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Soil Sampling and 
Methods of Analysis 

Second Edition 

Edited by 

M.R. Carter 
E.G. Gregorich 

Canadian Society of Soil Science 

Soil Sampling and 
Methods of Analysis 

Second Edition 

In physical science the first essential step in the direction of learning any subject is to find 
principles of numerical reckoning and practicable methods for measuring some quality 
connected with it. I often say that when you can measure what you are speaking about, 
and express it in numbers, you know something about it; but when you cannot measure it, 
when you cannot express it in numbers, your knowledge is of a meagre and unsatisfactory 
kind; it may be the beginning of knowledge, but you have scarcely in your thoughts 
advanced to the state of science, whatever the matter may be. 

Lord Kelvin, Popular Lectures and Addresses (1891-1894), 
vol. 1, Electrical Units of Measurement 

Felix qui potuit rerum cognoscere causas. 
Happy the man who has been able to learn the causes of things. 

Virgil: Georgics (II, 490) 

Soil Sampling and 
Methods of Analysis 

Second Edition 

Edited by 

M.R. Carter 
E.G. Gregorich 

Canadian Society of Soil Science j^e^Zzm 

CRC Press 

Taylor & Francis Group 

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Boca Raton, FL 33487-2742 

roup, LLC 

du P , an Informa business 

No claim to original U.S. Government works 

Printed in the United States of America on acid-free paper 

10 987654321 

International Standard Book Number-13: 978-0-8493-3586-0 (Hardcover) 

This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted 
with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to 
publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of 
all materials or for the consequences of their use. 

No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or 
other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any informa- 
tion storage or retrieval system, without written permission from the publishers. or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 
978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For orga- 

Library of Congress Cataloging-in-Publication Data 

Soil sampling and methods of analysis / edited by M.R. Carter and E.G. Gregorich. 
P- cm- 
Includes bibliographical references and index. 
ISBN-13: 978-0-8493-3586-0 (alk. paper) 
ISBN-10: 0-8493-3586-8 (alk. paper) 
1. Soils-Analysis. 2. Soils-Sampling. I. Carter, Martin R. II. Gregorich, E. G. I 

Visit the Taylor & Francis Web site al 


This volume is an update of the book, Soil Sampling and Methods of Analysis, first published 
in 1993. The aims of this second edition remain the same as those of the earlier edition — to 
provide a compilation of soil analytical and sampling methods that are commonly used, 
straightforward, and relatively easy to use. The materials and procedures for these methods 
are presented with sufficient detail and information, along with key references, to charac- 
terize the potential and limitation of each method. 

As methods develop, so do their degree of sophistication. Taking these developments into 
account, the second edition includes several chapters that serve as "primers," the purpose of 
which is to describe the overall principles and concepts behind a particular type or types of 
measurement, rather than just methods alone. 

All of the chapters retained from the earlier edition have been modified and updated. The 
second edition also introduces new chapters, particularly in the areas of biological and 
physical analyses, and soil sampling and handling. For example, the "Soil Biological 
Analyses" section contains new chapters to reflect the growing number and assortment of 
new microbiological techniques and the burgeoning interest in soil ecology. New chapters 
are offered describing tools that characterize the dynamics and chemistry of soil organic 
matter. A new section devoted to soil water presents up-to-date field- and laboratory-based 
methods that characterize saturated and unsaturated soil hydraulic properties. 

This second edition of Soil Sampling and Methods of Analysis comprises 7 sections and a 
total of 85 chapters and 2 appendices written by 140 authors and co-authors. Each section is 
assembled by two section editors and each chapter reviewed by at least two external 
reviewers. We are grateful to these people for their diligent work in polishing and refining 
the text and helping to bring this new volume to fruition. We particularly thank Elaine Nobbs 
for her support in working with the many authors involved in writing this book. 

We offer this new edition of Soil Sampling arid Methods of Analysis in the belief that it will 
continue as a useful tool for researchers and practitioners working with soil. 

M.R. Carter and E.G. Gregorich 



The Canadian Society of Soil Science is a nongovernmental, nonprofit organization for 
scientists, engineers, technologists, administrators, students, and others interested in soil 
science. Its three main objectives are 

• To promote the wise use of soil for the benefit of society 

• To facilitate the exchange of information and technology among people and 
organizations involved in soil science 

• To promote research and practical application of findings in soil science 

The society produces the international scientific publication, the Canadian Journal of Soil 
Science, and each year hosts an international soil science conference. It sponsored the first 
edition of So I ' ai ipli) ■ ai d W hods of Analysis (Lewis Publishers, CRC Press, 1993) and 
also promoted the publication of the popular reference book Soil and Environmental Science 
Dictionary (CRC Press, 2001). The society publishes a newsletter to share information and 
ideas, and maintains active liaison and partnerships with other soil science societies. 

For more information about the Canadian Society of Soil Science, please visit 


M.R. Carter holds degrees in agriculture and soil science from the University of Alberta and 
obtained a PhD in soil science from the University of Saskatchewan in 1983. Since 1977, he 
has held agricultural research positions with Agriculture and Agri-Food Canada (AAFC) 
and is currently a research scientist at the AAFC Research Center, Charlottetown, Prince 
Edward Island. Dr. Carter is a fellow and past-president of the Canadian Society of Soil 
Science, and past editor of the Canadian Journal of Soil Science. He edited the first edition 
of Soil Sampling and Methods of Analysis, (CRC Press, 1993) and also edited Conservation 
Tillage in Temperate Agroecosystems (CRC Press, 1994) and Structure and Organic Matter 
Storage in Agricultural Soils (CRC Press, 1996). In collaboration with Dr. Gregorich, he 
edited Soil Quality for Crop Production and Ecosystem Health (Elsevier, 1997) and Soil & 
Environmental Science Dictionary (CRC Press, 2001). Dr. Carter presently serves as editor- 
in-chief for the international scientific journal Agriculture Ecosystems & Environment. 

E.G. Gregorich is a research scientist with Agriculture and Agri-Food Canada at the Central 
Experimental Farm in Ottawa, Canada. His work focuses on soil biochemistry, particularly 
carbon and nitrogen cycling in soil. He is a fellow and past-president of the Canadian Society 
of Soil Science, and has served the Soil Science Society of America as chair of the soil 
biology and biochemistry division. Dr. Gregorich has been a member of the International 
Panel on Climate Change, has conducted field studies in Scotland, New Zealand, and 
Antarctica, and directs a Canadian international development project in Vietnam. He has 
served as associate editor for the Journal of Environmental Quality; Agriculture, Ecosystems 
& Environment: European Journal of Soil Science: and the Canadian Journal of Soil 
Science. This is the third book on which he and Dr. Carter have collaborated as editors. 


D. Acosta-Mercado 

Department of Biology 
University of Puerto Rico 
Mayaguez, Puerto Rico 

J.A. Addison 

School of Sustainability and En 

Royal Roads University 

Victoria, British Columbia, Canada 

S.M. Adl 

Department of Biology 
Dalhousie University 
Halifax, Nova Scotia, Canada 

B.C. Ball 

Scottish Agricultural College 
Edinburgh, Scotland, United Kingdom 

M.H. Beare 

New Zealand Institute for Crop and Food 

Christchurch, New Zealand 

E.G. Beauchamp 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

D.W. Anderson 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

Denis A. Angers 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 

V.M. Behan-Pelletier 
Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

N. Belanger 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

H. Antoun 

Department of Soils and Agrifood Engineering 
Laval University 
Quebec, Quebec, Canada 

J.M. Arocena 

College of Science and Management 
University of Northern British Columbia 
Prince George, British Columbia, Canada 

V.L. Bailey 

Biological Sciences Division 

Pacific Northwest National Laboratory 

Richland, Washington, United States 

G.H. Baker 


Commonwealth Scientific and Industrial 

Research Organization 
Glen Osmond, South Australia, Australia 

J.A. Baldock 

Land and Water 

Commonwealth Scientific and Industrial 

Research Organization 
Glen Osmond, South Australia, Australia 

Normand Bertrand 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 

R.P. Beyaert 

Agriculture and Agri-Food Canada 
London, Ontario, Canada 

H. Bolton, Jr. 

Biological Sciences Division 

Pacific Northwest National Laboratory 

Richland, Washington, United States 

Jeff Braidek 

Saskatchewan Agriculture and Food 
Saskatoon, Saskatchewan, Canada 

E. Bremer 

Symbio Ag Consulting 
Lethbridge, Alberta, Canada 

J.A. Brierley 

Agriculture and Agri-Food Canada 
Edmonton, Alberta, Canada 

P.C. Brookes 

Agriculture and Environment Division 

Rothamsted Research 

Harpenden, Hertfordshire, United Kingdom 

M.S. Bullock 
Holly Hybrids 
Sheridan, Wyoming, United States 

B.J. Cade-Menun 

Department of Geological and 

Environmental Sciences 
Stanford University 
Stanford, California, United States 

C.A. Campbell 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

J. Caron 

Department of Soils and Agrifood 

Laval University 
Quebec, Quebec, Canada 

M.R. Carter 

Agriculture and Agri-Food Canada 
Charlottetown, Prince Edward Island 

H.P. Cresswell 

Land and Water 

Commonwealth Scientific and Industrial 

Research Organization 
Canberra, Australian Capital Territory 


J.A. Crumbaugh 

Canadian Forest Service 
Natural Resources Canada 
Edmonton, Alberta, Canada 

J.L.B. Culley 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

M.P. Curran 

British Columbia Ministry of Forests 
Nelson, British Columbia, Canada 

Denis Curtin 

New Zealand Institute for Crop and Food 

Christchurch, New Zealand 

Y. Dalpe 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

Martin H. Chantigny 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 

M.J. Clapperton 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

F.J. Cook 

Land and Water 

Commonwealth Scientific and Industrial 

Research Organization 
Indooroopilly, Queensland, Australia 

Pauline Defossez 

French National Institute for Agricultural 

Laon, France 

J.R. de Freitas 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

C.F. Drury 

Agriculture and Agri-Food Canada 
Harrow, Ontario, Canada 

F. Courchesne 

Department of Geography 
University of Montreal 
Montreal, Quebec, Canada 

K.E. Dunfield 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

M. Duquette 
Montreal, Quebec, Canada 

B.H. Ellert 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

J.A. Elliott 

Environment Canada 

Saskatoon, Saskatchewan, Canada 

D.E. Elrick 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

R.E. Farrell 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

Ty P.A. Ferre 

Department of Hydrology and Water 

University of Arizona 
Tucson, Arizona, United States 

C.T. Figueiredo 

Department of Renewable Resources 
University of Alberta 
Edmonton, Alberta, Canada 

T.A. Forge 

Agriculture and Agri-Food Canada 
Agassiz, British Columbia, Canada 

C.A. Fox 

Department of Renewable Resources 
Agriculture and Agri-Food Canada 
Harrow, Ontario, Canada 

CD. Grant 

School of Earth and Environmental Sciences 

University of Adelaide 

Glen Osmond, South Australia, Australia 

E.G. Gregorich 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

M. Grimmett 

Agriculture and Agri-Food Canada 
Charlottetown, Prince Edward Island 

P.H. Groenevelt 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

Umesh C. Gupta 

Agriculture and Agri-Food Canada 
Charlottetown, Prince Edward Island 

C. Hamel 

Agriculture and Agri-Food Canada 
Swift Current, Saskatchewan, Canada 

X. Hao 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

S.C. Hart 

School of Forestry and Merriam-Powell 

Center for Environmental Research 
Northern Arizona University 
Flagstaff, Arizona, United States 

A. Hartmann 

National Institute of Agronomic Research 
Dijon, France 

J.J. Germitla 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

W.H. Hendershot 

Department of Renewable Resources 

McGill University 

Sainte Anne de Bellevue, Quebec, Canada 

Tee Boon Goh 

Department of Soil Science 
University of Manitoba 
Winnipeg, Manitoba, Canada 

Ganga M. Hettiarachchi 

School of Earth and Environmental Sciences 

University of Adelaide 

Glen Osmond, South Australia, Australia 

D.W. Hopkins 

Scottish Crop Research Institute 
Dundee, Scotland, United Kingdom 

H.H. Janzen 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

R.G. Kachanoski 

Department of Renewable Resources 
University of Alberta 
Edmonton, Alberta, Canada 

Klaus Kaiser 

Soil Sciences 

Martin Luther University 

Halle-Wittenberg, Halle, Germany 

Karsten Kalbitz 

Soil Ecology 
University of Bayreuth 
Bayreuth, Germany 

Y.P. Kalra 

Canadian Forest Service 
Natural Resources Canada 
Edmonton, Alberta, Canada 

A. Karam 

Department of Soils and Agrifood 

Laval University 
Quebec, Quebec, Canada 

Thomas Keller 

Department of Soil Sciences 

Swedish University of Agricultural Sciences 

Uppsala, Sweden 

J. Kimpinski 

Agriculture and Agri-Food Canada 
Charlottetown, Prince Edward Island 

Peter J.A. Kleinman 

Pasture Systems and Watershed 
Management Research Center 
U.S. Department of Agriculture 
University Park, Pennsylvania 
United States 

C.G. Kowalenko 

Agriculture and Agri-Food Canada 
Agassiz, British Columbia, Canada 

D. Kroetsch 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

H. Lalande 

Department of Renewable Resources 

McGill University 

Sainte Anne de Bellevue, Quebec, Canada 

David R. Lapen 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

F.J. Larney 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

R. Lessard 

Environmental Division 
Bodycote Testing Group 
Edmonton, Alberta, Canada 

B.C. Liang 

Environment Canada 
Gatineau, Quebec, Canada 

N.J. Livingston 

Department of Biology 
University of Victoria 
Victoria, British Columbia, Canada 

D.H. Lynn 

Department of Integrative Biology 
University of Guelph 
Guelph, Ontario, Canada 

J.D. MacDonald 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 

D.G. Maynard 

Pacific Forestry Centre 
Natural Resources Canada 
Victoria, British Columbia, Canada 

R.A. McBride 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

W.B. McGill 

College of Science and Management 
University of Northern British Columbia 
Prince George, British Columbia 


D.C. Oik 

U.S. Department of Agriculture 
Agriculture Research Service 
National Soil Tilth Laboratory 
Ames, Iowa, United States 

D. Pare 

Natural Resources Canada 
Canadian Forest Service 
Quebec, Quebec, Canada 

G.R. Mehuys 

Department of Renewable Resources 

McGill University 

Sainte Anne de Bellevue, Quebec, Canada 

A.R. Mermut 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

L.E. Parent 

Department of Soils and Agrifood 

Laval University 
Quebec, Quebec, Canada 

G.W. Parkin 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

J.C. Michel 

INH-INRA-University of Angers 
Angers, France 

G.T. Patterson 

Agriculture and Agri-Food Canada 
Truro, Nova Scotia, Canada 

Jim J. Miller 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

J.O. Moir 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

D.D. Myrold 

Department of Crop and Soil Science 
Oregon State University 
Corvallis, Oregon, United States 

Dan Pennock 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

Caroline Preston 

Pacific Forestry Centre 
Natural Resources Canada 
Victoria, British Columbia, Canada 

D. Prevost 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 

R. Naasz 

Department of Soils and Agrifood 

Laval University 
Quebec, Quebec, Canada 

LP. O'Halloran 

University of Guelph 
Ridgetown, Ontario, Canada 

P. Qian 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

D. Reyes 

Department of Renewable Resources 

McGill University 

Sainte Anne de Bellevue, Quebec, Canada 

W.D. Reynolds 

Agriculture and Agri-Food Canada 
Harrow, Ontario, Canada 

Guy Richard 

French National Institute for Agricultural 

Olivet, France 

Philippe Rochette 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 

L. Rock 

Agriculture and Agri-Food Canada 
Lethbridge, Alberta, Canada 

P.M. Rutherford 

College of Science and Management 
University of Northern British Columbia 
Prince George, British Columbia, Canada 

S. Sauve 

Department of Chemistry 
University of Montreal 
Montreal, Quebec, Canada 

J.J. Schoenau 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

Andrew N. Sharpley 
Crop, Soil and Environmental Sciences 
University of Arkansas 
Fayetteville, Arkansas, United States 

S.C. Sheppard 

ECOMatters Inc. 

W.B. Lewis Business Centre 

Pinawa, Manitoba, Canada 

B.C. Si 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

Myrna J. Simpson 

Department of Physical and Environmental 

University of Toronto 
Toronto, Ontario, Canada 

J.O. Skjemstad 

Land and Water 

Commonwealth Scientific and Industrial 

Research Organization 
Glen Osmond, South Australia, Australia 

J.L. Smith 

U.S. Department of Agriculture 
Agriculture Research Service 
Washington State University 
Pullman, Washington, United States 

Y.K. Soon 

Agriculture and Agri-Food Canada 
Beaverlodge, Alberta, Canada 

P. St-Georges 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

C. Swyngedouw 

Environmental Division 
Bodycote Testing Group 
Calgary, Alberta, Canada 

M. Tenuta 

Department of Soil Science 
University of Manitoba 
Winnipeg, Manitoba, Canada 

Y.-C. Tien 

Agriculture and Agri-Food Canada 
London, Ontario, Canada 

H. Tiessen 

Inter- American Institute for Global 

Change Research 
Sao Jose dos Campos 
Sao Paulo, Brazil 

E. Topp 

Agriculture and Agri-Food Canada 
London, Ontario, Canada 

G. Clarke Topp 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

T. Sen Tran 

Institute of Research and Development 

Quebec, Quebec, Canada 

M.-C. Turmel 

Department of Geography 
University of Montreal 
Montreal, Quebec, Canada 

A.J. VandenBygaart 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

Ken C.J. Van Rees 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

R.P. Voroney 

Department of Land Resource Science 
University of Guelph 
Guelph, Ontario, Canada 

C. Wang 

Agriculture and Agri-Food Canada 
Ottawa, Ontario, Canada 

Jennifer L. Weld 

Department of Crop and Soil Sciences 
The Pennsylvania State University 
University Park, Pennsylvania, United States 

G. Wen 

Lemington, Ontario, Canada 

O.O.B. Wendroth 

Department of Plant and 
Soil Sciences 
University of Kentucky 
Lexington, Kentucky, United States 

J.P. Winter 

Nova Scotia Agricultural College 
Truro, Nova Scotia, Canada 

N. Wypler 

Leibniz-Centre for Agricultural 

Landscape Research 
Institute for Soil Landscape Research 
Muncheberg, Germany 

X.M. Yang 

Agriculture and Agri-Food Canada 
Harrow, Ontario, Canada 

Thomas Yates 

Department of Soil Science 
University of Saskatchewan 
Saskatoon, Saskatchewan, 

N. Ziadi 

Agriculture and Agri-Food Canada 

Quebec, Quebec, Canada 



Section Editors: G.T. Patterson and M.R. Carter 

1. Soil Sampling Designs 

Dun Pen nock, Thomas Yates, and Jeff Braidek 

2. Sampling Forest Soils 

N. Be'langer and Ken C.J. Van Rees 

3. Measuring Change in Soil Organic Carbon Storage 

B.H. Ellert, H.H. Janzen, A.J. VandenBygaart, and E. Bremer 

4. Soil Sample Handling and Storage 
S.C. Sheppard and J. A. Addison 

5. Quality Control in Soil Chemical Analysis 
C. Swyngedouw and R. Lessard 



Section Editors: J.J. Schoenau and LP. O'Halloran 

6. Nitrate and Exchangeable Ammonium Nitrogen 
D.G. Maynard, Y.P. Kalra, and J. A. Crumbaugh 

7. Mehlich 3-Extractable Elements 
\ Ziadi and I V n Trail 

8. Sodium Bicarbonate-Extractable Phosphorus 
J.J. Schoenau and LP. O'Halloran 

9. Boron, Molybdenum, and Selenium 

Ganga M. Hettiarachchi and Umesh C. Gupta 

10. Trace Element Assessment 

W.H. Hendershot, H. Lalande, D. Reyes, and J.D. MacDonald 

1 1 . Readily Soluble Aluminum and Manganese in Acid Soils 
Y.K. Soon, N. Be'langer, and W.H. Hendershot 

12. Lime Requirement 

\ Ziadi and I V >; I ran 

13. Ion Supply Rates Using Ion-Exchange Resins 
P. Qian, J.J. Schoenau, and N. Ziadi 

14. Environmental Soil Phosphorus Indices 

Andrew N. Sharpley, Peter J. A. Kleinman, and Jennifer L. Weld 

15. Electrical Conductivity and Soluble Ions 
Jim J. Miller and Denis Curtin 


Section Editors: Y.K. Soon and W.H. Hendershot 

16. Soil Reaction and Exchangeable Acidity 

U './/. Hcndcrshol, II. Lalande, and M. Duquette 

Collection and Characterization of Soil Solutions 

J .[). MilcDoihild, N. Bc'langer, S. Sauve', F. Courcliesne, 

and W.H. Hendershot 

Ion Exchange and Exchangeable Cations 

W.H. Hendershot, H. Lalande, and M. Duquette 

Nonexchangeable Ammonium 

Y.K. Soon and B.C. Liang 


Tee Boon Goh and A.R. Mermut 

Total and Organic Carbon 

J.O. Skjemstad and J A. Baldock 

Total Nitrogen 

P.M. Rutherford, W.B. McGill, J.M. Arocena, and C.T. Figueiredo 

Chemical Characterization of Soil Sulfur 

C.G. Kowalenko and M. Grimmett 

Total and Organic Phosphorus 

LP. O'Halloran and BJ. Cadc-Menutn 

Characterization of Available P by Sequential Extraction 

H. Tiessen and J.O. Moir 

Extractable Al, Fe, Mn, and Si 

F. Courchesne and M.-C. Turmel 

Determining Nutrient Availability in Forest Soils 

N. Be'langer, D. Pare', and W.H. Hendershot 

Chemical Properties of Organic Soils 

. \. Karuru 


Section Editors: E. Topp and C.A. Fox 

29. Cultural Methods for Soil and Root-Associated Microorganisms 
/./. Germida and J.R. de Freitas 

30. Arbuscular Mycorrhizae 
Y. Dalpe' and C. Hamel 

31. Root Nodule Bacteria and Symbiotic Nitrogen Fixation 
D. Pre'vost and H. Antoun 

32. Microarthropods 

J. P. Winter and V.M. Behan-Pelletier 

33. Nematodes 

T.A. Forge and J. Kimpinski 

34. Earthworms 

M.J. Clapperton, G.H. Baker, and C.A. Fox 

35. Enchytraeids 
S.M. Adl 

36. Protozoa 

S.M. Adl, D. Acosta-Mercado, and D.H. Lynn 

37. Denitrification Techniques for Soils 

C.F. Drury, D.D. Myrold, E.G. Beauchamp, and W.D. Reynolds 

38. Nitrification Techniques for Soils 
C.F. Drury, S.C. Hart, andX.M. Yang 

39. Substrate-Induced Respiration and Selective Inhibition as Measures 
of Microbial Biomass in Soils 

V.L. Bailey, H. Bolton, Jr., and J.L. Smith 

40. Assessment of Soil Biological Activity 
R.P. Beyaert and C.A. Fox 

41. Soil ATP 

R.P. Voroney, G. Wen, and R.P. Beyaert 

42. Lipid-Based Community Analysis 
K.E. Dunfield 

43. Bacterial Community Analyses by Denaturing Gradient Gel 

E. Topp, Y.-C. Tien, and A. Harinianii 

44. Indicators of Soil Food Web Properties 
T. A. Forge and M. Tenuta 


Section Editors: E.G. Gregorich and M.H. Beare 

Carbon Mineralization 

D.W. Hopkins 

Mineralizable Nitrogen 

Denis Curtin and C.A. Campbell 

Physically Uncomplexed Organic Matter 

E.G. Gregorich and M.H. Beare 

Extraction and Characterization of Dissolved Organic Matter 

Martin H. Chantigny, Denis A. Angers, Klaus Kaiser, and Karsten Kc 

Soil Microbial Biomass C, N, P, and S 

R.P. Voroney, P.C. Brookes, and R.P. Beyaert 


Martin H. Chantigny ami Denis A. Angers 

Organic Forms of Nitrogen 

D.C. Oik 

Soil Humus Fractions 

D.W. Anderson and J.J. Schoenau 

Soil Organic Matter Analysis by Solid-State 13 C Nuclear Magnetic 

Resonance Spectroscopy 

Mxnni .1 . Simpson and Caroline Preston 

54. Stable Isotopes in Soil and Environmental Research 
B.H. Ellert and L.Rock 


Section Editors: Denis A. Angers and F.J. Larney 

55. Particle Size Distribution 
D. Kroetsch and C. Wang 

56. Soil Shrinkage 
CD. Grant 

Soil Density and Porosity 

X. Hao, B.C. Ball, J.L.B. Culley, M.R. Carter, and G.W. Parkin 

Soil Consistency: Upper and Lower Plastic Limits 

R.A. McBride 

Compaction and Compressibility 

Pauline Defossez, Thomas Keller, and Guy Richard 

Field Soil Strength 

G. Clarke Topp and David R. Lapen 

Air Permeability 

CD. Grant and P.H. Groenevelt 

Aggregate Stability to Water 

Denis A. Angers. M.S. Bullock, and G.R. Mehuys 

Dry-Aggregate Size Distribution 

F.J. Larney 

Soil Air 

R.E. Farrell and J. A. Elliott 

Soil-Surface Gas Emissions 

Philippe Pochette and Normand Bertrand 

Bulk Density Measurement in Forest Soils 

D.G. Maynard and M.P. Curran 

Physical Properties of Organic Soils and Growing Media: Particle Size 

and Degree of Decomposition 

L.E. Parent and J. Caron 

Physical Properties of Organic Soils and Growing Media: Water and Air 

Storage and Flow Dynamics 

/. Caron, D.E. Elrick, J.C. Michel, and R. Naasz 


Section Editors: W.D. Reynolds and G. Clarke Topp 

69. Soil Water Analyses: Principles and Parameters 
W.D. Reynolds and G. Clarke Topp 

70. Soil Water Content 

G. Clarke Topp, G.W. Parkin, and Ty PA. Ferre 
1 1 . Soil Water Potential 

N.J. Livingston and G. Clarke Topp 

72. Soil Water Desorption and Imbibition: Tension and Pressure 

W.D. Reynolds and G. Clarke Topp 

73. Soil Water Desorption and Imbibition: Long Column 
W.D. Reynolds and G. Clarke Topp 

74. Soil Water Desorption and Imbibition: Psychrometry 
W.D. Reynolds and G. Clarke Topp 

75. Saturated Hydraulic Properties: Laboratory Methods 
W.D. Reynolds 

76. Saturated Hydraulic Properties: Well Permeameter 
W.D. Reynolds 

Saturated Hydraulic Properties: Ring Infiltrometer 

W.D. Reynolds 

Saturated Hydraulic Properties: Auger Hole 

G. Clarke Topp 

Saturated Hydraulic Properties: Piezometer 

G. Clarke Topp 

Unsaturated Hydraulic Conductivity: Laboratory Tension Infiltrometer 

F.J. Cook 

Unsaturated Hydraulic Properties: Laboratory Evaporation 

O.O. B. Wendroth and N. Wypler 

82. Unsaturated Hydraulic Properties: Field Tension Infiltrometer 
W.D. Reynolds 

83. Unsaturated Hydraulic Properties: Instantaneous Profile 
W.D. Reynolds 

Estimation of Soil Hydraulic Properties 
F.J. Cook and H.P. Cresswell 
Analysis of Soil Variability 
B.C. Si. R.G. Kachanoski, and W.D. Reynolds 

Site Description 

G.T. Patterson and J. A. Brierley 

General Safe Laboratory Operation Procedures 

P. St-Georges 


Section Editors: G.T. Patterson and M.R. Carter 

Chapter 1 
Soil Sampling Designs 

Dan Pennock and Thomas Yates 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

Jeff Braidek 

Saskatchewan Agriculture and Food 

Saskatoon, Saskatchewan Canada 


Sampling involves the selection from the total population of a subset of individuals upon 
which measurements will be made; the measurements made on this subset (or sample) will 
then be used to estimate the properties (or parameters) of the total population. Sampling is 
inherent to any field research program in soil science because the measurement of the total 
population is impossible for any realistic study. For example, even a single 10 ha field 
contains about 100,000 1 m 2 soil pits or 1 x 10 7 10 cm 2 cores, and sampling of the entire 
population would be more of an unnatural obsession than a scientific objective. 

Sampling design involves the selection of the most efficient method for choosing 
the samples that will be used to estimate the properties of the population. The definition 
of the population to be sampled is central to the initial formulation of the research study 
(Eberhardt and Thomas 1991; Pennock 2004). The sampling design defines how specific 
elements will be selected from the population, and these sampled elements form the 
sample population. 

There are many highly detailed guides to specific sampling designs and the statistical 
approaches appropriate for each design. The goal of this chapter is to present the issues 
that should be considered when selecting an appropriate sampling design. In the final section, 
specific design issues associated with particular research designs are covered. Suggested 
readings are given in each section for more in-depth study on each topic. 


1.2.1 Haphazard, Judgment, and Probability Sampling 

Sample locations can be chosen using (a) haphazard sampling, (b) judgment sampling, or 
(c) probability sampling. Haphazard, accessibility, or convenience sampling involves a series 
of nonreproducible, idiosyncratic decisions by the sampler and no systematic attempt is 
made to ensure that samples taken are representative of the population being sampled. This 
type of sampling is antithetical to scientific sampling designs. Judgment sampling (also 
termed purposive sampling [e.g., de Gruijter 2002]) involves the selection of sampling points 
based on knowledge held by the researcher. Judgment sampling can result in accurate 
estimates of population parameters such as means and totals but cannot provide a measure 
of the accuracy of these estimates (Gilbert 1987). Moreover the reliability of the estimate is 
only as good as the judgment of the researcher. Probability sampling selects sampling points 
at random locations using a range of specific sample layouts, and the probability of sample 
point selection can be calculated for each design. This allows an estimate to be made of the 
accuracy of the parameter estimates, unlike judgment sampling. This allows a range of 
statistical analyses based on the estimates of variability about the mean to be used, and is by 
far the most common type of sampling in soil science. 

1.2.2 Research Designs Using Judgment Sampling 

Pedogenetic and soil geomorphic studies focus on determining the processes that formed the 
soil properties or landscapes under study and the environments that controlled the rates of 
these processes. Pedon-scale studies are closely associated with the development of soil 
taxonomic systems, and focus on vertical, intrapedon processes. Soil geomorphic studies are 
the interface between quaternary geology and soil science, and soil geomorphologists focus 
on lateral transfer processes and the historical landscape evolution. 

Both types of studies involve the identification of soil and/or sediment exposures that are 
highly resolved records of the sequence of processes that have formed the soil landscape. 
The researcher locates these exposures by using his judgment as to the landscape positions 
where optimum preservation of the soil-sediment columns is most likely. The development 
of the chronological sequence can be done with a detailed analysis of a single exposure; no 
replication of exposures is required. 

