however, methods for viable cell plate counts have been published for use in pure cultures and acid mine drainage water (Kuenen & Tuovinen, 1981; Tuovinen & Kelly, 1973; Mishra et a1., 1983). 46-2.1.2 Materials The medium of Silverman and Lundgren (1959) (9K medium) and the medium of Tuovinen and Kelly (1973) (TK medium) or Kuenen and Tuovinen (1981) (KT medium) are suitable for cultivating Fe-oxidizing thiobacilli and leptospirilla and similar organisms (Ehrlich, 1990). 9K medium contains in giL: FeS04·7H20, 44.22; (NH4)S04, 3.0; KCI, 0.1; K2HP0 4, 0.5; MgS0 4·7H20, 0.5; Ca(N0 3 h, 0.01. Dissolve the FeS04·7H20 in 300 mL of distilled water; adjust to pH~2 with 1 mL of 10 N H 2S0 4; autoclave, cool, and mix with filtered FeS0 4·7H20 solution. The TK medium contains in giL: FeS04·7H20, 33.3; (NH4) S04' 0.4; K2HP0 4, 0.5; MgS0 4·7H20, 0.4. Dissolve salts in 1 L of O.IL N of H 2S04. Sterilize by passage through a 0.2 !lm filter. KT medium contains in g·L -1: K2HP0 4, 0.5; (NH4hS04' 0.5; MgS0 4·7H20, 0.5; and 1 N H 2S04, 5.0 mL (Solution I). Solution II contains in gIL: FeS0 4·7H20, 167; 1 N H 2S04, 50 mL. Solutions I and II are sterilized by autoclaving. Mix 4 parts Solution I and 1 part Solution II for the final medium, pH~2. The pH may be varied by altering the amount of H 2S0 4. Lowering the pH to 1.3 will avoid formation of a ferric iron precipitate that forms at higher pH, but growth may be inhibited. Media can be solidified by adding 1% or less electrophoresis-grade agarose (Holmes & Yates, 1990; Mishra et a1., 1983) or

1088

GmORSE

washed high-purity agar (Tuovinen & Kelly, 1973; Kuenen & Tuovinen, 1981) to the liquid media. For solid 9K medium, add 10 g of solidifying agent to the 700 mL of salts solution before autoclaving. For solid TK medium, autoclave 10 g of agar or agarose in one-half of the H 2S04 solution, dissolve salts in the remaining 500 mL; mix after autoclaving. For KT medium mix 4 parts Solution I, 2 parts Solution II, and 4 parts 1% agar or agarose after autoclaving. 46-2.1.3 Procedure

Enrichment cultures are prepared by adding l00-mL of sterile 9K or TK medium to 300-mL sterile Erlenmeyer flasks and inoculating with 1 g of soil or sediment. Flasks are incubated at 25 to 30°C on a rotary shaker operated at 120 rpm. Aeration can be improved by using baffle flasks. Carbon dioxide can be supplied by bubbling glass-wool filtered 1% CO2 air mixture in glass tubes inserted through a rubber stopper. Ferrous iron oxidation is indicated by the formation of a yellow-orange mixed precipitate consisting of Fe2(S04h that forms above pH 2. The enrichment cultures should last at least 14 to 21 d. Ferrous Feoxidizing autotrophs grow slowly. Thiobacilli growing on Fe2+ and CO2 may oxidize 100 mol of Fe2+ for each mole of CO2 fixed (Ehrlich, 1990). Therefore, Fe precipitates may be visible in a culture long before the cells causing the precipitate are seen; the number of cells produced is very low. Acidophilic hetereotrophs may appear first. Thiobacillus ferrooxidans is a short gram-negative rod; Leptospirillum ferrooxidans is a slender vibrioid gram-negative rod. Both are motile with polar flagella. Consult original papers listed in Ehrlich (1990) and Norris (1990) for description of these and other acidophiles. Isolation can be achieved by streaking enrichment cultures in the usual manner on solidified media. Bind the plates with parafilm and incubate for 14 to 21 d at 25 to 30°C. Check for growth of rust-colored colonies. Streak colonies to purify them taking care to note colony and cellular morphology. Look for morphologically different satellite bacteria in the microscope. The satellite bacteria are acidophilic heterotrophs living from organic acids and alcohols excreted by the autotrophs (Ehrlich, 1990); therefore, they are never present in high numbers. These heterotrophs will grow when organic energy sources are supplied, but they will not oxidize Fe2+ (Norris, 1990). Conversely, true Fe2+ oxidizing autotrophs will grow best when supplied Fe2+ and CO2 in the absence of organic energy sources. Enumeration can be achieved with a standard MPN technique using the 9K, TK, or KT media under conditions described for enrichment cultures. Tubes that show the formation of the yellow-orange precipitate are scored positive. Viable cell plate counts may be done by the dilution spread plate technique (described above) using solid 9K, TK, or KT medium. Count Prussian Blue-positive rust-colored colonies after 14 to 21 d of incubation. Growth on agar media may be enhanced by low oxygen tension (Kuenen & Tuovinen, 1981).

OXIDATION AND REDUCTION OF FE AND MN

1089

46-2.2 Neutrophiles (Enrichment, Isolation, and Enumeration) 46-2.2.1 Principles A large number of neutrophilic (pH 5-8) Fe-depositing bacteria are included in a group known classically as iron bacteria (Hanert, 1981a,b; Mulder & Deinema, 1981; Ghiorse, 1984a; Jones, 1986). This group includes many genera known to deposit Fe in morphologically distinctive structures (Pringsheim, 1949). It is questionable whether any of these bacteria are truly chemolithoautotrophic as defined originally by Winogradsky (1888) (see Ghiorse, 1984, for discussion). An exception appears to be Gallionella ferruginea, which has long been suspected to be a chemolitho autotrophic bacterium using Fe2+ oxidation as an energy source to fix CO2 (Wolfe, 1964; Hanert, 1981a; Ghiorse, 1984). Recently, G. ferruginea has been shown to possess key enzymes for CO2 fixation (Liithers & Hanert, 1989) and carboxysome-like structures containing ribulose bis-phosphate carboxylase found in T. ferrooxidans and other obligately autotrophic bacteria. Gallionella ferruginea is not common in agricultural soil. It prefers a sessile life style in aqueous environments containing gradients of Fe and O 2 such as Fe seep areas, drainage ditches, water wells, and the like. It can be a major cause of well clogging and it usually is a component of Fe-depositing bacterial consortia. It also occurs in water distribution systems where low concentrations of O 2 mix with CO2- and Fe2+-rich waters encourages their chemolithoautotrophic growth (Hanert, 1981a; Ghiorse, 1986; Ghiorse & Ehrlich, 1992). It is important to note that other classical iron bacteria such as Leptothrix ochracea (Mulder & Deinema, 1981) also may be capable of chemolithoautotrophic growth; however, these bacteria have so far resisted cultivation (Ghiorse & Ehrlich, 1992). Reliable enrichment, isolation, and enumeration procedures have been published for G. ferruginea (Hanert, 1981a). 46-2.2.2 Materials 1. 2. 3. 4. 5. 6.

FeS suspension. Modified Kucera-Wolfe medium. Carbon dioxide. Screw capped test tubes (16 x 150 mm). Low temperature incubator (15-20 0c). Phase contrast microscope.

46-2.3.3 Procedures Detailed instructions on the procedures for enrichment and isolation of G. ferruginea are given by Hanert (1981a). FeS suspension is prepared by mixing 78 g of (NH4h Fe(S04h·6H20 with 44 g of Na2S·9H20 or 140 g of Fe(S04)z-7H20 with 120 g of Na2S·9H20 in boiled deionized water cooled to 50°C with constant stirring in a nearly full glass-stoppered 1-L

1090

GHIORSE

bottle. The bottle is filled with boiled deionized water and stoppered while the FeS precipitate settles. The FeS precipitate is washed by decanting and replacing the supernatant with boiled deionized water at 50°C several (5-10) times at minimum of 4-h intervals until the pH in the FeS precipitate remains unchanged. Successive stable pH readings in the neutral range indicates that S2-ions are no longer being released from the FeS precipitate. Precipitation of FeS may be accelerated by adding a few drops of a saturated FeCl3 solution. The FeS suspension is stored in a completely filled glass-stoppered bottles to prevent oxidation. Smaller (100 mL) quantities are removed for autoclaving in 100-mL milk dilution bottles. Modified Kucera-Wolfe mineral medium is prepared following the procedures of Kucera and Wolfe (1957) as modified by Hanert (1981a). The medium contains per liter of distilled water: 1.0 g of NH4 Cl, 0.2 g of MgS0 4 ·H20, 0.1 g of CaCI2·2H20, and 0.05 g of K2 HP0 4 ·3H20. A phosphate concentration of 1I1Oth the original Kucera-Wolfe concentration is used to prevent precipitation of Ca, Mg, and Fe phosphates during autoclaving (Hanert, 1981a). Enrichment cultures are set up by adding 9 mL of cold (5-10 0c) mineral medium to 16 x 125 mm sterile screw-capped tubes. The medium is bubbled with cotton-filtered CO 2 for 5 s through a sterile Pasteur pipette inserted to the bottom of the tube. One milliliter of sterile FeS suspension is carefully added to the bottom of each tube. Enrichment cultures should be inoculated immediately, with material known from microscopic examination to contain stalks of Gallionella (see Hanert, 1981a). Incubate at 15 to 20°C and look for growth on the sides of the tubes in 3 to 5 d or longer. Fluffy yellow-white to orange colonies will grow in a discrete zone where the proper O 2 and Fe2+ concentrations exist in the gradient established in the tube. The presence of G. ferruginea in colonies is confirmed in a phase contrast microscope at 400 to 1000 x. Actively growing cultures of G. ferruginea are identified by the occurrence of characteristic bean-shaped cells at the end of twisted or convoluted stalks (Fig. 46-1). Isolation may be achieved by the end-point dilution method and enumeration by the dilution MPN method using variations of the same procedure. A series of tubes is prepared to the FeS addition stage and held at low temperature. This helps to keep more CO 2 in solution until the tubes are inoculated. Care must be taken that the inocula are well dispersed so that individual cells can be diluted away from colonies or clumps of flocculent material.

46-3 IRON- AND MANGANESE-REDUCING HETEROTROPHS 46-3.1 Enumeration of Non-Enzymatic Iron- and Manganese-Reducers 46-3.1.1 Principle Simple enumeration procedures for Fe- and Mn-reducing heterotrophic microorganisms in soil and sediment have been developed using a

OXIDATION AND REDUCTION OF FE AND MN

1091

Fig. 46-1. Phase contrast photomicrograph showing four bean-shaped Gallionella ferruginea cells at the ends of Fe (III) oxide impregnated stalks (center). The cells are best observed at the edges of a micro-colony such as this one growing in modified Kucera-Wolfe medium. Magnification 2000 x .

differential plate count procedure with agar media contammg hydrous FellI or Mn(III, IV) oxides (Ehrlich, 1990; Lovley, 1991). Non-enzymatic Mn reducers may be detected in this way under aerobic or anaerobic conditions. In principle, non-enzymatic Fe reducers could be detected in this way only under anaerobic conditions; however, because of the rapid reoxidation of Fe z + at circumneutral pH in the presence of Oz. Plate counts depend on the production of clear zones of Fe- or Mn-oxide reduction that form around colonies growing on organic substrates in the medium. The zones of reduction are thought to result from reactions of extracellular excretion products of metabolism (usually acidic products of sugar metabolism) with the oxides to reduce them non-enzymatically in the vicinity of the colony . This has been termed indirect reduction (Ehrlich, 1990) . Although Fe- and Mn-respiring bacteria have been detected in this way (Myers & Nealson, 1988; Ehrlich, 1990), as a rule neither direct enzymatic reduction for Fe or Mn respiration can be inferred from the formation of a clear zone around a colony (Ghiorse, 1988a; Lovley, 1991). Nevertheless, the Fe- and Mn-reducing bacteria and fungi detected in this way may be important members of the total Fe- and Mn-reducing community. The accumulation of metabolic products and acidic conditions, especially in microenvironments of soil and sediments, may encourage indirect reduction mechanisms.

1092

GIllORSE

46-3.1.2 Materials 1. Nutrient agar. 2. 20 gIL of KMn04 solution or sonicated ()Mn02 powder (Myers & Nealson, 1988). 3. Fe (III) oxide (46-3.2.3). 4. Spot test reagents for Fe an Mn (46-1.1.2).

46-3.1.3 Procedure Prepare and sterilize nutrient agar medium according to manufacturers instructions and dispense molten agar into petri dishes (20 mL). Prepare and sterilize overlay agar tubes containing 5 mL of the same medium. Cool plates to solidify agar. Keep overlay agar molten at 50 °C. Add 0.05 mL of 20 glKg of K2Mn04 each of the molten overlay agar tubes and mix to form an even suspension of brown Mn oxide in the agar. Alternatively, 0.5 mL of a sterile concentrated suspension of ()Mn02 (Myers & Nealson, 1988) or hydrous Fe(III) oxide (Lovley, 1991) can also be mixed into the overlay agar. Pour the molten suspension over the solidified agar in petri dishes. Rotate plates to disperse the molten overlay suspension evenly. Allow top agar to solidify and dry. Plates are inoculated and spread as described above (46-1.2) with 0.1 or 1 mL samples from standard dilution series. Incubate at 20 to 25 °C noting growth and clear zones around colonies. The clear zones may be tested for the absence of Fe(III) and Mn(III, IV) oxide with leuco crystal violet or Prussian blue spot tests (46-1.1.2).

46-3.2 Enrichment and MPN Enumeration of Iron- and Manganese-Respiring Bacteria 46-3.2.1 Principles Recently Lovley and Phillips (1988) and Myers and Nealson (1988) showed that Fe- and Mn-respiring bacteria can be obtained from sediments by anaerobic enrichment with a nonfermentable electron donor (e.g., acetate and lactate) and hydrous Fe or Mn oxide as the sole electron acceptor. Enrichment methods have been developed for anoxic lake and river sediments, but these are applicable to soil and subsurface sediments (Lovley et aI., 1990; D.R. Lovley, 1991, personal communication). Dissimilatory Fe and Mn respiration is now accepted as an important biological process in the oxidation of organic matter under anaerobic conditions; characterization of these dissimilatory Fe- and Mn-reducing bacteria is progressing rapidly (Ehrlich, 1990; Myers & Nealson, 1990; Lovley, 1991; Nealson & Myers, 1992). Enumeration of Fe- and Mn-respiring bacteria is best achieved using a standard MPN method and strictly anaerobic cultural procedures. Tubes are scored on the basis of a color change of the medium. Bacterial Fe reduction results in a color change from reddish brown to black as amor-

OXIDATION AND REDUCTION OF FE AND MN

1093

phous Fe(III) oxide is reduced to magnetite, the principal product of Fe reduction. Reduction of brown Mn(III, IV) oxide results is the formation of a whitish precipitate of rhodochrosite (MnC03). 46-3.2.2 Materials 1. 2. 3. 4. 5.

Basal salts medium. Fe (III) or Mn (IV) oxide slurry. Trace element solution. Vitamin solution. Belleo anaerobic pressure tubes with butyl rubber stoppers and aluminum crimp seals. 6. 02-free N 2/C0 2 (80:20) gas mixture. 7. Tubes and syringes for anaerobic dilution. 46-3.2.3 Procedure Prepare basal salts medium by the following constituents in g/900 mL of deionized wateriNaHC03, 2.5; CaCI2·2H20, 0.1; KCI, 0.1; NH4Cl, 1.5; NaH2P04·6H20, 0.6; a nonfermentable C-energy source, 3.0 (for discussion, see Myers & Nealson, 1990 and Lovley, 1991). Sodium salts of acetate, lactate, pyruvate, succinate or a mixture of these compounds are the best C-energy substrates for enrichment cultures. Add 10 mL each of trace element and vitamin solutions. Mix constituents and adjust pH to 6.5 to 7.0. The trace element solution contains in grams per liter: NTA, 1.5; MgS0 4, 0.3; MnS04, 0.5; NaCl, 1.0; FeS04, 0.1; CaCI2, 0.1; ZnCI, 0.13; CUS04, 0.01; AIK(SO)4h. 0.01; H3B02' 0.01; NaMo0 4, 0.025; NiCI2·6H20, 0.024; Na2W04, 0.025. The vitamin solution contains in milligram per liter: biotin, 2; folic acid, 2; pyridoxine HCI, 10; riboflavin, 5; thiamine, 5; nicotinic acid, 5; pantothenic acid, 5; vitamin B 12 , 0.1; p-aminobenzoic acid, 5; thioctic acid, 5. Add 1 or 2 mL of Fe (III) oxide slurry or 0.5 mL of Mn oxide slurry to the anaerobic pressure tubes containing 9 mL of the basal salts medium. Bubble the mixture with the 02-free N 2/C0 2 gas mixture for 10 min using a stainless steel canula to reach the bottom of each tube. Cap each anaerobically under a flow of 02-free N2 using thick butyl rubber stoppers and seal with an aluminum crimp. Sterilize by autoclaving. Fe (III) oxide slurry is prepared by dissolving 108 g of FeCl3 in 1 L of distilled water and slowly raising the pH to 7 with drops of 10 N of NaOH. Do not allow bulk pH to go above 7 at any time during the neutralization process. Wash the resultant floc by centrifugation to remove excess salt. Store concentrated suspension in distilled water at 4 dc. Manganese(IV) oxide slurry is prepared by dissolving 3.16 g of KMn04 and 3.2 g of NaOH in 1 L of distilled water, dissolve 5.94 g of MnCl2 in another liter of water. Add the MnC0 2 solution slowly to the basic KMn04 solution while mixing constantly with a magnetic stir bar. Wash the resultant floc to remove salts. Store the concentrated suspension in distilled water at 4 dc.

GmORSE

1094

For enrichment cultures and MPN enumeration, dilute samples anaerobically in an appropriate diluent (e.g., basal salts medium without C-energy source). Set up anaerobic MPN tubes following standard procedures. Use syringes and needles flushed with filter sterilized 02-free N2 or Ar gas to transfer dilution samples and inoculate tubes. Incubate at an environmentally relevant temperature or 20 to 25°C. Iron(III) reduction is indicated by conversion of the non-magnetic, orange-red Fe(III) oxide to a black magnetic precipitate (magnetite). Manganese(IV) reduction is indicated by conversion of dark brown Mn(IV) oxide to a whitish rhodochrosite (MnC03 ) precipitate. A wide variety of electron donors including aromatic compounds and H2 can yield successful enrichments (Lovley, 1991; Myers & Nealson, 1990); but it is not always certain whether the organisms metabolizing the added electron donors are the Fe(III) or Mn(IV)-reducers (Lovley, 1991). This determination requires isolation and study of the individual members of the Fe-Mn-reducing community. It is advisable to use only nonfermentable compounds such as those listed above for MPN determination in the enrichment cultures.

ACKNOWLEDGMENT Derek R. Lovley kindly provided guidance for section 46-3.2. Thanks are due to Patti Lisk for help in preparing the manuscript.

REFERENCES Alexander, M. 1977. Introduction to soil microbiology. 2nd ed. John Wiley and Sons, New York. Bromfield, S.M. 1978. The oxidation of manganous ions under acid conditions by an acidophilous actinomycete from acid soil. Aust. J. Soil. Res. 16:91-100. Bromfield, S.M., and V.B.D. Skerman. 1950. Biological oxidation of manganese in soils. Soil Sci. 69:337-348. Chapnick, S.D., W.S. Moore, and K.H. Nealson. 1982. Microbially mediated manganese oxidation in a freshwater lake. Limnol. Oceanogr. 27:1004-1014. Cullimore, D.R., and A.E. McCann. 1977. The identification, cultivation, and control of iron bacteria in ground water. p. 219-261. In F.A. Skinner and J.M. Shewan (ed.) Aquatic microbiology. Academic Press, London. Eagiesham, B.S., R.E. Garen, and W.C. Ghiorse. 1986. Ultrastructure of Gallionella ferruginea. p. 100. In Abstract, Annu. Meet. ASM, J-11. Am. Soc. for Microbiol., Washington, DC. Ehrlich, H.L. 1987. Manganese oxide reduction as a form of anaerobic respiration. Geomicrobiol. J. 5:423-431. Ehrlich, H.L. 1990. Geomicrobiology. 2nd ed. Marcel Dekker, New York. Ehrlich, H.L., and C.L. Brierley. 1990. Microbial mineral recovery. McGraw-Hili Publ. Co., New York. Ehrlich, H.L., W.J. Ingiedew, and J.C. Salerno. 1991. Iron and man~anese-oxidizing bacteria. p. 147-170. In J.M. Shively and L.L. Barton (ed.) Variation in autotrophic life. Academic Press, London. Emerson, D., R. Garen, and W.C. Ghiorse. 1989. Formation of Metallogenium-like structures by a manganese-oxidizing fungus. Arch. Microbiol. 151:223-231.

OXIDATION AND REDUCTION OF FE AND MN

1095

Ghiorse, W.C. 1984. Biology of iron- and manganese-depositing bacteria. Ann. Rev. MicrobioI. 38:515-550. Ghiorse, W.C. 1986. Biology of Leptothrix, Gallionella, and Crenothrix relationship to plugging. p. 97-108. In R. Cullimore (ed.) Proceedings of international symposium on biofouled aquifers: Prevention and restoration. Am. Water Resourc. Assoc., Bethesda, MD. Ghiorse, W.C. 1988a. Microbial reduction of manganese and iron. p. 305-331. In A.J.B. Zehnder (ed.) Biology of anaerobic microorganisms. John Wiley and Sons, New York. Ghiorse, W.e. 1988b. The biology of manganese transforming microorganisms in soil. p. 75-85. In R.D. Graham et al. (ed.) Manganese in soils and plants. Kluver, Dordrecht, Netherlands. Ghiorse, W.C. 1989. Manganese and iron as physiological electron donors and acceptors in aerobic-anaerobic transition zones. p. 163-169. In Y. Cohen and B. Rosenberg (ed.) Microbial maps: Physiological ecology of benthic microbial communities. Am. Soc. for Microbiol., Washington, DC. Ghiorse, W.C., and H.L. Ehrlich. 1994. Microbial biomineralization of iron and manganese. p. 75-99. In R.W. Fitzpatrick and H.C.W. Skinner (ed.) Iron and manganese biomineralization processes in modern and ancient environments. CATENA, West Germany. Ghiorse, W.C., and P. Hirsch. 1978. Iron and manganese deposition by budding bacteria. p. 897-909. In W.E. Krumbein (ed.) Environmental biogeochemistry and geomicrobiology. Ann Arbor Science, Ann Arbor, MI. Hackett, G., and J.H. Lehr. 1986. Iron bacteria occurrence, problems and control methods in water wells. Natl. Water Well Assoc., Worthington, OH. Hanert, H.H. 1981a. The genus Gallionella. p. 509-516. In M.P. Starr et al. (ed.) The prokaryotes. Vol. I. Springer-Verlag, Berlin. Hanert, H.H. 1981b. The genus Siderocapsa (and other iron- and manganese-oxidizing eubacteria). p. 1049-1059. In M.P. Starr et al. (ed.) The prokaryotes. Vol. I. SpringerVerlag, Berlin. Holmes, D.S., and J.R. Yates. 1990. Basic principles of genetic manipulation of Thiobacillus ferrooxidans by biohydrometallurgical applications. p. 29-54. In H.L. Ehrlich and C.L. Brierley (ed.) Microbial mineral recovery. McGraw-Hill Publ. Co., New York. Jones, J.G. 1986. Iron transformations by freshwater bacteria. Adv. Microb. Ecol. 9: 149-185. Kessick, M.A., J. Vuceta, and J.J. Morgan. 1972. Spectrophotometric determination of oxidized manganese with leuco crystal violet. Environ. Sci. Technol. 6:642-644. Kucera, S., and R.S. Wolfe. 1957. A selective enrichment method for Gallionellaferruginea. J. Bacteriol. 74:344-349. Kuenen, J.G., and O.H. Tuovinen. 1981. The genus Thiobacillus and Thiomicrospira. p. 1023-1036. In M.P. Starr et al. (ed.) The prokaryotes. Vol. I. Springer-Verlag, Berlin. Lovley, D. R. 1987. Organic matter mineralization with the reduction of ferric iron: A review. Geomicrobiol. J. 5:375-399. Lovley, D.R. 1991. Dissimilatory Fe(III) and Mn(IV) reduction. Microbiol. Rev. 55:259287. Lovley, D.R., P.H. Chapelle, and E.J.P. Phillips. 1990. Fe(III)-reducing bacteria in deep buried sediments of the Atlantic Coastal Plain. Geology 18:954-957. Lovley, D.R., S.J. Giovannoni, D.e. White, J.E. Champine, E.J.P. Phillips, Y.A. Gorby, and S. Goodwin. 1993. Geobacter metallireducens gen. nov. sp. nov., a microorganism capable of coupling the complete oxidation of organic compounds to the reduction of iron and other metals. Arch. Microbiol. 159:336-344. Lovley, D.R., and E.J.P. Phillips. 1988. Novel mode of microbial energy metabolism: Organic carbon oxidation coupled to dissimilatory reduction of iron and manganese. Appl. Environ. Microbiol. 54:1472-1480. Liithers, S., and H.H. Hanert. 1989. The ultrastructure of chemolithoautotrol?hic Gallionella ferruginea and Thiobacillus ferrooxidans as revealed by chemical fixatIon and freezeetching. Arch. Microbiol. 151:245-251. Mishra, A.K., P. Roy, and S.S.R. Mahapatra. 1983. Isolation of Thiobacillus ferrooxidans from various habitats and their growth pattern on solid medium. Curro Microbiol. 8:147-152. Mulder, E.G., and M.H. Deinema. 1981. The sheathed bacteria. p. 425-440. In M.P. Starr et al. (ed.) The prokaryotes. Vol. I. Springer-Verlag, Berlin. Myers, C.R., and K.H. Nelson. 1988. Bacterial manganese reduction and growth with manganese oxide as the sole electron acceptor. Science 240:1319-1321.

1096

GmORSE

Myers, C.R., and K.H. Nealson. 1990. Iron mineralization by bacteria: Metabolic coupling of iron reduction to cell metabolism in Alteromonas putrefaciens strain MR-1. p. 131-149. In R.B. Frankel and R.P. Blakemore (ed.) Iron biominerals. Plenum Press, New York. Nealson, K.H., and C.R. Myers. 1992. Microbial reduction of manganese and iron: new approaches to carbon cycling. Appi. Environ. Microbioi. 58:439-443. Nealson, K.H., R.A. Rosson, and C.R. Myers. 1989. Mechanisms of oxidation and reduction of manganese. p. 383-411. In R.J. Beveridge and R.J. Doyle (ed.) Metal ions and bacteria. John Wiley and Sons, New York. Nealson, K.H., B.M. Tebo, and R.A. Rosson. 1988. Occurrence and mechanism of microbial oxidation of manganese. Adv. Appi. Microbioi. 33:279-318. Norris, P.R. 1990. Acidophilic bacteria and their activity in mineral sulfide oxidation. p. 3-27. In H.L. Ehrlich and C.L. Brierley (ed.) Microbial mineral recovery. McGraw-Hill Pubi. Co., New York. Paul, E.A., and F.E. Clark. 1989. Soil microbiology and biochemistry. Academic Press, New York. Pringsheim, E.G. 1949. Iron bacteria. BioI. Rev. 24:200-245. Seeley, H.W., Jr., and P.J. VanDemark. 1981. Microbes in action. W.H. Freeman and Co., San Francisco. Silverman, M.P., and D.G. Lundgren. 1959. Studies on the chemoautotrophic iron bacterium Ferrobacillus ferrooxidans. I. An improved medium and a harvesting procedure for securing high cell yields. J. Bacterioi. 77:642-647. Staley, J.T. 1968. Prosthecomicrobium and Ancalomicrobium: New prosthecate freshwater bacteria. J. Bacterioi. 95:1921-1942. Tuovinen, O.H., and D.P. Kelly. 1973. Studies on the growth of Thiobacillus ferrooxidans. I. Use of membrane filters and ferrous iron agar to determine viable numbers and comparison with 14C02 fixation and iron oxidation as measures of growth. Arch. Mikrobioi. 88:285-298. Uren, N.C., and G.W. Leeper. 1978. Microbial oxidation of divalent manganese. Soil BioI. Biochem. 10:85-87. Winogradsky, S. 1888. Ueber Eisenbacterien. Bot. Ztg. 46:262-270. Wolfe, R.S. 1958. Cultivation, morphology, and classification of the iron bacteria. J. Am. Water Works Assoc. 50:1241-1249. Wolfe, R.S. 1964. Iron and manganese bacteria. p. 82-97. In H. Heukelebian and N.C. Dondero (ed.) Principles and applications in aquatic microbiology: Proceedings of the Rudolfs Research Conference. John Wiley and Sons, New York.