Surveys are designed to define the extent of spatial units. Soil surveyors map the distribution 
of soil taxonomic units and provide descriptive summaries of the main properties of the soils. 
In soil survey the association between soil classes and landscape units is established in the 
field by judicious selection of sampling points (termed the free survey approach). This type 
of judgment sampling can be an extremely efficient way of completing the inventory. 
Contaminant surveys are most typically undertaken by private-sector environmental con- 
sultants, and the specific objective may range from an initial evaluation of the extent of 
contamination to the final stage of remediation of the problem. Laslett (1997) states that 
consultants who undertake these surveys almost always employ judgment sampling and 
place their samples where their experience and prior knowledge of site history suggest the 
contamination might be located. In many jurisdictions the sampling design may also be 
constrained by the appropriate regulatory framework. 

1.2.3 Research Designs Using Probability Sampling 

Inventory studies share the common goal of measuring the amount of a property or 
properties under study and the uncertainty surrounding our estimate of the amount. For 
example, in agronomic sampling we may wish to estimate the amount of plant-available 
nutrients in a given field; in contaminant sampling the goal may be to estimate the 
amount of a contaminant present at a site. In comparative mensurative experiments, 
comparisons are drawn among classes that the researcher defines but cannot control — 
for example, sampling points grouped by different soil textures, landform positions, soil 
taxonomic classes, and drainage class. Their location cannot be randomized by the 
researcher, unlike imposed treatments such as tillage type or fertilizer rates where 
randomization is essential. In manipulative experiments the treatments can be directly 
imposed by the researcher — ideally as fixed amounts that are applied precisely. Many 
studies are hybrid mensurative-manipulative designs — for example, the measurement of 
yield response to different fertilizer rates (imposed treatment) in different landform 
positions (characteristic or inherent treatment). The role of sampling in inventory, men- 
surative, and manipulative designs is very similar — to allow statistical estimation of the 
distribution of the parent population or populations. In inventory studies the statistical 
estimates may be the end point of the study. 

Pattern studies are undertaken to assess and explain the spatial or temporal pattern of proper- 
ties. Two main types of pattern studies exist: (a) the quantification of the spatial and temporal 
variability in properties and (b) hypothesis generation and testing using point patterns. In 
pattern studies the initial goal may be a visual assessment of the pattern of observations in time 
or space, and statistical estimation of the populations may be a secondary goal. 

Geostatistical and other spatial statistical studies are undertaken to model the spatial pattern 
of soil properties, to use these models in the interpolation of values at unsampled locations, 
to assess the suitability of different spatial process models, or to assist in the design of 
efficient sampling programs. 


1.3.1 Measures of Central Tendency and Dispersion 

The key characteristics of the distribution of attributes are measures of its central tendency 
and the dispersion of values around the measure of central tendency. In the initial stage of 
study formulation the researcher defines the population, which is composed of the sampling 
units and one or more attributes measured on these sampling units. Each attribute has a 
distribution of values associated with it, which can be characterized by parameters such as 
the population mean (/jl) and variance (a 2 ). A sample of the sampling units is drawn from the 
population, and statistics such as the sample mean (x) and variance (s 2 ) are calculated, which 
serve as estimates of the population parameters. Calculations of these statistics are readily 
available and will not be repeated here. The number of samples taken is denoted as n. For 
sample populations that are more or less normally distributed the arithmetic mean (x) is an 
appropriate measure of central tendency. The variance (s 2 ) is a common measure of the 
deviation of individual values from the mean and its square root; the standard deviation (s) 

reports values in the same units as the mean. The coefficient of variation (CV) is 
normalized measure of the amount of dispersion around the mean, and is calculated by 

Sample populations in the soil science commonly show a long tail of values to the right of the 
distribution (i.e., they are right-skewed). In this case a log normal or other right-skewed 
distribution should be used. 

The mathematical properties of the normal distribution are well understood and the prob- 
ability that the true population mean lies within a certain distance of the sample mean can be 
readily calculated. For sample populations the estimated standard error of the sample mean is 

s(x) = s/V7i (1.2) 

For a sample population that has a large sample size or where the standard error is known and 
that approximates a normal distribution, the true mean will be within +1.96 standard errors 
of the sample mean 95 times out of 100 (i.e., where the probability P = 0.05). The range 
defined these limits are the 95% confidence interval for the mean and these limits are the 
95% confidence limits. The value 1.96 is derived from the t distribution, and values of t can 
be derived for any confidence limit. For sample populations based on a small sample size or 
where the standard error is not known the value of 1 .96 must be replaced by a larger r-value 
with the appropriate degrees of freedom. A probability of exceeding a given standard error 
(a) may be selected for any sample distribution that approximates the normal distribution 
and the appropriate confidence limits calculated for that distribution. 

1.3.2 Independence, Randomization, and Replication 

The goal of sampling is to produce a sample that is representative of the target population. If 
the choice of samples is not probability based then a strong likelihood exists that the sample 
will not be representative of the population. For example, selection of sampling locations 
convenient to a farmyard (instead of distributed throughout the field) may lead to overesti- 
mates of soil nutrients due to overapplication of farmyard manure near the source of the 
manure through time. The use of probability-based sampling designs (i.e., the designs 
discussed in Section 1.4) confers a design-specific independence on the sample selection 
process, which satisfies the need for independence of samples required by classical statistical 
analysis (a theme developed in great detail by Brus and de Gruijter 1997). 

Replication is an important consideration in mensurative and manipulative experiments. In a 
manipulative study, replication is the repeated imposition of a set of treatments (e.g., 
fertilizer or pesticide rates). In a pattern or mensurative study, replication is the repeated, 
unbiased selection and sampling of population elements in a particular class — for example, 
the selection of multiple 5 x 5 m slope elements in a field that have markedly convex 
downslope curvatures. Replication provides an estimate of the experimental error, and 
increasing replication improves precision by reducing the standard error of treatment or 
class means (Steel and Torrie 1980). Correct identification and sampling of replicates is 
critical for estimating the parameters of the class the sample is drawn from and is required for 
statistically correct procedures. Pseudoreplication (Hurlbert 1984) occurs when a researcher 
assumes a very general effect from a limited sampling and often occurs because the target 
population has not been clearly defined at the outset of the research. 

Randomization is a consideration in manipulative designs. Steel and Torrie (1980, p. 135) 
summarizes the need for randomization: 

"... it is necessary to have some way of ensuring that a particular treat- 
ment will not be consistently favored or handicapped in successive repli- 
cations by some extraneous sources of variation, known or unknown. In 
other words, every treatment should have an equal chance of being 
assigned to any experimental unit, be it unfavorable or favorable." 

Randomization is implemented by ensuring the random placement of treatment plots within a 
field design; the repeated imposition of the same sequence of treatments in a block of treat- 
ments may cause an erroneous estimate of the experimental error. The random order of treatment 
placement is achieved using random number tables or computer-generated randomizations. 


Although many types of sampling designs exist (reviewed in Gilbert 1987; Mulla and 
McBratney 2000; de Gruijter 2002) only two main types (random and systematic) are 
commonly used in the soil and earth sciences. Inventory studies can be completed using 
any of the designs discussed in the following two sections. Pattern and geostatistical studies 
typically use transect or grid designs, as is discussed in more detail in Section 1.5. 

1.4.1 Simple Random and Stratified Random Sampling 

In simple random sampling all samples of the specified size are equally likely to be the one 
chosen for sampling. In stratified random sampling, points are assigned to predefined groups 
or strata and a simple random sample chosen from each stratum. The probability of being 
selected can be weighted proportionally to the stratum size or the fraction of points sampled 
can vary from class to class in disproportionate sampling. Disproportionate sampling would 
be used if the degree of variability is believed to vary greatly between classes, in which case 
a higher number of samples should be drawn from the highly variable classes to ensure the 
same degree of accuracy in the statistical estimates. 

Stratified sampling (correctly applied) is likely to give a better result than simple random 
sampling, but four main requirements should be met before it is chosen (Williams 1984): 

1 Population must be stratified in advance of the sampling. 

2 Classes must be exhaustive and mutually exclusive (i.e., all elements of the population 
must fall into exactly one class). 

j Classes must differ in the attribute or property under study; otherwise there is no gain 
in precision over simple random sampling. 

4 Selection of items to represent each class (i.e., the sample drawn from each class) 
must be random. 

The selection of random points in a study area has been greatly facilitated by the widespread 
use of Global Positioning System (GPS) receivers in field research. The points to be sampled 
can be randomly selected before going to the field, downloaded into the GPS unit, and then 
the researcher can use the GPS to guide them to that location in the field. 

TABLE 1 .1 Sample Sizes Required for Estimating the True Mean fx. Using a Prespecified 

Relative Error and the Coefficient of Variation 
Confidence level Relative error, d r Coefficient of variation (CV), % 

0.80 0.10 2 7 27 42 165 370 

0.25 6 7 27 60 

0.50 2 7 15 

1.0 2 4 

0.90 0.10 2 12 45 70 271 609 

0.25 9 12 45 92 

0.50 2 13 26 

1.0 2 8 

0.95 0.10 4 17 63 97 385 865 

0.25 12 17 62 139 

0.50 4 16 35 

1.0 9 16 

Source: Adapted from Gilbert, R.O., in Statistical Methods for Environmental Pollution 

Monitoring, Van Nostrand, Reinhold, New York, 1987, 320 pp. 

Determination of Sample Numbers in Inventory Studies 

A necessary and important step in the planning stages of a project is to determine the number 
of samples required to achieve some prespecified accuracy for the estimated mean. One 
approach is to use prior knowledge about the CV of the property under study to estimate 
sample numbers required to achieve a certain prespecified relative error. The relative error 
(d T ) is defined as 

d T = | sample mean — population mean | /population mean (1.3) 

The sample numbers required to achieve a specified relative error at a selected confidence 
level can be estimated from Table 1.1. For example, at a confidence level of 0.95 and a 
relative error of 0.25, 16 samples are required if the CV is 50% and 139 samples are required 
if the CV is 150%. Estimates of CV for different soil properties are widely available, and are 
summarized in Table 1.2. 

1 .4.2 Systematic Sampling 

The most commonly used sampling design for many field studies is systematic sampling using 
either transects or grids. Systematic sampling designs are often criticized by statisticians 
but the ease with which they can be used and the efficiency with which they gather information 
makes them popular in the field of earth sciences. Ideally the initial point of the transect or grid 
and/or its orientation should be randomly selected. The major caution in the use of systematic 
sampling with a constant spacing is that the objects to be sampled must not be arranged in 
an orderly manner which might correspond to the spacing along the transect or the grid. 

The choice of a transect or a grid depends on several factors. Certain types of research 
designs require particular types of systematic designs — as discussed below, wavelet analysis 
requires long transects whereas geostatistical designs more typically use grid designs. Grids 
are often used for spatial pattern studies because of the ease with which pattern maps can be 
derived from the grids. The complexity of landforms at the site is also a consideration. 

TABLE 1.2 Variability of Soil Properties 

Coefficient of variation 



Very high 

Low (CV <15%) 


(CV 35%-75%) 

(CV 75%-150%) 

Soil hue and value 3 

Sand content 3 

Solum thickness 3 

Nitrous oxide flux b 

pH a 

Clay content 3 

Ca, Mg, K 3 

Electrical conductivity 13 

A horizon 

CEC a 

Soil nitrate N b 

Saturated hydraulic 
conductivity' 3 

Thickness 3 

% BS" 

Soil-available P b 

Solute dispersion 
coefficient 15 

Silt content 3 

CaC0 3 equivalent 3 

Soil-available K b 

Porosity b 

Crop yield b 

Bulk density b 

Soil organic C b 

Adapted from Wilding, L.P. and Drees, L.R., in L.P. Wilding, N.E. Smeck, and G.F. Hall, (Eds.), 
Pedogenesis and Soil Taxonomy. I. Concepts and Interactions, Elsevier Science Publishing, 
New York, 1983, 83-116. 

Adapted from Mulla, D.J. and McBratney, A.B., in M.E. Sumner (Ed.), Handbook of Soil 
Science, CRC Press, Boca Raton, Florida, 2000, A321-A3S2. 

For level and near-level landscapes either a transect or a grid can be used (Figure 1.1). The 
appropriateness of transects in sloping terrain depends in part on the plan (across-slope) 
curvature. Where no significant across-slope curvature exists each point in the landscape 
receives flow from only those points immediately upslope and a single transect can 
adequately capture the variations with slope position (Figure 1.2). A single transect will 
not, however, be sufficient if significant plan curvature exists. In this case a zigzag design or 
multiple, randomly oriented transects could be used, but more typically a grid design is used 
(Figure 1.3). It is important to ensure that all slope elements are represented in the grid 

FIGURE 1.1. Example of a grid sampling layout composed of four parallel transects on a near- 
level surface form. Soil samples would be taken at each point labeled with a 
diamond shape. 

FIGURE 1.2. Example of a transect sampling layout on a sloping surface with no significant 
across-slope (plan) curvature. Soil samples would be taken at each point labeled 
with a diamond shape. 

FIGURE 1.3. Example of a grid sampling layout composed of six parallel transects on a sloping 
surface form with pronounced across-slope curvature. The arrow-oriented down- 
slope delineates the minimum downslope length of the long axis of the grid, and the 
arrow across the slope indicates the minimum length of the short axis of the grid. 
Soil samples would be taken at each point labeled with a diamond shape. 

design. A rule of thumb is that the grid should extend from the level summit of the slope to 
the toeslope along the long axis of the slope and along at least one complete convergent- 
divergent sequence across the slope. 

The distance between sampling points in either a transect or a grid should be smaller than the 
distance required to represent the variability in the field. For example, if the study area 
contains landforms whose tops and bottoms are equally spaced at 30 m, then a transect 
crossing these landforms should have sample locations spaced much shorter than this (e.g., 
5 or 10 m). It is desirable to base sample spacing on prior knowledge of the area. 


1.5.1 Sampling Designs for Mensurative and Manipulative Experiments 

In mensurative and manipulative designs a typical goal is to assess if the attributes sampled 
from different classes have different distributions or the same distribution, using difference 
testing. In the simplest type of hypothesis testing, two hypotheses are constructed: the null 
hypothesis (Ho) of no difference between the two groups and the alternative hypothesis of a 
significant difference occurring. The researcher chooses an a level to control the probability 
of rejecting the null hypothesis when it is actually true (i.e., of finding a difference between 
the two groups when none, in fact, existed in nature or a Type I error). Peterman (1990) states 
that the consequences of committing a Type II error (i.e., of failing to reject the null 
hypothesis when it is, in fact, false) may be graver than a Type I error, especially in 
environmental sampling. The probability of failing to reject the Hq when it is, in fact, false 
is designated as /3 and the power of a test equals (1-/3). Calculation of power should be done 
during the design stage of a mensurative or manipulative experiment to ensure that sufficient 
samples are taken for a strong test of differences between the groups. 

The use of nonstratified, systematic designs may be very inefficient for mensurative experi- 
ments. For example, in a landscape where 60% of the site is classified as one class of 
landform element and 5% is classified into a second class, a 100-point grid should yield 
approximately 60 points in the major element and 5 points in the second. The dominant 
element is probably greatly oversampled and the minor element undersampled. Appropriate 
sample numbers can be efficiently gathered by stratified sampling by a priori placement of 
points into the relevant groups or strata, and then a random selection of points is chosen 
within each stratum until the desired number is reached. 

In manipulative designs the treatments are commonly applied in small strips (or plots). If the 
experimental unit is believed to be homogenous then the treatments can be randomly 
assigned to plots in a completely random design. More typically some degree of heterogen- 
eity is believed to occur — for example, a slight slope or a gradient in soil texture exists across 
the plot. In this case the treatments are assigned to square or rectangular blocks. Each block 
typically contains one of each of the treatments being compared in the experiment, and the 
sequence of treatments in each block is randomly determined. This is termed as a random- 
ized complete block design (RCBD), and is the most commonly used manipulative design. 
Many other types of manipulative designs have been developed for field experimentation 
(Steel and Torrie 1980) and the advice of a biometrician is invaluable for the design of these 
types of experiments. 


1.5.2 Soil Sampling for Nutrient Inventories 

These are a particular type of inventory study that are undertaken to provide average values 
of soil nutrient properties over a field or field segment (more commonly called soil testing). 
This average value is then often used as the basis for fertilizer recommendations in the next 
growing season. The accuracy with which soil test results reflect the true condition of soils in 
the field is more dependent on the way in which the sample is collected and handled rather 
than on error associated with the laboratory analysis (Cline 1944; Franzen and Cihacek 
1998). As such, the sample used for laboratory analysis must be representative of the field 
from which it was taken and sample collection and sample handling must not cause a change 
to the soil properties of interest before the laboratory analysis. 

The development of a sampling procedure must address the following points. 

Division of the Field into Different Sampling Units 

The farm operator must decide what level of detail is relevant to his or her field operations. 
Are there parts of the field that have different fertility patterns? Are these areas large enough 
to be relevant? Does the operator want to engage in site-specific management? Has the 
operator has the ability to vary fertilizer application rates to accommodate the field subsec- 
tions identified? 

Subsections of a field would commonly be identified by differences in topography (termed 
landscape-directed soil sampling), parent material, management history, or yield history. It 
may be impossible to subdivide a field into smaller units if the farm operator has no prior 
knowledge of the field, or if there is no obvious topographic or parent material differences. 
Under these conditions a grid sampling design has the potential to provide the greatest 
amount of spatial detail. However, a grid is also the most expensive sampling method and is 
not typically economically feasible for routine soil testing. 

Where landscape-directed soil sampling can be implemented it has been shown to provide 
superior information on nutrient distribution and the identification of separate management 
units than that obtained via grid sampling. Landscape-directed soil sampling is particularly 
effective at assessing patterns of mobile soil nutrients. 

Selection of Sampling Design and Sample Numbers 

For each field or field subsection samples can be taken using a random sampling design, a 
grid sampling design, or a benchmark sampling design. 

In random sampling individual samples are collected from locations that are randomly 
distributed across the representative portion of the field. These random locations can be 
generated with a GPS. A zigzag sampling pattern (Figure 1.4) is often used for field 
sampling. The sampler should avoid sampling atypical areas such as eroded knolls, 
depressions, saline areas, fence lines, old roadways and yards, water channels, manure 
piles, and field edges. Typically, all samples are combined and a composite sample is 
taken and submitted for laboratory analysis. Composite sampling is comparatively 
inexpensive since only one sample from each field or subsection of a field is sent for 
laboratory analysis. However, this design provides no assessment of field variability, and 
relies on the ability of the farm operator to identify portions of the field that may have 
inherently different nutrient levels. 

FIGURE 1.4. Example of a zigzag sampling layout on a near-level surface. Soil samples would be 
taken at each point labeled with a diamond shape. 

Soil-testing laboratory guidelines consistently suggest that on average 20 samples be col- 
lected for each field or subsection of a field regardless of the actual area involved. 

Grid Sampling 

In this sampling design a grid system is imposed over each field or subsection of a field. One 
composite sample from each grid node is sent for laboratory analysis. The grid sampling design 
is the most expensive method employed in soil sampling but it can provide highly detailed 
information about the distribution of nutrient variability if the grid size is small enough. 

Benchmark Sampling 

In this design a single representative site (benchmark) is selected for each field or subsection 
of a field. The benchmark site should be approximately 1/4 acre or 30x30 m. Twenty or 
more samples should be randomly taken from within the benchmark and then composited. 
The farm operator can return to the same benchmark site in subsequent years for repeated 
testing. The advantage of this design is that year to year changes in nutrient status are more 
accurately reflected. 

1.5.3 Sample Timing, Depth of Sampling, and Sample Handling 

As a general rule, sampling for mobile nutrients should be taken as close to seeding as possible 
or when biological activity is low. Fall sampling should generally start after the soil tempera- 
ture is less than 10°C at which time no further changes in the soil nutrient levels are expected. 
Spring sampling, before seeding, can be done as soon as the soil frost is gone. 

Commonly used sample depth combinations are to 15 cm (0"-6") and 15 to 60 cm (6"-24"), 
or to 30 cm (0"-12") plus 30 to 60 cm (12"-24"). However, if the soil nutrient of interest 
is expected to be stratified by depth, as with water-soluble highly mobile nutrients, then 
additional sampling increments would help ensure accurate recommendations. If organic 
matter and/or pH measurements are of importance (particularly when evaluating potential 
herbicide residue carryover) then a to 15 cm (0"-6") sample should be taken. 


To ensure that a uniform volume of soil is taken through the full depth of each sampling 
increment samples should be collected using soil probes and augers designed for this purpose. 
A wedge-shaped sample like that collected using a spade will not give consistent results. All 
probes should be kept clean and rust free. Avoid contamination at all stages of sample 

In many situations, a lubricant will need to be applied to the soil probe to prevent the soil 
sticking inside the probe. This lubricant will help to prevent compaction of the soil as the probe 
is pressed into the ground, and it will facilitate emptying the collected sample from the probe. 
Research by Blaylock et al. (1995) suggests that the commonly used lubricants will not affect 
soil test results other than the case of the micronutrients iron, zinc, manganese, and copper. The 
most commonly used lubricants include WD-40 lubricant, PAM cooking oil, and Dove dish- 
washing liquid. 

1.5.4 Sampling for Geostatistical, Spectral, and Wavelet Analysis 

The choice of geostatistical techniques over the approaches discussed above involves a 
fundamental decision about whether the sampling is design based or model based; potential 
users of the geostatistical approach are referred to Brus and de Gruijter (1997) (and the 
discussion papers following their article) and de Gruijter (2002) for a comprehensive 
discussion of the difference between the two approaches. 

Geostatistics, spectral analysis, and wavelet analysis all address the spatial dependence in 
soil properties between locations. Thus the location of each sample point in space using 
GPS-determined spatial coordinates is critical information. Sample programs where this type 
of analysis is intended should include a topographic survey and generation of digital 
elevation model. 

Sampling for Geostatistics 

Spatial variability in soil properties can be separated into random and nonrandom compon- 
ents (Wilding and Drees 1983). The nonrandom variability is due to the gradual change of a 
soil property over distance. Knowledge of this nonrandom variation gained through the 
application of geostatistics can be useful in the design of efficient sampling programs and the 
estimation of the value of a soil property at unsampled locations. There are comprehensive 
discussions of geostatistics in Webster and Oliver (1990), Mulla and McBratney (2000), and 
Yates and Warrick (2002). 

Geostatistics assume that the value of a soil property at any given location is a function of the 
value of that same property at locations nearby (spatial dependence). The distance and 
direction between locations determine the degree of spatial dependence between values of 
a soil property at those locations. The use of geostatistics thus requires that not only the value 
of a soil property be known, but the location as well. The primary geostatistical tools are the 
semivariogram and kriging. The semivariogram provides a measure of spatial dependency, 
the range, which can be used to determine optimum sample spacing or the extent of soil 
unit boundaries. Kriging is used to estimate the value of a soil property at a location where 
the value is unknown by using the known values at locations about the point of interest. 
Spatial dependence between two different soil properties can be explored using cross- 
semivariograms and cokriging techniques. 

A common sample design to determine optimal sample spacing and soil boundary definitions 
is the linear transect. Calculations are simplest if equal spacing is maintained between 


sample points; however, unequal spacing can be accommodated with more complicated 
mathematics. If the study area has recognizable topographic features then the transect should 
be directed perpendicular to the trend of these features. 

Kriging techniques require that sample locations are taken on a grid. Sample locations are 
typically chosen by random selection from a set of predetermined grid intersections. In this 
case distances between locations are not equal. Efficient grid design and kriging may be based 
on a semivariogram constructed from preliminary sampling along a transect in the same area. 

Geostatistics require the assumption of stationarity. Stationarity assumes that all values of a soil 
property within an area are drawn from the same distribution. This assumption is not always 
valid. As well, variation in a soil property may occur at more than one scale. For scale analysis 
and nonstationarity more advanced statistical techniques must be used. 

Sampling for Spectral Analysis 

In landscapes where landforms are repetitive such as a hummocky, rolling, or undulating 
terrains the continuous variation of soil properties may result in a data series with a repetitive 
cycle of highs and lows. The periodicity may be examined in the frequency domain using 
techniques referred collectively as spectral analysis (see McBratney et al. 2002 for a recent 
discussion of these techniques). The total variance of a data series is partitioned by fre- 
quency. The soil property is considered to cycle at a particular period if a significant portion 
of the variance is associated with the frequency represented by that period. Period is 
comparable to scale or distance much like the range from a semivariogram. Unlike a 
semivariogram, more than one scale can be identified. A cross spectrum can identify soil 
properties that cycle together and the coherency spectrum can identify scales at which two 
properties may be positively or negatively correlated in the same area. 

The linear transect is the most common sample design used to amass a data series for spectral 
analysis. Sample spacing must be consistent. As for geostatistical methods the number of 
samples, the spacing, and the direction of the transect should be chosen to best represent the 
landscape features of the site. 

Sampling for Wavelet Analysis 

Both geostatistics and spectral analysis require the assumption of stationarity. Nonstationar- 
ity can occur, for example, due to changes in land use or geomorphology across the site, 
resulting in more than one population of values. A method of analysis that does not require 
the assumption of stationarity is wavelet analysis (see McBratney et al. 2002; Si 2003 for 
recent summaries of developments in this technique). A wavelet is a mathematical function 
that yields a local wavelet variance for each point in a data series. Like spectral analysis, 
wavelets portion the total variance of a data series according to frequency (scale), but unlike 
spectral analysis the total variance is also portioned according to space (location). A wavelet 
approach allows the ability to discern between multiple processes occurring in the field, the 
scale at which the processes are operating, and the location or distribution of these processes 
along the data series. 

Like spectral analysis, wavelet analysis requires a data series collected from locations 
spaced equally along a linear transect. Wavelets are rescaled by powers of two and 
thus transects that contain a power of two data points (64, 128, 256, ...) are best for 
computational speed (Si 2003). As a result, large transects are common when using 

wavelet analysis. In cases where the number of transect locations is not a power of two, the 
data series can be padded with zero values to the nearest power of two. Transects of 128 
points are large enough for detailed scale analysis, yet may be manageable by most research 

Blaylock, A.D., Bjornest, L.R., and Lauer, J.G. 
1995. Soil probe lubrication and effects on soil 
chemical-composition. Commun. Soil Sci. Plant 
Anal. 26: 1687-1695. 

Brus, D.J. and de Gruijter, J.J. 1997. Random 
sampling or geostatistical modelling? Choosing 
between design-based and model-based sampling 
strategies for soil (with discussion). Geoderma 

Methods. Soil Science Society of America, Inc., 
Madison, WI, pp. 159-200. 

Mulla, D.J. and McBratney, A.B. 2000. Soil spa- 
tial variability. In M.E. Sumner, Ed. Handbook of 
Soil Science. CRC Press, Boca Raton, FL, 
pp. A321-A352. 

Pennock, D.J. 2004. Designing field studies in 
soil science. Can. J. Soil Sci. 84: 1-10. 

Cline, M.G. 1944. Principles of soil sampling. 
Soil Sci. 58: 275-288. 

de Gruijter, J.J. 2002. Sampling. In J.H. Dane and 
GC. Topp, Eds. Methods of Soil Analysis, Part 
4 — Physical Methods. Soil Science Society of 
America, Inc., Madison, WI, pp. 45-79. 

Eberhardt, L.L. and Thomas, J.M. 1991. Designing 
environmental field studies. Ecol. Monogr. 6: 53-73. 

Franzen, D.W. and Cihacek, L.J. 1998. Soil 
Sampling as a Basis for Fertilizer Application. 
North Dakota University Extension Service 
Publication SF990. Available at: http://www.ext. 
(July 2006). 

Gilbert, R.O. 1987. Statistical Methods for Envir- 
onmental Pollution Monitoring. Van Nostrand 
Reinhold, New York, NY, 320 pp. 

Hurlbert, S.H. 1984. Pseudoreplication and the 
design of ecological field experiments. Ecol. 
Monogr. 54: 187-211. 

Laslett, G.M. 1997. Discussion of the paper by 
D.J. Brus and J.J. de Gruijter. Random sampl- 
ing or geostatistical modeling? Geoderma 80: 

McBratney, A.B., Anderson, A.N., Lark, R.M., 
and Odeh, I.O. 2002. Newer application tech- 
niques. In J.H. Dane and GC. Topp, Eds 
Methods of Soil Analysis, Part 4— Physical 

Peterman, R.M. 1990. Statistical power analysis 
can improve fisheries research and management. 
Can. J. Fish. Aquat. Sci. 47: 2-15. 

Si, B. 2003. Scale and location dependent soil 
hydraulic properties in a hummocky landscape: 
a wavelet approach. In Y. Pachepsky, 
D. Radcliffe, and H.M. Selim, Eds. Scaling 
Methods in Soil Physics. CRC Press, Boca 
Raton, FL, pp. 169-187. 

Steel, R.G.D. and Torrie, J.H. 1980. Principles 
and Procedures of Statistics. A Biometrical 
Approach. McGraw-Hill, New York, NY, 633 pp. 

Webster, R 
Methods ii 
Oxford Uni 

and Oliver, M.A. 1990. Statistical 
Soil and Land Resource Survey. 
ersity Press, Oxford, 316 pp. 

Wilding, L.P. and Drees, L.R. 1983. Spatial vari- 
ability and pedology. In L.P. Wilding, N.E. 
Smeck, and G.F. Hall, Eds. Pedogenesis and 
Soil Taxonomy. I. Concepts and Interactions. 
Elsevier Science Publishing Company, New York, 
NY, pp. 83-116. 

Williams, R.B.G. 1984. Introduction to Statistics 
for Geographers and Earth Scientists. Macmillan 
Publishers Ltd., London, 349 pp. 

Yates, S.R. and Warrick, A.W. 2002. Geostatistics. 
In J.H. Dane and G.C. Topp, Eds. Methods of 
Soil Analysis, Part 4 — Physical Methods. Soil 
Science Society of America, Inc., Madison, WI, 
pp. 81-118. 