Published 1994

SUBJECT INDEX Acanthamoeba, 494 Acetobacter, 179 Acetylene side effects, 181 sources, 1050-1051 Acetylene inhibition method, denitrification, 1048 Acetylene reduction test, N2 fixation, 216, 1019-1025 calibration, 1042-1043 free-living bacteria, 180-181 grass systems, 1023-1025 nodulated root system, 1020-1023 Achromadora, 486 Achromobacter, 247 Acid hydrolysis, soils, 896-897 Acid methyl-green stain, 502 Acid phosphatase, 777, 802, 807-809 Acid-insoluble nitrogen, 899 Acidophiles actinomycetes, 281-282 Fe-oxidizing, 1086-1089 Aconitase, 568 Acridine orange, 84-85 Acrylamide gel, 624-627 Actinomadura, 269, 271, 277, 285 isolation, 278 Actinomyces, 269, 271, 282 catalase-negative, 282 Actinomycetes, 269-287 cell wall analysis, 284-286 culture medium, 276-277 detection rRNA probes, 273 in situ hybridization, 273-274 differentiating from bacteria, 132 diversity, 270-272 enrichment, 270-279 enumeration, 270-279 extraction from soil, 274 fixation, 274 Frankia, 291-322 identification, 284 isolation, 270-279 acidophiles and alkalophiles, 281-282 antibiotic-producing strains, 283-284 autotrophs, 280 chemical disruption of soil, 275-276 physical disruption of soil, 275 physiological groups, 279-284 strains with biodegradative activity, 282-283 thermophiles, 280-281 light microscopy, 273-274, 285-287 maintenance of isolates, 286

microaerophiles, 282 morphology, 285-287 reduction of background microorganisms heat treatment, 277 selective inhibitors, 277-278 scanning electron microscopy, 272-273 thermophilic, 270 Actinoplanes, 271, 285 baiting techniques, 278 Actinorhizal plants attributes, 293-294 Frankia symbiosis, 291-322 geographic distribution, 294 nutrient solutions for growing seedlings, 312 propagation, 311 Adenylate kinase, 568 Adhesive, 214 Aeration, C mineralization experiments, 839 Aerobic incubation method inorganic N production, 960, 962-967 N transformations, 988, 1006-1009 Aeroponic system, VAM fungi, 369-371 Agar, 119, 139 Agar film method, mycelial length, 96-97 Agar well method, antibiotic-producing microorganisms, 391 Agar-diffusion method, intrinsic antibiotic resistance strains, 583-584 Agar-dilution method, intrinsic antibiotic resistance strains, 584-585 Agarose gel, 650, 655-656 Aggregation, soil, hyphal development in VAM fungi, 357-358 Agrobacterium, 202, 264, 638 Agromyces, 269,271,285 Alcaligenes, 179, 247, 261-262, 264, 411, 643-644 Alcohol dehydrogenase, 568 Algae, 427-455 collection of soil veneers, 432-433 colonial, 430 culture, 444-452 culture media, 448-452 endolithic, 439-440 enumeration cell counting, 433-434 chlorophyll autofluorescence, 434-435 chlorophyll extraction and quantitation, 441-444 colony-forming units, 440 direct methods, 432-441 implanted slide method, 435 indirect methods, 440-444 most probable number method, 440-441

1097

1098

Algae (cont.) filamentous, 430 green, 429, 431 identification, 428-431 isolation and purification centrifugation method, 446-447 repeated washings, 445-446 streak plate method, 446 zoospore method, 447 light microscopy, 431, 433-434 photosynthesis estimation, 453-454 siphonous, 430 storage and preservation, 452-453 yellow-green, 429, 431 Alkaline lysis miniprep, 642-643 Alkaline phosphatase, 612, 614, 777, 802, 807-809 Alkaline phosphatase-linked immunosorbent assay, 611-613 Alkalophiles, actinomycetes, 281-282 Amidase, 790-801 Amidohydrolase, 790-801 Amino acid-nitrogen, 899 Amino sugar-nitrogen, 900 (Ammonia + amino sugar)-nitrogen, 899 Ammonia-nitrogen, 899, 939 Ammonia-oxidizing nitrifying bacteria, 159176 Ammonium, production during waterlogged incubation, 960-962 Ammonium method, inorganic N automated test, 975-977 manual test, 977-979 Ammonium persulfate, 624-627 Amoeba, 492, 499, 510 Ampicillin resistance, 578 AmpliTaq polymerase, 708 Ampullariella, 271 Amycolata, 271,281, 285 Amycolatopsis, 271,285 Anabaena, 189-190,429-430, 454 Anacystis, 430 Anaerobe culture system, 233-235 Anaerobic bacteria, 121, 179,223-241 aerotolerant anaerobes, 225 anaerobic incubations of soil, 240-241 culture conventional anaerobic methods, 233 redox indicators, 231-232 removal of O 2 , 226-229 strict anaerobic method, 233-235 culture media, 229-231, 236-238, 240 diversity, 224-226 enumeration, 235-240 CO2 reducers, 239-240 clostridia, 237-238 heterotrophs, 236-237 sulfate reducers, 238-239 facultative anaerobes, 223

SUBJECT INDEX

obligate anaerobes, 223-224 strict anaerobes, 225 Anaerobic conditions, 182-183,223 Anaerobic jar, 233 Analog enrichment, 408 Andersen sampler, 280-281 Aniline blue, phenolic, 97-98 Animal wastes, lsN-labeling, 917-918 Antagonist. See Antibiotic-producing microorganisms Antibiotic classes, 578 Antibiotic resistance markers, 575-589, See also Intrinsic antibiotic resistance strains; Spontaneous antibiotic resistance strains R plasmids, 637 Antibiotic-producing microorganisms actinomycetes, 283-284 culture media, 382-384 detection agar well method, 390 biomass method, 392-393 cross streak method, 388-389 culture filtrate methods, 389-393 dual culture methods, 385-389 fungal disk methods, 387-388 microbial lawn methods, 385-387 paper disk method, 390 radial growth method, 391-392 isolation, 379-403 assay standardization, 381-382 method selection, 380-381 sampling strategies, 380 methods for selected compounds, 396-402 preparation of inocula bacterial, 384-385 fungal,385 screening, 393-396 multiple agar layer method, 395-396 single agar layer method, 394-395 Antibodies, 592-616 against microorganisms, 597-600 conjugation, 592-616 with fluorescein isothiocyanate, 604-606 cross-reactive, adsorption from polyclonal antiserum, 601-602 monoclonal, 596-597, 599-600 polyclonal, 596-597, 599 procedures for utilization, 601 production, 599-600 purification from antiserum, 602-604 specificity, 599 storage, 606 structure, 594-595 Antigens bacterial, 598 fungal,598 internal, 598 microbes as, 594

SUBJECT INDEX Antigens (cont.) somatic, 598 viral, 598 API system, 549 Apparent lag, 416 Aquaspirillum, 179,261-262 Araeolaimida, 485-486 Arbuscules, 353-354 Arginine glycerol salts agar, 276 Arthrobacter, 271 selective medium, 137-138 Arthropods archiving, 533-535 assigning morphospecies to functional ecological groupings, 533-534 transforming census data, 534-535 evaluation in field destructive sampling of soil, 518-519 pitfall trapping, 519-521 extraction based on cuticle, 528 centrifugation method, 528 elutriation, 527-528 physical methods, 527-528 through behavior modification, 522-527 flotation in salt solution, 527 functions in soil, 517-518 identification, 531-532 preservation, 532-533 processing extracted biota sample, 528-531 rearing, 535-537 sampling soil cores, 521-527 distribution in cores by microscopic methods, 521-522 passive extraction, 521 statistical methods to analyze diversity, 538-539 taxonomy, 531-532 Arylsulfatase, 777, 814-818 L-Asparaginase, 790-801 Assimilatory nitrate reduction, 245-246 Asymmetry, statistics as indicators, 17 Atom percent nitrogen-15, 888 Atomic mass, 1030 ATP analysis, biomass determination, 753, 767-769 Aureobacterium, 271 Autoclave, 42-43 Autoradiography, detection of nucleic acid probes, 684-685 Autotrophs actinomycetes, 280 nitrifying bacteria, 159-176 Availability ratio, N, 891-892 A-value method, 1031 Azide sterilization, 48-49 Azolla, 454, 1040-1042 Azomonas, 179, 188 detection, 184-185

1099 Azorhizobium, 199-218 Azospirillum, 179-195,247,261-262,264 Azotobacter, 179-195 detection, 184-185 Azotobacter medium, 184 Azotobacteraceae, 179-195 N2 fixation, 183-186

Bacillariophyceae, 431 Bacillus, 179-195,247-248,264 Bacitracin resistance, 578 Bacteria. See also specific groups and genera antibiotic resistance markers, 575-589 biomass, 101 C source utilization, 543-551 DNA. See also Bacterial community DNA fingerprinting, 647-662 isolation and purification from soil, 727750 efficiency of recovery from soil, 91-92 extraction from soil blending and differential centrifugation, 722-723 calcium chloride and sucrose density centrifugation, 721-722 fatty acid analysis, 551-553 Fe oxidation and reduction, 1079-1094 fluorescently labeled oligonucleotides directed at 16S rRNA, 101-103 identification, 644 Mn oxidation and reduction, 1079-1094 monitoring populations in environment, 727-729 plasmid profiles, 635-644 population genetics, 557-572 RFLP analysis, 647-662 S oxidation and reduction, 1067-1076 separation from soil, 82-87 taxonomy, 543-553 total, enumeration in soil, 82-92 consecutive extraction, 92 density gradient centrifugation, 90-91 dispersion method, 83 extraction and flocculation method, 89-90 filtration and staining methods, 83-87 immunofluorescence microscopy, 87-89 whole-cell protein profiles, 619-633 Bacteria, viable cell elongation assay, 92-94 culture media, 134-139 enumeration, 119-142 plate count, 119-121 data handling, 139-141 diluents, 125-126 dilution plate method, 121 drop plate method, 133 limitations, 121-122 materials and equipment, 122-123

SUBJECT INDEX

1100 Bacteria (cont.) pour-plate method, 121, 130-131 samples, 123-124 serial dilutions, 127-129 spread-plate method, 121, 131-133 recovery, 119-142 reduction of tetrazolium dyes, 94-95 release from soils, 124-125 Bacterial community DNA desalting and concentration, 745-746 fractionation of DNA gradients, 743-746 isolation and purification bacterial biomass and, 729-730 bacterial fractionation approach, 731-733,736-741 direct lysis approach, 733-736, 741-743 soil clay content and, 730-731 soil organic matter and, 730 quantitation spectrophotometry, 746-747 UV fluorescence in presence of ethidium bromide, 747-750 Bacterial fractionation approach, bacterial community DNA, 731-733, 738-741 Bacteriocin-producing microorganisms, 397399 Bacteriophage. See Phage Baermann funnel arthropod extraction, 526 nematode extraction from plant material, 478 from soil, 470-473 Baiting techniques Actinopianes, 278

arthropod, 519-520 BAP medium, 301 Basidiomycetes, 332 Becquerel, 874 Beggiatoa, 179 Beijerinckia, 179, 188

detection, 185 Beijerinckia medium, 185 Bennett's agar, 300 Benzene, 411, 414 Berlese funnel arthropod extractor, 523524, 526 Berthelot reaction, 975 BG-11 medium, 449 Bioassay, Frankia, 316-319 Biodegradation research, 407-424 actinomycetes, 282-283 Cl determination, 423-424 enrichment culture, 408-412 growth in liquid cultures, 415-417 isolation of pure cultures, 412-413 maintenance of cultures, 413-415 mineralization, 835 O 2 consumption studies, 421-423 using cell-free extracts, 419-421 using washed cell suspensions, 418-419

Biodiversity studies, PCR in, 724 Biological tracer, antibiotic resistance markers, 575-576 Biomass, 753-770 active, Frankia, 305 ATP analysis, 753, 767-769 bacterial, 101 chloroform fumigation extraction method, 753,763-767 chloroform fumigation incubation method, 753-760 determining weight, 100-101 fungal,101 labeling with 14C, 869-871 with 15N, 889-890 light microscopy to determine, 770 soil sampling, preparation, and storage, 754 substrate-induced respiration method, 753,760-763 total, Frankia, 305 Biomass carbon calculation, 757-759, 762-763, 765-766 determination, 764-765 Biomass nitrogen calculation, 759-760, 766-767 determination, 765 Biomass method, antibiotic-producing microorganisms, 392-393 Biotin-labeled DNA, 673 Biotinylated antibody, 613 Biovolume, microscopic estimates, 100-101 Biovolume conversion factor, 100 Biphenyl, 411, 414-415 Bisacrylamide, 624 Blending, release of bacteria from soils, 124 Biepharisma, 495

Bold's basal medium, 449-450 Botrydium, 430

Bouin's fixative, 532 Bouin-Hollande fixative, 502 Bradyrhizobium, 199-218, 264, 565, 575, 580, 593, 599, 621, 630 Bromothymol blue, 545 Broth dilution method, intrinsic antibiotic resistance strains, 585 BTEX utilization, 414 Buffers gel electrophoresis, 652-653 MLEE analysis, 560-561, 565 Bulking, sample, 26-27, 32, 148 Bumilleria, 431

Bunsen absorption coefficients, 254, 1053 Buried-bag method, field estimation of N transformations, 988, 1004-1006 CAHs. See Chlorinated aromatic hydrocarbons Calcofluor M2R, 97-98

SUBJECT INDEX Campylobaeter, 179 Candle jar, 229 Capillary pipet, preparing fine pipets, 444445 Carbon, biomass calculation, 757-759, 762-763, 765-766 determination, 764-765 Carbon dioxide alkali trapping and detection, 836-837 14C-labeled, collection and counting, 871875 detection by soda lime absorption, 841-844 detection in gaseous samples, 837-838 determination by gas chromatography, 844-849 dynamic laboratory methods, 850-855 release from metabolizing organisms, 835836 scrubbing system, 851-852 static laboratory methods, 855-859 static incubation-gas chromatographic analysis, 855-858 static incubation-titrimetric determination, 858-859 trapping system, 851-852 Carbon dioxide-reducing bacteria, 225 anaerobic, enumeration, 239-240 Carbon-14 labeled organic matter commercial sources of labeled materials, 868 decomposition, 866-875 extraction from soil, 894-896 incubation of labeled materials, 871-875 labeling biomass and microbial products, 869-871 labeling plant material, 867-869 Carbon mineralization, 835-859 determination, 756-757 experimental principles, 836-841 aeration, 839 alkali trapping and detection of CO 2 , 836-837 chambers, 839 CO 2 detection in gaseous samples, 837838 O 2 detection, 838-839 field methods, 836, 841-850 CO 2 detection by soda lime absorption, 841-844 soil CO 2 and O 2 by gas chromatography, 844-849 laboratory methods, 836, 850-859 dynamic method for CO 2 , 850-855 static method for CO 2 and O2 , 855-859 Carbon-13 natural abundance technique, 875-876 apparatus and reagents, 877-888 calculating sources of soil organic matter, 884-885 conversion of organic C to CO 2 for mass