Chapter 2 
Sampling Forest Soils 

N. Belanger and Ken C.J. Van Rees 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 


The causes for forest soil variability are many. Spatial variability is a function of bedrock 
type and parent material, climate, tree species composition and understory vegetation, 
disturbances (e.g., harvesting, fire, windthrow), and forest management activities (e.g., site 
preparation, thinning, pruning, fertilization, vegetation management). For example, a second 
generation 50-year-old Radiata pine plantation grown on plowed alluvial sands in Australia 
would have lower spatial variability compared to mixed hardwoods developed from a 
shallow rocky till of the Precambrian (Canadian) Shield after harvest. The mixed hardwoods 
would likely show high variability in forest floor properties such as forest floor thickness due 
to tree fall (Beatty and Stone 1986; Clinton and Baker 2000) and the influence of different 
tree species (Finzi et al. 1998; Dijkstra and Smits 2002). Moreover, the fact that the soil is 
plowed in the pine plantation would likely reduce some of the soil variability that could have 
been created by the previous plantation (e.g., changes in soil properties when sampling away 
from the stem). In the mineral soil, it would be more difficult to assess nutrient pools 
compared to the pine plantation because of the problem of measuring bulk density and 
percentage of coarse fragments in the rocky till (Kulmatiski et al. 2003). It would also be 
more problematic to develop a replicated sampling scheme by depth in the natural forest 
because horizon thickness across the landscape evolves as a continuum with complex spatial 
patterns (e.g., Ae pockets along old root channels and thick FH material in pits). 

All these sources of spatial variability must be considered in efforts to systematically sample 
and describe forest soil properties. This is why sampling strategies and methodologies must 
be selected with care and this chapter is dedicated to that goal; however, information 
regarding field designs and plot establishment can be found in Pennock (2004) or Pennock 
et al. (see Chapter 1). 


Developing a sampling scheme that represents the inherent variability and true value of the 
population mean in forest floor chemistry may require many sampling points. Calculating the 
sample size is important because a sample size that is too large leads to a loss of time, human 


resources, and money, whereas a sample size that is too small leads to erroneous statistical 
testing. The margin of error (d) is the maximum difference between the observed sample 
mean and the true population mean. It can be calculated according to the following equation 
(Snedecor and Cochran 1980): 

where t a is the Student t factor for a given level of confidence (generally 95%) and s is the 
coefficient of variation (CV) as a percentage of the mean value. The equation can be 
rearranged to solve the sample size needed to produce results to a specified p and margin 
of error: 

In a field study designed to test the spatial variability of nutrient concentrations and pools in 
the forest floor, Arp and Krause (1984) sampled the forest floor at 98 locations in a 900 m 2 
plot. They showed that concentrations and pools of KC1 extractable NO3-N and NH4-N and 
extractable P on field-moist soils had the highest CV values and required as many as 1371 
samples (i.e., KC1 extractable NO3-N pool) to decrease the margin of error on the population 
mean to 10% at a confidence level of 95% and t a = 1 .96 (a = 0.05). An accurate estimate of 
the mean content of a nutrient required more samples than that for measuring its mean 
concentration. This was due mostly to the large variation in forest floor weight and thickness 
in the study. Figure 2.1 shows margins of error obtained using CV values in Arp and Krause 
(1984) with 10, 15, and 20 sampling points and confidence level set at 95%. This simple 
exercise demonstrates that a margin of error of 5% is generally not possible using 10 
sampling points, except for total C concentration and soil pH. For nutrient concentrations 
(except for NO3-N, NH4-N, and P on field-moist soils) and physical properties (i.e., 
moisture, thickness, and weight), a margin of error between 31% and 9.9%, 26% and 
8.0%, and 22% and 7.0% is possible with 10, 15, and 20 sampling points, respectively, 
with forest floor weight having the highest margin of error and total N having the lowest. 
However, 20 sampling points are required to obtain a margin of error between 19% and 29% 
when these concentrations are transformed as pools. Similarly, McFee and Stone (1965) 
found that it was necessary to have 50 sampling points to have a 10% margin of error 
(confidence level of 95%) on the calculated mean of forest floor weight and thickness for 
forest plots in the Adirondacks. This supports the idea that the problem of assessing forest 
floor nutrient pools with a high level of confidence comes in large part from the high 
variability in forest floor weight and thickness. Results also show that it is not financially 
and logistically feasible to develop replicated field design testing treatment effects on 
concentrations and pools of KC1 extractable NO3-N and NH4-N as well as water-extractable 
P pools on field-moist samples. 

The number of sampling points required for a reliable representation of a plot's mean does 
not appear to be related to its size. Quesnel and Lavkulich (1980) and Carter and Lowe 
(1986) had smaller study plots (300 and 400 m 2 , respectively) than Arp and Krause (1984), 
but the intensities of sampling required for obtaining a reasonable estimate of the plot's mean 
were similar. Interestingly, Carter and Lowe (1986) conducted the study with LF and H 
horizons as distinct samples and found that the LF horizons required fewer samples (3 to 10) 
than the H horizons (3 to 38 samples) for a reliable estimate of the population mean for total 
C, N, P, and S concentrations and pH (margin of error of 10% at a confidence level of 95%). 


110 points □ 15 points □ 20 points 

_. 10 ° " 

t 80 - 

■s 60 " 
§ 40 - 

20 - 




o^ oC> j,?" ^ 

FIGURE 2.1. Margins of error of the population mean (forest floor (a) weight, moisture, pH and 
extractable nutrient, total C (Ct), and total N (Nt) concentrations as well as 
(b) extractable nutrient, Ct and Nt pools) obtained using coefficients of variation 
in Arp and Krause (1984) with 10, 15, and 20 sampling points with the level of 
confidence set at 0.95. 

The results also suggested that 15 sampling points should be enough to characterize the 
population mean of total Mg, K, N, P, C, Cu, and Zn concentrations, lipid concentrations, pH 
and bulk density in LF, and H material within a margin of error of 20% at a confidence level 
of 95%. However, a more intensive sampling strategy was required for obtaining similar 
margins of error on the population mean of total Ca and Mn concentrations in the H material 
(81 and 47 samples, respectively) and total Al and Fe concentrations in LF material (41 and 
50 samples, respectively). 

In the mineral soil, the intensity of sampling required to obtain a reliable estimate of the 
population mean also appears to depend on the variable tested. Studying the variability of 
organic matter in the forest floor and mineral soil in a Tuscany forest, Van Wesemael and 
Veer (1992) sampled six 2500 m 2 plots and found that between 17 and 80 sampling points 
were required to have a 10% margin of error on the plots' population means (confidence 
level of 95%) of organic matter content in the first 5 cm of mineral soil compared to 33 to 
235 sampling points for organic matter content in LF or FH horizons. This appears to fit with 

the values of Arp and Krause (1984) who found that 114 samples were required to arrive 
at the same level of confidence for total C content in the forest floor. An accurate measure of 
the mean for soil pH, particle size, and moisture appears to be considerably easier: Ike and 
Clutter (1968) demonstrated that 1 to 12 sampling points in forest plots of the Georgia Blue 
Ridge Mountains were necessary to obtain a 10% margin of error on the population mean of 
pH, separate sand, silt and clay fractions, and available water and moisture. However, 
available P and exchangeable K concentrations required 15 to 32 samples per plot for the 
same margin of error, 14 to 76 samples per plot for exchangeable Mg concentration, and 153 
to 507 for exchangeable Ca concentration. 


There are two generally accepted techniques for sampling the forest floor: soil cores or a 
square template. McFee and Stone (1965) used a sharp-edged steel cylinder with a diameter 
of 8.7 cm (59 cm 2 ) for coring the forest floor to quantify the distribution and variability of 
organic matter and nutrients in a New York podzol. Similarly, Grier and McColl (1971) used 
a steel cylinder with a diameter of 26.6 cm (556 cm 2 ). As an alternative to soil corers, Arp 
and Krause (1984) used a square wooden sampling template of 25 x 25 cm (625 cm 2 ) placed 
on the surface of the forest floor as a cutting guide. Others have used smaller or larger cutting 
templates (225 to 900 cm 2 ) and Klinka et al. (1981) suggested using a 10 x 10 cm template. 
A corrugated knife used on the inside edge of the frame will generally cut through the forest 
floor material with no difficulty and once the sample is cut on all sides, it is relatively simple 
to partition it from the mineral soil. Square sampling templates can also be constructed with 
heavier gauge metal and sharp edges can be added to the bottom of the frame in order to push 
or hammer (use hard plastic hammers or mallets) the frame into the forest floor until the 
mineral soil is reached. The litter can then be pulled from the frame. In some cases, a wooden 
cap can be built for the metal frames to assist in hammering into the forest floor. We believe 
this a convenient way of sampling the forest floor as it allows at the same time, after the 
measurement of thickness and determination of wet and dry mass, a measure of bulk density 
and water content. 

The general rule of thumb for sampling the forest floor is that the larger the surface area 
being sampled, the greater chances you have of reducing microsite variability in the sample 
once it is air-dried, cleaned for roots and other woody material, and mixed in the laboratory. 
Therefore, it is recommended to use a sampling scheme that will cover, individually or 
bulked, at least 200 cm 2 . 


Sampling of forest floor horizons varies among soil scientists and there are no accepted 
standards for how horizons should be sampled. Generally, LFH horizons are sampled as 
a whole (Bock and Van Rees 2002) or samples are taken from individual (i.e., L or F or H 
horizon) or combinations of horizons (i.e., FH horizon) (Olsson et al. 1996; Hamel et al. 2004), 
depending on the objective of the study. Normally, all layers are collected together (LFH) or 
the litter is collected individually (L + FH) for nutrient cycling studies or individually if one 
is investigating specific processes such as decomposition (e.g., Cade-Menun et al. 2000). 

Sampling problems can occur when trying to distinguish between H horizons and Ah horizon 
sequences. In forest soils with an abrupt transition between the forest floor and the mineral 

soil such as those classified as mor forest floors, it is relatively simple to distinguish the 
forest floor from the mineral soil. However, in forest soils with Mull and sometimes Moder 
forest floors (i.e., Chernozems and Melanic Brunisols), the F or H horizons are often not 
easily discernible from the mineral Ah horizon, thus making it more difficult to sample the 
forest floor layers separately. The incorporation of organic matter in the mineral soil 
therefore introduces a bias in forest floor sampling as some of the Ah material can be 
incorporated in the forest floor samples. The Expert Committee on Soil Survey (1987) 
defines the Ah horizon as "A horizon enriched in organic matter, it has a color value one 
unit lower than the underlying horizon or 0.5% more organic C than the IC or both. It 
contains less than 17% organic C by weight." If correct sampling of the forest floor is an 
important issue for the study, then the most appropriate way to distinguish between the FH 
and Ah horizons is to carry out a presampling campaign and then conduct C analyses on the 
samples. Running a quick and fairly reliable loss-on-ignition (LOI) test should be very 
informative and allow separation between forest floor and mineral soil material: organic C 
constitutes 58.3% of the soil organic matter content and thus, LOI should not exceed 30% 
on Ah samples, whereas an LOI of 30% or more is expected from forest floor material 
depending on the amounts of mineral soil particles, coarse fragments, and charcoal 
incorporated in the material. If the cost for accessing the study site is high and there is 
no possibility for presampling and returning to the site after LOI testing, then a second 
option for separating FH horizons from Ah material is to rely on color and feel. Humus 
forms do vary and their taxonomy can be quite complex. In this respect, the reader is 
directed to Klinka et al. (1981) and/or Green et al. (1993) for an in-depth description of 
these horizons. 


Soil bulk density is a commonly measured parameter in forest soil studies to assess harvest- 
ing effects on forest soil quality such as compaction induced by logging or site preparation 
practices (e.g., Powers 1991; Aust et al. 1995). For forests growing on glacial till of the 
Precambrian Shield or other rocky soils, however, the presence of large rocks and coarse 
fragments makes it difficult to measure soil bulk density with standard techniques. In 
addition, quantifying the amount of coarse fragments is important for accurately calculating 
nutrient pools in soils (Palmer et al. 2002; Kulmatiski et al. 2003). There are a variety of 
forest soil sampling techniques to assess coarse fragments and bulk density ranging from the 
clod, core, pit, to the sand cone technique (i.e., Page-Dumroese et al. 1999; Kulmatiski et al. 
2003). The intensive approach is to excavate a sample that is larger than the largest rock in 
the sample (see Chapter 66 of this book for a detailed description of the excavation and sand 
replacement method) while the extensive approach is to collect smaller sized samples over a 
large area using a corer. 

Page-Dumroese et al. (1999) conducted a study where two different size cores (183 and 
2356 cm 3 ) were compared to two pit excavation methods and one nuclear source mois- 
ture gauge for calculating bulk density. They found that bulk densities measured with the 
two excavation methods were 6% to 12% lower than those measured with the two core 
measurements and the nuclear gauge method. The nuclear gauge method gave the highest 
values of total and fine bulk densities and the small corer method produced the most variable 
results. Sampling with a corer produces higher values compared to the pit methods because 
compaction may occur during sampling. This was more apparent at the greater depth 
increments, probably because some compaction likely occurred during core insertion (Lichter 
and Costello 1994). To prevent this, it was suggested to remove the top mineral soil with an 


auger or shovel and then hammering the corer to the desired soil depth. On the other hand, 
Page-Dumroese et al. (1999) also argued that the smaller corer may have provided samples 
too small to be representative of overall soil conditions: it is possible that the small core 
technique underestimates total bulk density because it does not account for large rocks with 
high densities. The larger size corer generally produced intermediate bulk density values, 
although estimates were low at the greater depths sampled because of incomplete filling or 
soil loss at the bottom of the core sampler. The accuracy of this method is likely increased for 
greater soil depths as rock fragments usually augment with depth. 

Similarly, Kulmatiski et al. (2003) compared the ability of the core and excavation methods 
for detecting a 10% change in total C and N pools in forest soils of southern New England. 
They found that mean total C and N contents measured from the extensive core techniques 
were 7% higher than those measured from the intensive pit approach, but these differences 
were not statistically significant. The core techniques produced lower estimates of percent- 
age C and N and bulk densities compared to the pit technique, but the core techniques also 
produced lower estimates of coarse fragments and higher soil volume values. Consequently, 
both techniques produced very similar estimates of total N and C soil pools. The 7% 
divergence between mean total C pools measured using the two techniques was reduced 
when coarse roots were added in the calculations, whereas coarse roots were not a significant 
portion of the total N pools and had no impact on estimates. The results also showed little 
variability of total C and N pools at a depth greater than 15 cm (assessed by the pit 
technique), meaning that deeper nutrient pools are insensitive to environmental factors 
such as tree species composition and topography. Moreover, Kulmatiski et al. (2003) 
suggested that the extensive core approach required less than one-half of the sampling 
time for determining the population mean (i.e., N and C pools) compared to the intensive 
pit approach and that a smaller number of samples was required for a low margin of error of 
the population mean. They recommended the use of the core techniques to calculate total N 
and C contents in the upper mineral soil horizons. However, one advantage of the pit 
technique is that it allows direct measurement of large rock fragments in the soil. For 
calculating total C and N pools in deeper soils with generally greater rock fragments, 
Kulmatiski et al. (2003) therefore recommended to extrapolate data from the upper mineral 
horizons to deeper soil by building regression models developed from a few local soil pits. 


Obtaining a reliable estimate of the population mean of a specific nutrient concentration in 
the mineral soil probably requires less sampling points than that in the forest floor (e.g., 
organic matter content in Van Wesemael and Veer (1992)). The number of sampling points 
is also probably less if the soil is sampled by diagnostic horizon compared to sampling by 
depth. More variability in soil properties is expected from sampling by depth because the 
sample is a mixture of soil material with different properties. For example, sampling Bhf 
horizons of sandy Ferro-Humic Podzols means that the soil material has at least 5% organic 
C and 0.4% pyrophosphate-extractable Fe and Al. However, if the mineral soil is sampled by 
depth, e.g., 20 cm increments, then Ae material (higher in Si and lower in Al, Fe, and C than 
the Bhf, see Table 2.1) is bound to be incorporated with Bhf material in the first increment 
and Bhf and Bf/BC material will be bulked in the second increment. In a study on jack pine 
growth, Hamilton and Krause (1985) showed a negative relationship between the depth of 
the eluvial material and tree growth. In podzols, roots develop most of their biomass in the 
forest floor and upper B horizons and not in the Ae material (e.g., Cote et al. 1998). Sampling 
by 20 cm increments in well-drained forest soils with a fully developed Ae horizon means 

TABLE 2.1 Total Elemental Composition (Given as Percentage of Total Soil Matrix) of Ae and Bf 
Horizons of Podzols Developed under Balsam Fir in the Gaspe Peninsula of Quebec 
(Mean + Standard Deviation with n = 6) 

Ae horizon Podzolic B horizon 

Si0 2 84.5±4.18 53.3±7.56 

Ti0 2 1.17 + 0.16 0.68±0.18 

Al 2 3 4.98±1.08 11.2 + 1.99 

Fe 2 3 0.62 ±0.1 5 7.06 ±1.79 

MgO 0.24±0.07 0.90±0.35 

CaO 0.08±0.02 0.12±0.05 

Na 2 0.69±0.09 0.83 ±0.1 8 

K 2 0.92±0.24 1.34 + 0.33 

P 2 O s 0.0S±0.01 0.24±0.08 

LOI a 6.59±3.05 24.5±7.52 

a LOI is loss-on-ignition. Total elemental composition does not sum up to 100% as trace 
elements are not shown here. 

Note: Total iron present has been recalculated as Fe 2 3 . In cases where most of the iron was 
originally in the ferrous state, a higher total is the result. 

that the arbitrary differences in soil morphology will govern the results of the chemical 
analyses. In this respect, significant correlation between tree nutrition/growth and mineral 
soil chemistry may be masked by the fact that the sampling scheme used is not representative 
of the capacity factor of the actual mineral soil to provide nutrients to the trees. Also, an 
admixture of soil material with different properties may camouflage the response of specific 
soil horizons to harvesting, acid deposition, etc., as some of the material incorporated in the 
sample may be in steady-state with the conditions created by the disturbance whereas 
the other material may not. 

Note that there are also clear advantages of sampling soil by depth when conducting studies on 
soil changes over time. One of the best conceptual examples for demonstrating the benefits of 
sampling by depth is a study comparing soil C pools in a natural forest with a plantation 
established close by. The plantation is building a new forest floor (as it was plowed) and is 
likely shallower than that of the natural forest. Also, the natural sequence of horizons in the 
plantation is obviously different from that of the natural forest to a depth of about 5-8 cm. 
Therefore, as the sequencing of diagnostic horizons differs between the plantation and natural 
forest, sampling by depth is the best option for comparing soil C pools. Due to the horizontal 
variability, it is strongly recommended to sample the soil evenly across the whole sampling 
increment: sampling only a part of the full increment will indisputably result in artifacts. 
Examples of studies on long-term changes in forest soil properties that required this sampling 
strategy can be found in Eriksson and Rosen (1994), Parfitt et al. (1997), and Belanger et al. 
(2004). Moreover, the reader will find a thorough discussion on sampling strategies to study 
temporal changes in soil C for agricultural soils in Ellert et al. (see Chapter 3). 


In some forests, soil variability can be enhanced by forest processes such as tree falls to 
create "pit and mound" topography. These kinds of sites need different types of sampling 
strategies to account for changes in microtopography. In a study on "pits and mounds" in 
New York state hardwoods, Beatty and Stone (1986) made a composite sample from four 
4.5 cm or five 2 cm diameter cores (total surface area 64 and 16 cm 2 , respectively) at 0.5 or 
1 m intervals across the microsites. Although these samples have a small surface area, the 


sampling procedure is quite accepted considering that the study is conducted at 
scale and that more or larger samples were likely not needed over such a small area to 
calculate a valid population mean. Similarly, forest soil scientists are bulking forest floor 
samples for studies conducted at the plot scale, i.e., a set of samples coming from the same 
population (plot) are carefully mixed together so that they are equal in terms of weight or 
volume. Obviously, this is a tedious task to do in the field and unfortunately, it is often 
unclear whether proper mixing is done. Preferably, samples should be stored separately and 
bulking should be done in the laboratory after they have been air-dried and sieved. 

A disadvantage of bulking the samples in a plot is that it does not allow for the calculation of 
the standard deviation or CV values. In an effort to assess the precision of the variables 
measured by bulking forest floor samples, Carter and Lowe (1986) compared the mean 
nutrient contents weighted by depth and bulk density using the 15 sampling points within a 
plot to the values obtained from analyzing a single sample obtained by bulking these 15 
samples (as a function of depth and bulk density). Values from composite samples were all 
within one standard deviation of the mean, except for total P and Cu concentrations in LF 
material. Moreover, they investigated the relationships between the weighted means and the 
composite sample values across the six study plots and found that they were quite strong for 
most variables, suggesting that bulking samples can provide good estimates of the real 
population mean (r > 0.90, except for Ca and Al concentrations in LF, and Mn and C in 
LF and H horizons). Similarly, Bruckner et al. (2000) investigated the impact of bulking soil 
samples on microarthropod abundance on a Norway spruce plantation in Austria. It was 
assumed that the grinding action of soil particles during mixing would injure or kill part of 
the population and thus underestimate the population relative to a mean weighted from 
samples of the population analyzed individually. However, using a special mixing procedure 
of the extracts, Bruckner et al. (2000) came to the conclusion that no microarthropod was lost 
or damaged because a large number of samples were bulked in a systematic manner and 
mixed in equal amounts. 

Arp, P.A. and Krause, H.H. 1984. The forest 
floor: lateral variability as revealed by systematic 
sampling. Canada. Can. J. Soil Sci. 64: 423-437. 

Aust, W.M., Tippett, M.D., Burger, J.A., and 
McKee, W.H. Jr. 1995. Compaction and rutting 
during harvesting aiicci better drained soils more 
than poorly drained soils on wet pine flats. South. 
J. Appl. Forest 19: 72-77. 

Beatty, S.W. and Stone, E.L. 1986. The variety of 
soil microsites created by tree falls. Can. J. Forest 
Res. 16: 539-548. 

A case study with Norway spruce. Can. J. Forest 
Res. 34: 560-572. 

Bock, M.D. and Van Rees, K.C.J. 2002. Forest 
harvesting impacts on soil properties and vegetation 
communities in the Northwest Territories. Can. 
J. Forest Res. 32: 713-724. 

Bruckner, A., Barth, G., and Scheibengraf, M. 
2000. Composite sampling enhances the 
confidence of soil microarthropod abundance 
and species richness estimates. Pedobiologia 

44: 63-74. 

Belanger, N., Pare, D. 
Daoust, G. 2004. Is the usi 
superior growth a threat to soil 

Bouchard, M., and Cade-Menun, B.J., Berch, S.M., Preston, CM., 

se of trees showing and Lavkulich, L.M. 2000. Phosphorus forms 

'ailability? and related soil chemistry of Podzolic soils on 


northern Vancouver Island. I. A comparison of 
two forest types. Can. J. Forest Res. 30: 

Carter, R.E. and Lowe, L.E. 1986. Lateral vari- 
ability of forest floor properties under second- 
growth Douglas-fir stands and the usefulness of 
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Clinton, B.D. and Baker, C.R. 2000. Cata- 
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A.G., Bradley, R., Biron, P.M., and Courchesne, F. 
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Dijkstra, F.A. and Smits, M.M. 2002. Tree species 
effects on calcium cycling: the role of calcium 
uptake in deep soils. Ecosystems 5: 385-398. 

Eriksson, H.M. and Rosen, K. 1994. Nutrient 
distribution in a Swedish tree species experiment. 
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Hamilton, W.N. and Krause, H.H. 1985. Relation- 
ship between jack pine growth and site variables 
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1981. Ta.wiioiiiic classification of humus forms in 
ecosystems of British Columbia. First approxima- 
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forms. Forest Science Monograph 29. Society of 
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level. Environ. Pollut. 116: S209-S219. 

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Chapter 3 

Measuring Change in Soil 

Organic Carbon Storage 

5.H. Ellert and H.H. Janzen 

Agric allure and Agri-rood Canada 
Lethbridge, Alberta, Canada 

A.J. VandenBygaart 

Agric ullure and Agri-rood Canada 
Ottawa, Ontario, Canada 

E. Bremer 

Symbio Ag Consulting 
Lethbridge, Alberta, Canada 


Organic carbon (C) must be among the most commonly analyzed soil constituents, starting with 
the earliest soil investigations. Already in the nineteenth century, chemists were routinely 
analyzing soil C (e.g., Lawes and Gilbert 1885). Initially, these analyses were done to investi- 
gate pedogenesis and to assess soil productivity, both of which are closely linked to organic C 
(Gregorich et al. 1997). But more recently, scientists have been analyzing soil organic C (SOC) 
for another reason: to measure the net exchange of C between soil and atmosphere (Janzen 
2005). Indeed, building reserves of SOC has been proposed as a way of slowing the rising 
atmospheric CO2 concentrations caused by burning fossil fuel (Lai 2004a,b). 

Measuring SOC to quantify soil C "sinks" requires more stringent sampling and analyses 
than measuring SOC to evaluate productivity. Where once it was sufficient to measure 
relative differences in concentration over time or among treatments, now we need to know 
the change in amount of C stored in Mg C per ha. Reviews of SOC measurement typically 
focus on the chemical methods of determining the SOC concentrations after samples have 
been brought to the laboratory. Here we emphasize soil sampling procedures and calculation 
approaches to estimate temporal changes in SOC stocks. Uncertainties along the entire chain 
of procedures, from designing the soil sampling plan, to sampling in the field, to processing 
and storing the samples, through to chemical analysis and calculating soil C stocks need to be 
considered (Theocharopoulos et al. 2004). 


SOC is dynamic: newly photosynthesized C is added regularly in the form of plant litter, 
and existing SOC is gradually decomposed back to CO2 by soil biota. Management or 
environmental conditions that change the relative rates of inputs and decomposition will 
effect a change in the amount of SOC stored. Rates of change in SOC (typically less than 
0.5 Mg C ha~' year ) are quite small, however, compared to the large amounts of SOC 
often present (as high as 100 Mg C ha~', or more, in the top 30 to 60 cm soil layer). Thus 
changes in SOC can only be reliably measured over a period of years or even decades (Post 
et al. 2001). Since the distribution of SOC in space is inherently variable, temporal changes 
(e.g., attributable to management practices, environmental shifts, successional change) must be 
distinguished from spatial ones (e.g., attributable to landform, long-term geomorphic processes, 
nonuniform management). 

Temporal changes in SOC can be defined in two ways (Figure 3.1): as an absolute change in 
stored C (SOC at / = x minus SOC at t = 0), or as a net change in storage among treatments 
(SOC in treatment A minus SOC in treatment B, after x years). The former provides 
an estimate of the actual C exchange between soil and atmosphere; the latter provides an 
estimate of the C exchange between soil and atmosphere, attributable to treatment A, relative 
to a control (treatment B). Both expressions of temporal change may be available from 
manipulative experiments with appropriate samples collected at establishment (assesses 
spatial variability) and at various intervals (say 5 to 10 years) thereafter. 

This chapter provides selected methods for measuring the change in C storage, either 
absolute or net, typically for periods of 5 years or more. To be effective, the method 
needs to: measure organic (not total) C, provide estimates of C stock change (expressed in 
units of C mass per unit area of land to a specified soil depth and mass), be representative 
of the land area or management treatment under investigation, and provide an indication of 
confidence in the measurements. These methods are applicable, with minor modification, 
to a range of scales and settings, including benchmarks sites and replicated research 

Net change in treatment A"' 
Absolute change in treatment A 

Time (years) 

n of hypothetical changes in soil organic C in two tr 
For treatment A, the absolute change is the difference in SOC at time = x, compared to 
that at time = 0. The net change is the difference between SOC in treatment A and that 
in treatment B, at time = x, assuming that SOC was the same in both treatments at 
time = 0. The latter approach is often used to measure the effect on SOC of a proposed 
treatment (e.g., no-till) compared to a standard "control" (e.g., conventional tillage). 



Determining the optimum number and spatial arrangement of sampling points to estimate 
SOC storage remains as much an art as a science. Nevertheless, careful study of the site, 
along with clearly articulated objectives can improve the cost-effectiveness and precision of 
the estimates (VandenBygaart 2006). 

3.2.1 Materials 

7 Descriptions of soil properties, landscape characteristics, and agronomic history 
at the study site, from sources such as: soil maps and reports, aerial photos, 
scientific publications, cropping records, and yield maps. 

3.2.2 Procedure 

Two general approaches can be used in sampling a study area (e.g., a plot, field, watershed): 

a Nonstratified sampling, where the entire study area is considered to be one unit, and 
sampled in a systematic or random manner. 

b Stratified sampling, where the study area is first subdivided into relatively homo- 
geneous units, based on factors such as topography (e.g., slope position), and each 
unit is sampled separately. 

3.2.3 Nonstratified Sampling 

7 Obtain an estimate of the likely sample variance and required accuracy for SOC 
at the study site, based on previously compiled information. 

2 Using as much information as available, calculate the number of samples required 
using Equation 3.1. The required number of samples will increase as variability 
and the required accuracy increase (Figure 3.2) (Garten and Wullschleger 1999; 
Wilding et al. 2001 ). Required accuracy is expressed as in the same units used for 
the sample mean, and often is less than 10% of that value because even small 
changes in the mean imply appreciable pedosphere-atmosphere C exchange over 
large tracts of land. 

3 Select an appropriate grid or linear sampling pattern, suited to the study site and 
sampling equipment. 

3.2.4 Stratified Sampling 

7 Subdivide the study site into areas likely to have similar SOC stocks, based on 
factors such as topography or management history. 

2 Select the number of sampling sites within each subarea, using Equation 3.1, or 
Figure 3.2 as a guide, or by fixed allotment. In the latter case, for example, one 
or several sampling sites may be designated for each of three slope positions within 
a large research plot. 


. -•- a 2 =4, cv 

= 5% 

-'- \ -D- a 2 = 17, c 

v= 10% 

o \ -*- a 2 =34, c 

v = 1 5% 

-- \\ ^^ a 2 =67, c 

v = 20% 

\1 -■- a 2 = 100, 

cv = 25% 


i . TTrf; 1 , .7** T8 

Number of soil samples 

FIGURE 3.2. Decrease in the minimum detectable difference (MDD) between mean soil Cat two 
sampling times for contrasting levels of variance as the number of samples collected 
at each time doubles (4, 8, 1 6, . . .). The MDD was calculated for a = 0.05 signifi- 
cance and (1— p) = 0.90 statistical power (i.e. probability of rejecting the null 
hypothesis when it really is false and should be rejected). The lines correspond to 
increasing variance (a 2 ) selected for a hypothetical soil layer containing a mean of 
40 Mg C ha -1 with the coefficient of variation (cv) increasing from 5% to 25%. 
(Adapted from Garten, C.T. and Wullschleger, S.D., J. Environ. Quai, 28, 1359, 
1999. With permission.) 

3.2.5 Calculations 

(d x mean) 

where « req is the required number of samples, t is the Student's f-value, at the desired 
confidence level (typically 1 — a = 0.90 or 0.95), s 1 is the sample variance, d is the required 
accuracy or maximum acceptable deviation from the mean (e.g. <i = 0.10), and mean is the 
arithmetic sample mean. 


Sampling patterns and intensiti 

other factors, notably economi 

achieve the desired sensitivity 

is somewhat arbitrarily reduced. As well, sampl: 

plots, such as long-term experiments, where 

es will vary widely, depending on site characteristics and on 

considerations. Often, the number of samples required to 

exceedingly expensive, and the number of sampling points 

intensity may have to be reduced in small 

al may disturb the site to the 

extent that future research is jeopardized. But such compromises, if carried too far, may reduce 
the chance of measuring any differences with reasonable reliability. Studies with insufficient 
sampling points typically lack statistical power to assess treatment effects. Consequently, the 
' 'cost' ' of erroneous conclusions drawn from such data (when the data really are inconclusive) 
may greatly exceed the "savings" provided by reduced sample numbers. 


Precisely measuring temporal changes in SOC first depends on identifying or minimizing 
spatial changes. Spatial changes can be minimized by pairing sampling locations in space 
(Ellert et al. 2001, 2002; VandenBygaart 2006). This approach allows for effective meas- 
urement of SOC changes in time at comparatively few sampling points, but measured 
C stock change values at these points are not necessarily representative of the entire 
study site. Conant and Paustian (2002) and Conant et al. (2003) have evaluated similar 
sampling strategies. 


The following procedure is intended for the extraction of soil cores, from agricultural plots or 
landscapes, for subsequent organic C analysis. It is provided as an illustration, recognizing 
that individual studies may require modification to satisfy specific objectives and local 

3.3.1 Materials 

7 Truck-mounted hydraulic soil coring device. 

2 Soil coring tube, with slots 1 cm wide by 30 cm long, and a cutting bit with inside 
diameter of about 7 cm. The bit usually has slightly smaller diameter (by 1 to 4 mm) 
than the tube; this difference should be small enough to avoid soil mixing, but large 
enough to prevent sticking. In dry, coarse-textured soils with weak consolidation 
this difference should be reduced so there is enough friction to hold the core when 
the tube is pulled from the soil. The diameter of the coring bit should be measured 
accurately and recorded for future calculations of soil core density. 

3 Piston to push the soil core out of tube. A simple piston can be constructed by 
attaching a rubber stopper to the end of a wooden dowel. 

4 Knife, steel ruler, scissors, wire brush. 

5 Aluminum foil trays (~24 x 30 x 6 cm, used in steam tables for serving food), 
coolers for transporting trays from field, and heavy polyethylene bags 
(^30 x 50 cm) to contain trays of field-moist soil. 

6 Analytical balance (3000 g capacity, resolution to 0.01 g), moisture tins (8 cm 
diameter x 6 cm tall), drying oven (105°C). 

j Paper "coffee" bags with plastic lining and attached wire ties (e.g., Zenith 
Specialty Bag Co., 11 x 6 cm base x23 cm height). 

g "Rukuhia" perforated drum grinder, with 2 mm perforations (Waters and 
Sweetman 1955); or another coarse soil grinder and a 2 mm soil sieve. 

9 Equipment to measure soil sampling locations. This may be a simple surveyor's 
tape to measure locations relative to permanent marker stakes in long-term field 
experiments, or a Global Positioning System (GPS) receiver. For precise pairing (in 
space) of samples collected at sequential time intervals of several years, a two- 
stage measuring approach may be useful: the general location is measured 
relative to permanent reference points or is recorded using a simple GPS receiver, 


and the position of the initial cores is marked by burying an electromagnetic 
marker originally developed to identify underground utilities (Whitlam 1998). 
Alternatively, high-resolution GPS is available in many regions. 

3.3.2 Procedure 

/ Before sampling, label paper bags with name, sampling date, location, and soil 
depth. These bags, eventually to be used for storing the air-dried soils, also serve 
as labels throughout the sampling process. Weigh the aluminum trays, one for 
each sample, and record the weight on the tray. 

2 In the field, for each sampling point, lightly brush away surface residue and 
extract a core to a depth of at least 60 cm. Move the core from the vertical to a 
horizontal position (e.g., in a sectioning trough made of 10 to 15 cm diameter 
pipe cut lengthwise), and measure the depths of any visible discontinuities (e.g., 
depth of A p horizon). Be prepared to discard cores that are unrepresentative (e.g., 
excessively compacted during sampling, evidence of atypical rodent activity, 
gouged by a stone pushed along the length of the core during sampling). It may 
prove useful to push the core (from the deepest end) out in increments, using the 
top end of the tube as a guide to make perpendicular cuts. Cut the core into 
carefully measured segments (for example: to 1 0, 1 to 20, 20 to 30, 30 to 45, 
and 45 to 60 cm), and place segments into aluminum trays, avoiding any loss of 
soil. Repeat the procedure for a second core, about 20 cm apart, and composite 
with the first core segments. Place aluminum trays inside a polyethylene bag, 
along with the labeled paper bag, fold over polyethylene bag, and store in cooler 
before subsequent processing indoors. 

j In the laboratory, remove aluminum trays from the polyethylene bags and air-dry at 
room temperature. Except for very sandy soils, itwill be much easierto grind the soils 
if the field-moist soil cores are broken apart by hand before air drying and subsequent 
grinding. Great care is required to avoid sample losses during processing and 
contamination by dust, plant material, paper, or other C-rich contaminants during 
drying. Wear rubber gloves when handling soil to avoid contamination. 

4 Once samples are air-dry, record weight of sample + aluminum tray. Remove a 
small, representative subsample (e.g., 50 to 80 g, excluding stones and large 
pieces of plant residue), and determine air-dry moisture content by oven-drying 
for 48 h at 105°C. Alternatively, the weights of field-moist cores plus trays 
may be recorded immediately after removal from the polyethylene bag and before 
they are broken apart and air-dried. In this case, accurate field moisture contents 
are crucial to estimate the densities of core segments, but spillage when cores are 
broken apart and mixed may be less consequential than the case when cores 
are dried before weighing. Thoroughly mix soils before subsampling to deter- 
mine field moisture content and possibly to retain a field-moist subsample for 
biological analyses. 

5 Crush or grind entire samples to pass a 2 mm sieve, and screen out gravel >2 mm 
in diameter. All organic material in the sample should be included; if necessary, 
separately grind roots and other large organic debris to <2 mm, and mix into the 
sample. A "Rukuhia" perforated drum grinder (Waters and Sweetman 1955) 

allows efficient, effective grinding of soil samples for SOC analysis. For each 
sample, remove and record the air-dry weight of gravel >2 mm in diameter. 

6 Place coarsely ground samples in labeled "coffee" bags for storage under cool, 
dry conditions, before analysis. For permanent storage (longer than 1 year), soil 
samples should be placed in sealed glass or plastic jars, and kept under cool, dry, 
and dark conditions. If finely ground soil is required (e.g., for elemental micro- 
analysis), the coarsely ground (<2 mm) soil should be thoroughly mixed and 
subsampled before bagging. 

3.3.3 Calculations 

7 Air-dry moisture content 

W s = (Mad - Mod)/(M d - M tin ) (3.2) 

where W s is the water content of air-dry soil, by weight (g g _1 ), Mad is the mass of 
air-dry soil and tin (g), M OD is the mass of oven-dry soil and tin (g), and M tin is the 
mass of tin (g). 

2 Density of core segment 

The following calculation provides an estimate of the density of the soil core 
segments. This may not be identical to more exacting estimates of soil bulk 
density, because compaction or loose surface layers may thwart efforts to collect 
samples of a uniform volume without altering the original mass in situ. Despite 
this, core segment density is preferred over a separate bulk density measurement 
for calculating SOC stocks. 

D cs = [(M cs - A4 g )/(1 + W s )]/[L cs ttR£] (3.3) 

where D cs is the density of core segment (g cm' 3 ), stone-free mass averaged over the 
entire sample volume, M cs is the total mass of air-dry soil in the core segment, M g is 
the mass of gravel (g), L cs is the length of core segment (cm), and R^ is the core 
radius (cm), i.e., inside diameter of coring bit/2. If the sample is a composite of more 
than 1 core segment, then L cs is the cumulative length. For example, if the sample 
contains two segments from 1 to 20 cm depth, then L cs = 20 cm. 


The procedure described above may be modified to make it applicable to individual study 
sites and objectives. Some of the important considerations include: 

a Sampling depth 

The sampling depth should, at minimum, span the soil layers significantly affected by 
the management practices considered. For example, it should include the entire depth 
of soil affected by tillage. The preferred depth may also vary with crop type; for 
example, studies including perennial forages may require deeper samples than those 
with only shallow-rooted annual crops. As the number of sampling depths increases, 
so does the effort and cost of sampling, processing and analysis. Detection of a given 


change in soil C (e.g., 2Mg C ha~ 1 ) becomes more difficult as the change is averaged 
over increasingly thick soil layers containing increasing soil C. In such instances, it 
may be reasonable to calculate changes for a layer thinner (to a minimum of perhaps 
30 cm) than that sampled, although it might have been preferable to shift resources 
from sampling deeper layers to sampling at more points. The best compromise may be 
to sample to below the zone of short-term agricultural influence, but not much 
deeper. Usually, the sampling depth should be at least 30 cm for annual vegetation 
and 60 cm or more for perennial vegetation. 

b Division of cores into segments 

The number and length of core segments depends on the vertical heterogeneity of 
SOC in the profile. Generally, the greater the gradient, the shorter should be the core 
segments. Often, the length of segments increases with depth because the SOC is 
less dynamic and more uniform at depth. Where possible, core segments might be 
chosen to correspond roughly to clear demarcations in the profile, such as tillage 
depth or horizon boundary. To facilitate comparisons among a fixed soil volume it is 
preferable to have at least one common sampling depth, but this is not essential for 
comparisons among a fixed soil mass. 

c Core diameter and number per sampling point 

The preferred core diameter and number of cores per sampling point depend on the 
sensitivity required and the amount of soil needed for analysis. Sampling larger 
volumes of soil makes the sample more representative, but also increases cost and 
disturbance of the experimental area. Soil coring may not be feasible in stony soils 
that are impenetrable, but larger cores may effectively sample profiles containing 
some gravel. 

d Core refilling 

The soil void left after removing the sample can be filled by a soil core from an 
adjacent area (e.g., plot buffers), thereby preserving the physical integrity of the 
sampling site. This replacement, however, is labor-intensive and introduces soil 
from outside the treatment area which could affect subsequent samplings. Without 
intentional replacement, core voids become filled by adjacent topsoil, so subse- 
quent cores should be positioned far enough away to avoid areas most affected by 
removal of previous cores, but close enough to exclude excessive spatial variations. 

e Core location relative to plants 

Proximity to plants may affect sample SOC contents, especially at the soil surface 
where plant C is concentrated at the crowns and under perennial or tap-rooted 
vegetation with localized plant C inputs to soil. Cores should be positioned to avoid 
bias, for example, when about 1/3 of the soil surface area is occupied by plants, 
three cores could be collected: one beneath plants, and two more between plant 
rows or crowns. Often basal areas occupied by the crowns of crops planted in 
rows are small (<30%) relative to the interrow areas, so samples are collected 
exclusively from the interrow. In other cases, such approximations may introduce 
considerable bias. 


f Measuring total soil C stocks 

In earlier studies of SOC, largely from the perspective of soil fertility, recent plant 
litter in the sample was often removed by sieving and discarded. In studies of C 
sinks, however, the total C stock should be measured. The procedure described 
above includes recent litter directly in the sample. An alternative approach is to 
analyze the plant debris separately, but include it in the calculation of C stocks. 
Above-ground residue, if present in significant amounts, may also need to be 
considered in calculating total C stocks (Peterson et al. 1998). 

g Contamination from other C sources 

Care should be taken to avoid introducing extraneous C from oil used as lubricant in 
soil coring tubes, wax in sample bags, and coatings on foil trays. The sample drying 
area should be free of dust (e.g., from plant sample processing), insects, and rodents. 
Cross contamination (e.g., between carbonate-rich subsoil and organic matter-rich 
surface soil) should be avoided during processing. 

h Repeated measurements of SOC over time 

Temporal changes in SOC can be measured with higher sensitivity if successive 
samples are removed from close proximity to (though not directly on) previous soil 
cores (Ellert et al. 2001; Conant et al. 2003; VandenBygaart 2006). To do that, the 
original sampling locations can be recorded using the GPS receiver, or by burying 
an electronic marker in one of the voids left by core removal. At subsequent 
sampling times, soil cores can then be taken immediately adjacent to previous 
cores, often in a grid pattern within "microplots" (Figure 3.3). The pattern may 
be modified to accommodate additional sampling times or other site conditions 







% Cores at 7=0 year 

© Electromagnetic markers 

O Cores at T= 6 years 

FIGURE 3.3. An example of the arrangement of soil cores within 4 x 7 m microplots intended 
for measuring temporal change in SOC stocks. (Adapted from Ellert, B.H., 
Janzen, H.H., and McConkey, B.G. in R. Lai, J.M. Kimble, R.F. Follett, and 
B.A. Stewart, (Eds.), Assessment Methods for Soil Carbon, Lewis Publishers, 
Boca Raton, Florida, 2001.) 


(Conant et al. 2003; VandenBygaart 2006). To most efficiently assess temporal 
changes in soil C stocks, the number of cores within each microsite and of 
microsites within a field or plot may be adjusted for differences in variability at 
the microsite and field levels (Bricklemyer et al. 2005). 


3.4.1 Materials 

7 Fine soil grinder and small test sieves (No. 60 with 250 |jim openings and No. 1 00 
with 150 |xm openings). 

2 Carbon analyzer, using dry combustion and subsequent analysis of C0 2 . (For 
information on analysis of total and organic C see Chapter 21 .) 

3.4.2 Procedure 

7 Obtain a representative subsample of the previously stored air-dry soil samples, 
ideally using "drop through" sample riffles or centrifugal sample dividers, as 
needed to avoid a biased subsample. Variability introduced by simpler, more 
expedient approaches (e.g., small scoops from six distinct areas within a 
thoroughly mixed tray of air-dried, <2 mm soil) is easily quantified by collecting 
multiple subsamples from a few samples. Scooping from the tops of sample bags 
or jars is not recommended, because soil constituents tend to separate during bag 
or jar filling and sample handling. 

2 For most microanalytical techniques the coarsely ground (<2 mm) sample will 
have to be finely ground using a roller or jar mill, ball-and-capsule mill, shatter- 
box or ring-and-puck mill, or a mortar and pestle (e.g., Kelley 1994; Rondon and 
Thomas 1994; McGee et al. 1999; Arnold and Schepers 2004). The preferred 
fineness depends on the amount of sample analyzed. If less than 0.1 g is to be 
combusted, the sample should be ground to pass through a 150 |xm sieve. The 
entire subsample should be ground to pass through the designated sieve (verified 
by testing a representative subset of samples rather than every sample). Finely 
ground samples can be stored in glass vials. 

3 Dry samples and standards at 60°C to 70°C for 18 h, and determine the 
organic C concentration (g C kg~ 1 soil) (see Chapter 21). It is critical that 
inorganic C be completely removed before analysis by addition of acid, or 
that inorganic C be analyzed separately and then subtracted from total C 
concentration to estimate organic C concentration (see Chapter 21 ). Ideally certified 
reference materials should be used to verify analytical accuracy, but standard 
soils with certified values for total and organic C remain rare (Boone et al. 1999). 
At minimum, standard soils prepared in-house or obtained from a commercial 
supplier should be used to calibrate analyses and monitor analytical precision. 

4 Express the concentration in units of mg C g~ 1 dry soil (=kg C Mg~ 1 = % x 10). 

3.4.3 Calculations 

The SOC stock is the amount of organic C in a fixed layer of soil per unit area of land. 
Typically, it is expressed in units of Mg C ha~' to a specified depth. Alternative units 
include kg C m~ 2 = Mg C ha x 0.100. The simplest way to calculate SOC stocks is 
to accumulate the products of concentration and core density to a fixed soil depth and 
volume (see calculation below). But this approach is subject to bias when comparing SOC 
across space or time if core density varies even slightly (Ellert and Bettany 1995). For 
example, when comparing SOC stocks in two treatments, if the average core density to the 
specified depth is 1.10 Mg m~ 3 in treatment A and 1.00 Mg m~ 3 in treatment B, then 
the SOC stocks in treatment A will be biased upward because it has 10% more soil in the 
layers compared. For that reason, SOC stocks should be calculated on an "equivalent mass" 
or "fixed mass" basis (see calculation below), unless core densities are very uniform. 

SOC Stocks (Fixed Depth) 

SOC fd = XAsC cs L cs x0.1 (3-4) 


where SOCfd is the SOC stock to a fixed depth (Mg C ha~' to the specified depth), £> cs is 
the density of core segment (g cm -3 ), C cs is the organic C concentration of core segment 
(mg C g _1 dry soil), and L cs is the length of core segment (cm). 

SOC Stocks (Fixed Mass) 

1 For all samples, calculate the mass of soil to the designated depth: 

A/J soi | = Y^ D CS L CS x 1 00 (3.5) 

where A^ soi | is the mass of soil to a fixed depth (Mg ha~'). 

2 Select, as the reference, the lowest soil mass to the prescribed depth from all 
sampling sites (Kef)- 

3 Calculate the soil mass to be subtracted from the deepest core segment so that 
mass of soil is equivalent in all sampling sites: 

A/f ex = Mjoii - Kef (3.6) 

where M ex is the excess mass of soil, to be subtracted from deepest core segment. 

4 For each sampling site, calculate SOC stock to fixed mass: 

SOCfm = SOCfd - Me, x C sn /1 000 (3.7) 

where SOCfm is the SOC stock for a fixed mass of M K \ and C sn is the SOC 
concentration in deepest soil core segment (mg C g~ 1 dry soil) (core segment = n). 


Sample Calculations 

Given the following three hypothetical soil cores: 

Depth (cm) 

SOC concentration (g C kg soil) 
Core 1 Core 2 Core 3 


SOCfd to 40 cm is 

78.3, 85.9, and 74.1 Mg C ha~' for cores 1, 2, and 3, respectively. 

For SOC FM : 

M soU = 4810, 5070, and 4690 Mg ha~' to 40 cm, for cores 1, 2, and 3, respectively. 


M re f = 4690 Mg ha~' (mass of soil core 3), and 

M ex = 120, 380, and Mg ha -1 , for cores 1, 2, and 3, respectively. 


For core 1, SOC FM = 78.3 - 120 x 14.3/1000 = 76.6 Mg C ha~\ 
Similarly, SOCfm = 80.1 and 74.1 Mg C ha -1 , for cores 2 and 3, respectively. 
Thicknesses of the fixed masses = 40 - M ex /(D CS x 100) = 39.1, 37.2, and 40.0 cm 
for cores 1, 2, and 3, respectively. 


The approach described to estimate SOC stocks is applicable to sites where temporal changes 
are attributable to biological processes (chiefly the balance between soil C inputs and 
outputs), rather than geomorphic processes (soil erosion and deposition). The fundamental 
assumption is that soil mass is largely conserved among sampling times. At sites where this 
does not hold, other approaches are required to estimate lateral soil redistribution or net soil 
imports or exports, before temporal changes in SOC may be estimated. For example at sites 
with considerable mass additions or removals (e.g. waste application or soil export) survey 
techniques that enable sampling to a fixed subsurface elevation might be appropriate (Chang 
et al. 2007). 

Numerous variations are possible in the calculation of SOC stocks by the "fixed mass" 
approach. For example, instead of using the SOC concentration of layer n in the correction 
(Equation 3.7), it may be more appropriate to use the weighted mean concentration in layers 
n and n + 1 . Or, rather than subtracting SOC in the correction, some researchers select a 
reference mass and add SOC, based on the SOC concentration of the layer below. In all 
cases, the method assumes that concentration value used is representative of the layer added 
or subtracted. For that reason, some researchers have used core configurations with a short 
segment just below the depth of interest. For example, if C stocks are to be estimated for the 
to 20 cm layer, a 20 to 25 cm segment is isolated to be used for the "fixed depth" 


Whether comparisons are based on a fixed soil depth or mass is immaterial for situations 
with soil redistribution, accumulation, or export. In such situations, it is practically impos- 
sible to distinguish between the effects of geomorphological processes (soil redistribution) 
and biological processes (plant C inputs and SOC decay). Only in rare instances (e.g., soils 
with a persistent and uniform marker layer, such as a fragipan) can soil deposition or erosion 
be inferred from routine soil sampling. 

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Chapter 4 
Soil Sample Handling and Storage 

S.C. Sheppard 

ECOMatters Inc. 
Pinawa, Manitoba, Canada 

J.A. Addison 

Royal Roads University 
Victoria, British Columbia, Canada 


This chapter deals with soil samples between when they are sampled and when they are 
analyzed. The key message is that sample handling and storage can profoundly affect 
analysis results, and no one way is suitable for all analytes. The issues related to soil sample 
handling and storage relate to the management of sample clump size, moisture content, 
temperature, and storage time. 

With the increased availability of software to gather and interpret spatial information, there 
have been important advances in the past decade on methods to sample soils. Similarly, 
analytical capabilities have been remarkably enhanced, with greater sensitivity and more 
analytes. This includes notable advances in the characterization of soil organisms and 
biological attributes. However, there has been much less research and practical emphasis 
on the effects of handling and storage of soil samples. Nonetheless, there is abundant 
evidence that differences in handling and storage can profoundly affect the interpretation 
of results. 

Perhaps the single most important role of analysis in soil science is to move beyond the 
reporting of absolutes, and toward the reporting of environmentally relevant measures. 
Absolute quantities, such as total elemental composition, total organic matter content, and 
even total porosity, are relatively simple to measure, and are relatively insensitive to effects 
related to sample handling and storage. However, these quantities are only partially relevant 
to what many researchers want to measure. Often, the more important measures are attributes 
such as the bioavailable or leachable elemental composition, and functional and biotic 
properties of the soil. For these more subtle measures, methods of sample handling and 
storage become critical. Examples from the literature include: 


• Plant-available nitrogen (Craswell and Waring 1972; Wang et al. 1993; Verchot 1999; 
Fierer and Schimel 2002; Magesan et al. 2002; Riepert and Felgentreu 2002), phos- 
phorus (Potter et al. 1991; Grierson et al. 1998; Turner and Haygarth 2003; Worsfold 
et al. 2005), potassium (Luo and Jackson 1985), and sulfur (Chaudhry and Cornfield 
1971; David et al. 1989; Comfort et al. 1991) 

• Speciation of metals and soil solution composition (Leggett and Argyle 1983; Lehmann 
and Harter 1983; Haynes and Swift 1985, 1991; Walworth 1992; Neary and Barnes 
1993; Meyer and Arp 1994; Simonsson et al. 1999; Ross et al. 2001) 

• Soil biological activity (Ross 1970; Zantua and Bremner 1975; Ross 1989; Van Gestel 
et al. 1993; Stenberg et al. 1998; Mondini et al. 2002; Allison and Miller 2005; 
Goberna et al. 2005) 

• Studies of soil organic matter (Kaiser et al. 2001) 

• Extraction of organic contaminants (Belkessam et al. 2005) 

Without doubt, researchers must refer to the primary literature to identify the requirements 
and limitations for sample handling and storage specific to the analysis they undertake. It is 
not a default process; the researcher must be able to defend the sampling handling and 
storage decisions. Unfortunately, several researchers have shown that the effects of sample 
preparation and storage are not similar from soil to soil, so that inappropriate handling 
can jeopardize interpretation of results among different soils (e.g., Brohon et al. 1999; 
Neilsen et al. 2001). 

The objective of this chapter is to provide guidance on sample handling, including compo- 
siting, reduction in clump size, and management of soil moisture. Table 4.1 gives an 
overview. The chapter also discusses two aspects of sample storage; storage between 
sampling and the primary analysis, and the long-term storage or archive of samples. 
Handling of samples of soil constituents separated in the field, such as soil pore water 
collected in lysimeters (e.g., Derome et al. 1998) is not discussed. 


The requirements for each sampling campaign will differ, but a typical sequence is as follows: 

Collect composite sample in the field or from the experimental system. 

If the sample is too large, reduce clump size, mix and package a portion of the 
composite to transport to the laboratory. 

Collect a subsample for determination of moisture content, the subsample is weighed, 
dried at 105°C, and reweighed. 

Dry remaining sample to a moisture content suitable for further sample handling. 
If appropriate and required, further reduce clump size, such as by grinding. 

TABLE 4.1 Typical Attributes for Handling and Storage of Soil Samples 

Compositing and clump 

Storage before analysis 

So/7 fauna: earthworms, 

Avoided, generally use 



nematodes, other 

minimally disturbed soil 


cores or clods (point 
samples, not composites) 

Microbial activities: 

May be minimally disturbed 


loist or workable 

respiration, functionality 

point samples or composites 

ire content 


of gently ground soil 

Microbial populations: 

Need for aseptic conditions 


loist or workable 

enumeration, population 

often results in point samples 

ire content 


(not composites) 

Microbial attributes: PLFA, 

May be minimally disturbed 

Field-moist or workable 


point samples or composites 
of gently ground soil 

ire content 

So/7 organic matter. 

Bioavailability and chemical 

Bulk physical properties: pore 
size distribution, bulk 

Physical: granulometry, tc 
organic matter content 

Elemental analysis: total and 
strong-acid extractable 

Moderately aggressive 
grinding may be acceptable 

Moderately aggressive 
grinding may be acceptable 

Avoided, generally use 
minimally disturbed soil 
cores or clods (point 



Varies with analysis, may 

include oven drying 
Workable moisture content 

Field-moist or workable 
moisture content, but 
results reported on oven- 

Aggressive grinding 
acceptable as long a 3 j,,, b ,c 

Aggressive grinding 
acceptable as long as single 

Aggressive grinding 

orally reported o 
n-dried basis 

Generally reported on a 
oven-dried basis 

Generally reported on a 
oven-dried basis 

Minimal time, refrigerated 
but not frozen 

Minimal time, refrigerated 
but not frozen 

Minimal time, refrigerated 
but not frozen 

Varies with analysis, 
freezing may be 

Minimal time, refrigerated 
may be ideal 

Indefinite if refrigerated, may 
change upon freezing 

Indefinite in dried st 

Indefinite in dried state 

Indefinite as long as 
contamination avoided 

Not possible for primary 
analytes, suitable for some 
ancillary measurements 

Not possible for primary 
analytes, suitable for some 
ancillary measurements 

Not possible for primary 
analytes, suitable for some 

Varies with analyte, 
extremely low temperature 
freezing (-80°C) may be 

Varies with analysis 

Not possible for primary 
analytes, suitable for some 

Indefinite if refrigerated, may 
change upon freezing 

Indefinite in dried state 

Indefinite in dried state 

Indefinite as long as 
contamination avoided 


• Subsample as required for analysis. 

• Prepare an archive sample. 


The intended outcome of compositing and reduction in clump size is to ensure the sample 
represents the whole. Compositing involves the gathering and mixing of a series of indivi- 
dual samples, typically from a series of sampling points across the landscape. Reduction in 
clump size is often required so that both compositing and subsampling for analysis represent 
a uniform material. See Schumacher et al. (1990) for detailed discussion of methods of 
sample splitting and subsampling. 

One key issue is that the clumps be small enough that the composite sample or subsample 
contains a large number of them. This is a statistical issue. Allison and Miller (2005) 
described how variability in biological assays increased as the size of the analyzed sub- 
samples decreases, and Liggett et al. (1984) commented that the size of subsample required 
to obtain consistent measurements of plutonium in soils was too large to be practical (in their 
case, variability among subsamples always dominated over field variation). As a general 
guideline, if a required composite sample is 1 kg of soil, a reasonable clump size might be 
~ 5 g (5 cm 3 ) or less. If a required subsample is 0.5 g, then the ' 'clump" size might better be 
described as powder, ground as fine as practical within the limits required by the analysis. 
For example, Neary and Barnes (1993) and Wang et al. (1993) both recommended grinding 
to pass a <0.5 mm mesh if subsamples were to be <1 g. 

The other key issue is that the process of breaking up the clumps does not disrupt the 
analytes. Some of this is self-evident; if one is sampling to measure soil macropore 
properties or soil fauna, then breaking up of clumps should be minimal and not aggressive. 
Craswell and Waring (1972) showed that grinding affected microbial mineralization rates in 
soil, and Neary and Barnes (1993) found that grinding, and especially mechanical grinding, 
affected extractable iron and aluminum concentrations. In contrast, if the analyte is total 
elemental concentration, quite aggressive grinding (hammer mill, mortar, and pestle) may be 
acceptable, as long as the grinder itself does not introduce contamination. 

More controversial is the degree of grinding appropriate for measures of bioavailable 
element composition, or microbial attributes. As an example, tests of soil nutrient availabi- 
lity (soil fertility testing) were originally calibrated with soils that had very specific prepar- 
ation, typically air-dried, hand-sieved to pass a 2 mm mesh, followed by volumetric (as 
opposed to mass-based) sampling for analysis. More aggressive drying and grinding affects 
the amount of nutrient removed by the selective extractants employed, increasing the 
extractable P by up to 165% in some soils (Turner and Haygarth 2003). Unfortunately, 
gentle manual preparation is expensive and, with the commercialization of soil fertility 
testing, more rapid and more aggressive grinding is now the norm. It is not clear if the 
underlying test response data have been recalibrated accordingly. 

Another difficult issue in soil sample preparation is the decision of what to do with pebbles, 
roots, and anything else that behaves differently during sample preparation than the bulk soil 
matrix. Many researchers simply remove these nonconforming materials, but obviously their 
presence can significantly affect the interpretation of analytical results back to the field, if 
for no other reason than they represent a volumetric dilution of the soil matrix. As a default, 


it may be an appropriate rule to remove pebbles larger than the required mesh size, but record 
their mass relative to the mass of the whole soil. This implies the full sample, apart from the 
pebbles, is ground to pass the mesh. For roots and organic debris, it may be appropriate to 
simply remove these as they could be considered ephemeral to the soil. For some analytes, 
the organic debris might be considered an important secondary subsample. This might be the 
case for analysis of lipophilic compounds or of fungal activities. 

Subsampling organic soils and horizons can also be problematic, especially when materials 
such as decaying woody plants are present within the soil profile. Knife mills may be useful 
for grinding fibrous organic soils, if appropriate for the intended analysis. 


The soil moisture content of stored samples is not only of importance for issues related to 
sample preparation (e.g., reduction of clump size) but can also profoundly affect the results 
of subsequent analyses. Many soils are physically impossible to handle when they are too 
wet, and clay soils can be very difficult to grind if they become too dry. One argument in 
deciding how much to dry the sample is that soils in their native setting are usually subject to 
wetting and drying processes, and so drying in the laboratory to moisture contents that can be 
found in the field seems defensible for many analytes. 

The standard for measurement of soil mass is dried at 105°C for as long as required to reach a 
constant weight. For analyses of soil properties reported on a dry weight basis, this basis 
should be, and is usually assumed to be, the weight after drying at 105°C. 