1101 spectrometry, 880-883 field sampling, 878-879 kinetics of soil organic matter turnover, 885-886 mass spectrometric analysis and isotopic indices, 883-884 natural BC and measurement of organic matter dynamics, 877 preparation of soil samples, 879-880 stable C isotope ratio of plants and soils, 876 stable isotope terminology, 875-876 Carbon source utilization bacteria, 543-551 chemical indicators of growth, 545 compounds in root exudates, 547 microtiter plate method, 546-552 possible C and energy sources, 547 substrate selection, 545 Carnoy's fixative, 502, 532 Catalase, 777 Catechol dioxygenase, 419-421 Catellatospora, 271 Cell counting, algae and cyanobacteria, 433434 Cell elongation assay, viable bacteria, 92-94 Cell smear, preparation, 102-103 Cell suspension, biodegradation research, 418-419 Cell wall analysis actinomycetes, 284-286 Frankia, 306 Cell-free extract, biodegradation research, 419-421 Cellulomonas, 271 Cellulose degradation, 270 Centrifugation method arthropod extraction, 528 isolation of algae and cyanobacteria, 446447

Cephalothin resistance, 578 Cerenkov counting, 685-686 Cesium chloride balance solution, 739, 742 Cesium chloride gradient, fractionation of DNA bands, 744 Cesium-137 irradiation, 44-45 Characterization medium, 546 Chasmolithic community, 428 Checkerboard titration, 613 Chemodenitrification, 250 Chitin determination filamentous fungi, 345 VAM fungi, 356, 358 Chlamydomonas, 430 Chloramphenicol resistance, 578 Chloride determination, 423-424 Chlorinated aromatic hydrocarbons (CAHs), 407-424 Chlorobenzene, 412 3-Chlorobenzoate, 411

1102

4-Chlorobiphenyl,414 Chloroform fumigation extraction method, biomass determination, 753, 763-767 Chloroform fumigation incubation method, biomass determination, 753-760 Chlorophyceae, 429, 431 Chlorophyll autofluorescence, enumeration of algae and cyanobacteria, 434-435 cell density/pigment correlation in soil, 441-442 extraction DMSO method, 443-444 ethanol method, 442-443 quantitation for enumeration of algae and cyanobacteria, 441-444 Chloroplast, 429 Chlorosarcinopsis, 430 Chromadorida, 485-486 Chromobacterium, 264 Ciliates, 492, 499, 510-511 Ciliophora, 492 Citrobacter, 248 Clostridium, 179-195,224,248 enumeration, 237-238 N2 fixation, 192-194 Cluster analysis, 544-545, 562 Cobalt-60 irradiation, 44-45 Cobb sieving, nematode extraction from soil, 469-471 Coefficients of similarity, 546 Coenobium, 430 Coenocytic organism, 430 Cofactor, enzyme, 784-785 Coliform bacteria, 145-156 culture medium, 151 Escherichia coli direct methods for detection, 156 rapid test, 154-155 fecal, 146 confirmation, 152-153 diluents, 150 multiple-tube fermentation method, 149-152 soil sampling, 147-149 temporal variations, 147 total, 146 Collection of soil, 148-149 Colloidal chitin agar, 276 Colonial algae, 430 Colony hybridization, 666, 680-682 Colony-forming units, algae and cyanobacteria, 440 Combined C medium, 188 Cometabolism, 408 Commensal, 408 Competitive inhibition, 786-787 Composite samples, 6-7 Computer programs statistical tests, 37-38

SUBJECT INDEX USDA Technology Transfer documents, 38 COMTESA, 534 Conductimetric analysis, 836 Confidence factor, 61-62 Confidence limits, 23-25 factors for calculating, 37 mean, 24 median, 24-25 most probable number methods, 61-62, 75 Conjugative plasmid, 637 Consortium, 412 Coomassie brilliant blue R, 628-629, 632 Copper-cadmium reduction method, inorganic N, 968-974 Coralloid root, 455 Core, soil, 149, 754, 929 arthropods, 521-527 collection, 1051 denitrification studies, 1054 double-cylinder hammer-driven core sampier, 521-522 Corn meal agar, 383 Corynebacterium, 179,247,271 Coulter counting, protozoa, 509 Counting procedure, light microscopy, 85-86 Coverslip well technique, filamentous fungi, 342-344 Cross streak technique, antibiotic-producing microorganisms, 388-389 Cross-feeding, 412 Cross-reactive antibody, adsorption from polyclonal antiserum, 601-602 Cryoprotectant, 205 Cryptic plasmid, 637 Cryptoendolithic community, 428 Cryptogamic crust, 428 Crytolophosis, 494 Culture filtrate method antibiotic-producing microorganisms, 389-393 filtrate preparation, 389-390 Culture media actinomycetes, 276-277 algae and cyanobacteria, 448-452 anaerobic bacteria, 229-231, 236-238, 240 antibiotic-producing microorganisms, 382-384 bacteria, 134-139 coliforms, 151 filamentous fungi, 330-333 Frankia, 299-302 inhibitors added to, 136 reduction, 229-231 rhizobia, 203-204, 206 selective, 136-138, 206 specific culture media Azotobacter, 184 BAP,301

SUBJECT INDEX Culture media (cont.) Beijerinckia, 185 Bennett's, 300 BG-11,449 Bold's basal, 449-450 colloidal chitin, 276 Czapek-Dox, 330, 383 dextrose-peptone, 331 EC,151 Frankia, 300 glucose nutrient, 383 9K,1087 Keyser-Munns, 204 King's B, 383, 399, 695 KT,1087 Kucera-Wolfe mineral, 1090 U2,301 lauryl tryptose, 151, 154 Luria-Bertani,695 M3 ,276 M9 minimal, 135,696 malt extract, 331, 383 Melin-Norkrans, 333 mineral salts, 410 minimal lactose X-gal, 695 N-deficiency combined C, 182 N-deficient lactate, 194-195 N-deficient malate, 191 NPCC, 136-137 nutrient, 135 nutrient poor, 138-139 nutrient rich, 138-139 oatmeal, 276 peptone yeast extract, 135, 203-204 peptone-water, 126, 150 PTYG,135 Qmod,3oo S,3OO

Sl, 137 soil agar film, 112-113 soil extract, 134, 451 soil solution equivalent, 135-136 soybean-casein digest, 383 starch casein, 276 TK,1087 tryptic soy, 134-135 water, 139 Winogradsky's salt solution, 126, 736737 X-gal, 695 yeast extract -dextrose-carbonate, 383-384 yeast-extract mannitol, 203, 384 strength, 138-139 Cyanide production, 401-402 Cyanobacteria, 179,427-455 collection of soil veneers, 432-433 culture, 447-452 culture media, 448-452 endosymbiotic, cycad roots, 454-455

1103

enumeration cell counting, 433-434 chlorophyll autofluorescence, 434-435 chlorophyll extraction and quantitation, 441-444 colony-forming units, 440 direct methods, 432-441 implanted-slide method, 435 indirect methods, 440-444 most probable number method, 438, 440-441 filamentous, 430 identification, 428-431 isolation and purification centrifugation method, 446-447 repeated washings, 445-446 streak plate, 446 light microscopy, 431,433-434 N- and organic C-deficient media, 190 N2 fixation, 189-190,428,454 photosynthesis estimation, 453-454 rice fields, 428, 437-439 sample collection, serial dilution, and plating, 438 visual estimation of coverage, 438-439 storage and preservation, 452-453 Cyanophyceae, 431 Cycad roots, endosymbiotic cyanobacteria, 454-455 Cyclohexane, 408 Cycloheximide, 136, 277 Cyst Heterodera, 475-477 protozoa, 498-499 Cysteine reducing agent, 229-231 Cysteine-sulfide reducing agent, 239 Cytophaga, 264 Czapek-Dox medium, 330, 383 D' Agostino test, 19 Dactyiosporangium, 270-271,277,285 DAPI. See Diphenylamidino indophenol DDT (dichlorodiphenyltrichloroethane), 408 DEA. See Denitrifier enzyme activity Decomposition, 835 isotopic methods for study of organic matter, 865-901 DECORANA, 538 Degradative plasmid, 637-638 Dehalogenation, 423-424 Dehydrogenase, 777, 820-823 013CPDB ' 875 Dendrogram, 538 Denitrification, 245, 756 field measurements, 1047-1062 acetylene inhibition method, 1048 closed chamber field gas flux method, 1054-1060 emission from 15N-labeled fertilizer, 1060-1061

1104 Denitrification (cont.) gas diffusion problems, 1062 gas sampling and storage problems, 1061 15N method, 1049 static core protocol, 1049-1054 nonrespiratory, 249-250 respiratory, 246-251, 259-265 Denitrifier(s), 245-265. See also Nitrate reduction confirmation of respiratory denitrification, 259-265 DNA probes, 263-265 enumeration, 251-255 DNRA strains, 255-256 most probable number method, 251-255 isolation, 257-258 respiratory, physiological and ecological features, 250-251 taxonomy, 264-265 Denitrifier enzyme activity (DEA), 256-257, 263-265 Density centrifugation extraction of bacteria from soil sample, 721-722 Frankia, 297 nematode extraction from soil, 470, 473474 protozoa, 506-507 total bacteria, 90-91 Derxia, 179-195 Desmodorida, 485-486 Destaining, 85 Desulfotomaculum, 179, 194-195 Desulfovibrio, 179, 194-195,248 Dextrose-peptone agar, 331 Diatom, 429, 436-437 2,4-Dichlorophenoxyacetic acid, 411 Differential centrifugation, extraction of bacteria from soil sample, 722-723 Digoxigenin-labeled nucleic acid probe, 674 Diluent, 126-127, 150 Dilution spread plate count bacteria, 121 Fe-depositing heterotrophs, 1085-1086 Mn-oxidizing heterotrophs, 1085-1086 Dinitrogen emissions, from 15N-labeled fertilizer,1060-1061 Dinitrogen fixation, 1019-1043 acetylene reduction test, 1019-1025 calibration, 1042-1043 grass systems, 1023-1025 nodulated root system, 1020-1023 N difference methods, 1025-1030 controlled environment, 1025-1028 field experiments, 1028-1030 15N methods, 1030-1038 isotope dilution method, 1030-1036 15N natural abundance method, 10361038

SUBJECT INDEX using dinitrogen-15 gas, 1038-1042 Dinitrogen-15 gas calibration of acetylene reduction test, 1042-1043 measuring N2 fixation, 1038-1042 Dinitrogen-fixing bacteria actinorhizal symbionts. See Frankia free-living, 179-195 acetylene reduction assay, 180-181 Azotobacteraceae, 183-186 classification into genera, 183 clostridia, 192-193 cyanobacteria, 189-190, 428, 454 enrichment procedures, 180 H-using, 188-189 methanotrophs, 186-187 methods for N2 fixers in general, 181183 most probable number method, 180 photosynthetic purple nonsulfur bacteria, 191-192 sulfate-reducing bacteria, 194-195 nodule symbionts. See Rhizobia 2,6-Dinitrophenol, 411 Diphenylamidino indophenol (DAPI), 87 Diphenylmethane,411 Direct count enteric viruses, 114-115 phages, 110-112 Direct lysis approach, recovery of bacterial community DNA, 733-736, 741-743 Disinfection, soil, 366-367 Dispens-O-Disc kit, 700 Dispersion, bacteria in soil, 83 Dissimilatory nitrate reduction, 246 Dissimilatory nitrate reduction to ammonium (DNRA), 248-249 enumerations of strains, 255-256 Distilled water, as diluent, 150 DMSO extraction method, chlorophyll, 443444 DNA. See also Restriction fragment length polymorphism bacterial community. See Bacterial community DNA detection of specific sequences in environmental samples, 707-725 fingerprinting, 647-656, 700 gel electrophoresis, 650-652, 655-656 isolation of DNA fragments from gel, 669-671 isolation and purification, 649 large-scale, 653-654 small-scale, 654-655 minipreps, 654-655 quantitation spectrophotometry, 746-747 UV fluorescence in presence of ethidium bromide, 747-750 Southern transfer, 659-660

SUBJECT INDEX DNA probe. See also Nucleic acid probes denitrifiers, 263-265 Frankia, 308-309, 319-321 radioactive hybridization of Southern blot to, 660661 preparation, 660 DNNRNA assays, Frankia nodule, 314-316 DNA-based microbial monitoring, 727-729 DNRA. See Dissimilatory nitrate reduction to ammonium Dormant organisms, 156 Dorylaimida, 483-485 Dot blot, 607, 680-682 Double PCR, 722 Doubling time, 416-417 DPM medium, 301 Drinking water, coliform content, 146-147 Drop plate method, count of bacteria, 133 Dry combustion, 939-941 Dry funnel method, arthropod extraction from soil, 522-527 Dry heat sterilization, 43-44 Dry weight, cellular water content and, 100 Dual culture method, antibiotic-producing microorganisms, 385-389 Dumas technique, 939-941 Durham tube, 152-153 Eadie-Hofstee transformation, 781 EC medium, 151 Ectomycorrhizal roots, isolation of filamentous fungi, 341 Electric field method, protozoa, 509 Electrophoresis gel. See Gel electrophoresis MLEE analysis. See Multilocus enzyme electrophoresis analysis Electrophoresis duplicating paper, 630 ELISA. See Enzyme-linked immunosorbent assay Elutriation arthropod extraction, 527-528 nematode extraction, 470, 474-475 Emission spectrometry, measuring N-isotope ratio, 942, 1035 End labeling, oligonucleotide probes, 678679 Endolithic algae, 439-440 Endolithic habitat, 427 Endosymbiotic cyanobacteria, cycad roots, 454-455 Enoplida, 485-486 Enrichment culture biodegradation research, 408-412 free-living N2-fixing bacteria, 180 Enrichment procedures, phages, 112-113 Enteric Differential system, 550 Enteric virus, direct count, 114-115 Enterobacter, 145-156, 179-195,248