However, the 105°C temperature and the resultant low moisture content are very disruptive 
to many soil properties. It kills meso- and microbiota, denatures organic entities including 
soils enzymes, oxidizes some inorganic constituents, collapses clay interlayers, and can 
modify other soil solids. It is a suitable dryness for absolute measures such as total elemental 
composition and granulometric composition, and is suitable for some levels of grinding for 
some soils. For many other analytes, and for successful grinding of clay or organic soils, it is 
better to allow the soil to retain more moisture. 

Nonetheless, if soil samples are not dried to 105°C and the results are to be presented per unit 
of soil dry weight, then the researcher should measure the soil moisture content of the soil 
"as analyzed," and convert the results to the 105°C-dry basis. Very often, there is little 
difference in moisture content between air-dried and 105°C-dried, but they cannot be 
assumed to be equivalent. 

Typical target moisture contents are: 

• Field moist or "as is" moisture content, which can be extremely variable but neces- 
sary to avoid disruption if living organisms are to be extracted. 

• Workable, a judgment by the researcher where the soil is allowed to dry to a moisture 
content that is typically between field capacity and air-dry, and the soil is just dry 
enough to allow gentle grinding, such as sieving, with no dust production. Microbial 
activity will be present, seeds may germinate, and refrigerated and dark storage should 
be considered. As the soil still contains living organisms, allowance for gas-exchange 
may be required, but the sample should be protected against excessive moisture loss. 

Polyethylene bags may be suitable as they allow diffusion of oxygen and limit water 
loss. The actual moisture content should be confirmed whenever analyses are 

• Air-dried, where the soil is allowed to equilibrate with humidity in the air, resulting in 
soil that is nearly as dry as oven dry and can be aggressively ground (if required). Soils 
at this moisture content can be stored in water-permeable containers (e.g., cardboard 
boxes). Microbial activity is minimal and a flush of microbial activity is expected 
when the soil is rewetted. This is the most convenient moisture content, as long as it is 
consistent with the intended analyses (see examples in Table 4.1). 

• Oven-dried at 105°C, where the soil is dry enough that it will accumulate moisture 
from the air. Soils at this moisture content must be stored in sealed containers or 
desiccators, and it may be necessary to redry the soils to assure they are at the required 
moisture content when used. The advantage of this moisture content is that it is the 
reference standard. 

• Oven-dried to a temperature intermediate between air-dry and 105°C, which is gener- 
ally a compromise between the rather slow process of air-drying and the damaging 
effects of 105°C. Temperatures of 30°C-40°C are arguably in the range of temperat- 
ures experienced at the soil surface in the field. Temperatures of 50°C-80°C are 

Drying a soil, even at room temperature, causes a number of reactions. Living organisms 
either pass into a resting stage, or die. Dissolved inorganic materials will become 
more concentrated in the remaining pore water, and ultimately will form precipitates 
or perhaps gel-phase materials. Dissolved organic materials probably coagulate, both because 
they become concentrated and because the salt concentration of the pore water increases. Solid 
organic materials will deform when dry, uncover underlying mineral surfaces and may 
become very hydrophobic. Mineral-phase materials are generally resistant to modification 
until the soil becomes extremely dry or excessive heat is used. 

Given these changes, it is obvious that moisture management must vary according to the 
required analysis (Table 4.1). Storage of air-dried or oven-dried samples is very convenient, 
and although dry storage will introduce gradual changes in some soil attributes, at least for 
the measurement of some soil chemical and physical properties these changes may be 
minimal. However, some types of chemical analyses are affected by drying. For example, 
some soil nitrogen fertility tests are influenced by drying, and as a result some commercial 
laboratories request soils not be dried before being sent to the laboratory. For most other 
large-scale operations, such as other soil fertility testing where large numbers of samples are 
required, air-dried or a low temperature oven-dried samples are the norm, for convenience as 
well as reasonable consistency. 

An approach used by some to overcome the effects of drying is to rewet and incubate soil 
samples before analysis. The rationale is that air-drying and rewetting are natural occur- 
rences, and so rewetting may be appropriate mitigation for the temporary effects of air- 
drying. Lehmann and Harter (1983) noted some recovery of copper sorption when soils were 
rewetted and incubated for 1 month. Haynes and Swift (1991) noted that extractability of 
metals could be restored with rewetting, whereas effects of drying on extractability of 
organic matter "was only slowly reversed following rewetting." 


For biological, microbial, and enzyme assays, drying should generally be avoided or 
restricted to drying to a workable moisture content. Numerous studies have shown that 
drying and then rewetting the soil has a tremendous impact on biological properties, 
including microbially mediated soil chemical transformations (Van Gestel et al. 1993; 
Riepert and Felgentreu 2002). Although some studies have shown that rewetting and 
incubation of dried soil restores biological activity to at least some degree, it is also clear 
that different segments of the microbial population respond in different ways. Consequently 
the degree of recovery and the time taken for microbial population and functions to 
reestablish differs for different soils and for different microbial groups (e.g., Fierer and 
Schimel 2002; Pesaro et al. 2004). 


As indicated in the introduction, there is no default storage method for all analytes and each 
researcher must be able to defend decisions made about sample storage. Any analysis of 
biological attributes or biologically mediated activities, and any analysis of volatile or labile 
constituents obviously require minimal storage time and specific conditions of temperature, 
moisture content, and container type. Analysis of nitrogen compounds and organic chemicals 
subject to biodegradation are notably among those where storage conditions are an issue 
(Stenberg et al. 1998; Rost et al. 2002). 

In situations where a living soil fauna is of interest, soil samples should be stored at 5°C rather 
than frozen. The ability to withstand freezing temperatures in soil invertebrates is determined by 
a complex set of physiological and behavioral adaptations that are time dependent, so it is 
generally not reasonable to assume that soil samples can be safely frozen simply because the 
sample comes from an area subject to seasonal freezing. Edwards and Fletcher (1971) concluded 
that soil storage up to a week at 5°C should not cause any serious changes in the numbers of 
individuals or groups of soil fauna extracted from soil samples, but that after 28 day storage at 
5°C, or even earlier at higher temperatures, significant changes were to be expected. 

The appropriate temperature for storing soil samples required for determining microbial 
parameters, including the potential of the indigenous microbial flora to degrade contamin- 
ants, is controversial. Stenberg et al. (1998) concluded it was acceptable to store soils for 
microflora analyses at — 20° C if the soils were from areas where they were normally frozen 
in winter. Indeed some test guidelines that measure microbial activity (e.g., OECD 2000) 
agree that if soils are collected from areas where they are frozen for at least 3 months of the 
year, then storage at — 18°C for 6 months "can be considered." However several other 
authors, including some working on soils from northern areas, stress that freezing soil 
samples causes significant and long-term changes in microbial abundance and activity and 
that certain groups are particularly sensitive to the effects of freezing (Zelles et al. 1991; 
Shishido and Chanway 1998; Pesaro et al. 2003). On the other hand other microbial assays 
(e.g., phospholipid fatty acid [PLFA]) generally require samples to be stored in a frozen state 
in order to minimize degradation of the fatty acids during storage. 


Archival storage is intended to serve a number of objectives. The most immediate is to allow 
reanalysis of samples where the primary results are questioned. This is a form of replication 
of analysis. Relatedly, it is sometimes important to measure other attributes of a specific 


sample in order to explain the primary results. For example, retrospective analysis of trace 
element content may confirm a hypothesis about differences in the initial analyses. 

However, both these objectives relate to the initial reason to collect the samples. Archive 
samples serve other objectives as well, related to future research. An improved analytical 
method may become available, and reanalysis of archived samples is one way to validate the 
new method and relate the new and old methods. Alternatively, another research project may 
require a suite of soils with the specific attributes available in the archive samples. 

Another key role for archived soils samples is to provide reference standards, and in the case 
of ecotoxicology assays to provide a diluent soil (Sheppard and Evenden 1998). Ehrlichmann 
et al. (1997) commented that in their reference soils, the toxicity of organic contaminants 
decreased with storage time, whereas the toxicity of metals increased with storage time. 
Riepert and Felgentreu (2002) investigated soils stored as reference soils for plant ecotoxicity 
bioassays, and concluded that "soil kept as a laboratory standard under air-dried conditions 
over a long time period is not suitable [ . . . ] due to the [ ... ] microbial situation," especially 
as related to nitrogen mineralization. 

There is not a lot of information on how long an archive sample remains valid. Certainly 
samples lose biological validity fairly quickly, but will retain physical attributes such 
as granulometry indefinitely. In contrast, Bollen (1977) found that samples stored dry for 
54 years retained their ability to respire and oxidize sulfur, some more and some less than 
when the samples were originally collected. 

Perhaps the single most important aspect of archived soil samples, just as with any kind of 
archive, is the documentation. This must include provenance of the sample, collection 
details, preparation and storage conditions, and ideally the linkage to the researcher, and 
the primary analysis the researcher completed on the samples. 


A review of the literature will immediately indicate that artifacts have been shown to arise 
from all types of soil sample handling and storage. No one protocol is suitable for all 
analytes. Convenient protocols such as air-drying and grinding have profound effects 
on physical, chemical, and biological attributes of soils. Even soil fertility testing for 
phosphorus and metals can be jeopardized by subtle differences in sample handling. Soil is 
a living material, and perhaps soil samples need the same care in handling that is afforded to 
tissue samples. 

The most important message of this chapter is that sample handling and storage protocols are 
not by default. It is the responsibility of the researcher to consider and be prepared to defend 
the decisions taken. 

Allison, V.J. and Miller, R.M. 2005. Soil grinding Belkessam, L., Lecomte, P., Milon, V., and 

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Chapter 5 

Quality Control in Soil 

Chemical Analysis 

C. Swyngedouw 

Bodycote Testing Group 
Calgary, Alberta, Canada 

R. Lessard 

Bodycote Testing Group 
Edmonton, Alberta, Canada 


In analytical work, quality can be defined as the "delivery of reliable information within an 
agreed span of time, under agreed conditions, at agreed costs, and with the necessary 
aftercare" (FAO 1998). The agreed conditions include specifications as to data quality 
objectives (DQOs), which include precision, accuracy, representativeness, completeness, 
and comparability. These objectives are directly related to "fitness of use" of the data and 
they determine the degree of total variability (uncertainty or error) that can be tolerated in the 
data. The DQOs ultimately determine the necessary quality control (QC). 

Quality management systems have been developed for analytical laboratories (USEPA 2004) 
and there are examples of these systems in the literature (CAEAL 1999). More information 
can be obtained from the International Organization for Standardization (ISO 17025). 

Implementation of quality management implies the next level of quality — quality 
assurance (QA), defined as the "assembly of all planned and systematic actions necessary to 
provide adequate confidence that an analytical result will satisfy given quality objectives or 
requirements" (FAO 1998). The use of QA guarantees that the delivered product is commen- 
surate with the intended use and ensures that data have scientific credibility, and thus permits 
statistical interpretations as well as management decisions to be made (AENV 2004). 

All sampling and laboratory activities have one target: the production of quality data that is 
reliable, consistent, and has a minimum of errors. Thus, to ensure the integrity of QA a 
system of checks are needed to establish that quality management systems are maintained 
within prescribed limits providing protection against "out of control" conditions and 
ensuring that the results are of acceptable quality. To achieve this, an appropriate program 


of QC is needed. QC includes "the operational techniques and activities that are used to 
satisfy the quality requirements or DQOs" (FAO 1998). Producing quality data is a major 
enterprise requiring a continuous effort. Approximately 20% of the total costs of analysis are 
spent on QA and QC. 

This chapter focuses on some pertinent aspects of QC in soil chemical analysis. QA topics 
are not discussed but QA information can be found in CCME (1993), FAO (1998), Taylor 
(1990), IUPAC (1997), and ISO 17025 (2005). 


Determining a property or a concentration of an analyte in a soil sample follows four general 

j Sample collection and handling 

2 Sample shipping and transport 

j Sample preparation and analysis 

4 Results data entry, handling, and reporting 

Each of these steps has the potential to introduce errors into the final estimate of a property 
or a concentration. The careful use of tested and established protocols at each step, along 
with careful tracking of the samples, can help minimize, but not eliminate the errors. 
Table 5.1 outlines field and laboratory sources of error, while Table 5.2 indicates some 
corrective actions to counteract specific laboratory errors. 

5.2.1 Sample Collection and Handling 

Bias caused by sampling is often difficult and expensive to measure. Field spikes (samples of 
analyte-free media such as clean soil or sand fortified or spiked with known amounts of the 
target analytes) are sometimes used to assess sampling bias. Sampling errors are usually 
much larger than analytical errors (Jenkins et al. 1997; Ramsey 1998; IAEA 2004). 

5.2.2 Contamination 

Contamination is a common source of error in soil measurements (Lewis 1988; USEPA 
1989). Field blanks (analyte-free media) are the most effective tools for assessing and 
controlling contamination. In addition, equipment rinsate blanks may also be used. Field 
blanks are not effective for identifying matrix interferences or for spotting noncontaminant 
error sources (such as analyte loss due to volatilization or decomposition). Field spikes, 
however, can be used for noncontaminant sources. 

5.2.3 Soil Sample Storage/Preservation 

Physical and chemical changes to soil samples can occur between collection and analysis. 
Physical changes include volatilization, adsorption, diffusion, and precipitation, while chem- 
ical changes include photochemical and microbiological degradation (Maskarinic and 
Moody 1988). 


TABLE 5.1 Field and Laboratory Sources of Uncertainty in Chemical Analysis Data and 
Their Assessment 

Nonrandom spatial 


Sample handling 



Arises from the complexity 
of the soil (clay, silt, and 
sand). The error inherent i 
using a portion to 
represent the whole 

Error caused by sampling, 
sample handling, and 

Error from analytical 
measurements, includin 
sample preparation 

Faulty data handlin 
transcription error 

required to build a 
representative sample. 
Take replicates from 
spatially distinct poii 
and take a larger nui 
of samples. Use a less 
expensive and less precis 
analytical method 
Increase amount of sampli 
taken (sample mass) to 
represent the matrix 

Make several large 
composites and split them 
into replicates. Also, take a 
larger number of samples 

Split samples into replicates 

just before sample 

preparation. Splits may 

be sent to another 

laboratory for confirmatory 

Automate data transfer, 

perform data verification 

The following QC practices are helpful to store and preserve soil samples: 

• Seal sample containers to reduce contamination and prevent water loss. 

• Minimize sample container headspace to reduce loss of volatiles. 

• Refrigerate or freeze samples during storage and transportation to reduce loss of 
volatiles and minimize biodegradation. 

• Carry out extractions and digestions as soon as possible. This keeps the analyte in the 
resulting extraction phase (e.g., solvent or acid), thereby stabilizing the analyte. As a 
result, a sample extract can be held for a longer time, up to the maximum limits as 
specified by the method. 

Analyze samples as soon as possible. 


TABLE 5.2 Corrective Action for Laboratory Sources of Error 

Corrective action 

Segregation or stratification of soils o 


Sample or equipment c 

the laboratory e 
Sample carryover on extraction vessels or 

Samples weighed, processed, or analyzed 

out of order 
Inaccurate o 

r calibration solution mismatch 


Drift in instrument response 
Poor instrument sensitivity or high detectioi 
Faulty data handling or human transcriptio 

Rehomogenize before subsampling for 

Store samples, reagents, equipment 

Rinse with cleaning solution between 

Run a known reference sample at a 

regular interval 
tions Check new standards against old 

before use 
Make up standards in e> 

used for soil samples 
Use frequent calibration/QC checks 
imits Optimize all operating parameters 

Proofread input, automate data transfer 

ling solution 

Source: From Hoskins, B. and Wolf, A.M., 
for the North Central Region, Missou 
65-69. With permission. 

n Recommended Chemical Soil Test Procedures 
i Agricultural Experiment Station, Columbia, 1 998, 

5.2.4 Sample Holding Times 

Holding time is the storage time between sample collection and sample analysis, in 
conjunction with designated preservation and storage techniques (ASTM 2004). Usually 
microbiological and volatiles analyses have short holding times. A holding time study involves 
storing replicate spiked samples for a period of time and periodically (e.g., once a day) analyz- 
ing three replicates for a specific characteristic (e.g., toxicity). The holding time is established 
as the time when the concentration or characteristic drops below the criterion set by the DQOs 
(e.g., a 10% drop). For more information, see Chapter 4 and USACE (2005). 

Maximum holding times for soil samples depend on the soil type, the analyte or the 
characteristic being determined, storage conditions, and loss of sample integrity (Maskarinic 
and Moody 1988). 

Results of samples not analyzed within the specific holding time are considered "compro- 
mised" (see Section 5.5). The actual result (e.g., concentration) is usually assumed to be 
equal or greater than the result determined after the holding period has expired. 

5.2.5 Subsampling the Soil Sample 

In most cases, the soil sample that arrives in the laboratory is not analyzed entirely. Usually 
only a small subsample is analyzed, and the analyte concentration of the subsample is 
assumed representative of the sample itself (see Figure 5.1). A subsample cannot be perfectly 
representative of a heterogeneous sample, and improper subsampling may introduce signifi- 
cant bias into the analytical process. Bias that occurs as a result of subsampling may be 
improved by procedures such as grinding and homogenizing the original samples (Gerlach 
et al. 2002). One way to detect errors due to subsampling would be to set up an experiment 
where one subsamples a reference material, or a material that is already well characterized. 


Preparation: digestion, 
extraction, filtration 

S' Within 

<^ calibration 







M naysis 



Measured result 




Reported result 

FIGURE 5.1. Laboratory sample pro 

Once the sample enters the laboratory, it undergoes established procedures from sample 
preparation to final analysis. After the sample extract is introduced into the analytical 
instrument, the analyte is sensed by the detector and that information is converted into an 
electronic signal. The intensities of these electronic signals are converted into c 

5.2.6 Detection Limits 

Detection limits are estimates of concentrations where one can be fairly certain that the 
compound is present. The USEPA in 40 CFR136 (USEPA 1984) defines the method detection 
level (MDL) as "the minimum concentration of a substance that can be measured and 
reported with 99% confidence that the analyte concentration is greater than zero. ' ' Method 
detection limits are statistically determined values that define how measurements of an 
analyte by a specific method can be distinguished from measurements of a blank ("zero"). 

The MDL is a widely used precision-based benchmark of laboratory method performance 
determined during method validation (and periodically reevaluated). As a benchmark it 
compares the sensitivity and precision of various methods within and between laboratories 
under optimum conditions (assuming that all the laboratories determine the MDLs consist- 
ently), but it says little about the day-to-day performance of a method. 

Detection limits are usually determined by analyses of replicate low-level spiked samples or 
blanks. A detection limit is laboratory specific as it is determined in a particular laboratory 
with its reagents, equipment, and analysts. Each sample will have its own detection limit, 


determined by the matrix of the sample. The more the matrix interferences in the sample, the 
higher the sample detection limit. 

One procedure to determine the MDL for an analyte is by performing seven or eight replicate 
analyses (n = 7 or 8) of the analyte at low concentration. The MDL is defined as t x sigma, 
where sigma is the standard deviation and / is the Student's t factor for a 99% probability 
level (t = 3 for n = 8). It can be reasoned that at 3 sigma concentration there is only about a 
1% chance of a false positive (assuming normal distribution). Still, at the concentration of 
3 sigma, there is about a 50% chance of a false negative if data are censored below that level 
and are treated as nondetections (see Section 5.3.2). 

Interpretation of data on trace constituents (e.g., metals, organics, and pesticides) is further 
complicated by data censoring (not reporting concentrations below a designated limit), 
nondetections, and variability and bias (less than 100% recovery). 

Other benchmarks besides MDL are discussed in the following sections. 

Reliable Detection Limit 

The reliable detection limit (RDL) is the lowest true concentration in a sample that can be 
reliably detected (Keith 1991). The most common definition is based on the same statistical 
principles as the MDL and is often defined as 6 sigma (2 x MDL), assuming sigma is 
constant. At this true concentration, the theoretical expected frequency of false negatives 
is reduced to 1% if measured values were censored at the MDL. Again, the RDL will vary 
from matrix to matrix and from sample to sample. For a different perspective, consult AOAC 
(1985), where the limit of reliable measurement is introduced. 

Limit of Quantification 

The concept of the limit of quantification (LOQ) is that measurements reported at or above 
this level meet a high standard for quantification, not just detection. Various multiples 
of sigma have been suggested; the higher the multiple, the greater the confidence in 
concentrations reported at or above this value. Commonly, the LOQ is defined at 10 sigma 
or 3.33 x MDL. At 10 sigma, the true concentration is within +30% of the reported 
concentration. The LOQ is equivalent to the practical quantitation limit (PQL). 

Caution is advised in using method-reporting limits, because many were established using 
the best estimates of the analytical chemists many years ago and may have little or no 
statistical basis. Reporting an MDL and a limit of quantitation limit along with low-level data 
alerts data users of the uncertainties and limitations associated with the data. A better way 
would be to report Y + U at any concentration Y found (i.e., no data censoring), where U is 
the calculated uncertainty at that concentration. 

5.2.7 Reporting Results and Estimates of Uncertainty 

A reported value from the laboratory analysis is an estimate of the true concentration in the 
sample at the time of collection. Thus, this measurement has variability associated with it 
referred to as measurement uncertainty. This uncertainty in the concentration of an analyte in a 
soil sample can be categorized into three general types of errors (Taylor 1988; Swyngedouw 
et al. 2004): 


Random errors that affect the precision of the results 
c errors that affect the bias 

• Blunders (mistakes that result in gross errors or lost samples — unpredictable and often 
yield unknown errors, i.e., the errors cannot be measured) 

Although errors due to blunders are mostly controlled through proper education and training, 
some will always occur. Data verification and validation attempt to detect and reduce these 
blunders. QC samples may also detect some types of blunders. 

Sampling and analytical errors do occur but are independent of each other. Therefore, 
sampling-related errors cannot be compensated for by the laboratory (AENV 2004). Thus, 
the limit of uncertainty for data on samples includes both the uncertainty of the sampling and 
of their measurement (Taylor 1988, 1997; Bevington and Robinson 2003) as indicated by the 
following equation: 

Estimates of uncertainty are obtained by a four-step process (Eurachem 2000): 

7 Specification of the analyte 

2 Identification of the uncertainty sources 

j Quantification of these uncertainty sources and 

4 Calculation of the combined uncertainty 

By combining uncertainty sources, only duplicate variance, long-term variance, and uncer- 
tainties in bias, calibration, and reference material need to be considered. These sources can 
be obtained from existing laboratory data, thus they are more easily quantified (Swyngedouw 
et al. 2004). 

An advantage of reporting realistic estimates of uncertainty together with measurements of 
concentration (Y ± U) is that end users of the analysis can consider the implications of the 
uncertainty in their use of the data. The traditional deterministic approach is to compare the 
measured concentration values with an appropriate regulatory threshold value. With this 
approach, any sampling point that has a reported concentration value below the threshold is 
classified as "uncontaminated" and those above as "contaminated." This approach does 
not account for uncertainty in the data. 


5.3.1 Overview 

DQOs specify requirements for analytical data that are clear and unambiguous concerning 
the intent of an investigation and the data parameters necessary to achieve that intent. These 
objectives are stated in both qualitative terms concerning the intended end use of the data 


as well as in quantitative terms with respect to precision, accuracy, representativeness, 
comparability, and completeness (USEPA 2000a). 

DQOs ensure that the proper methods and procedures (including method modifications) 
are in place with respect to MDLs, LOQs, or PQLs, applicable requirements, action 
limits, analyte specificity, analyte selectivity, reproducibility, false positives, and false 

The following issues or stages are important for developing DQOs: 

State the precise problem to be resolved. 

Identify all the decisions needed to resolve the problem. 

Identify all the inputs needed to make the decisions. 

NaiTow the boundaries of the project. 

Develop a decision rule. 

Develop uncertainty constraints. 

Optimize the design for obtaining data. 

These issues are often termed the "seven stages of DQO planning." Some of these 
stages can be further expanded as follows. Stage 1 asks "Are the analyses primarily 
for characterizing the soil (e.g., pH, organic matter, texture), or for determining contam- 
inant concentrations (e.g., metals, hydrocarbons, salts)?" or "Is the purpose of the soil 
analysis for screening or is it determinative?" or "Are average values of the chemicals 
of concern allowed?" Chemical analyses are conducted for a purpose; hence, decisions 
will be made based on the analytical results. Here, one needs to consider the general 
kind of decisions that will be made (Stage 2). Decisions involving health and safety of 
the public, impacts of pollutants on the environment, regulatory compliance, and other 
aspects need to be considered. In Stage 3 one needs to know what analytes need to be 
analyzed (i.e., what are the chemicals of concern), what the associated action levels are 
with the decisions of Stage 2, and what detection levels need to be achieved with each 

Since methods are specific for target analytes, a decision is required as to whether a 
particular method is appropriate or whether it will need to be modified to make it acceptable. 
Questions that need to be addressed involve the requirements for detection levels, method 
selectivity, accuracy, precision, and reproducibility (Table 5.3). These questions are 
addressed in the following sections. 

Method Sensitivity (Detection Levels) 

Estimating the lowest concentration levels needed to be achieved affects the available 
methods to choose from, the rates of false positive and false negative data, the ability to 
composite samples, and the number of samples required to meet the project DQOs. 


TABLE 5.3 Recommended Method Selection and Quality Control for Different Situations 

Method selection 

Alternative procedui 

Action level (or desired 
sensitivity) is close to the 

Matrix effects, 
contamination, and 

Contamination, procedural 
losses, need bias-free dat£ 

Need precision data 
(replicate agreement) 

Multiple operators, 

Need to ir 
confidence levels by 

Blanks for contamination, 
spikes, or surrogates for 
matrix effects and 

Spikes (spiked samples 
analyte recovery) 

Need to ir 
confidence or decrease the 
standard deviations 

Interlaboratory studies 

a bias-free method 

below the action level 

Choose a method with a 
specific detector that is not 
influenced by 

Choose a bias-free method 

Select a precise method 

Choose an accredited and 
audited method 

Increase the number of 
samples and field 

Run more blanks, 
laboratory control 
samples, or standard 
reference materials 

Increase replicates 

Choose another laboratory 


Method Selectivity 

Method selectivity directly affects the probability of detecting interferences in samples, 
especially in complex environmental samples. Interferences may cause an increase or 
decrease in signals of target analytes and thus lead to false positive or false negative 
conclusions. The tolerance for false positives and/or false negatives in the data is closely 
related to sample characteristics and method selectivity. 


Accuracy is a measure of how close an analytical result is to its true value. It has two 
components, bias and precision. 


To obtain overall precision (i.e., both sampling and analysis), field replicate samples need 
to be analyzed. Field replicate samples are two or more portions of a sample collected as 
close as possible at the same point in time and space to be considered identical. These samples 
are used to measure imprecision caused by inhomogeneity of the target analytes distributed in 
the soil. As imprecision increases, the relative standard deviation (RSD) will also increase. It is 
not unusual for the overall RSD to be larger than those of laboratory values. 


Reproducibility is the precision of measurements for the same sample at different labora- 
tories, or at the same laboratory but determined by a different analyst. Reproducible results 
are those that can be reproduced within acceptable and known limits of deviation and 
therefore demonstrate correct and consistent application of standard methodologies. 

5.3.2 Decision Errors 

As mentioned above, two potential decision errors are identified based on interpreting 
sampling and analytical data. 

False Positives (Decision Error B or False Acceptance) 

An important criterion in chemical analytical data is ensuring that a detected parameter is 
present. Equally important is determining whether the mean concentration in the study area 
is statistically significantly higher than the action level. In either of these situations, when 
incorrect conclusions are made, the result is a false positive, i.e., the wrong analytes are 
concluded to be present. Method blanks are used to demonstrate the absence of false 
positives. The consequences of decision error B would result in needless expenditure of 
resources to pursue additional actions and assessments. 

False Negatives (Decision Error A or False Rejection) 

Correctly concluding from analytical data that analytes are absent from samples is also 
important. Failing to detect a parameter when it is present is a false negative. Similarly, 
concluding that a mean analyte concentration in the study area is not statistically signifi- 
cantly higher than the action level, when it actually is, is also a false negative. False 
negatives are often the result of poor recovery of analytes from soil matrices or are caused 

by interferences that mask the analyte response. Method spikes (matrix spikes) are used to 
demonstrate the absence of false negatives. Minimization of false negatives is important with 
risk assessment and regulatory agencies. The consequences of decision error A would result 
in, for example, a health risk going undetected and unaddressed. 

Both decision errors need to be examined and a decision made as to which error poses the 
more severe consequence. As an example, the planning team may decide that the decision 
error A (false negative) poses more severe consequences, because the true state of soil 
contamination could go undetected and may cause health risks to neighborhood residents. 

Stage 6 of DQO planning sets acceptable limits for precision, accuracy, rates of false 
positives and/or false negative decision errors and for confidence levels in the sampling, 
and analytical data that relate to the DQOs. These decision error limits are set relative to the 
consequences of exceeding them (IAEA 2004). One could initially set the allowable decision 
errors to be at 1% (i.e., P = 0.01). This means that enough samples need to be collected and 
analyzed so that the chance of making either a false rejection (alpha) or a false acceptance 
(beta) decision error is only one out of a hundred. 


The type of QC samples to select depends on the DQOs of the site being investigated. 
Selections should be made depending on the following conditions (see Table 5.4): 

• Whether bias-free and/or precision data are required. 

• Whether differentiation between laboratory or sampling sources of error is needed. 

TABLE 5.4 Types of Quality Control Samples Used in the Field and Laboratory 
Purpose QC to use 

Field Check representativeness Field duplicates (precision) 

Check for matrix effects Surrogates, spikes, duplicates 

Check for contamination Blanks (field blanks, 

rinsate blanks) 
Slowdown the chemistry Holding times, lower 

temperature, appropriate 
containers, preservatives 

Laboratory Check representativeness Laboratory duplicates (from 

Check method bias Laboratory control samples, 

reference materials 
Check regulations (bias) Method detection levels 

(MDL), practical 
quantitation limits (PQL) 
Check comparability (with other Outside QC samples, e.g., 

laboratories) performance test (PT) 


Source: British Columbia Ministry of the Environment (BCME), 2003. 


• Whether the degree of error to be estimated is relatively small (e.g., from typical 
contamination type sources) or large (e.g., from operator and/or procedural sources). 

The methods selected need to be validated on soil matrices typical of those being received 
for analysis. Such validation does not guarantee that the methods will perform equally well 
for other soil types. In addition to unanticipated matrix effects, sampling artifacts, equipment 
malfunctions, and operator errors can also cause inaccuracies. Table 5.2 lists some sources of 
error that contribute to the uncertainty (variability) in analytical data. 

5.4.1 Impact of Bias on Test Results 

Bias is defined as the difference between the expected value of a statistic (e.g., sample 
average) and a population parameter (e.g., population mean). The need to take fewer 
replicates to reliably determine the mean value is an advantage in terms of cost and time. 
If no adjustment for bias is made, then for many purposes, the less biased, more vari- 
able method is preferable. However, by proper bias adjustment, the more precise method 
becomes the preferred method. Such adjustment can be based on QC check sample results 
(USEPA 2000b). 

5.4.2 Field Control Samples 

Field replicate, background, and rinsate (i.e., analyte-free water) blank samples are the most 
commonly collected field QA/QC samples for soil analysis. These are described in the 
following sections and are summarized in Table 5.4. 

Field Replicates 

Field replicates are field samples obtained from one location, homogenized and divided into 
separate containers and treated as separate samples throughout the remaining sample hand- 
ling and analytical processes. These samples are used to assess errors associated with sample 
heterogeneity, sample methodology, and analytical procedures. 

Equipment Rinsate Blanks 

A rinsate blank is a sample of analyte-free water run over or through decontaminated field 
sampling equipment before collection of the next sample. It is used to assess the adequacy of 
cleaning or decontamination processes in the sampling procedure. The blank is placed in 
sample containers for handling, shipment, and analysis identical to the field samples. 

Field Blanks 

A field blank is a sample of analyte-free media, similar to the sample matrix, which is 
transferred from one vessel to another or exposed to the sampling environment at the 
sampling site, and shipped to the laboratory with the field samples. It is used to evaluate 
contamination error associated with field operations and shipping, but may also be used to 
evaluate contamination error associated with laboratory procedures. 


Background Samples 

Background samples determine the natural composition of the soil, and are considered 
"clean" samples. Although background samples are not considered QC samples per se, 
they are best planned for along with the QC samples. They provide a basis for comparison of, 
for example, contaminant concentration levels with naturally occurring levels of target 
analytes in the soil samples collected on site. Again, if the objective does not involve 
whether a site is contaminated or not, then background samples are not needed. If back- 
ground samples are needed, they are collected first. 

Computer expert systems are available that help researchers collect the proper type of QC 
samples and then calculate how many of each sample type are needed to meet the stated 
DQOs (Keith 2002; Pulsipher et al. 2003). 

5.4.3 Laboratory QA and QC Procedures 

Internal QC monitors the laboratory's current performance versus the standards and criteria 
that have been set, normally at the time of method development or validation. 

To ensure that quality data are continuously produced during all analyses and to allow eventual 
review, systematic checks are performed to show that the test results remain reproducible. 
Such checks also show if the analytical method is measuring the quantity of target analytes 
in each sample within acceptable limits for bias (Environment Canada 2002a,b; USEPA 
2003; IUPAC 2005). Analytical QC procedures that determine whether the sample handling 
procedures and laboratory methods are performing as required are presented in Table 5.5. 

External laboratory QC involves reference help from other laboratories and participation 
in national or international interlaboratory sample and data exchange programs such as 
proficiency testing (PT). Such programs may involve: 

• Exchange of samples with another laboratory. These samples would be prepared by a 
staff member other than the analyst or by the QC department. Similarly, samples 
prepared by the QC department can be used as internal check samples. 

• Participation in interlaboratory sample exchange programs (such as round robins 
and/or PTs). Often in a PT study, the laboratory is not aware of samples used, 
in-house, for external performance evaluation. 

The necessary components of a complete QA/QC program include internal QC criteria 
that demonstrate acceptable levels of performance, as determined by a QA review 
(audit). External review of data and procedures is accomplished by the monitoring 
activities of accreditation organizations (SCC 2005). This includes laboratory evaluation 
samples (PT samples, see above) and a periodic (normally every 2 years) on- 
of all QA/QC procedures, performed by external assessors from the accreditin 


Data verification occurs after the data analyses are completed. Data verification is a rigorous 
process whereby QC parameters are evaluated against a set of predetermined criteria or 
functional guidelines. 

TABLE 5.5 Data Verification Checklist and Suggested Procedures 


What to check 

How to check 

Holding time (HT) 

Matrix spike (MS)/matrix 
spike duplicates (MSD)/ 
duplicates (DUP) 

Holding time 

:t for the data to be c 

Normally, only method blanks and any specific blank 
submitted with the samples will be reported. No blank 
should have a reportable concentration of any compound o 
interest above the reporting limit. Exceptions are the 
common laboratory contaminants' 1 

Surrogates only apply to organics at this time. Surrogates are 
compounds that are spiked (i.e., added at a known 
concentration) into every organic sample. A surrogate is a 
compound that is not found in nature and is not a "normal' 

Spikes, spike duplicates, and duplicates are used for both 
organic and inorganic data. Spikes are used to check for 
accuracy, while duplicates are a check for precision 

Reporting li 

i\ the method detectio 

Look at the chain of custody (COO attached to the report, 
check the sampling date and compare this to the 
extraction/digestion date (or just the analysis date if no 
preparation step is performed) given in the report. The 
number of days must be less than or equal to the 
required HT 

Look at the blank reports. Any compound that has a 
concentration reported above the reporting limit in the blank 
and is present in any sample must be considered estimated 
or a nondetect at concentrations up to five times the level in 
the blank (up to 10 times for the common laboratory 

Check the report for surrogate recoveries. They should appear 
at the end of the analytical compounds list for a method. The 
recoveries should be 30%-1 50% to be acceptable. If the 
surrogate recovery is low, then flag positive values reported 
and reject nondetects. If it is high, then nondetect data are 
considered acceptable and positive data are flagged as 

The MS/MSD/DUP results should appear at the end of the 
compound list. Verify that the recoveries are reasonable. 
Some values are: organic analysis (30%-150% recovery); 
inorganic analysis (80%-120% recovery); and duplicates 
(<50% relative percent difference 13 ) 

Look at the reporting limits. The limits should meet the 
requirements for the site. The limits for soils vary 
considerably depending on the method 

Reporting limits are used by laboratories as a level of 
confidence in reporting a concentration. Sometimes the 
practical quantitation limit (PQL) is used, which is 2 to 10 

times the MDL 

Common laboratory contaminants often include phthalates, dichloromethane, acetone, 2-butanone, hexanone, zinc, and iron. 
Relative percent difference (RPD) = \X - Y\ x 200/(X + Y), where X and Y are the concentrations of each duplicate. 


Data quality can be measured in several ways and these form the basis of deciding whether 
the DQOs have been met: 

• Rates (%) of false positive and negatives in the analytical data 

• Precision (closeness of values from repeat analyses — expressed as standard deviation) 

• Bias (i.e., accuracy) 

• Estimation of the uncertainty of the results 

This type of information can be used to improve the quality of data interpretation. It is useful 
to analyze the QC data first and then review the sample data. Typical practices for analyzing 
QC data are presented in Table 5.5. More information on data verification is available in the 
literature (e.g., USEPA 1996). 

5.5.1 Statistical Control 

Besides documenting uncertainty, descriptive statistics from an established QA program can 
be used to determine if a methodology is in "statistical control," i.e., whether QC criteria 
are being met over the long term. Check sample statistics are also used as daily decision- 
making tools during sample analysis to determine if expected results are being generated and 
if the analytical system is functioning properly (AOAC 1985). As described earlier, QC 
provides information to determine sample and laboratory data quality using data trend 
analysis (i.e., statistical process control). Statistical reports that evaluate specific anomalies 
or disclose trends in many areas are commonly generated (AOAC 1985; Kelly et al. 1992; 
FAO 1998; Garfield et al. 2000). 

These trend analysis techniques are used to monitor the laboratory's performance over time, 
to detect departures of the laboratory's output from required or desired levels of QC, and to 
provide an early warning of QA or QC problems that may not be apparent from the results of 
an individual case. 

Trend analyses also provide information needed to establish performance-based criteria for 
updated analytical protocols, in cases where advisory criteria were previously used (control 

5.5.2 Control Charts 

Quality assessment statistics can be presented graphically through control charts for ease of 
interpretation. These charts can be used to present both bias and precision data. Repeated 
measurements of external or internal reference or QC samples are graphed on a time line. 
Superimposed on the individual results is the cumulative mean or the known value. Control 
levels which typically represent ±2 sigma (upper and lower warning limits, UWL and 
LWL) and +3 sigma (upper and lower control limits, UCL and LCL) from the mean are 
also included (see Figure 5.2). In a normally distributed sample population, the warning levels 
represent a 95% confidence interval, while the control limits correspond to a 99% confidence 
interval. As an example, a single value outside the UCL or LCL is considered unacceptable. If 
statistical control is considered unacceptable, all routine sample unknowns between the 
unacceptable check sample(s) and the last check sample that was in control should be rerun. 


F1 (C6-C10) Hydrocarbons control chart 

FIGURE 5.2. Example of a control chart. (UCL, upper control level, mean +3 x standard devi- 
ation of values; UWL, upper warning level, mean +2 x standard deviation of 
values; mean, average of values; LWL, lower warning level, mean -2 x standard 
deviation of values; LCL, lower control level, mean -3 x standard deviation 
of values.) 

5.5.3 Trace of Test 

When data quality is not achieved, a "trace of test" is a good verification tool. A systematic 
approach is applied in this test, starting with a check for calculation and typing errors. Items 
that are checked include samples, standards, reagents, equipment, glassware, and the ana- 
lytical instruments and their calibrations. Then the method itself is checked, focusing on 
method validation factors such as sensitivity (detection limits), precision, recovery, 
and interferences. Batch control is also checked including laboratory control samples and 
reference materials used, and inspection of control charts and feedback logs (e.g., complaints). 

The order of e 
as follows: 

n the investigation is the n 

e of that given in Figure 5.1 and could be 

I Confirm that the results were correctly reported and correctly associated to the specific 

Recheck the results and confirm that they have been calculated correctly. 

Verify analytical QC associated with the test to ensure the measurement process was it 
statistical control. 

Investigate deviations from the routine procedure and the data record. 


Investigate any nonconformance relating to the sample such as matrix effects and 
holding times. 

Determine whether the results make sense: compare the results to other analyses, 
compare to historical data (if known), and/or communicate with the data user. 
Computer data checks can be built-in functions of laboratory databases, models, or 
spreadsheets. Automated QA/QC can be used to facilitate peer review or, in some 
cases, manual checks. 

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American Society for Testing and Materials 
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Samples Containing Organic Constituents. 
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Sciences, 3rd edn. McGraw-Hill, Toronto. 

British Columbia Ministry of the Environment 
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Water, Air and Climate Change Branch 2003. 
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labsys/field_man_03.html#pdf (verified February 
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(verified March 2005). 

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Environment Canada. 2002a. Contaminated Site 
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[verified March 2005]. This Webpage shows a list 
of Technical Assistance Bulletins. See Section 5: 
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Environment Canada. 2002b. Guidance Docu- 
ment for the Sampling and Analysis of Metal 
Mining Effluents. [Online] Available at: http:// 
(verified October 2004). 

Eunichcm. 2(100. Euniclicm Guide: Qualifying 
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Garfield, F.M., Klesta, E„ and Hirsch, J. 2000. 
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Gerlach, R., Dobb, D.E., Raab, G.A., and Nocerino, 
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International Atomic Energy Agency (IAEA). 
2004. Soil Sampling for Environmental Con- 
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Atomic Energy Agency, Vienna. Worked DQOs 
examples p. 39. 

International Organization for Standardization 
(ISO). 2005. ISO/IEC 17025: 2005 General 
Requirements for the Competence of Testing and 
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3&ICS2 = 120&ICS3 = 20 (01 March 2006). 

International Union of Pure and Applied Chemis- 
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[Online] Available at: 
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International Union of Pure and Applied Chemis- 
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Jenkins, T.F., Walsh, M.E., Thorne, P.G., 
Thiboutot, S., Ampleman, G., Ranney, T.A., and 
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Associated with Collection and Analysis of Soil 
Samples at a Firing Range Contaminated with 
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Keith, L.H. 1991. Environmental Sampling and 
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Keith, L.H. 2002. Environmental Monitoring 
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Section Editors: J.J. Schoenau and LP. O'Halloran 

Chapter 6 

Nitrate and Exchangeable 

Ammonium Nitrogen 

D.G. Maynard 

Natural Resources Canada 
Victoria, British Columbia, Canada 

Y.P. Kalra and J.A. Crumbaugh 

Natural Resources Canada 
Edmonton, Alberta, Canada 


Inorganic N in soils is predominantly in the form of nitrate (NO3) and ammonium (NH4). 
Nitrite is seldom present in detectable amounts, and its determination is normally unwar- 
ranted except in neutral to alkaline soils receiving NH4 and NH 4 -producing fertilizers 
(Keeney and Nelson 1982). Soil testing laboratories usually determine NO3 to estimate 
available N in agricultural soils, while laboratories analyzing tree nursery and forest soils 
often determine both NO3 and NH4. 

There is considerable diversity among laboratories in the extraction and determination 
of NO3 and NH 4 . In addition, incubation methods (both aerobic and anaerobic) have 
been used to determine the potentially mineralizable N (see Chapter 46) and nitrogen 
supply rates using ion exchange resins (see Chapter 13). 

Nitrate is water-soluble and a number of solutions including water have been used as 
extractants. Exchangeable NH4 is defined as NH4 that can be extracted at room temp- 
erature with a neutral K salt solution. Various molarities have been used, such as 
0.05 M K2SO4, 0.1 M KC1, 1.0 M KC1, and 2.0 M KC1 (Keeney and Nelson 1982). The 
most common extractant for NO3 and NH4, however, is 2.0 M KC1 (e.g., Magill and Aber 
2000; Shahandeh et al. 2005). 

The methods of determination for NO3 and NH4 are even more diverse than the 
methods of extraction (Keeney and Nelson 1982). These range from specific ion electrode 
to manual colorimetric techniques, microdiffusion, steam distillation, and continuous 
flow analysis. Steam distillation is still sometimes employed for 15 N; however, for routine 


analysis automated colorimetric techniques using continuous flow analyzers are preferred. 
Segmented flow analysis (SFA) and flow injection analysis (FIA) are continuous flow 
systems that are rapid, free from most soil interferences, and very sensitive. 

The methods for the most commonly used extractant (2.0 M KC1) and SFA methods for the 
determination of NO3 and NH4 are presented here. The FIA methods often use the same 
chemical reactions but with different instruments (e.g., Burt 2004). The steam distillation 
methods for determination of NO3 and NH 4 have not been included, since they have not 
changed much over the last several years. Detailed description of these methods can be found 
elsewhere (Bremner 1965; Keeney and Nelson 1982). 


6.2.1 Principle 

Ammonium is held in an exchangeable form in soils in the same manner as exchange- 
able metallic cations. Fixed or nonexchangeable NH4 can make up a significant portion 
of soil N; however, fixed NH4 is defined as the NH4 in soil that cannot be replaced by a 
neutral K salt solution (Keeney and Nelson 1982). Exchangeable NH4 is extracted by shaking 
with 2.0 M KCI. Nitrate is water-soluble and hence can also be extracted by the same 
2.0 M KCI extract. Nitrite is seldom present in detectable amounts in soil and therefore is 
usually not determined. 

6.2.2 Materials and Reagents 

/ Reciprocating shaker. 

2 Dispensing bottle. 

2 Erlenmeyer flasks, 125 ml_. 

4 Nalgene bottles, 60 ml_. 

5 Filter funnels. 

5 Whatman No. 42 filter papers. 

7 Aluminum dishes. 

8 Potassium chloride (2.0 M KCI): dissolve 149 g KCI in approximately 800 ml_ 
NH 3 -free deionized H 2 in a 1 L volumetric flask and dilute to volume with 
deionized H 2 0. 

6.2.3 Procedure 

A. Moisture determination 

7 Weigh 5.00 g of moist soil in a preweighed aluminum dish. 


2 Dry overnight in an oven at 1 05°C. 
j Cool in a desiccator and weigh. 
B. Extraction procedure 

1 Weigh (5.0 g) field-moist soil (or moist soil incubated for mineralization 
experiments) into a 125 mL Erlenmeyer flask. In some instances air-dried soil 
may also be used (see Comment 1 in Section 6.2.4). 

2 Add 50 mL 2.0 M KCI solution using the dispensing bottle. (If the sample is 
limited, it can be reduced to a minimum of 1 .0 g and 1 mL to keep 1 :1 ratio.) 

j Carry a reagent blank throughout the procedure. 

4 Stopper the flasks and shake for 30 min at 1 60 strokes per minute. 

5 Filter through Whatman No. 42 filter paper into 60 mL Nalgene bottles. 

g Analyze for NO3 and NH 4 within 24 h (see Comment 3 in Section 6.2.4). 

1 Significant changes in the amounts of NO3 and NH 4 can take place with 
prolonged storage of air-dried samples at room temperature. A study conducted 
by the Western Enviro-Agricultural Laboratory Association showed that the NO3 
content of soils decreased significantly after a 3-year storage of air-dried samples 
at room temperature (unpublished results). Increases in NH 4 content have also 
been reported by Bremner (1965) and Selmer-Olsen (1971). 

2 Filter paper can contain significant amounts of NO3 and NH 4 that can potentially 
contaminate extracts (Muneta 1980; Heffernan 1985; Sparrow and Masiak 1987). 

3 Ammonium and NO3 in KCI extracts should be determined within 24 h of 
extraction (Keeney and Nelson 1982). If the extracts cannot be analyzed imme- 
diately they should be frozen. Potassium chloride extracts keep indefinitely when 
frozen (Heffernan 1985). 

4 This method yields highly reproducible results. 




6.3.1 Principle 

Nitrate is determined by an automated spectrophotometric method. Nitrates are reduced to 
nitrite by a copper cadmium reductor coil (CRC). The nitrite ion reacts with sulfanilamide 


under acidic conditions to form a diazo compound. This couples with AM-naphthyl- 
ethylenediamine dihydrochloride to form a reddish purple azo dye (Technicon Instrument 
Corporation 1971). 

6.3.2 Materials and Reagents 

/ Technicon AutoAnalyzer consisting of sampler, manifold, proportioning pump, 
CRC, colorimeter, and data acquisition system. 

2 CRC— activation of CRC (O.I. Analytical 2001a)— Refer to point 5 in this 
section for CRC reagent preparation. This procedure must be performed before 
connecting the CRC to the system. Do not induce air into CRC during the 
activation process (see Comment 6 in Section 6.3.5 regarding the efficiency 
of the CRC). 

a. Using a 1 mL Luer-Lok syringe and a 1 /4"-28 female Luer-Lok fitting, slowly 
flush the CRC with 1 mL of deionized H 2 0. If any debris is seen exiting the 
CRC, continue to flush with deionized H 2 until all debris is removed. 

b. Slowly flush the CRC with 1 mL of 0.5 M HCI solution. Quickly proceed to 
the next step as the HCI solution can cause damage to the cadmium surface if 
left in the CRC for more than a few seconds. 

c. Flush the CRC with 1 mL of deionized H2O to remove the HCI solution. 

d. Slowly flush the CRC with 10 mL of 2% cupric sulfate solution. Leave this 
solution in the CRC for approximately 5-10 min. 

e. Forcefully flush the CRC with 1 mL of NH 4 CI reagent solution to remove any 
loose copper that may have formed within the reactor. Continue to flush until 
all debris is removed. 

f. The CRC should be stored and filled with deionized H 2 when not in use. 

Note: Solution containing Brij-35 should not be used when flushing or storing 
the CRC. 

Nofe: Do not allow any solutions other than deionized H 2 and reagents to 
flow through the CRC. Some solutions may cause irreversible damage to the 

a. Stock solution (100 juLg NO3-N mL -1 ): dissolve 0.7218 g of KNO3 (dried 
overnight at 105°C) in a 1 L volumetric flask containing deionized H 2 0. Add 
1 mL of chloroform to preserve the solution. Dilute to 1 L and mix well. 

b. Working standards: pipet 0.5, 1.0, 1.5, and 2.0 mL of stock solution into a 
1 00 mL volumetric flask and make to volume with 2.0 M KCI solution to obtain 
0.5, 1.0, 1.5, and 2.0 |xg NO3-N mL~ 1 standard solution, respectively. 


4 Reagents 

a. Dilute ammonium hydroxide (NH 4 OH) solution: add four or five drops of 
concentrated NH 4 OH to approximately 30 mL of deionized H 2 0. 

b. Ammonium chloride reagent: dissolve 10 g NH 4 CI in a 1 L volumetric flask 
containing about 750 mL of deionized H 2 0. Add dilute NH 4 OH to attain a pH 
of 8.5, add 0.5 mL of Brij-35, dilute to 1 L, and mix well. {Note: it takes only 
two drops of dilute NH 4 OH to achieve the desired pH.) 

c. Color reagent: to a 1 L volumetric flask containing about 750 mL of deionized 
H 2 0, carefully add 100 mL of concentrated H3PCU (see Comment 2 in 
Section 6.3.5) and 10 g of sulfanilamide. Dissolve completely. Add 0.5 g of N- 
1-naphthyl-ethylenediamine dihydrochloride (Marshall's reagent), and dis- 
solve. Dilute to 1 L volume with deionized H 2 and mix well. Add 0.5 mL 
of Brij-35. Store in an amber glass bottle. This reagent is stable for 1 month. 

5 Reagents for CRC 

a. Cupric sulfate solution (2% w/v): dissolve 20 g of CuS0 4 ■ 5H 2 in approxi- 
mately 900 mL of deionized H 2 in a 1 L volumetric flask. Dilute the solution 
to 1 L with deionized H 2 and mix well. 

b. Hydrochloric acid solution (0.5 M): carefully add 4.15 mL of concentrated 
HCI to approximately 70 mL of deionized H 2 in a 100 mL volumetric 
flask (see Comment 2 in Section 6.3.5). Dilute to 100 mL with deionized 
H 2 and mix well. 

6.3.3 Procedure 

7 If refrigerated, bring the soil extracts to room temperature. 

2 Shake extracts well. 

j Set up AutoAnalyzer (see Maynard and Kalra 1993; Kalra and Maynard 1991). 
Allow the colorimeter to warm up for at least 30 min. 

4 Place all reagent tubing in deionized H 2 and run for 10 min. 

5 Insert tubing in correct reagents and run for 20 min to ensure thorough flushing of 
the system (feed 2.0 M KCI through the wash line). 

6 Establish a stable baseline. 

7 Place the sample tubing in the high standard for 5 min. 

8 Reset the baseline, if necessary. 

g Transfer standard solutions to sample cups and arrange on the tray in descending 


10 Transfer sample extracts to sample cups and place in the sample tray following the 

11 Begin run. 

12 After run is complete, rerun the standards to ensure that there has been no drifting. 
Reestablish baseline. 

13 Place tubing in deionized H2O, rinse and run for 20 min before turning the 
proportioning pump off. 

6.3.4 Calculation 

Prepare a standard curve from recorded readings (absorption vs. concentration) of standards 
and read as |xg N0 3 -N mL -1 in KC1 extract. Results are calculated as follows: 

t soil (g) 


Moist soil (g) 

Moisture factor = — — - =■ — (6.2) 

Oven-dried soil (g) 

NO3-N in oven-dried soil (jjug g _1 ) = NO3-N in moist soil (|xg g _1 ) x moisture factor 


There are data collection software packages associated with the data acquisition systems and 
these will automatically generate calculated concentration values based on intensities 
received from the colorimeter and inputs of the appropriate information (e.g., sample weight, 
extract volumes, and moisture factor). 


1 Use deionized H 2 throughout the procedure. 

2 Warning: Mixing concentrated acids and water produces a great amount of heat. 
Take appropriate precautions. 

j All reagent bottles, sample cups, and new pump tubing should be rinsed with 
approximately 1 M HCI. 

4 Range: 0.01-2 |uug NO3-N ml_~ 1 extract. Extracts with NO3 concentrations 
greater than the high standard (2.0 |xg NO3-N ml_~ 1 ) should be diluted with 
2.0 M KCI solution and reanalyzed. 

5 Prepared CRCs can be purchased from various instrument/parts supplies for SFA 
systems. Previously, the method called for preparation of a cadmium reductor 


column. However, preparation was tedious and time consuming and cadmium 
granules are no longer readily available. 

6 Reduction efficiency of the CRC (O.I. Analytical 2001 a). 

a. In the CRC, nitrate is reduced to nitrite. However, under some conditions, 
reduction may proceed further with nitrite being reduced to hydroxylamine 
and ammonium ion. These reactions are pH-dependent: 

N0 3 + 2H+ + 2e^ N0 2 + H 2 (6.4) 

N0 2 + 6H+ + 6e^ H 3 NOH + H 2 (6.5) 

N0 2 + 8H+ + 6e^ NH+ + 2H 2 (6.6) 

At the buffered pH of this method, reaction 6.4 predominates. However, if the 
cadmium surface is overly active, reaction 6.5 and reaction 6.6 will proceed 
sufficiently to give low results of nitrite. 

b. If the cadmium surface is insufficiently active, there will be a low recovery of 
nitrate as nitrite. This condition is defined as poor reduction efficiency. 

c. To determine the reduction efficiency, run a high-level nitrite calibrant fol- 
lowed by a nitrate calibrant of the same nominal concentration. The reduction 
efficiency is calculated as given below. 

PR= (N3/N2) x 100 (6.7) 

where PR is the percent reduction efficiency, N 3 is the nitrate peak height, and 
N 2 is the nitrite peak height. 

d. If the response of the nitrite is as expected but the reduction efficiency is less 
than 90%, then the CRC may need to be reactivated. 

j The method includes NO3-N plus NO2-N; therefore, samples containing signifi- 
cant amounts of NO2-N will result in the overestimation of NO3-N. 

g The method given in this section outlines the configuration of the Technicon 
AutoAnalyzer. However, the cadmium reduction method can be applied to 
other SFA and FIA systems. 

6.3.6 Precision and Accuracy 

There are no standard reference samples for accuracy determination. Precision measure- 
ments for NO3-N carried out for soil test quality assurance program of the Alberta Institute of 
Pedology (Heaney et al. 1988) indicated that NO3-N was one of the most variable parameters 
measured. Coefficient of variation ranged from 4.8% to 30.4% for samples with 67.3 + 3.2 
(SD) and 3.3 + 1.0 (SD) (jug NO3-N g -1 , respectively. 





6.4.1 Principle 

Ammonium is determined by an automated spectrophotometric method utilizing the 
Berthelot reaction (Searle 1984). Phenol and NH 4 react to form an intense blue color. 
The intensity of color is proportional to the NH4 present. Sodium hypochlorite 
and sodium nitroprusside solutions are used as oxidant and catalyst, respectively 
(O.I. Analytical 2001b). 

6.4.2 Materials and Reagents 

1 Technicon AutoAnalyzer consisting of sampler, manifold, proportioning pump, 
heating bath, colorimeter, and data acquisition system. 

2 Standard solutions: 

a. Stock solution #1 (1000 |xg NH 4 -N mL~ 1 ):ina1 L volumetric flask containing 
about 800 ml_ of deionized H 2 dissolve 4.71 70 g (NH 4 ) 2 S0 4 (dried at 
105°C). Dilute to 1 L with deionized H 2 0, mix well, and store the solution 
in a refrigerator. 

b. Stock solution #2 (100 (jug NH 4 -N ml_~ 1 ): dilute 10 ml_ of stock solution #1 to 
100 ml_ with 2.0 M KCI solution. Store the solution in a refrigerator. 

c. Working standards: transfer 0, 1 , 2, 5, 7, and 1 ml_ of stock solution #2 to 1 00 
ml_ volumetric flasks. Make to volume with 2.0 M KCI. This will provide 0, 1 , 2, 
5, 7, and 10 jjLg NH 4 -N ml_~ 1 standard solutions, respectively. Prepare daily. 

j Complexing reagent: in a 1 L flask containing about 950 ml_ of deionized H 2 0, 
dissolve 33 g of potassium sodium tartrate (KNaC 4 H 4 6 ■ H 2 0) and 24 g of sodium 
citrate (HOC(COONa)(CH 2 COONa) 2 • H 2 0). Adjust to pH 5.0 with concentrated 
H 2 S0 4 , add 0.5 ml_ of Brij-35, dilute to volume with deionized H 2 0, and mix well. 

4 Alkaline phenol: using a 1 L Erlenmeyer flask, dissolve 83 g of phenol in 50 mL of 
deionized H 2 0. Cautiously add, in small increments with agitation, 1 80 mL of 20% 
(5 M) NaOH. Dilute to 1 L with deionized H 2 0. Store alkaline phenol reagent in an 
amber bottle. (To make 20% NaOH, dissolve 200 g of NaOH and dilute to 1 L with 
deionized H 2 0.) 

5 Sodium hypochlorite (NaOCI): dilute 200 mL of household bleach (5.25% 
NaOCI) to 1 L using deionized H 2 0. This reagent must be prepared daily, 
immediately before use to obtain optimum results. The NaOCI concentration in 
this reagent decreases on standing. 

6 Sodium nitroprusside: dissolve 0.5 g of sodium nitroprusside (Na 2 Fe(CN)s 
NO • 2H 2 0) in 900 mL of deionized H 2 and dilute to 1 L. Store in dark-colored 
bottle in a refrigerator. 


6.4.3 Procedure 

Follow the procedure (6.3.3) outlined for NO3-N (see Kalra and Maynard 1991; Maynard 
and Kalra 1993). 

6.4.4 Calculation 

The calculations are the same as given in 6.3.4. 


1 Use NH 4 -free deionized H 2 throughout the procedure. 

2 All reagent bottles, sample cups, and new pump tubing should be rinsed with 
approximately 1 M HCI. 

3 Range: 0.01-10.0 |xg NH 4 -N ml_~ 1 extract. Extracts with NH 4 concentrations 
greater than the high standard (10.0 (jug NH 4 -N mL~ 1 ) should be diluted with 
2.0 M KCI solution and reanalyzed. 

4 It is critical that the operating temperature is 50°C + 1°C. 

5 The method given in this section outlines the configuration of the Technicon 
AutoAnalyzer (Technicon Instrument Corporation 1973). However, the phenate 
method can be applied to other SFA and FIA systems. 

6.4.6 Precision and Accuracy 

There are no standard reference samples for accuracy determination. Long-term analyses of 
laboratory samples gave coefficient of variations of 21%-24% for several samples over a 
wide range of concentrations. 

Bremner, J.M. 1965. Inorganic forms of nitrogen. 
In C.A. Black, D.D. Evans, J.L. White, 
E. Ensminger, and F.E. Clark, Eds. Methods 
of Soils Analysis. Part 2. Agronomy No. 9. 
American Society of Agronomy, Madison, WI, 
pp. 1179-1237. 

Burt, R. (Ed.) 2004. Soil Survey Laboratory 
Methods Manual. Soil Survey Investigations 
Report No. 42, Version 4.0. United States Depart- 
ment of Agriculture, Natural Resources Conser- 
vation Service, Lincoln, NE, 700 pp. 