1105

Enterobacteriaceae, 145-156, 179,248 Enterobacterial repetitive intergenic consensus (ERIC) sequences, 647 Entero-Set 20, 549 Enterotube system, 550 Enzyme(s). See also Multilocus enzyme electrophoresis analysis acid phosphatase, 777, 802, 807-809 aconitase, 568 activation energy, 783 adenylate kinase, 568 alcohol dehydrogenase, 568 catalase, 777 catechol dioxygenase, 419-421 cofactors, 784-785 concentration, 778-782 detection of marker enzyme synthesis for bioassay, 723-724 extracellular, 777 extraction from soils, 777 factors affecting rates, 778-790 ~ galactosidase, 568, 691 glucose 6-phosphate dehydrogenase, 568 a glucosidase, 777 glucuronidase, 154-155 hexokinase, 568 L-histidine ammonia-lyase, 826 hydrogenase, 188-189, 1020 hydroxybutyrate dehydrogenase, 568-569 hypoxanthine dehydrogenase, 569 indophenol oxidase, 569 inhibition, 784-787 competitive, 786-787 noncompetitive, 786-787 uncompetitive, 786 invertase, 777, 826 ionic environment, 784-785 isocitrate dehydrogenase, 569 lactose permease, 691 leucine aminopeptidase, 569 leucine dehydrogenase, 569 leucyl-DL-alanine peptidase, 569 leucyl-glycyl-glycine peptidase, 569-570 lysozyme, 739 malate dehydrogenase, 570 mechanism of action, 778 meta-pyrocatechase, 419-421 monooxygenase, 408 nitrogenase, 188-189, 1019-1025 nitrous oxide reductase, 247-248, 263 nucleoside phosphorylase, 570 peroxidase, 609 pH dependence, 783-784 phenylalanyl-L-leucine peptidase, 570--571 phosphatase, 801-814 phosphoglucomutase, 570 6-phosphogluconate dehydrogenase, 570 phosphoglucose isomerase, 570 polymorphic, 561 pronase E, 739

SUBJECT INDEX

1106

Enzyme(s) (cont.) ortho-pyrocatechase, 419-421 soil, 775-826 amidase, 790-801 arylsulfatase, 777, 814-818 asparaginase, 790-801 dehydrogenase, 777, 820-823 ~-glucosidase, 823-826 L-glutaminase, 790-801 804, inorganic pyrophosphatase, 810--812 phosphodiesterase, 801-814 phosphomonoesterase, 801-814 rhodanese, 818-820 trimetaphosphatase, 805, 812-813 urease, 777, 790-801 substrate concentration, 778-782 Taq polymerase, 707-708, 710, 717 temperature dependence, 782-783 Enzyme-linked immunosorbent assay (ELISA), 593, 601, 606-616 alkaline phosphatase-linked, 611-613 biotinylated second antibody for enhanced ELISA,613 checkerboard titration, 613 direct, 606-607 endogenous enzymes interfering with, 609 indirect, 607-608 peroxidase-linked, 609-611 Epifluorescence microscopy chlorophyll determination, 434 VAM fungi, 369-370 Epoxy embedding, 521-522 Ergosterol determination filamentous fungi, 345 VAM fungi, 356 ERIC sequences. See Enterobacterial repetitive intergenic sequences Erwinia, 179-195 Erythromycin resistance, 578 Escherichia coli, 145-156 direct methods for detection, 156 rapid test, 154-155 Ethanol extraction method, chlorophyll, 442-443 Ethidium bromide, 739 extraction from DNA with isopropanol, 744-745 quantitation of DNA, 747-750 Ethylbenzene, 414 Ethylene oxide sterilization, 45-47 Euglena, 453 Euglenophyceae, 431 Extracellular enzymes, 777 Extraction and flocculation method, total bacteria, 89-90 Facultative anaerobes, 223 Fasiculochloris, 430

Fatty acid analysis, bacteria, 551-553

Fecal coliforms. See Coliform bacteria, fecal Fecal contamination, 145-146 Fenwick can method, extraction of Heterodera cysts, 476-477 Fertilizer 15N-Iabeled conventional fertilizers, 916-917 enrichment level and form, 915-916 N2 emissions, 1060-1061 organic residues, 917-918 slow-release formulations, 917 S,1067 VAM inocula production, 367-368 Feulgen stain, 502 Field chambers, C mineralization, 839 Field plot, large, N tracer studies, 924 Field-inversion gel electrophoresis (FIGE), 650-651 FIGE. See Field-inversion gel electrophoresis Filamentous algae, 430 Filamentous cyanobacteria, 430 Filamentous fungi, 329-347 culture media, 330-333 isolation direct hyphal, 336-337 direct observation, 340-341 immersion methods, 335-336 root fragmentation, 340 root surface disinfection, 338-339 root washing, 338 selective methods, 341 soil dilution plate method, 333-334 soil plate methods, 334-335 soil washing, 337-338 quantitative methods chitin determination, 345 direct observation, 342 ergosterol determination, 345 membrane filtration, 344 soil-agar film, 342-344 substrate-induced respiration, 345-347 selective methods, 341 Filter blackening procedure, 84-85 Filter paper method, soil water potential, 53-57 Filter-sterilization, gases, 226-228 Filtration method Frankia, 297

protozoa, 507 total bacteria, 83-87 Filtration-gridline method, total hyphae of VAM fungi, 358-359 Finney's PSI function, 37-38 FITC. See Fluorescein isothiocyanate Fixation actinomycetes, 274 protozoa, 501-502 Flagellar stain, 502 Flagellates, 492, 499, 510

SUBJECT INDEX Flask method, extraction of Heterodera cysts, 475-476 Flavobacterium, 264, 415 Fleming's fixative, 502 Flexibacter, 264 Flotation method arthropods, 527 protozoa, 509 Flow cytometry, protozoa, 509 FLSD test, 31 Fluorescein diacetate method active hyphae of YAM fungi, 359 active mycelia, 98-99 Fluorescein filter set, 85 Fluorescein isothiocyanate (FITC), 95, 604606 Fluorescein-labeled immunoglobulin conjugate, 87 Fluorescence enhancer, 88 Fluorescence microscopy, algae and cyanobacteria, 434 Fluorescent antibody method Frankia, 319 nitrifying bacteria, 165-166 Fluorescent pseudomonads, 136-137, 689-705 Fortuitous metabolism, 408 Frankia, 271,291-322 cell wall chemistry, 306 characteristics, 291-294 in symbiosis, 309-316 culture media, 299-302 culturing, 302-303 decontamination, 302 production of clonal strains, 302-303 DNA sequencing and strain specific probes, 308-309 DNA:DNA hybridization, 308 DNA/RNA probes, assay in soil, 319-321 fluorescent antibodies against, 319 host specificity, 309-310 incubation and subculturing, 298 infective units, 317 isolation, 295-299 density fractionation, 297 differential filtration, 297 microdissection of nodule, 296-297 nodule sterilization, 296 maintenance of cultures, 303 most probable number method, 318-319 multi-locus allozyme electrophoresis, 307 N2 fixation, 310-313 nodulation capacity, 317-318 nodule DNA/RNA assays, 314-316 nodule metabolism, 310-313 nodule morphology, 313-314 PCR, 319-321 plant bioassay, 316-319 plasmids, 307 protein patterns, 306-307

1107

quantification, 316-320 active biomass, 305 total biomass, 305 RFLP analysis, 308 sample collection, 295-296 serology, 306 sporulation, 313-315 strain differentiation, 306-309 strain registry, 303-304 taxonomy, 303-305 Frankia medium, 300 Frustu1e, 429, 436-437 Fulvic acid, 895 Fumigant, 45-47 Fumigation, 754-760 Function probes, 665-666, 668 Fungal disk methods, antibiotic-producing microorganisms, 387-388 Fungi antibiotic-producing, 384 biomass, 101 filamentous, 329-347. See also Filamentous fungi metabolically active mycelia fluorescein diacetate hydrolysis method, 98-99 tetrazolium dye reduction method, 99-

100

mycelial length agar film method, 96-97 membrane filtration method, 97-98 Fusarium oxysporium, 249 B-Galactosidase, 568, 691 Gallionella ferruginea, 1089-1091 Gamma irradiation, sterilization of soil, 44-45 Gas(es) filter-sterilization, 226-228 soil. See Soil gases Gas chromatography, 1021 CO 2 in gaseous samples, 837-838, 844-849, 855-858 column packings, 849 denitrification studies, 1051-1053 detector 63Ni electron capture, 849 ultrasonic, 849 O 2 in gaseous samples, 838, 844-849, 855858 Gas collection vial, 844 Gas lines, O 2 removal, 226-228 Gas sample, 846 Gas-sampling well, 844-848 Gas-sampling device, 1061 Gassing manifold, 850, 852 Gas-storage device, 1061 Gastroenteritis, 145 Gel electrophoresis agarose, 650, 655-656

SUBJECT INDEX

1108 Gel electrophoresis (cont.) apparatus, 651-652 buffers, 652-653 DNA, 650-652, 655-656 isolation of DNA fragments from gel, 669-671 field.:tnversion, 650-651 one-dimensional,620 plasmid profiles, 638-643 protein profiles, 619-633 electrophoresis procedure, 623-628 gradient gel, 631 molecular weight markers, 627 principles, 620-621 storing and recording profiles, 630 visualizing proteins, 628-629, 632 pulsed-field,650-651 resolving gel, 621, 625 SDS-PAGE, 620 stacking gel, 621, 626-627 starch, 564-566 two-dimensional, 620 Gene probes, 665-686. See also Nucleic acid probes Gene transfer, horizontal, 724-725 Generation time, rhizobia, 202 Genetic distance dendrogram, 562 Genetic relatedness, 557-572 Genetic variation, 557-572 Genetically engineered microorganisms, lacZY-marking system, 689-705 Genetics, population, 557-572 Genius System (nonradioactive probe kit), 673-674 Genomic rearrangements, 724-725 Gentamycin resistance, 578 Geobacter metallireducens, 1079 Geodermatophilus, 271 Geometric mean, 25 Gigaspora, 363 Gloeocapsa, 190, 430 Glomus, 362 Glove box, anaerobic; 233-235 Gluconobacter, 264

Glucose nutrient agar, 383 Glucose 6-phosphate dehydrogenase, 568 a-Glucosidase, 777 ~-Glucosidase, 823-826 Glucuronidase, Escherichia coli, 154-155 L-Glutaminase, 790-801 Glutaraldehyde fixative, 502 Glycerol arginine agar, 383 Glycomyces, 271 Glycoside hydrolase. See ~-Glucosidase Gonostomum, 495

Goodness-of-fit tests, 19-21 Gordona, 271 Grass system, acetylene reduction test for N2 fixation, 1023-1025

Gravimetric method, soil water potential, 53-57 Green algae, 429, 431 Green photosynthetic sulfur bacteria, 179 Gridline-intersect method, colonized root length of VAM fungi, 355-356 Growth pouch, plastic, 207-210 Growth response, VAM fungi, 361-365 Growth-pouch infection assay, rhizobia, 216-218 Guillard's freshwater enrichment basal salt mixture, 450-451 Gum arabic, 214 Gunnera, 454

Hagem's agar, 332 Hanes-Wolf transformation, 780 Harvesting of cells, 417 Hauck technique, 15N-modified, 1055-1060 Headspace gas, 856-857 Helicotylenchus, 462 Herbaspirillum, 179 Heterodera cyst

extraction from soil Fenwick can method, 476-477 flask method, 475-476 recovery and counting, 477 Hexokinase, 568 High-copy-number plasmid, 636 High-gradient arthropod extractor, 524-525 L-Histidine ammonia-lyase, 826 Histogram, 17-19 Hoechst 33258, 87 Hollande's fixative, 502 Host plant, VAM fungi, 353, 366 Host specificity, Frankia, 309-310 Hot start method, PCR, 718 Housekeeping enzyme genes, 559-560 Humic acid, 895 Humic content, soil, 730 Humin, 895 HV agar, 276-277 Hybridization colony, 666, 680-682 direct DNA extracts, 666 DNA:DNA, Frankia, 308 nucleic acid probes, 679-684 in situ, detection of actinomycetes, 273-274 slot/dot blots, 680-682 Southern blot to radioactive DNA probes, 660-661 whole-cell, 102-103 Hybridoma, 596, 600 Hydrogenase 02-sensitive, 188-189 uptake, 1020 Hydrogen-using dinitrogen-fixing bacteria, 188-189

SUBJECT INDEX Hydrolyzable nitrogen, 940 Hydroxybutyrate dehydrogenase, 568-569 Hyperparasite, 367 Hyphae fungal, direct isolation, 336-337 YAM fungi, quantification, 357-359 Hypothesis testing, 28-31 Hypoxanthine dehydrogenase, 569 IAR strains. See Intrinsic antibiotic resistance strains Immersion methods, isolation of filamentous fungi, 335-336 Immobilization, N, 985-1016 Immunoassay, 606-616 Immunoblot, 607, 613-616 Immunofluorescence microscopy hyphae of YAM fungi, 359 nitrifying bacteria, 165-166 total bacteria, 87-89 Immunoglobulin G, 594-595, 602 Immunoglobulin M, 594-595 Implanted slide method, algae and cyanobacteria, 435 Incompatible plasmids, 637 IncP plasmids, 637 IncQ plasmids, lacZY-marking system, 689705 Index of abundance method, visual estimation of cyanobacteria in rice fields, 439 Indicator organism coliform bacteria, 145-156 isolation of antibiotic-producing microorganisms, 380-381 Indirect reduction, 1091 Indophenol blue test, 975-979 Indophenol oxidase, 569 Infection-unit method, YAM fungi, 353 Infective unit, Frankia, 317 Infectivity assay, YAM fungi, 353 Infrared gas analyzer, 837-838 Inoculant enumeration of rhizobia, 206-210 field experimentation, 210-216 inoculation of seed, 213-215 Inoculant carrier, 211 Inoculating stamp, multipoint, 549 Inorganic nitrogen, 940 ammonium method automated test, 975-977 manual test, 977-979 nitrate method automated test, 968-972 manual test, 972-974 production during aerobic incubation, 960, 962-967 Inorganic pyrophosphatase, 804, 810-812 Insects, 517-542. See also Arthropods