Heaney, D.J., McGill, W. 
1988. Soil test quality 

and Nguyen, C 

Unpublished report. Alberta Institute of 
Pedology, Edmonton, AB, Canada. 

Heffernan, B. 1985. A Handbook of Methods of 
Inorganic Chemical Analysis for Forest Soils, 
Foliage and Water. Division of Forest Research, 
CSIRO, Canberra, Australia, 281 pp. 

Kalra, Y.P. and Maynard, D.G. 1991. Methods 
Manual for Forest Soil and Plant Analysis. Infor- 
mation Report NOR-X-319. Northern Forestry 
Centre, Northwest Region, Forestry Canada. 
Edmonton, AB, Canada, 116 pp. Access online 
(July 2006). 


Keeney, D.R. and Nelson, D.W. 1982. Nitrogen 
in organic forms. In A.L. Page, R.H. Miller, and 
D.R. Keeney, Eds. Methods of Soil Analysis. 
Part 2. Agronomy No. 9, American Society of 
Agronomy, Madison, WI, pp. 643-698. 

Magill, A.H. and Aber, J.D. 2000. Variation in soil 
net mineralization rates with dissolved organic car- 
bon additions. Soil Biol. Bi i < ., :n '■ 1 97-601. 

Maynard, D.G. and Kalra, Y.P. 1993. Nitrate and 
extractable ammonium nitrogen. In M.R. Carter, 
Ed. Soil Sampling and Methods of Analysis. 
Lewis Publishers, Boca Raton, FL, pp. 25-38. 

Muneta, P. 1980. Analytical errors resulting from 
nitrate contamination of filter paper. ./. Assoc. Off. 
Anal. Chem. 63: 937-938. 

O.I. Analytical. 2001a. Nitrate plus nitrite nitro- 
gen and nitrite nitrogen in soil and plant extracts 
by segmented flow analysis (SFA). Publication 
No. 15300301. College Station, TX, 27 pp. 

O.I. Analytical. 2001b. Ammonia in soil and plant 
extracts by segmented flow analysis (SFA). Publi- 
cation No. 15330501. College Station, TX, 17 pp. 

Searle, P.L. 1984. The Berthelot or indophenol 
reaction and its use in the analytical chemistry of 
nitrogen: a review. Analyst 109: 549-568. 

Selmer-Olsen, A.R. 1971. Determination of am- 
monium in soil extracts by an automated indophe- 
nol method. Analyst 96: 565-568. 

Shahandeh, H., Wright, A.L., Hons, F.M., and 
Lascano, R.J. 2005. Spatial and temporal vari- 
ation in soil nitrogen parameters related to soil 
texture and corn yield. Agron. J. 97: 772-782. 

Sparrow, S.D. and Masiak, D.T. 1987. Errors in 
analysis for ammonium and nitrate caused by 
contamination from filter papers. Soil Sci. Soc. 
Am. J. 51: 107-110. 

Technicon Instrument Corporation 1971. Nitrate 
and Nitrite in Water. Industrial method No. 32- 
69W. Technicon Instrument Corporation, Tarry- 
town, New York, NY. 

Technicon Instrument Corporation 1973. Ammo- 
nia in Water and Seawater. Industrial method 
No. 154-71W. Technicon Instrument Corporation, 
Tarrytown, New York, NY. 

Chapter 7 
Mehlich 3-Extractable Elements 

N. Ziadi 

Agrk allure and Agri-rood Canada 
Quebec, Quebec, Canada 

T. Sen Tran 

Institute of Research and Development 

in Agroenvironment 

Quebec, Quebec, Canada 


During the past few years, numerous techniques and methods have been developed to 
estimate soil nutrient availability. Among these methods, the Mehlich 3 (M3) is considered 
an appropriate and economic chemical method since it is suitable for a wide range of soils 
and can serve as a "universal" soil test extractant (Sims 1989; Zbiral 2000a; Bolland et al. 
2003). M3 was developed by Mehlich (1984) as multielement soil extraction and is widely 
used, especially in agronomic studies, to evaluate soil nutrient status and establish fertilizer 
recommendations mainly for P and K in humid regions. The following elements can be 
successfully analyzed using M3 extracting solution: P, K, Ca, Mg, Na, Cu, Zn, Mn, B, Al, 
and Fe. The extracting solution is composed of 0.2 M CH 3 COOH, 0.25 M NH4NO3, 
0.015 M NH 4 F, 0.013 M HNO3, and 0.001 M ethylene diamine tetraacetic acid (EDTA). 
M3-extractable phosphorus (M3-P) is obtained by the action of acetic acid and fluoride 
compounds, while K, Ca, Mg, and Na (M3-K, M3-Ca, M3-Mg, and M3-Na, respectively) are 
removed by the action of ammonium nitrate and nitric acid. The Cu, Zn, Mn, and Fe (M3-Cu, 
M-Zn, M3-Mn, and M3-Fe) are extracted by NH 4 and the chelating agent EDTA. 

Many studies have compared the M3 method to other chemical and nonchemical methods 
and reported significant correlations between tested methods (Zbiral and Nemec 2000; Cox 
2001; Bolland et al. 2003). Indeed, M3-P is closely related to P extracted by M2, Bray 1, 
Bray 2, Olsen, strontium chloride-citric acid, and water (Mehlich 1984; Simard et al. 1991; 
Zbiral and Nemec 2002). In a study conducted in Quebec, Tran et al. (1990) reported that the 
amount of M3-P is approximately the same as that determined by the Bray 1 method on most 
noncalcareous soils. Recently, Mallarino (2003) concluded that M3 test is more effective 
than the Bray test for predicting corn (Zea mays L.) response to P across many Iowa soils 
with pH values ranging from 5.2 to 8.2. A good correlation was also obtained between M3-P 
and P desorbed by anionic exchange membranes and electroultrafiltration (EUF) techniques 


(Tran et al. 1992a,b; Ziadi et al. 2001). Many studies reported a strong correlation between 
M3-P and plant P uptake or between M3-P and relative plant yield in a wide range of soils 
(Tran and Giroux 1987; Ziadi et al. 2001; Mallarino 2003). Others, however, have indicated 
that some alkaline extractants (i.e., NaHCC>3) are superior to acidic extractants (M3) when 
used to evaluate plant P availability (Bates 1990). Depending on the determination method 
used, the critical level of M3-P for most common crops is about 30 to 60 (xg g~ : (Sims 1989; 
Tran and Giroux 1989; Bolland et al. 2003). 

In addition to its value in agronomic studies, M3-P has also been used in environmental 
studies as an agrienvironmental soil test for P (Sims 1993; Sharpley et al. 1996; Beauchemin 
et al. 2003). The concept of P saturation degree was developed and successfully used in 
Europe and North America to indicate the potential desorbability of soil P (Breeuwsma and 
Reijerink 1992; Beauchemin and Simard 2000). In the mid- Atlantic USA region, Sims et al. 
(2002) reported that the M3-P/(M3-A1 + M3-Fe) can be used to predict runoff and leachate 
P concentration. In a study conducted in Quebec, Khiari et al. (2000) reported that the 
environmentally critical (M3-P/M3-A1) percentage was 15%, corresponding to the critical 
degree of phosphate saturation of 25% proposed in Netherlands using oxalate extraction method 
(Van der Zee et al. 1987). In Quebec, the ratio of M3-extractable P to Al (M3-P/M3-A1) 
has been recently introduced in the local recommendation in corn production (CRAAQ 
2003). The reader is referred to Chapter 14 for a more complete description of environmental 
soil P indices. 

In addition to P, significant correlations have been obtained between the other nutrients 
(K, Ca, Mg, Na, Cu, Zn, Mn, Fe, and B) extracted by the M3 solution and other methods 
currently used in different laboratories (Tran 1989; Cancela et al. 2002; Mylavarapu et al. 
2002). Furthermore, Michaelson et al. (1987) reported significant correlation between the 
amounts of K, Ca, and Mg extracted by M3 and by ammonium acetate. Highly significant 
correlations have also been reported between M3-extractable amounts of Cu, Zn, Mn, Fe, and 
B and those obtained by the double acid, diethylene triamine pentaacetic acid-triethanolamine 
(DTPA-TEA),or0.1MHCl,Mehlich 1 (Sims 1989; Simsetal. 1991; Zbir aland Nemec 2000). 

The use of automated methods to quantify soil nutrients has expanded rapidly since the early 
1990s (Munter 1990; Jones 1998). The inductively coupled plasma (ICP) emission spectros- 
copy is becoming one of the most popular instruments used in routine soil testing labora- 
tories. The ICP instruments (optical emission spectroscopy [OES] or mass spectroscopy 
[MS]) are advantageous because they are able to quantify many nutrients (P, K, Ca, Mg, and 
micronutrients) in one analytical process. However, there has been criticism on the adoption 
of ICP, especially for P, instead of colorimetric methods which have been historically used in 
soil test calibrations for fertilizer recommendations (Mallarino and Sawyer 2000; Zbiral 
2000b; Sikora et al. 2005). Because of observed differences between P values obtained by 
ICP and by colorimetric methods, some regions in the United States do not recommend the 
use of ICP to determine P in any soil test extracts (Mallarino and Sawyer 2000). Zbiral 
(2000b) reported a small, but significant difference (2% to 8%) for K and Mg determined by 
ICP-OES and flame atomic absorption. In the same experiment, the amount of P determined 
by ICP-OES was higher by 8% to 14% than that obtained by the spectrophotometric method. 
Recently, Sikora et al. (2005) confirmed these results when they compared M3-P measured 
by ICP with that by colorimetric method, and concluded that further research is needed to 
determine if the higher ICP results are due to higher P bioavailability or analytical interfer- 
ences. Eckert and Watson (1996) reported that P measured with ICP is sometimes up to 50% 
higher than P measured with the colorimetric methods. The reason for such differences is 


explained by the fact that the spectrophotometry method determines only the orthophosphate 
forms of P, whereas the ICP determines the total P content (i.e., organic P as well as total 
inorganic P forms not just orthophosphate) present in the soil extract (Zbiral 2000a; 
Mallarino 2003). Mallarino (2003) reported a strong relationship between P determined by 
ICP method and the original colorimetric method (R 2 = 0.84) and concluded that M3-P as 
determined by ICP should be considered as a different test and its interpretation should be 
based on field calibration rather than conversion of M3-P measured by colorimetric method. 
Since automated systems are frequently employed to measure the concentration of nutrient 
ions in the extract and specific operating conditions and procedure for the instrument 
are outlined in the manufacturer's operating manual, only a manual method is described in 
this chapter. 


7 Reciprocating shaker 

2 Erlenmeyer flasks 125 ml_ 

3 Filter funnels 

4 Filter paper (Whatman #42) 

5 Disposable plastic vials 

g Instrumentation common in soil chemistry laboratories such as: spectrophoto- 
meter for conventional colorimetry or automated colorimetry (e.g., Technicon 
AutoAnalyzer; Lachat Flow Injection System); flame photometer; or ICP-OES or 

j M3 extracting solution: 

a. Stock solution M3: (1 .5 M NH 4 F + 0.1 M EDTA). Dissolve 55.56 g of ammo- 
nium fluoride (NH 4 F) in 600 mL of deionized water in a 1 L volumetric flask. 
Add 29.23 g of EDTA to this mixture, dissolve, bring to 1 L volume using 
deionized water, mix thoroughly, and store in plastic bottle. 

b. In a 1 L plastic carboy containing 8 L of deionized water, dissolve 200.1 g of 
ammonium nitrate (NH4NO3) and add 1 00 mL of stock solution M3, 1 1 5 mL 
concentrated acetic acid (CH 3 COOH), 82 mL of 10% v/v nitric acid (10 mL 
concentrated HNO3 in 100 mL of deionized water), bring to 10 L with 
deionized water and mix thoroughly. 

c. The pH of the extracting solution should be 2.3 + 0.2. 

g Solutions for the manual determination of phosphorus: 

a. Solution A: dissolve 1 2 g of ammonium molybdate ((NH 4 ) 6 Mo7024 • 4H 2 0) in 
250 mL of deionized water. In a 1 00 mL flask, dissolve 0.2908 g of potassium 
antimony tartrate in 80 mL of deionized water. Transfer these two solutions 


into a 2 L volumetric flask containing 1000 ml_ of 2.5 M H 2 S0 4 (141 mL 
concentrated H2SO4 diluted to 1 L with deionized water), bring to 2 L with 
deionized water, mix thoroughly, and store in the dark at 4°C. 

b. Solution B: dissolve 1 .056 g of ascorbic acid in 200 mL of solution A. Solution 
B should be fresh and prepared daily. 

c. Standard solution of P: use certified P standard or prepare a solution of 
100 |j,g mL -1 P by dissolving 0.4393 g of KH 2 P0 4 in 1 L of deionized water. 
Prepare standard solutions of 0, 0.5, 1,2,5, and 1 |xg mL -1 P in diluted M3 

g Solutions for K, Ca, Mg, and Na determination by atomic absorption: 

a. Lanthanum chloride (LaCI 3 ) solution: 10% (w/v). 

b. Concentrated solution of cesium chloride (CsCI) and LaC^: dissolve 3.16 g of 
CsCI in 100 mL of the 10% LaCI 3 solution. 

c. Combined K and Na standard solutions: use certified atomic absorption stand- 
ard and prepare solutions of 0.5, 1 .0, 1 .5, 2.0 and 0.3, 0.6, 0.9, 1 .2 |xg mL~ 1 of 
K and Na, respectively. 

d. Combined Ca and Mg standard solutions. Prepare 2, 4, 6, 8, 10 and 0.2, 0.4, 
0.6, 0.8, 1 .0 (i,g mL~ 1 of Ca and Mg, respectively. 

1q Standard solution for Cu, Zn, and Mn determination by atomic absorption: 

a. Combined Cu and Zn standard solution: 0, 0.2, 0.4, 0.8, 1 .2 to 2.0 |xg mL~ 1 of 
Cu and of Zn in M3 extractant. 

b. Mn standard solutions: prepare 0, 0.4, 0.8, 1 .2 to 4 |xg mL~ 1 of Mn in diluted 
M3 extractant. 

7.3.1 Extraction 

7 Weigh 3 g of dry soil passed through a 2 mm sieve into a 1 25 mL Erlenmeyer flask. 

2 Add 30 mL of the M3 extracting solution (soiksolution ratio 1:10). 

j Shake immediately on reciprocating shaker for 5 min (120 oscillations min -1 ). 

4 Filter through M3-rinsed Whatman #42 filter paper into plastic vials and store at 
4°C until analysis. 

5 Analyze elements in the filtrate as soon as possible using either an automated or 
manual method as described below. 


7.3.2 Determination of P by Manual Colorimetric Method 

1 Pipet 2 ml_ of the clear filtrate or standard (0 to 10 (jug ml_~ 1 ) P solution into a 
25 ml_ volumetric flask. The sample aliquot cannot contain more than 1 |xg of 
P and dilution of the filtrate with M3 maybe required. 

2 Add 15 ml_ of distil led water and 4 ml_ of solution B, make to volume with distilled 
water and mix. 

3 Allow 10 min for color development, and measure the absorbance at 845 nm. 

7.3.3 Determination of K, Ca, Mg, and Na by Atomic Absorption 
or by Flame Emission 

Precipitation problems can result from the mixture of the CsCl-LaCb solution with the M3 
extract. It is therefore recommended that the extracts be diluted (at least 1:10 final dilution) 
as indicated below to avoid this problem. 

7 Pipet 1 to 5 ml_ of filtrate into a 50 ml_ volumetric flask. 

2 Add approximately 40 ml_ of deionized water and mix. 

3 Add 1 ml_ of the CsCI-LaCb solution, bring to volume with deionized water 
and mix. 

4 Determine Ca, Mg by atomic absorption and K, Na by flame emission. 

7.3.4 Determination of Cu, Zn, and Mn by Atomic Absorption 

The Cu and Zn concentrations in the extract are determined without dilution while the Mn 
concentration is determined in diluted M3 extract. 


1 Filter paper can be a source of contamination which may affect the end results, 
especial ly for Zn,Cu, and Na.Mehlich (1984) proposed to use 0.2% AICI3 as a rinsing 
solution for all labware including qualitative filter paper. Based on local tests, we 
suggest the use of M3 extracting solution as a rinsing solution for filter paper. 

2 Because of Zn contamination, Pyrex glassware cannot be used for extraction or 
storage of the M3 extractant and laboratory standards. 

j Tap water is a major source of Cu and Zn contamination. 


The M3 extractant is widely used as ' 'universal extractant' ' in North America, Europe, and 
Australia (Zbiral and Nemec 2000; Cox 2001; Bolland et al. 2003). Jones (1998) reported that 
M3 is becoming the method of choice since many elements can be determined with this 


extractant. In Canada, it is used in the soil testing program in the provinces of Quebec and 
Prince Edward Island (CPVQ 1989; CRAAQ 2003). Many studies have been conducted over 
the world comparing the M3 method to the commonly used methods (ammonium acetate for 
K and DTPA for micronutrients) and report in general highly significant relationships between 
these methods. Some comments on relative amounts of elements extracted are provided below. 


The amounts of K and Na extracted by M3 are equal to those determined by 
ammonium acetate (Tran and Giroux 1989). 

The amounts of Ca and Mg extracted by M3 are about 1 .1 times more than those 
extracted by ammonium acetate method (Tran and Giroux 1989). 

The amount of Zn extracted by M3 is about one half to thre 
amount extracted by DTPA (Lindsay and Norvell 1978). 

The amount of Cu extracted by M3 is about 1 .8 times more than that extracted by 
DTPA (Makarim and Cox 1 983; Tran 1 989; Tran et al. 1 995). 

Bates, T.E. 1990. Prediction of phosphorus avail- 
ability from 88 Ontario soils using five phos- 
phorus soil tests. Commun. Soil Sci. Plant Anal. 
21: 1009-1023. 

Beauchemin, S. and Simard, R.R. 2000. Phos- 
phorus status of intensively cropped soils of the 
St-Lawrence lowlands. Soil Sci. Soc. Am. J. 64: 

Beauchemin, S., Simard, R.R., Bolinder, M.A., 
Nolin, M.C., and Cluis, D. 2003. Prediction of phos- 
phorus concentration in tile drainage water from the 
Montreal lowlands soils. Can. J. Soil Sci. 83: 73-87. 

Bolland, M.D.A., Allen, D.G., and Walton, K.S. 
2003. Soil testing for phosphorus: comparing the 
Mehlich 3 and Colwell procedures for soils of 
south-western Australia. Aust. J. Soil Res. 41: 

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Jones, J.B. Jr. 1998. Soil test methods: past, pre- 
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by ICP-AES. Rostlinna-Vyroba 46 (4), 141-146. 167-174. 

Chapter 8 

Sodium Bicarbonate-Extractable 


J.J. Schoenau 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

LP. O'Halloran 

University of Guelph 
Ridgetown, Ontario, Canada 


Sodium bicarbonate (NaHC03)-extractable phosphorus, commonly termed Olsen-P (Olsen 
et al. 1954), has a long history of worldwide use as an index of soil-available P on which to 
base P fertilizer recommendations (Cox 1994). It has been successfully used as a soil test for 
P in both acid and calcareous soils (Kamprath and Watson 1980). As a soil test, Olsen-P is 
sensitive to management practices that influence bioavailable soil P levels, such as fertilizer 
(O'Halloran et al. 1985) or manure (Qian et al. 2004) additions, although it is not suitable for 
P extraction from soils amended with relatively water-insoluble P materials such as rock 
phosphate (Mackay et al. 1984; Menon et al. 1989). 

As an extractant, NaHC03 acts through a pH and ion effect to remove solution inorganic 
P (Pj) plus some labile solid-phase P ; compounds such as phosphate adsorbed to free lime, 
slightly soluble calcium phosphate precipitates, and phosphate loosely sorbed to iron and 
aluminum oxides and clay minerals. Sodium bicarbonate also removes labile organic P 
(Bicarb-P ) forms (Bowman and Cole 1978; Schoenau et al. 1989) that may be readily 
hydrolyzed to P; forms and contribute to plant-available P (Tiessen et al. 1984; O'Halloran 
et al. 1985; Atia and Mallarino 2002) or be reassimilated by microorganisms (Coleman et al. 
1983). Although these labile P fractions once mineralized may play an important role in 
the P nutrition of crops, most regions using the Olsen-P soil test only consider the P; fraction. 
A modification of the Olsen-P method is one of the extraction steps used in the sequential 
extraction procedure for soil P outlined in Chapter 25. In this method, the NaHC03- 
extractable Pi (Bicarb-P;) and Bicarb-P are determined after a 16 h extraction. If the 
researcher is interested in a measure of the impact of treatments or management on these 
labile Pi and P fractions, one can simply follow the NaHC03 extraction and analysis 
procedure outlined in Chapter 25, ignoring the initial extraction using exchange resins. 


As with many soil tests for P, the Olsen-P test has been used as a surrogate measure of 
potential P loss through runoff (Pote et al. 1996; Turner et al. 2004) and in regions using the 
Olsen-P as the recommended soil P test it is often a criterion in soil P indices for assessing 
risk of P loss and impact on surface waters (Sharpley et al. 1994). The reader is referred to 
Chapter 14 for a more comprehensive discussion of methods for determining environmental 
soil P indices. Owing to its widespread use as an extractant for assessing P availability and its 
utilization in environmental P loading regulations, this chapter covers methodology for 
measurement of Olsen-P as a soil test. 


In this extraction, a soil sample is shaken with 0.5 M NaHCOi adjusted to a pH of 8.5, 
and the extract filtered to obtain a clear, particulate-free filtrate. The filtrate is usually a 
yellowish to dark brown color, depending upon the amount of organic matter removed 
from the soil. When relatively small amounts of organic matter are removed (pale 
yellowish-colored filtrates) it is possible to simply correct for its presence by using a 
blank correction (i.e., measure absorbance of a suitably diluted aliquot without color- 
developing reagent added). Presence of higher concentrations of organic matter can 
interfere with the color development in some colorimetric methods, or result in the 
precipitation of organic materials. Several options exist for the removal of the organic 
material in the extracts such as the use of charcoal (Olsen et al. 1954) and polyacrylamide 
(Banderis et al. 1976). 

8.2.1 Extraction Reagents 

7 Sodium bicarbonate (NaHCOs) extracting solution, 0.5 M adjusted to pH 8.5. 
For each liter of extracting solution desired, dissolve 42 g of NaHCC>3 and 0.5 g of 
NaOH in 1000 ml_ of deionized water. The NaHCC>3 extracting solution should 
be prepared fresh each month and stored stoppered since changes in pH of 
solution may occur that can affect the amount of P extracted. 

2 If using charcoal to remove organic material from the extracting solution: prepare 
by mixing 300 g of phosphate-free charcoal with 900 ml_ of deionized water (see 
Comment 2 in Section 8.2.3). 

j If using polyacrylamide to remove organic material from the extracting solution: 
dissolve 0.5 g of polyacrylamide in approximately 600 ml_ of deionized water in a 
1 L volumetric flask. This may require stirring for several hours. When the polymer 
has dissolved, dilute to volume with distilled water. 

8.2.2 Procedure 

1 Weigh 2.5 g sample of air-dried (ground to pass through a 2 mm sieve) soil into a 
125 ml_ Erlenmeyer flask. Include blank samples without soil. 

2 Add 50 ml_ of 0.5 M NaHC0 3 extracting solution at 25°C. 

If using charcoal to remove dissolved soil organic matter from the extracting 
solution: add 0.4 ml_ of the charcoal suspension. 

If using polyacrylamide to remove dissolved soil organic matter from extracting 
solution: add 0.25 ml_ of the polyacrylamide solution. 

Shake for 30 min on a reciprocating shaker at 120 strokes per minute. 

Filter the extract into clean sample cups using medium retention filter paper (i.e., 
VWR 454 or Whatman No. 40). If the filtrate is cloudy, refilter as necessary. 

See Section 8.3 for the determination of Olsen-P in the filtrates. 


7 The conditions under which the extraction is conducted can influence the amount 
of P extracted from the soil. Increasing the speed and time of the shaking will 
usually result in greater amounts of P being extracted (Olsen and Sommers 1982). 
Limiting extraction times to 30 min have been adopted for most soil testing 
purposes although a more complete and reproducible extraction may be obtained 
with a 1 6 h extraction. Increasing temperature of extraction will also increase the 
amount of P extracted. Olsen et al. (1 954) reported that extracted Pj increased by 
0.43 mg P kg -1 soil for each 1°C increase in temperature between 20°C and 30°C 
in soils testing between 5 and 40 mg P kg~ soil. It is therefore important that if 
the results are to be interpreted in terms of regional management recommenda- 
tions, the conditions of extraction must be similar to those used for the calibration 
of the soil test. If the results are for a comparative purpose between samples, then 
uniformity of extraction conditions between sample extractions is of greater 
importance than selecting a specific shaking speed, duration, and temperature 
of extraction. 

2 Most commercially available sources of charcoal or carbon black are contamin- 
ated with P. It is strongly recommended that the charcoal be washed with 6 M HCI 
to remove the P, followed by repeated washings with deionized water. Analysis of 
sample blanks of NaHCC>3 extracting solution with and without the charcoal 
solution will indicate if P removal from the charcoal has been successful. 

j The NaHC0 3 extracts should be analyzed as soon as possible, as microbial 
growth can proceed very rapidly, even under refrigeration. One can add one or 
two drops of toluene to inhibit microbial activity, although this increases the 
biohazard rating of the filtrates for handling and disposal. Preferably, the filtrates 
should be stored under refrigeration and analyzed within 5 days if they cannot be 
analyzed immediately. 


The amount of orthophosphate in the NaHCC>3 extractions is usually determined color- 
imetrically and various methods, both manual and automated, are available. The manual 


method described here is based on one of the most widely used procedures, the ammonium 
molybdate-antimony potassium tartrate-ascorbic acid method of Murphy and Riley 
(1962). This method is relatively simple and easy to use and the manual method described 
is adaptable to automated systems. The addition of antimony potassium tartrate eliminates 
the need for heating to develop the stable blue color. The phosphoantimonylmolybdenum 
complex formed has two absorption maxima; one at 880 nm and the other at 710 nm 
(Going and Eisenreich 1974). Watanabe and Olsen (1965) suggest measuring absorbance 
at 840 to 880 nm utilizing the greater of the two absorbance maxima, while Chapter 25 
suggests using 712 nm to reduce possible interference from traces of organic matter in 
slightly colored e 

8.3.1 Reagents for P Measurement 

1 Ammonium molybdate solution: dissolve 40 g of ammonium molybdate 
((NH 4 ) 6 Mo 7 024 • 4H 2 0) in 1 000 ml_ of deionized water. 

2 Ascorbic acid solution: dissolve 26.4 g of L-ascorbic acid in 500 ml_ of deionized 
water. Store under refrigeration at ~2°C. Prepare fresh if solution develops a 
noticeable color. 

j Antimony potassium tartrate solution: dissolve 1.454 g of antimony potassium 
tartrate in 500 ml_ of deionized water. 

4 Sulfuric acid (H 2 S0 4 ), 2.5 M: slowly add 278 ml_ concentrated H 2 S0 4 to a 2 L 
volumetric flask containing ~1 L of deionized water. Mix and allow to cool before 
making to volume with distilled deionized water. 

5 Sulfuric acid (H 2 S0 4 ), -0.25 M: slowly add -14 ml_ concentrated H 2 S0 4 to a 
100 ml_ volumetric flask containing -75 ml_ of distilled water. Mix well and 
make to volume with distilled water. 

6 p-nitrophenol solution, 0.25% (w/v): dissolve 0.25 g of p-nitrophenol in 100 mL 
of distilled water. 

y Standard P stock solution: prepare 1 00 mL of a base P standard with concentration 
of 5 jULg P ml_~ 1 . 

g Making the Murphy-Riley color-developing solution: using the above 
reagents, prepare the Murphy-Riley color-developing solution in a 500 mL 
flask as follows: add 250 mL of 2.5 M H 2 S0 4 , followed by 75 mL of ammo- 
nium molybdate solution, 50 mL of ascorbic acid solution, and 25 mL of 
antimony potassium tartrate solution. Dilute to a total volume of 500 mL by 
adding 100 mL of deionized water and mix on a magnetic stirrer. The reagents 
should be added in proper order and the contents of the flask swirled after 
each addition. Keep the Murphy-Riley solution in an amber bottle in a 
dark location to protect from light. Fresh Murphy-Riley solution should be 
prepared daily. 


8.3.2 Procedure 

7 Pipette 10 mL or a suitable aliquot of the filtered NaHC03 extract into a 50 mL 
volumetric flask. Include both distilled water and NaHCCh blanks. (See Comment 
2 in Section 8.3.3). 

2 To prepare standards of desired concentration range: 0, 0.1, 0.2, 0.3, 0.4, and 
0.8 (jug P mL" 1 in NaHC0 3 matrix, add 0, 1, 2, 3, 4, 6, and 8 mL of the base P 
standard (5 (jug P mL~ 1 ) to 50 mL volumetric flasks. Then add 10 mL of 
0.5 M NaHC0 3 to each flask. 

j To adjust the pH of the solutions add one to two drops of p-nitrophenol to each 
flask, which should result in a yellow solution. Lower the pH by adding 
0.25 M H2SO4 until the solution just turns colorless. 

4 To each flask, add 8 mL of the Murphy and Riley color-developing solution 
prepared in Section 8.3.1. Make to volume (50 mL) with deionized water, shake 
and allow 15 min for color development. 

5 Measure the absorbance of the standards and samples on a suitably calibrated and 
warmed-up spectrophotometer set to either 71 2 or 880 nm. Construct a standard 
curve using the absorbance values from standards of known P concentration. 


7 The ammonium molybdate, ascorbic acid, and antimony potassium tartrate solutions 
are generally stable for 2 to 3 months if well stoppered and stored under refrigeration. 
If quality of the solutions or reagents is suspected, discard and prepare fresh, as 
deterioration and/or contamination is a common source of error in the analysis. 

2 Although several modifications of the Murphy and Riley procedure exist in the 
literature, when using reagents as originally described by Murphy and Riley 
(1962) the final concentration of P in the 50 mL volumetric flask should not 
exceed 0.8 (jug P mL~ 1 (Towns 1 986) as color development may not be complete. 
Thus, the suitable aliquot size for color development should contain <40 |xg P. 
See Chapter 24 (Section 24.5) for more discussion on color development using the 
Murphy and Riley reagents. 