1109 Internal antigen, 598 Intrinsic antibiotic resistance (IAR) strains, 575-576 evaluation, 585-589 cultural tests, 586-587 field studies, 587-589 growth chamber or greenhouse tests, 587 selection, 581-585 agar-diffusion method, 583-584 agar-dilution method, 584-585 broth-dilution method, 585 Invertase, 777, 826 Iodonitrotetrazolium method, active hyphae of YAM fungi, 359 Ionic environment, enzymes and, 784-785 Iron-depositing heterotrophs dilution spread plate counts, 1085-1086 enrichment, 1081-1085 microaerobic or O2 gradient methods, 1083 natural material based method, 1083 isolation, 1081-1085 Iron-hematoxylin stain, 502 Iron-oxidizing autotrophs acidophiles, 1086-1089 neutrophiles, 1089-1090 Iron-reducing heterotrophs, non-enzymatic, enumeration, 1090-1092 Iron-respiring bacteria enrichment, 1092-1094 most probable number method, 1092-1094 Irradiation, soil sterilization, 790 Isobutylidene diurea, 917 Isocitrate dehydrogenase, 569 Isopropanol extraction, ethidium bromide from DNA, 744-745 Isotope dilution method isotope dilution calculation, 911-915 N2 fixation, 1030-1036 Isotopic equilibria, 892 Isotopic indices, 883-884 Isotopic methods, 865-901 Jaccard coefficient, 546, 550 Judgmental sampling, 4-5, 148 9K medium, 1087 Kanamycin resistance, 578 Keyser-Munns agar, 204 Kibdellosporangium, 271 Kineosporia, 271, 285 King's B medium, 383, 399, 695 Kjeldahl method, 936 Klebshormidium, 431 Klebsiella, 145-156,179-195,248-249 KT medium, 1087 Kucera-Wolfe mineral medium, modified, 1090

1110

U2 medium, 301 Lactose, 691-692 Lactose permease, 691 lacZY-marking system, 689-705 attributes and deficiencies, 703-704 donor strain, 694 exconjugant type isolate vigor, 700-701 helper strain, 694 marker stability, 701 materials, 693-696 principles, 691-693 procedures, 696-700 confirmation of phenotype, 698-699 identification of exconjugants, 699-700 mating and selection, 696-698 recipient strain, 693-696 recovery of bacteria from nonsterile soil, 702-703 Lag phase, 416 Land's exact confidence limits of mean, 28-31 Lauryl tryptose broth, 151, 154 Leaching, enteric viruses, 109 Legume nodule symbionts, 199-218. See also Rhizobia Leonard jar, 1026-1027 Leptospirillum ferrooxidans, 1087-1088 Leptothrix ochracea, 1089 Leucine aminopeptidase, 569 Leucine dehydrogenase, 569 Leucyl-DL-alanine peptidase, 569 Leucyl-glycyl-glycine peptidase, 569-570 Light effect, VAM inocula production, 367 Light microscopy, 81-104 actinomycetes, 273-274, 285-287 algae and cyanobacteria, 431, 433-434 biomass determination, 770 nematodes, 479-482 protozoa, 498-499 sampling of soil, 81-82 Lignobacter, 179 Lineweaver-Burk transformation, 780 Linkage disequilibrium, 561 Linkage equilibrium, 561 Lipophilic extraction, arthropods, 528 Lipopolysaccharide (LPS), 598 Location parameter mean, 27-28 median, 26-27 selection, 25-28 Lognormality characteristics of lognormal distribution, 16-17 diagnosis, 17-25 Low-copy-number plasmid, 636 LPS. See Lipopolysaccharide Luria-Bertani medium, 695 Lysimeter, 932-934 Lysine dehydrogenase, 570 Lysozyme, 739

SUBJECT INDEX M3 agar, 276 M9 basal salts solution, 135, 696 M9 minimal medium, 135 Malate dehydrogenase, 570 Malt extract agar, 331, 383 Manganese extractable, 44, 47 oxidation and reduction, 1079-1094 Manganese-oxidizing heterotrophs dilution spread plate count, 1085-1086 enrichment, 1081-1085 microaerobic or O 2 gradient methods, 1083 natural material based methods, 1083 isolation, 1081-1085 Manganese-reducing heterotrophs, nonenzymatic, enumeration, 1090-1092 Manganese-respiring bacteria enrichment, 1092-1094 most probable number method, 10921094 Manure, 15N-labeled, 917-918 Mass spectrometry 13C analysis, 880-884 measuring N-isotope ratio, 941-942, 1035 Master mixes, PCR, 713 Matric potential, soil, 57 Matrix table, 537-538 Mean (population), 7-8, 16-17,21,25-26 confidence interval, 24 detecting difference, 28-30 effects of sample volume, 26 geometric, 25 as location parameter of choice, 27-28 sample number for estimating, 31 UMVU estimators, 21-23 Media. See Culture media Median (population), 16-17,21,25-26 confidence limits about, 24-25 detecting differences, 28 effects of sample volume, 26 as location parameter of choice, 26-27 UMVU estimators, 21-23 Megaplasmid, 635-636, 644 Melin-Norkrans medium, 333 modified, 333 Meloidogyne, 462, 465 Membrane filtration method filamentous fungi, 344 mycelial length determination, 97-98 protozoa, 502-503 Mercuric chloride sterilization, 47-48 Merthiolate, 87 Mesoarthropod extractor, 523-524 Metabolism cometabolism, 408 fortuitous, 408 Metal ions, effect on enzyme activity, 785 Methane, 849

SUBJECT INDEX Methanogens, 179,225 Methanotrophs, 179 most probable number method, 187 Nand C-deficient medium, 186-187 N2 fixation, 186-187 plate count, 186-187 Methyl bromide sterilization, 45-47 Methylene blue, 231-232 Methylobacter, 186-187 Methylococcus, 186-187 Methylocystis, 186-187 Methylomonas, 186-187 Methylosinus, 186-187 Methyltrophs, methane-using. See Methanotrophs Methylumbelliferone glucuronide, 154-155 Michaelis-Menten equation, 780 Microaerophiles, actinomycetes, 282 Microalgae. See Algae Microarthropod extractor, 524-526 Microbial inhibitors, 41 Microbial lawn methods, antibiotic-producing microorganisms, 385-387 Microbial monitoring, DNA-based, 727-729 Microbispora, 270-271,277,285 isolation, 278 Microcosm, for rearing arthropods, 535-536 Microdissection, isolation of Frankia from nodule, 296-297 Micro-ID system, 550 Micromonospora, 269-271,277,282,285 isolation, 278 Microplot, N tracer studies, 923-924 Microscopy. See Light microscopy Microstructure, soil, 521-522 Microsubplot, N tracer studies, 924-926 Microtetraspora, 271,277,285 isolation, 278 Microtiter plate method, C source utilization, 546-552 Microwave irradiation, sterilization of soil, 45 Mineral salts medium, 410 Mineralization, 835 C. See Carbon mineralization CO 2, determination, 756-757 N. See Nitrogen mineralization Minimal lactose X-gal medium, 695 Minitek system, 549-550 Mist chamber method, nematode extraction from plant material, 478 MLEE analysis. See Multilocus enzyme electrophoresis analysis Moist heat sterilization, 42-43 Molybdenum, 214 Monhysterida, 485-486 Monoclonal antibodies, 596-597, 599-600 Mononchida, 485-487 Monooxygenase, 408

1111 Most probable number (MPN) method, 59-78 algae and cyanobacteria, 438, 440-441 calibration of counts, 65 confidence limits, 61-62, 75 denitrifiers, 251-255 Fe-respiring bacteria, 1092-1094 Frankia, 318-319 free-living N2-fixing bacteria, 180 mathematical solution of most probable number, 60-61 methanotrophs, 187 methodology assigning tabular population estimates, 70 calculations, 75-77 correcting for initial dilution and inoculant volume, 71-72 experimental design, 65-66 materials, 66-67 preparation of dilution series and culture of inoculated units, 69 recording results, 69-70 soil sampling, 68 Mn-respiring bacteria, 1092-1094 nitrifying bacteria, 160-163 probability of occurrence of most probable number, 62-63 protozoa, 503-504 rhizobia, 207-210 tables, 71-74 tests of technique, 62-65, 75-77 theoretical assumptions, 60 VAM fungi, 352-353 MPN method. See Most probable number method Mucilage, 428 Multilocus allozyme electrophoresis, Frankia, 307 Multilocus enzyme electrophoresis (MLEE) analysis, 557-572 buffers, 560-561, 565 choice of enzymes, 560-561, 565 collecting and analyzing data, 571-572 electrophoresis procedure, 563-566 enzyme staining, 566-571 preparation of enzyme extracts, 562-563 principles, 559-572 Multiple agar layer technique, antibioticproducing microorganisms, 395-396 Multiple-tube fermentation method, fecal coliforms, 149-152 Multipoint inoculating stamp, 549 Mycelia actinomycetes, 269-270, 273 length determination agar film method, 96-97 membrane filtration method, 97-98 metabolically active

1112 Mycelia (cont.) fluorescein diacetate hydrolysis method, 98-99 tetrazolium dye reduction method, 99100 Mycobacterium, 179,269,271,285 Mycolytic enzymes, isolation of microorganisms producing, 400-401 Mycorrhizal dependency, VAM fungi, 361365 Mycorrhizal fungi, 332-333 NIF system, 550

Nalidixic acid, 93-94 Nalidixic acid resistance, 577-578 Naphthalene, 411, 414 Natamycin, 136 Nematodes, 459-487 biomass, 459-460 distribution in soil, 461-462 extraction from plant material Baermann funnel method, 478 direct examination, 477-478 mist chamber method, 478 root-incubation method, 478 extraction from soil Cobb sieving and decanting, 469-471 density centrifugation, 470, 473-474 elutriation, 470, 474-475 Heterodera cysts, 475-477 modified Baermann funnel method, 470-473 food items, 459 functions in soil, 460 identification, 479-487 length, 459 light microscopy, 479-482 permanent mounts, 480-482 sampling, 461-469 costs, 463, 467 ecological studies, 467-469 sample collection, 462-463 sample size, 467-468 sampling pattern, 463-466 stratification of sampling, 467 timing, 466 with stylets, 483-485 taxonomy, 482-487 temporary mounts, 479-480 trophic categories, 459-460 vertical migration, 465 without stylets, 484-487 Neomycin resistance, 578 Neospongiococcum, 430 Nephelometry flask, 411, 416 Neutrophiles, Fe-oxidizing, 1089-1090 Nfl kit, 699 Nick translation, 672, 674-676 Nigrosin, 502

SUBJECT INDEX

Nitrate method, inorganic N automated test, 968-972 manual test, 972-974 Nitrate nitrogen, 939 Nitrate reductase, 826 dissimilatory, 263 Nitrate reduction. See also Denitrifier(s) assimilatory, 245-246 chemodenitrification, 250 dissimilatory, 246 dissimilatory nitrate reduction to ammonium, 248-249 nitrate respiration, 249 nonrespiratory denitrification, 249-250 respiratory denitrification, 246-248 Nitrate respiration, 249 Nitrate test, soil preplant profile, 954 pre-sidedress, 954-955, 957-960 residual profile, 955-957 Nitric oxide reductase, 247-248, 263 Nitrification, 159,985-1016 gross, 987 laboratory methods, 1009-1015 net, 986-987 Nitrification potential, 171-172, 1009 shaken soil-slurry method, 988,1011-1015 Nitrifying bacteria ammonia-oxidizers, 159-176 autotrophic, 159-176 diversity, 160, 163-165 immunofluorescence microscopy, 165-166 isolation, 167-169 maintenance of pure cultures, 169-171 most probable number method, 160-163 nitrifying potential of soil, 171-172 nitrite-oxidizing, 159-176 short-term nitrifying activity and, 172-176 Nitrite nitrogen, 939 Nitrite reductase, 265 dissimilatory, 247 Nitrite-oxidizing nitrifying bacteria, 159-176 Nitrobacter, 159-176,264 Nitrogen acid-insoluble, 899 amino acid-N, 899 amino sugar-N, 900 (ammonia + amino sugar)-N, 899 ammonia-N, 899, 939 availability ratio, 891-892 biomass calculation, 759-760, 766-767 determination, 765 hydrolyzable, 940 immobilization, 985-1016 inorganic. See Inorganic nitrogen nitrate-N,939 nitrite-N,939 reimmobilization, 760

SUBJECT INDEX Nitrogen (cont.) (serine + threonine)-N, 900 total, 936-940 total hydrolyzable, 898-899 Nitrogen availability indices, 951-984 current status, 951-955 field methods, 954-960 laboratory methods, 960-968 biological, 961-967 chemical, 953, 967-968 methods for inorganic N, 968-979 undisturbed soil core incubation, 953-954 Nitrogen cycling, 985-986 Nitrogen difference methods, N2 fixation, 1025-1030 controlled environment, 1025-1028 field experiments, 1028-1030 Nitrogen-15 isotope techniques denitrification, 1049 dinitrogen-15 gas, 1038-1042 N2 fixation, 1030-1038 isotope dilution method, 1030-1036 15N natural abundance method, 10361038 Nitrogen-15 labeled materials. See also Nitrogen tracers commercial sources, 942-943 preparation determining 15N concentration needed, 911-915 fertilizers, 915-919 15N-labeled soil, 918-919 principles, 909-911 Nitrogen-15 labeled organic matter calculation of N availability ratio, 891-892 decomposition, 887-893 extraction from soil, 896-900 labeling biomass, 889-890 labeling organic matter, 887-890 labeling plants with 15N-fertilizer, 888-889 mineralization rate determination, 890-891 preparing samples for 15N analysis, 891 Nitrogen mineralization, 890-891, 953, 9851016 determination, 757 gross, 987 net, 986-987 rates, 890-891 Nitrogen-15 natural abundance method N2 fixation, 1036-1038 N transformations, 987-988 Nitrogen-13 studies, 907 Nitrogen tracers, 907-943 field studies, 919-935 application techniques, 921-923 confined microplots, 923-924 large plots, 924 managing field variability, 920-921 plot type and size, 923-926

1113 principles, 919-920 unconfined microsubplots, 924-926 15N supply and analytical service, 942-943 N transformation rates, 987-988 N-isotope ratio, 935-942 conversion of labeled N to ammonium, 936-939 conversion oflabeled N to N2, 939-941 measuring, 941-942 preparation, 909-911 fertilizers, 915-919 determining 15N concentration needed, 911-915 15N-labeled soil, 918-919 principles, 909-911 sampling plants, 926-927 soil, 928-930 soil gases, 934-935 soil water, 930-932 Nitrogen transformations, 985-1016 accumulation of inorganic N in closed-top solid cylinders, 988, 1002-1004 accumulation of inorganic N in polyethylene bags, 988, 1004-1006 aerobic incubation method, 988, 10061009 field methods, 999-1006 laboratory methods, 1006-1009 measurement of gross transformation rates, 987-999 15N isotope-dilution method, 988, 990-999 15N natural abundance method, 987-988 N tracer methods, 987-988 Nitrogenase, 1019-1025 02-sensitive, 188-189 Nitrogen-deficient combined C medium, 182 Nitrogen-deficient lactate medium, 194-195 Nitrogen-deficient malate agar medium, 191 Nitrogen-fixing bacteria. See Dinitrogen-fixing bacteria Nitrogen-isotope ratio, 935-942 conversion of labeled N to ammonium, 936-939 conversion of labeled N to N2, 939-941 measuring, 941-942 Nitrosococcus, 165 Nitrosolobus, 159-176 Nitrosomonas, 159-176,247,264 Nitrosovibrio, 165, 170 Nitrospira, 159-176 Nitrous oxide flux, 1055 Nitrous oxide reductase, 247-248, 263 Noble agar, 139 Nocardia, 269, 271, 285 Nocardioforms, isolation, 278-279 Nocardioides, 271, 285 Nocardiopsis, 271, 281, 285 alkalophilic, 282

1114 Nodulation capacity, Frankia, 317-318 Nodule, 199. See also Rhizobia acetylene reduction for nodulated root system, 1020-1023 collection, 200-202 disinfection, 200-201 Frankia, 291 DNA/RNA assays, 314-316 metabolism, 310-313 morphology, 313-314 Frankia isolation from microdissection, 296-297 sterilization of nodule, 296 Noncompetitive inhibition, 786-787 Non-conjugative plasmid, 637 Nonparametric statistical methods, 28-31 Nonrespiratory denitrification, 249-250 Normal distribution, 7 Normality assumption, 28 Nostoc, 189-190,429-430,454 Novobiocin resistance, 578 NPCC medium, 136-137 Nucleic acid probes. See also DNA probe detection, 684-686 autoradiography, 684-685 Cerenkov counting, 685-686 scintillation counting, 684-686 hybridization, 679-684 labeling, 671-679 end labeling of synthetic oligonucleotides, 678-679 nick translation, 672, 674-676 random primer fill-in, 672, 677 nonradioactive, 673 purification of DNA fragments, 669-671 selection, 667-669 single stranded, 672, 677 size, 669 Nucleic acid stains, 84-87 Nucleoside phosphorylase, 570 Numerical taxonomy, 543-553 Nutrient agar, 135 Nutrient cycling, 753-770, 775 Nutrient pools, 753-770 Nutrient-poor media, 138-139 Nutrient-rich media, 138-139 O-antigen, 598 Oatmeal agar, 276 Obligate anaerobes, 223-224 Oerskovia, 271 Oligonucleotide probe, 667 actinomycetes, 273 directed at 16S rRNA, 101-103 labeling, 678-679 Organic matter, 730, 865-901 C mineralization, 835-859 13C natural abundance technique, 875-886 apparatus and reagents, 877-888

SUBJECT INDEX calculating sources of soil organic matter, 884-885 conversion of organic C to CO 2 for mass spectrometry, 880-883 field sampling, 878-879 kinetics of soil organic matter turnover, 885-886 mass spectrometric analysis and isotopic indices, 883-884 natural 13 C and measurement or organic matter dynamics, 877 preparation of soil samples, 879-880 stable C isotope ratios of plants and soils, 876 stable isotope terminology, 875-876 14C-labeled decomposition, 866-875 extraction from soil, 894-896 incubation of 14C-labeled materials, 871-875 labeling biomass and microbial products, 869-871 labeling plant material, 867-869 15N-labeled calculation of N availability ratio, 891892 decomposition, 887-893 extraction from soil, 896-900 labeling biomass, 889-890 labeling organic matter, 887-890 labeling plants with 15N-fertilizer, 888889 mineralization rate determination, 890891 preparing samples for 15N analysis, 891 sources, 884-885 turnover kinetics, 885-886 Organic residues, 15N-labeling, 917-918 Oscillatoria, 190 Osmotic potential, soil, 57 Overlay techniques, protozoa, 507-508 Oxamide, 917 Oxi-Ferm system, 550 Oxygen consumption study, biodegradation research, 421-423 detection, 838-839 gas chromatography, 844-849 removal activated steel wool method, 229 candle jar method, 229 cold catalytic removal with H, 228-229 with cultures of fast-growing aerobes, 229 from gas lines, 226-228 pyrogallol method, 229 static laboratory methods, 855-859 static incubation-gas chromatographic analysis, 855-858

SUBJECT INDEX Oxygen (cont.) static incubation-titrimetric determination, 858-859 uptake, 835-836 Oxygen electrode, 838-839 Oxygen probe, 421-423 Oxygen scavenger, 627 Palmella, 430 Paper disk method, antibiotic-producing microorganisms, 390 Paracoccus, 247, 264 Parathion, 411 Pasteur pipet, preparing fine capillary pipets, 444-445 PathoTec Rapid I-D system, 550 PCB,408 PCR. See Polymerase chain reaction Peat inoculant, 206, 211 Penicillin, 136 Penicillin resistance, 578 Pentachlorophenol, 411, 415 Peptone yeast extract agar, 135,203-204 Peptone-water, 126, 150 Percoll gradient, 90-91 Permanent mount, nematodes, 480-482 Peroxidase, 609 Peroxidase-linked immunosorbent assay, 609-611 Persulfate digest, 764 Pesticide, VAM inocula production, 367-368 PFGE. See Pulsed-field gel electrophoresis PGPR. See Plant growth-promoting rhizobacteria pH soil, 802 enzyme activity and, 783-784 Phage, 107-115. See also Virus direct count, 11 0-112 enrichment procedures, 112-113 lysogenic, 108 principles of analysis, 109-110 purification and storage, 113-114 virulent, 108 Phage-typing system, 107 Phenol,411 Phenol red, 545 Phenosafranine, 232 Phenylalanyl-L-leucine peptidase, 570-571 p-Phenylenediamine, 88 Phosphatase, 801-814 Phosphate salt solution, 696 Phosphate-buffered saline, 126, 150, 601 Phosphodiesterase, 801-814 Phosphoglucomutase, 570 6-Phosphogluconate dehydrogenase, 570 Phosphoglucose isomerase, 570 Phosphomonoesterase, 801-814

1115

Phosphorus-response curve, VAM fungi, 361-365 Photosynthesis, 428 estimating in algae and cyanobacteria, 453-454 Photosynthetic purple nonsulfur bacteria, N2 fixation, 191-192 Phylogenetic probes, 665-667 Physiological saline, 126, 150 Pilimelia, 271 Pitfall trapping, arthropods, 519-521 Planachromat lens, 87 Planobispora, 271 Planomonospora, 285 Plant(s) growing with 15N-labeled fertilizer, 888889 labeling with 14C, 867-869 lsN-labeling, 917-918 Plant bioassay, Frankia, 316-319 Plant growth-promoting rhizobacteria (PGPR),689 Plant interactive plasmid, 638 Plaque, phage, 112-113 Plasmid conjugative, 637 copy number, 635 cryptic, 637 degradative, 637-638 Frankia, 307 functions, 637-638, 644 high-copy-number, 636 incompatible, 637 low-copy-number, 626 N2-fixation genes on, 199 non-conjugative, 637 plant interactive, 638 profile, 635 -644 alkaline lysis miniprep, 642-643 applications, 644 extraction of plasmids, followed by CsCI ultracentrifugation and gel electrophoresis, 640-642 gel electrophoresis, 638-643 lysis of bacteria containing plasmids greater than 20 kb, 639-640 relationship to chromosome. 635-638 relaxed, 636 resistance, 637 storage of plasmid DNA, 643 stringent, 637 types, 636 xenobiotic-degrading bacteria, 413-414 Plasmid pJP4, 638, 644 Plasmid pKT230, 692 Plasmid pMON5002, 692 Plasmid pMON5003, 692 Plasmid pMON7029, 692 Plasmid pMON7117, 699

1116

Plasmid pMON7197, 692-694 Plasmid pRK2013, 694 Plastic growth pouch, 207-210 Plate count bacteria, 119-121 data handling, 139-141 diluents, 126-127 dilution plate method, 121 drop plate method, 133 materials and equipment, 122-123 pour-plate method, 121, 130-131 serial dilutions, 127-129 spread-plate method, 121, 131-133 limitations, 121-122 methanotrophs, 186-187 Plectonema, 190 Pleuromonas, 494 Polyclonal antibody, 596-597, 599 Polyethylene bag method, field estimation of N transformations, 988, 1004-1006 Polymerase chain reaction (PCR), 707 amplification cycles, 714-715 amplification protocol, 712-714 applications, 708 detection of marker enzyme synthesis for bioassay, 723-724 detection of specific DNA sequences in soil, 721-723 evolutionary and biodiversity studies, 724 genomic rearrangements, 724-725 horizontal gene transfer, 724-725 detection of Escherichia coli, 156 double, 722 Frankia, 319-321 hot start method, 718 identification of amplified products, 717 master mixes, 713 Mg concentration, 715 nucleotide concentrations, 717 optimization of amplification, 714-717 primer annealing, 709, 716 primer concentration, 715 primer design, 710-712 primer dimer artifact, 711 primer extension, 709-710, 716 quality control, 717-719 ramp times, 716 RNA-PCR,723-724 sensitivity of amplification, 720 specificity of amplification, 719 strand separation, 709, 715-716 theory, 709-710 thermal cycling profiles, 714-716 Polymixin B, 136 Polyvinylpolypyrrolidone, acid-washed, 737 Population genetics bacterial, 557-572 statistics, 571-572 Porous ceramic cup, soil water sampling, 931

SUBJECT INDEX Pot culture, VAM inocula, 366-368 Potassium azide sterilization, 48-49 Potato dextrose agar, 383 Pour-plate technique, count of bacteria, 121, 130-131 Power of statistical test, 28-30, 32 Preplant soil profile nitrate tests, 954 Pre-sidedress soil nitrate test (PSNT), 954955, 957-960 Primer, 707. See also Polymerase chain reaction Primer dimer artifact, 711 Probit analysis, 19 Promicromonospora, 271 Pronase E, 739 Propionibacterium, 249 Propionispira, 179 Propylene oxide sterilization, 45-47 Protargol, 502 Protein profile, whole-cell disruption of cells, 622 Frankia, 306-307 gel electrophoresis, 619-633 procedure, 623-628 growth and collection of cells, 621-622 preparation of protein extracts, 622-623 principles, 620-621 storing and recording profiles, 630 visualizing proteins, 628-629, 632 Proton-reducing bacteria, 225 Protozoa, 491-511 abiotic factors and, 493-494 C:P ratio, 495 community structure, 495-496, 509 cyst, 498-499 definition, 491 as ecological indicators, 494-495, 511 ecological roles, 494-495 enumeration Coulter counting, 509 density centrifugation, 506-507 direct observation, 498-503 dried soil smears, 499-500 electric field method, 509 filtration method, 507 flotation method, 509 flow cytometry, 509 indirect methods, 503-509 membrane filtration method, 502-503 most probable number method, 503-504 overlay methods, 507-508 soil suspensions on agar, 505-506 soil-agar films, 500-501 staining and fixation methods, 501-502 Uhlig ice extraction method, 508 watered soil suspension plus MPN methods, 505 watered soil suspensions, 501 environmental selection, 493-494

SUBJECT INDEX Protozoa (cont.) escape from adverse conditions, 498 feeding groups, 496 functional groups, 494-496 identification, 509-511 light microscopy, 498-499 morphology, 492 N cycling and, 494-495 numbers, 496 in different ecosystems, 493, 497 versus function, 491-492 versus identification, 492 as plant pathogens, 495 as predators of bacteria, 494-495 rand K selected, 496 seasonality, 494 soil and types of protozoa, 493-496, 501 soil dilution and, 499-500 taxonomy, 509-510 Pseudomonad, fluorescent, 136-137, 399-400 lacZY-marking system, 689-705 Pseudomonas, 179, 247, 264, 382, 638-639, 643 Pseudonocardia, 270-271,281,285 PTYG medium, 135 Pulsed-field gel electrophoresis (PFGE), 650-851 Purified agar, 139 Purple photosynthetic nonsulfur bacteria, 179 Purple photosynthetic sulfur bacteria, 179 meta-Pyrocatechase, 419-421 ortho- Pyrocatechase, 419-421 Pyrogallol,229 Qmod medium, 300 Quality control, peR, 717-719 Radial growth method, antibiotic-producing microorganisms, 391-392 Random primer fill-in, 672, 677 Random samples, 148 simple, 5 stratified, 5 Range of transition (ROT), 63, 66 Rapid identification systems, 549-550 Recalcitrant compounds, 407-424 Receiver plant, VAM fungi, 357 Recombination, 561 Redox indicators, 231-232 Reducing agents, 229-230 Reduction, indirect, 1091 Relaxed plasmid, 636 REP sequences. See Repetitive extragenic palindromic sequences Repetitive extragenic palindromic (REP) sequences, 647 Resazurin, 231-232 Residual profile nitrate test, 955-957 Resistance plasmid, 637

1117

Resolving gel, 621, 625 Respiration nitrate, 249 rate, 843 soil, 835 studies dynamic systems, 839 static systems, 839 Respiratory denitrification, 246-251 confirmation, 259-265 Respiratory equivalents, arthropods, 535 Respiratory gas, soil, 838 Respirometry study, 421-423, 756, 836. 838 Resting cells, 418 Restriction enzymes digestion of DNA samples, 655 selection and use, 649-650 Restriction fragment length polymorphism (RFLP), 647-648, 656-662 Frankia, 308 RFLP. See Restriction fragment length polymorphism Rhabditida, 484-486 Rhizobia, 199-218 cultivation, 203-205 culture media, 203-204 enumeration, 206-210 in inoculants, 206-210 most probable number method, 207-210 field experimentation involving inoculation, 215-216 generation time, 202 growth-pouch infection assay, 216-218 inoculants for field experimentation, 210213 inoculation of seed, 213-215 isolation, 200-202 maintenance of cultures, 205-206 megaplasmids, 644 nodule collection, 200-202 plasmid profiles, 635-644 recommended strains for selected legumes, 212 selective media, 206 Rhizobium, 199-218, 247, 264, 559-560, 565,575-576,581-582,593,597-599, 615, 621, 630, 638-641 Rhizoplane soil, 9 Rhizosphere soil, 9, 82, 460 Rhodamine-gelatin conjugate, 88 Rhodanese, 818-820 Rhodobacter, 191-192,247 Rhodococcus, 271, 285 Rhodomicrobium, 191-192 Rhodopseudomonas, 191-192 Rhodospirillum, 191-192 Riboflavin, 627 Ribosomal RNA actinomycetes, 273 phylogenetic probes, 665-667

1118

Ribosomal RNA (cont.) 16S, tluorescently labeled oligonucleotides directed at, 101-103 Rice field, cyanobacteria, 428, 437-439 Rifampicin resistance, 577-578, 580 Ringer's solution, 97, 126 RNA-peR, 723-724 Root colonization by VAM fungi, 353-356 coralloid, 455 cycad, endosymbiotic cyanobacteria, 454455 fragmentation, isolation of filamentous fungi, 340 surface disinfection, 338-339 washing, isolation of filamentous fungi, 338 Root exudate, compounds in, 547 Root hair, 217 Root nodule, 199. See also Rhizobia Root-incubation method, nematode extraction from plant material, 478 Root-tip mark method, growth pouch infection assay for rhizobia, 216-218 ROT. See Range of transition Roundworms. See Nematodes S medium, 300 SI medium, 137 Saccharomonospora, 270-271,277,281,285 Saccharopolyspora, 270-271, 281, 285 Saccharothrix, 271, 281, 285 Sampling plants, N tracer studies, 926-927 soil, 1-13 algae and cyanobacteria, 432-433 antibiotic-producing microorganisms, 380 arthropods, 518-519 bacterial plate count, 123-124 bias, 3 biomass estimation, 754 bulk samples for isolation, 8-9 bulking. See Bulking for characterization studies, 9-10 coliform bacteria, 147-149 complete excavation, 928 composite samples, 6-7 depth, 82 Frankia, 295-296 judgement samples, 4-5, 148 light microscopy, 81-82 measurement error, 3, 12-13 most probable number methods, 68 N tracer studies, 928-930 nematodes, 461-469 nonrhizosphere soil, 8-9 number of samples to take, 7, 31-32 precision, 4

SUBJECT INDEX

processing samples for microbiological studies, 10-12 random, 5, 148 rhizosphere/rhizoplane soil, 9, 82 rice field, 438 sample volume, 26-27 sampling error, 6, 12 sampling plan, 9 sampling unit, 3-4 selection error, 12 small-core, 929 soil collection, 148-149 spiked sample, 91-92 storage of samples, 10-12, 754 subplot excavation, 928-929 systematic, 5-6, 148 time, 82 soil gases, 934-935 soil water, 930-932 lysimeter, 932-934 porous ceramic cup, 931 tile discharge, 932 trough extractor, 931-932 SAR strains. See Spontaneous antibiotic resistance strains Sarcodina, 492 Sarkosyl, 738 Scanning electron microscopy, actinomycetes, 272-273 Schaudin's fixative, 502 Scintillation counting, 684-686, 868, 871-875 Screening methods, antibiotic-producing microorganisms, 393-396 Scutellospora, 363 Sealed chambers, 855 Seed inoculation with rhizobia, 213-215 surface sterilization, 210 Serial dilution, 127-129 (Serine + threonine )-nitrogen, 900 Serology, 593-616 Frankia, 306 Sewage, 145 Shaken soil-slurry method, nitrification potential, 988, 1011-1015 Shaking, release of bacteria from soils, 124 Shapiro-Wilk test of normality, 19-21 coefficients, 34-35 quantiles, 36 Shewanella putrefaciens, 1079 Shikimate dehydrogenase, 571 Short-term nitrifying activity, 172-176 Siderophores, isolation of microorganisms producing, 399-400 Silver staining protein profile, 632 protozoa, 502 Similarity matrix, ET, 562 Simple matching coefficient, 546, 550

SUBJECT INDEX Simple random samples, 5 Single agar layer technique, antibiotic-producing microorganisms, 394-395 Single stranded probes, 672, 677 Single-spore culture, VAM fungi, 368 Siphonous microalgae, 430 SIR. See Substrate-induced respiration Skewed probability density model, 16 Slime molds, 510 Slot blot, 680-682 Soda lime, 837, 841-844 Sodium azide sterilization, 48-49 Sodium phosphate buffer, 742 Sodium pyrophosphate, 0.1 %, 126 Sodium sulfide nanohydrate, 229-231 Sodium thioglycollate, 229-231 Soft agar overlay method, 112-113 Soil dilution plate method, filamentous fungi, 333-334 Soil enzymes. See Enzyme(s) Soil extract, 126 Soil extract medium, 134, 451 Soil gases, sampling, 934-935 Soil hydrolysate, preparation and sampling, 898 Soil plate method, filamentous fungi, 334-335 Soil respiration, 835 Soil smear, protozoa, 499-500 Soil solution equivalent medium, 135-136 Soil suspensions on agar, protozoa, 505-506 Soil veneer, 432-433 Soil washing, isolation of filamentous fungi, 337-338 Soil water, sampling, 930-932 lysimeter, 932-934 porous ceramic cup, 931 tile discharge, 932 trough extractor, 931-932 Soil water tubes, 451 Soil-agar film filamentous fungi, 342-344 protozoa, 500-501 Soil-conditioning agent, 428 Soil-less media, VAM inocula, 368-369 Soluble organic carbon, 764 Somatic antigen, 598 Sonication, release of bacteria from soils, 124 Southern blot, 659-661 Soybean-casein digest agar, 383 Spatial variation, soil properties, 1 Spectinomycin resistance, 577 Spectrophotometric method, quantitation of DNA,746-747 Spiked sample, 91-92 Spirillospora, 272, 285 Spontaneous antibiotic resistance (SAR) strains, 575-576 evaluation, 585-589 cultural tests, 586-587

1119

field studies, 587-589 growth chamber or greenhouse tests, 587 identification, 580 isolation, 578-580 selection, 577-580 choice of antibiotics, 577-578 Spore(s) actinomycetes, 269 VAM fungi, 360 Spore cloud, 280 Sporichthya, 272, 285 Sporulation, Frankia, 313-315 Spread-plate method, count of bacteria, 121, 131-133 Square count method, visual estimation of cyanobacteria in rice fields, 439 Stable isotopes. See specific isotopes Stacking gel, 621, 626-627 Staines) acid methyl-green, 502 acridine orange, 84-85 alkaline phosphatase linked, 611-613 bromothymol blue, 545 calcofluor M2R, 97-98 coomassie brilliant blue R, 628-629, 632 ethidium bromide, 739, 744-750 Feulgen, 502 fluorescein diacetate, 98-99, 359 fluorescein isothiocyanate, 95, 604-606 iodonitrotetrazolium, 359 iron-hematoxylin, 502 lacZY-marking system, 689-705 methylene blue, 231-232 peroxidase-linked immunosorbent, 609-611 phenol red, 545 phenolic aniline blue, 97-98 phenosafranine, 232 rhodamine-gelatin conjugate, 88 rhodanese, 818-820 tetrazolium dye reduction, 94-95, 99-100 tetrazolium formazan,545, 549 Staining ELISA, 593, 601, 606-616 enzymes in MLEE analysis, 566-571 nucleic acids, 84-87 protozoa, 501-502 total bacteria, 83-87 Standard error of mean, 7 Starch, 429 Starch casein agar, 276 Starch gel electrophoresis, 564-566 Static core protocol, denitrification measurement, 1049-1054 Static incubation-gas chromatographic analysis, CO 2 and O 2 , 855-858 Static incubation-titrimetric determination, CO 2 and O 2 , 858-859

SUBJECT INDEX

1120

Statistical tables, 34-37 Statistical tests, 15-33 computer programs, 37-38 plate count data, 139-141 population genetics, 571-572 power, 28-30, 32 Type I error rates, 29-30 Steam distillation, 896-897 Steel wool, activated, 229 Stem nodule, 199. See also Rhizobia Sterilization, soil, 41-49, 789 methods dry heat, 43-44 gamma irradiation, 44-45 gaseous compounds, 45-47 microwave irradiation, 45 moist heat, 42-43 nongaseous compounds, 47-49 principles, 41-42 verification, 41-42 Stichococcus, 431 Strain registry, Frankia, 303-304

Stratified random samples,S Streak plate method, algae and cyanobacteria, 446 Streptoalloteichus, 272 Streptomyces, 269-270, 272, 281, 285

Streptomycetes, isolation, 278 Streptomycin resistance, 578 Streptosporangium, 270,272,277,285 isolation, 289 Strict anaerobes, 225 Stringent plasmid, 637 Substrate-induced respiration (SIR) biomass determination, 753, 760-763 filamentous fungi, 345-347 Sulfate oxidation of elemental S, 1068-1071 reduction, 1071-1076 methods, 1074-1076 principles, 1073-1074 Sulfate-reducing bacteria anaerobic, 224-225 enumeration, 238-239 N2 fixation, 194-195 Sulfur elemental, 1068-1071 marine sediments, 1072-1073 oxidation, 1068-1071 methods, 1070-1071 principles, 1070 Sulfur bacteria, 179 Sulfur-coated urea, 917 Sym plasmid, 638 Syntrophy, 412 Syringe, gas-tight, 845 Systematic sampling, 5-6, 148 TAE buffer, 748 Taq polymerase, 707-708, 710, 717

Taxonomy arthropods, 531-532 bacteria, 543-553 denitrifiers, 264-265 Frankia, 303-305

fungi, 360-363 nematodes, 482-487 protozoa, 509-510 VAM fungi, 360-363 TE solution, 738 TEMED, 624-627 Temperature effect enzyme activity, 782-783 VAM inocula production, 367 Temporal variation, soil properties, 1 Temporary mount, nematodes, 479-480 Test organism, isolation of antibiotic-producing microorganisms, 380-381 Tetracycline resistance, 578 Tetracystis, 430

Tetrazolium dye reduction method metabolically active mycelia, 99-100 viable bacteria, 94-95 Tetrazolium formazan, 545, 549 Thermal conductivity detector, 838 Thermal cycling profile, PCR, 714-716 Thermal shock, 129 Thermoactinomyces, 281 Thermomonospora, 270,272,277,281,285 Thermophiles, actinomycetes, 270, 280-281 Thiobacillus, 179,247,264, 1068-1069 Thiobacillus ferrooxidans, 1079, 1087-1089 Thiosphaera, 247

Thiosulfate, 1068, 1071 30 to 300 rule, 140-141 Ti plasmid, 638 Tile discharge, soil water sampling, 932 Titanium(III) citrate, 229-231 Titrimetric method, CO 2 and O 2 determination, 858-859 TK medium, 1087 Tn 7, 692-693, 700 TOL plasmid, 638 TOL-like plasmid, 413-414 Toluene, 411, 414 effect on microorganisms, 777, 789-790 Total coliforms presumptive test, 151 Total hydrolyzable nitrogen, 898-899 Total nitrogen, 936-940 Transduction, 107 Trap plant, Frankia, 316 Tribonema, 431

2,4,5-Trichlorophenoxyacetic acid, 411 Trimetaphosphatase, 805, 812-813 Triphenyl tetrazolium chloride (TIC), 545, 549 Tris/sucroselEDTA solution, 738 Tris-buffered saline, 612 Trough extractor, soil water sampling, 931932

SUBJECT INDEX Tryptic soy agar, 134-135 TIC. See Triphenyl tetrazolium chloride t-test, 28-31 TWINSPAN, 538 Tylenchida, 484-485 Tylocephalus, 486

Uhlig ice extraction method, protozoa, 508 Ultraviolet fluorescence, DNA in presence of ethidium bromide, 747-750 UMVU estimators. See Uniformly minimum variance unbiased estimators Uncompetitive inhibition, 786 Uniformly minimum variance unbiased (UMVU) estimators, 17,21-22 application, 22-23 computing, 37-38 mean, 21-23 median, 21-23 variance, 21-23 Ureaforms, 917 Urease, 777, 790-801 determination of urea remaining, 798-799 USDA Technology Transfer Documents, 38 Vacutainer, 1061 VAM fungi. See Vesicular-arbuscular mycorrhizal fungi Vancomycin, 136 Vancomycin resistance, 578 Variance, 21 UMVU estimators, 21-23 Vaucheria, 430, 453 Veneer, soil, 432-433 Vesicles, VAM fungi, 353-354 Vesicular-arbuscular mycorrhizal (VAM) fungi, 351-372 active hyphae, 359 aeroponic systems, 369-371 chitin determination, 356, 358 colonized root length estimation, 354-356 effective isolates, 365-366 ergosterol determination, 356 growth response, 361-365 identification, 360-363 infectivity assays, 353 monoxenic cultures, 372 most probable number method, 352-353 mycorrhizal dependency, 361-365 P-response curves, 361-365 quantification colonization in roots, 353-356 external hyphae, 357-359 propagules in soil, 352-357 spore recovery, wet sieving, 360 sterilization of soil, 44-45 taxonomy, 360-363 total hyphae direct methods, 358-359 indirect methods, 357-358 visualization in roots, 353-354

1121 Vesicular-arbuscular mycorrhizal (VAM) inocula, 366-372 application, 371-372 production fertilizers, pesticides, and pot size effects, 367-368 light, moisture, and temperature effects, 367 soil-based pot cultures, 366-368 soil-less media, 368-369 sheared-root, 369-371 storage, 371 Viability, 121 Viable-but-noncuIturable bacteria, 121, 156, 708,728 Vibrio, 179

Virus, 107-115. See also Phage enteric, direct count, 114-115 insect, 109 plant, 109 principles of analysis, 109-110 Volatile compounds isolation of microorganisms producing, 401-402 substrates for microorganisms, 411-415 Washed cell suspension, biodegradation research, 418-419 Water, soil. See Soil water Water agar, 139 Water potential, soil, filter paper method, 53-57 Watered soil suspension, protozoa, 501 Water-holding capacity, soil, 756 Waterlogged incubation, ammonium production during, 960-962 Waterlogged soils, 223 Wet funnel method, arthropod extraction from soil, 526-527 Wet sieving, spores of VAM fungi, 360 Whole-cell hybridization, 102-103 Wilsonema, 486

Winogradsky's salt solution, 126,736-737

Xanthobacter, 188-189

Xanthophyceae, 429, 431 Xenobiotic chemicals, 407-424 X-gal, 695 Xylene, 414

YCz medium, 301 Yeast extract-dextrose-carbonate agar, 383384 Yeast extract-mannitol agar, 203, 384 Yellow-green algae, 429, 431 Yield coefficient, 417 Zoomastigophorea, 492 Zoospore technique, algae, 447 Zygnema, 453