8.3.4 Calculation 

Using the concentrations of P suggested in Section 8.3.2, the standard curve should be linear. 
If the standard curve is constructed based on the |xg P contained in the 50 mL flask (i.e., 0, 5, 
10, 15, 20, 30, and 40 (JLg P) vs. absorbance, then the sample P content in mg P kg~' soil can 
be calculated using the following formula: 

1 50 mL (extraction volume) 1 

mg P kg" 1 soil = M,g P in flask x ^— - -x — — (8.1) 

mL aliquot g of soil 


Ada, A.M. and Mallarino, A.P. 2002. Agronomic 
and environmental soil phosphorus testing in 
soils receiving liquid swine manure. Soil Sci. 
Soc. Am. J. 66: 1696-1705. 

O'Halloran, LP., Kachanoski, R.G., and Stewart, 
J.W.B. 1985. Spatial variability of soil phos- 
phorus as influenced by soil texture and manage- 
ment. Can. J. Soil Sci. 65: 475^187. 

Banderis, A.S., Barter, D.H., and Henderson, K. Olsen, S.R., Cole, C.V., Watanabe, F.S., and 

1976. The use of polyacrylamide to replace car- Dean, L.A. 1954. Estimation of available plios 

bon in the determination of Olsen's extractable phorus in soils by extraction with sodium bicarbo 

phosphate in soil. J. Soil Sci. 27: 71-74. nate. US Dept. Agric. Circ. 939, Washington, DC. 

Bowman, R.A. and Cole, C.V. 1978. An explora- 
tory method for fractionation of organic phos- 
phorus from grassland soils. Soil Sci. 125: 95-101. 

Coleman, D.C., Reid, C.P., and Cole, C.V. 1983. 
Biological strategies of nutrient cycling in soil 
systems. In A. MacFayden and E.O. Ford, Eds. 
Advances in Ecological Research 13. Academic 
Press. New York, NY, pp. 1-56. 

Cox, F.R. 1994. Current phosphorus availability 
indices: characteristics and shortcomings. In 
J.L. Havlin et al., Eds. Soil Testing: Prospects 
for Improving Nutrient Recommendations. Soil 
Science Society of America Special Publication 
No. 40. SSSA-ASA, Madison, WI, pp. 101-114. 

Going, J.E. and Eisenreich, S.J. 1974. Spectro- 
photometric studies of reduced molybdoantimo- 
nylphosphoric acid. Anal. Chim. Acta 70: 95-106. 

Kamprath, E.J. and Watson, M.E. 1980. Conven- 
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phorus status of soil. In F.E. Khasawneh, E.C. 
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Phosphorus in Agriculture. American Society 
of Agronomy, Madison, WI, pp. 433^169. 

Mackay, A.D., Syers, J.K., Gregg, P.E.H., and 
Tillman, R.W. 1984. A comparison of three soil 
testing procedures for estimating the plant available 

phosphorus in soils using eithci superphosphate or 
phosphate rock. N. Z. J. Agric. Res. 27: 231-245. 

Menon, R.G., Hammond, L.L., and Sissingh, H.A. 
1989. Determination of plant-available phos- 
phorus by the iron hydroxide-impregnated filter 
paper (P ; ) soil test. Soil Sci. Soc. Am J. 53: 

Olsen, S.R. and Sommers, L.E. 1982. Phosphorus. 
In A.L. Page, R.H. Miller, and D.R. Keeney, Eds. 
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omy No. 9. American Society of Agronomy, 
Madison, WI, pp. 403^130. 

Pote, D.H., Daniel, T.C., Sharpley, A.N., Moore 
PA. Jr., Edwards, D.R., and Nichols, D.J. 1996. 
Relating extractable soil phosphorus to phosphorus 
losses in runoff. Soil Sci. Soc. Am. J. 60: 855-859. 

Qian, P., Schoenau, J.J., Wu, T., and Mooleki, P. 
2004. Phosphorus amounts and distribution in a Sas- 
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Schoenau, J.J., Stewart, J.W.B. , and Bettany, J.R. 
1989. Forms and cycling of phosphorus in prairie 
and boreal forest soils. Biogcot hemistry 8: 223-237. 

Sharpley, A.N., Chapra, S.C., Wedepohl, R., 
Sims, J.T., Daniel, T.C., and Reddy, K.R. 1994. 
Managing agricultural phosphorus for protection 
of surface waters: issues and options. /. Environ. 
Qual. 23: 437-441. 

Tiessen, H., Stewart, J.W.B., and Cole, C.V. 
1984. Pathways of phosphorus transformations 
in soils of differing pedogenesis. Soil Sci. Soc. 
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Chapter 9 

Boron, Molybdenum, 

and Selenium 

Ganga M. Hettiarachchi 

University of Adelaide 
Glen Osmond, South Australia, Australia 

Umesh C. Gupta 

Ay it ullure and Agri-rood Canada 
Charlottetown, Prince Edward Island, Canada 


Common features of B, Mo, and Se are that all three are nutrient elements that can be mainly 
found either in anionic or neutral form in soil solution and are relatively mobile in soils. 
Boron and Mo are essential elements for both plants and animals, while Se is an important 
element for humans and animals. Both B and Mo are essential micronutrients required for 
the normal growth of plants, with differences between plant species in the levels required 
for normal growth of plants. There is a narrow soil solution concentration range defining B 
or Mo deficiencies and toxicities in plants. 

Boron deficiencies can be found most often in humid regions or in sandy soils. Boron is 
subject to loss by leaching, particularly in sandy soils, and thus responses to B are common 
for sandy soils as summarized by Gupta (1993). Responses to B have been found on a variety 
of crops in many countries (Ericksson 1979; Touchton et al. 1980; Sherrell 1983). In 
contrast, B toxicity can be found mostly in arid and semiarid regions either due to high B 
in soils or high B containing irrigation water (Keren 1996). 

Responses to Mo have been frequently observed in legumes grown on soils that need lime. 
Elevated levels of Mo in soils and subsequent accumulations of Mo in plants, however, are of 
more concern than Mo deficiency in soils. High levels of Mo in plants eaten by ruminants 
can induce molybdenosis, a Mo-induced Cu deficiency (Jarrell et al. 1980). 

Yield responses to Se are generally not found. However, it is essential for livestock and is 
somewhat unique among the essential nutrients provided by plants to animals. In some areas, 
native vegetation can contain Se levels that are toxic to animals, whereas in other locations, 


vegetation can be deficient in Se, also causing animal health problems due to inclusion of 
low Se forage as part of animal diets (Mikkelsen et al. 1989). The Se concentration in soils in 
humid regions is generally inadequate to produce crops sufficient in Se to meet the needs of 
livestock. In acid soils, the ferric-iron selenite complex is formed, which is only slightly 
available to plants (NAS-NRC 1971). Selenium is generally present in excessive amounts 
only in semiarid and arid regions in soils derived from cretaceous shales, where it tends to 
form selenates (Welch et al. 1991). Selenium toxicity problems in the semiarid western 
United States are generally associated with alkaline soils where Se is present in the selenate 
form (Jump and Sabey 1989). 

9.2 BORON 

Boron in soils is primarily in the +3 oxidation state taking the form of the borate anion: 
B(OH) 4 ~. The two most common solution species of B are neutral boric acid (H3BO3) and 
borate anion (B(OH) 4 ~). Boron in soil can either be present in soil solution or adsorbed onto 
soil minerals such as clays. Below pH 7, H3BO3 predominates in soil solution, resulting in 
only a small amount of B adsorbed onto soil minerals. As the pH increases to about 9, the 
B(OH) 4 ~ increases rapidly, increasing B adsorption (Vaughan and Suarez 2003). Only the B 
in soil solution is important for plants. 

A number of extractants such as 0.05 M HC1 (Ponnamperuma et al. 1981), 0.01 M CaC^ + 
0.5 M mannitol (Cartwright et al. 1983), hot 0.02 M CaCl 2 (Parker and Gardner 1981), 
and 1 M ammonium acetate (Gupta and Stewart 1978) have been employed for deter- 
mining the availability of B in soils. One advantage of using CaCl2 is that it extracts little 
color from the soil, and predicted error due to this color is found to be low at 
0.00-0.07 mg kg~' (Parker and Gardner 1981). Such filtered extracts are also free of 
colloidal matter. 

Oyinlola and Chude (2002) reported that only hot water-soluble B correlated significantly 
with relative yields in Savannah soils of Nigeria, compared to several other extractants. 
Likewise Matsi et al. (2000) in northern Greece also noted that hot water-soluble B 
provided better correlation with yields than ammonium bicarbonate-diethylenetriamine- 
pentaacetic acid (AB-DTPA). Similar results were reported on some Brazilian soils where 
hot water-soluble B proved to be superior to HC1 and mannitol in predicting the B avail- 
ability for sunflower (Silva and Ferreyra 1998). Moreover, research work by Chaudhary 
and Shukla (2004) on acid soils of western India showed that both 0.01 M CaCl2 and hot 
water extractions were suitable for determining the B availability to mustard {Brassica 

Contrary to most other findings, Karamanos et al. (2003) concluded that hot water- 
extractable B was not an effective diagnostic tool for determining the B status of 
western Canadian soils. They, however, stressed that soil properties, especially organic 
matter, played an important role in determining the fate of applied B in the soil-plant 
system. Raza et al. (2002), on the other hand, found hot water-soluble B to be a good 
estimate of available B in the prairie soils of Saskatchewan. They further stated that soil 
cation exchange capacity appeared to be an important characteristic in predicting the B 

The most commonly used method is still the hot water extraction of soils as originally 
developed by Berger and Truog (1939) and modified by Gupta (1993). A number of 


modified versions of this procedure have since appeared. Offiah and Axley (1988) have 
used B-spiked hot water extraction for soils. This method is claimed to have an advantage 
over unspiked hot water extraction in that it removes from consideration a portion of the B 
fixing capacity of soils that does not relate well to plant uptake. A longer boiling time of 
10 min as opposed to the normally used 5 min boiling was found to reduce error for a Typic 
Hapludult soil by removing enough B to reach the plateau region of the extraction curve 
(Odom 1980). 

Once extracted from the soil, B can be analyzed by the colorimetric methods using 
reagents such as carmine (Hatcher and Wilcox 1950), azomethine-H (Wolf 1971), and 
most recently by inductively coupled plasma-atomic emission spectrometry (ICP-AES) 
(Keren 1996). 

9.2.1 Reagents 

7 Deionized water 
2 Charcoal 

9.2.2 Procedure (Gupta 1993) 

1 Weigh 25 g air-dried soil, screened through a 2 mm sieve, into a preweighed 
250 mL "acid-washed" beaker and add about 0.4 g charcoal and 50 ml_ deionized 
water and mix. The amount of charcoal added will vary with the organic matter 
content of the soil and should be in sufficient quantity to produce a colorless extract 
after 5 min of boiling (see Comments 2 and 3 in Section 9.2.5). A blank containing 
only deionized water and a similar amount of charcoal as used with the soil 
samples should also be included. 

2 Boil the soil-water-charcoal or water-charcoal mixtures for 5 min on a hotplate. 

3 The loss in weight due to boiling should be made up by adding deionized water 
and the mixture should be filtered while still hot through a Whatman No. 42 or 
equivalent type of filter paper. 

9.2.3 Determination of Boron by the Azomethine-H Method 


1 Azomethine-H: dissolve 0.5 g azomethine-H in about 10 mL redistilled 
water with gentle heating in a water bath or under a hot water tap at about 
30°C. When dissolved add 1.0 g L-ascorbic acid and mix until dissolved. Make 
the final volume up to 100 mL with redistilled water. If the solution is not clear, it 
should be reheated again till it dissolves. Prepare fresh azomethine-H solution for 
everyday use. 

2 Ethylene diamine tetraacetic acid (EDTA) reagent (0.025 M): dissolve 9.3 g EDTA 
in redistilled water and make the volume up to 1 L with redistilled water. Add 1 mL 
Brij-35 and mix. 

Buffer solution: dissolve 250 g ammonium acetate in 500 ml_ redistilled water. 
Adjust the pH to about 5.5 by slowly adding approximately 1 00 ml_ concentrated 
acetic acid, with constant stirring. Add 0.5 ml_ Brij-35 and mix. 

Standard solutions: prepare stock standard A by dissolving 1000 mg B 
(5.715 g H3BO4) in 1 L deionized water and prepare stock standard B by taking 
50 ml_ stock standard A and diluting it to 1 L with 0.4 M HCI. Prepare standard 
solutions from stock standard B by diluting a range of 2.5 to 30 ml_ stock 
standard B to 1 L with deionized water to give a range of 0.5 to 6.0 mg B L~ 1 in 
the final standard solution. 

7 Take 5 ml_ of the clear filtrate in a test tube and add 2 ml_ buffer solution, 2 mL 
EDTA solution, and 2 mL azomethine-H solution, mixing the contents of the test 
tube thoroughly after the addition of each solution. 

2 Let the solutions stand for 1 h and measure the absorbance at 430 nm on a 

j The color thus developed has been found to be stable for up to 3-4 h. 

4 The pH of the colored extract should be about 5.0. 

9.2.4 Determination of Boron by Inductively Coupled Plasma-Atomic 
Emission Spectrometry 

This technique has been found to be rapid and reliable for determining B in plant digests and 
soil extracts using the procedure described in Section 9.2.2 by Gupta (1993). An estimated 
detection limit by ICP-AES at wavelength of 249.77 nm is about 5 |jig L" 1 (APHA 1992) 
and therefore, it is reasonable to expect method detection limit to be about 100 |jig B kg~' 
soil. Care must be taken to filter samples properly as colloidal-free extracts are 
recommended for ICP-AES to avoid nebulizer-clogging problems. 


/ All glassware used in plant or soil B analyses must be washed with a 1 :1 mixture 
of boiling HCI acid with deionized water before use. Storage of the filtered 
extracts before the analysis of B must be in plastic sampling cups. 

2 Soils containing higher organic matter may require additional amount of charcoal to 
obtain a colorless extract, but the addition of excessive amounts of charcoal can 
reduce the amount of B in the extract. 

3 If the filtered solution is not colorless, the extraction may need to be repeated with 
a higher amount of charcoal. 

4 The use of azomethine-H is an improvement over those of carmine (Hatcher and 
Wilcox 1950), quinalizarin, and curcumin (Johnson and Ulrich 1959), since the 

procedure involving this chemical does not require use of a concentrated acid. 
This method has been found to give comparable results when compared to the 
carmine method (Gupta 1993). 

It is difficult to use an autoanalyzer because of its insensitivity at lower B 
concentrations generally found in the hot water extract of most soils. 


Molybdenum in soils is primarily in the +6 oxidation state taking the form of the molybdate 
anion, Mo0 4 2 ~ . The solution species of Mo, generally in the order of decrease in concen- 
tration, are Mo0 4 2 ", HM0O4", H 2 MoO 4 , Mo0 2 (OH)+, and Mo0 2 2+ , respectively. The 
latter two species can be ignored in most soils (Lindsay 1979). Molybdate is adsorbed by 
oxides, noncrystalline aluminosilicates, and to a lesser extent by layer silicates and adsorp- 
tion increases with decreasing pH. Therefore, Mo is least soluble in acid soils, especially acid 
soils containing Fe oxides. 

Studies on the extraction of available Mo from soils have been limited. Further, the 
extremely low amounts of available Mo in soils under deficiency conditions make it 
difficult to determine Mo accurately. The accumulation of Mo in plants mostly is not 
related to total concentrations of Mo in soils but rather to available Mo in soils. A variety 
of extractants have been used in attempts to extract available Mo in soils although no 
routine soil test for Mo is available. Molybdenum deficiencies are rare and are mostly of 
concern for leguminous crops. Since excessive Mo in forages can harm animal health, 
Mo fertilization is usually based on visual deficiency symptoms and/or history of crop 

Many extractants have been employed for the assessment of available Mo in soils. Those 
extractants are: ammonium oxalate, pH 3.3 (Grigg 1953); water (Gupta and MacKay 1965a); 
hot water, anion-exchange resin; AB-DTPA (Soltanpour and Workman 1980); ammonium 
carbonate (Vlek and Lindsay 1977); and Fe oxide strips (Sarkar and O'Connor 2001). 
However, most of those extractants are used to study the deficiency aspect rather than 
from consideration of toxic effects (Davies 1980). 

Despite its weaknesses, the most commonly used extractant for assessing Mo availability in 
soils has been ammonium oxalate, buffered at pH 3.3 (Grigg 1953). Examples for the 
successful use of acid ammonium oxalate in predicting Mo uptake by plants (Wang et al. 
1994) and its failures (Mortvedt and Anderson 1982; Liu et al. 1996) can be found in the 
literature. From studies that failed to predict plant uptake of Mo successfully with acid 
ammonium oxalate-extractable Mo, it appeared that plant Mo was more closely related to 
some soil property such as pH other than extractable Mo in soils. Some studies obtained a 
better regression between acid oxalate-extractable Mo in soil and plant Mo when soil pH 
was considered as a factor (Mortvedt and Anderson 1982). Sharma and Chatterjee (1997) 
stated that soil physical properties such as soil pH, organic matter, parent rock, and texture 
play an important role in determining the Mo availability in alkaline soils. Multiple- 
regression equations account for the contribution of the individual factors, which would 
make the critical limits more predictable. Moreover, Liu et al. (1996) found signifi- 
cant correlations (r 2 = 0.81) for soil Mo extracted with ammonium oxalate (pH 6.0) in a 
group of Kentucky soils with Mo uptake by tobacco (Nicotiana tabacom L.) growing in 


greenhouse. However, ammonium oxalate buffered at pH 3.3 was not statistically well 
correlated with Mo uptake. 

Some methods that have not been widely tested but appear to be promising are anion- 
exchange resin and AB-DTPA methods. Anion-exchange resins have been used with success 
to extract plant-available Mo in soils (Ritchie 1988). The AB-DTPA method (Soltanpour 
and Workman 1980; Soltanpour et al. 1982) has also been used successfully for alkaline and 
Mo-contaminated soils (Pierzynski and Jacobs 1986, Wang et al. 1994). Moreover, ammo- 
nium carbonate (Vlek and Lindsay 1977) also has shown good correlation with plant uptake 
of Mo, especially for soils that have Mo toxicity problems. This extraction followed by H2O2 
treatment leaves a decolorized extract that is useful for Mo analysis by colorimetric methods 
(Wang et al. 1994). 

To characterize the available Mo in biosolids-amended soils, Sarkar and O'Connor (2001) 
compared the potential of Fe-oxide impregnated filter paper with ammonium oxalate 
extraction method and total soil Mo. Their data showed that the best correlation between 
plant Mo and soil Mo was obtained using the Fe-oxide strip followed by ammonium oxalate 
extraction; while total soil Mo was generally not well correlated with plant Mo uptake. 
Sarkar and O'Connor (2001) further reported that Fe-oxide strips can serve as an analytically 
satisfactory and practical procedure for assessing available Mo, even in soils amended with 

Recently, McBride et al. (2003) found that dilute CaCl2 was found to be preferable to 
Mehlich 3 as a universal extractant for determining Mo and other trace metal availability 
in clover grown on near neutral soils amended with sewage sludge. Concentration of Mo in 
alfalfa {Medicago sativa L.) on soils treated with sewage sludge was well correlated to 
readily extractable Mo by 0.01 M CaCb in the soil. Total Mo and past Mo loading to soil 
were less reliable predictors of Mo concentration in alfalfa than the soil test for readily 
extractable Mo (McBride and Hale 2004). 

Two methods of extractions are outlined (1) ammonium oxalate, pH 3.0 (modified Grigg 
1953) and (2) AB-DTPA (Soltanpour and Schwab 1977). 

9.3.1 Extraction of Molybdenum by the Ammonium Oxalate, pH 3.0 
Method (Modified Grigg 1953) 

Reagents (Gupta and MacKay 1 966) 

7 Ammonium oxalate, 0.2 M buffered to pH 3.0: in a 1 L volumetric flask dissolve 
24.9 g of ammonium oxalate and 12.605 g of oxalic acid in approximately 800 ml_ 
deionized water. Make to volume with distilled water and mix well. 

Add 15 g air-dried soil, screened through a 2 mm sieve, to a 250 ml_ beaker or 
Erlenmeyer flask. 

Add 1 50 ml_ of the buffered (pH 3) 0.2 M ammonium oxalate solution and shake 
for 16 h at room temperature using an orbital shaker at 200 rpm. 


Filter the extraction through Whatman No. 42 filter paper or equivalent. Centri- 
fuge the filtrate for 20 min. 

Determine Mo concentration in the clear extract as described in Section 9.3.3. If 
required, the centrifuged extracts can be acidified to pH < 2 with HNO3 and 
stored in 1:1 HNO3 rinsed plastic or glass containers up to a maximum of 
6 months (APHA 1992). 

9.3.2 Extraction of Molybdenum by the Ammonium Bicarbonate- 

dlethylenetriaminepentaacetic acid solution method (soltanpour 
and Schwab 1977) 


7 Ammonium hydroxide (NH 4 OH) 1:1 solution. 

2 AB-DTPA solution (1 M NH4HCO3, 0.005 M DTPA buffered to pH 7.6): in a 1 L 
volumetric flask containing approximately 800 ml_ of distilled-deionized water, 
add 1.97 g of DTPA and approximately 2 ml_ of 1:1 NH 4 OH solution and 
mix. (The addition of the 1:1 NH 4 OH solution aids in the dissolution of 
DTPA and helps prevent frothing.) When most of the DTPA is dissolved, add 
79.06 g of NH4HCO3 and stir until all materials have dissolved. Adjust pH to 7.6 
by adding either NH 4 OH or HCI and then make to volume using distilled- 
deionized water. 

1 Weigh 1 g soil, screened through a 2 mm sieve, into a 1 25 ml_ Erlenmeyer flask 
and add 20 ml_ of AB-DTPA solution. 

2 Shake the mixture in open flasks on a reciprocal shaker at 1 80 rpm for 1 5 min and 
filter the extract using Whatman No. 42 filter paper or its equivalent. 

3 Determine Mo as described in Section 9.3.3. The filtered extracts can be pre- 
served until analysis as mentioned under Section 9.3.1 (Reagents (1)). 

9.3.3 Determination of Molybdenum 

Determine Mo concentration in extracts with graphite furnace atomic absorption spectrom- 
etry (GFAAS) or ICP-AES. The standards for GFAAS or ICP must be prepared in the 
extracting solution matrix. 

Since extractable Mo in normal situations is usually in the range of 10 to 50 (jug L _1 , 
analytical methods must be sensitive to measure low concentrations. Therefore, most 
suitable method is GFAAS (Mortvedt and Anderson 1982). It is recommended to use 
HNO3 as a matrix modifier (as enhancer); and pyrolytically coated tubes (to minimize 
problems due to carbide formation) for Mo determination in GFAAS. An estimated detection 


limit using pyrolytic graphite tubes is 1 |uug L _1 (APHA 1992). In situations where one could 
expect higher concentrations of Mo in the extracting solutions, flame atomic absorption 
spectrometry or atomic emission spectrometry (either direct or ICP-AES) can be used for Mo 
analysis (Soltanpour et al. 1996). An estimated detection limit using ICP-AES is 8 |xg L _1 
(APHA 1992) and therefore, it will be safer to assume method detection limits for ICP-AES 
for Mo to be 80 |xg L _1 or little lower. For spectrometry determinations standards must be 
made in AB-DTPA matrix solution. It has also been suggested to treat the extract with concen- 
trated HNO3 acid before determination of Mo by ICP-AES. After adding 0.5 mL con- 
centrated HNO3 acid to about 5 mL filtrate, mix it in a beaker on a rotary shaker for about 
15 min to eliminate carbonate species. 

Determination of Mo in soil extracts can also be done colorimetrically in laboratories that are 
not equipped with ICP-AES or GFAAS. Refer to Gupta and MacKay (1965b) for details of 
colorimetric determination of Mo. 


In general, ammonium oxalate shows greater ability to extract Mo from soils and mine spoils 
compared to AB-DTPA method (Wang et al. 1994). 


Soil Se forms include very insoluble reduced forms including selenium sulfides, elemental 
Se (Se°), and selenides (Se -2 ) and more soluble selenate (SeO/t 2- ), and selenite (HSe03~, 
Se03 2 ~). Elemental Se, sulfides, and selenides only occur in reducing environments. They 
are insoluble and not available for plants and living organisms (McNeal and Balistrieri 
1989). In alkaline, oxidized soils, selenates are the dominant forms while in slightly acidic, 
oxidized soils, selenites are dominant. Selenate and selenite precipitates and minerals are 
highly soluble in aerobic environments and therefore, the solubility of Se is controlled 
mainly by adsorption and complexation processes. Selenite is proven to be strongly adsorbed 
to soil surfaces while selenate is weakly adsorbed (Neal et al. 1987). 

The parent material has a significant effect upon the Se concentration in plants. For example, 
field studies conducted on wheat in west central Saskatchewan showed higher Se values in 
wheat plants grown on lacustrine clay and glacial till, intermediate in plants grown on 
lacustrine silt, and lowest on aeolian sand (Doyle and Fletcher 1977). A similar trend 
characterized the C horizon soil, with highest Se values associated with lacustrine clay and 
lowest with aeolian sand. The findings of Doyle and Fletcher (1977) pointed to the potential 
usefulness of information on the Se content of soil parent materials when designing sampling 
programs for investigating regional variations in plant Se content. 

Available Se in soils is highly variable. Although there were instances where a direct 
correlation between soil Se content and the plant grown on those soils existed (Varo et al. 
1988), more often the total Se in soil proved to be of little value in predicting plant uptake 
(Diaz-Alarcon et al. 1996). Selenium uptake by plants depends not only on the form and 
partitioning of Se species between solution and solid phases but also on the presence of other 
ions in soil solution (such as SC>4~ 2 ) and the species of plants (Bisbjerg and Gissel-Nielsen 
1969; Mikkelsen et al. 1989). Therefore, ideally extractants capable of predicting or evalu- 
ating plant-available Se should be capable of extracting Se in soil solution as well as Se 
associated with solid phases that would be potentially released into soil solution. The ability 


of an extractant to correlate significantly with plant uptake could vary depending on many 
factors, some of which are soil type, plant species, season, and location. Uptake of Se by 
plants and methods that can be used to predict and evaluate plant uptake of Se can be found 
in the literature (Soltanpour and Workman 1980; Soltanpour et al. 1982; Jump and Sabey 
1989; Mikkelsen et al. 1989). 

Soltanpour and Workman (1980) found a high degree of correlation between extracted Se by 
an AB-DTPA extraction procedure developed by Soltanpour and Schwab (1977) and Se 
uptake by alfalfa for five levels of Se(VI) in a greenhouse study. In addition, they found very 
high (r 2 = 0.99) correlation between AB-DTPA extractable and hot water-extractable Se 
(Black et al. 1965). The hot water-extractable Se soil test method is developed based on the 
assumption that soil and soil-like materials that contain appreciable amounts of water- 
soluble Se (majority as selenates) will give rise to Se-toxic vegetation (Black et al. 1965). 
Similarly, AB-DTPA should extract water-soluble Se as well as exchangeable selenate 
and/or selenite into solution due to bicarbonate anion. In addition, Soltanpour et al. (1982) 
found that the AB-DTPA-extractable Se in soil samples taken from a to 90 cm depth in the 
autumn before seeding winter wheat (Triticum aestivum L.) correlated well with Se in grain 
samples (r 2 = 0.82) that were taken in the following summer. 

Selenium in saturated paste extracts could also provide useful information about plant- 
available Se in soils (U.S. Salinity Laboratory Staff 1954) as mostly the soihwater ratio 
in these pastes can be related to field soil water content in a predictable manner. Using two 
Se-accumulating plant species, Jump and Sabey (1989) found that Se in saturated paste water 
extracts correlated highest with plant Se concentrations from a study that compared Se 
extracted from 18 different soils and mine-spoil materials by several different extractants 
(AB-DTPA, DTPA, hot water, saturated paste extract, and Na 2 C0 3 ). 

In addition to measuring total extractable Se, determination of Se species in soil solution, 
saturate paste extract, or any other extraction may also provide insight into potential for plant 
Se uptake. Mikkelsen et al. (1989) discussed the different mechanisms associated with 
energy-dependent uptake of Se(VI) and energy-independent uptake of Se(IV). They also 
discussed the variable uptake of Se by different plant species, which is an additional 
complication. Davis (1972a,b) demonstrated the variability for absorbing Se among different 
species within a single plant genus in two greenhouse experiments. All the above suggest 
that speciation information on Se(VI) and Se(IV) in extractions or soil solutions may also 
provide useful information on uptake of Se by plants. 

Relatively labile forms of Se in soils can be evaluated by using orthophosphate (PO4) as a 
soil extractant (Fujii et al. 1988). This is based on the assumptions that PO4 replaces 
adsorbed forms of Se and the dominant adsorbed species of Se in these soils is Se(IV). 
Fujii and Burau (1989) used 0.1 M PO4 solution adjusted to pH 8 and was able to extract 
89% to 103% of the sorbed Se(IV) for three surface soils. 

Sequential extraction procedures can also be used to identify fractions of Se in soils (Chao 
and Sanzolone 1989; Lipton 1991) and may be related to plant uptake. The sequential 
extraction method developed by Chao and Sanzolone (1989) fractionates soil Se into five 
operationally defined fractions (soluble, exchangeable, oxide bound, sulphide/organic mat- 
ter bound, and residual or siliceous material associated), whereas the Lipton (1991) method 
fractionates soil Se into nine operationally defined fractions (soluble, ligand exchangeable, 
carbonates, oxidizable, easily reducible oxides bound, amorphous oxide bound, crystalline 
oxide bound, alkali-soluble Al/Si bound, and residual). 


9.4.1 Extraction of Selenium in Soils 

We will outline five commonly used methods of extractions with appropriate references here. 

Five commonly used extractants for Se are given below: 

/ AB-DPTA (Soltanpour and Schwab 1 977): 1 g of air-dried soil, screened through a 
2mmsieve,isplacedina125mLErlenmeyerflask.Add20ml_of1 M NH4HCO3 + 
0.005 M DTPA (prepared as described in Section Reagents, p. 101) at pH 7.6. 
Shake the mixture in an open flask on a reciprocal shaker at 180 rpm for 15 min 
and filter the extract using Whatman No. 42 filter paper or its equivalent. 

2 Hot water (Black et al. 1 965): place 1 g of air-dried soil, sieved through a 2 mm 
sieve, in a 250 ml_ Erlenmeyer flask. Add 50 ml_ distilled water, and reflux over a 
boiling water bath for 30 min. Filter the soil suspension using Whatman No. 42 
filter paper or its equivalent. 

3 Saturated paste extractants (U.S. Salinity Laboratory Staff 1 954): weigh 200 to 400 g 
of air-dried soil, sieved through a 2 mm sieve into a plastic container with a lid. 
Weigh the container, and container plus soil. Add distilled water to the soil, while 
stirring, until soil is nearly saturated. Cover the container and allow the mixture to 
stand for several hours. Then add more water with stirring to achieve a uniformly 
saturated soi l-water paste. The criteria for saturation shou Id be checked as given here 
(soil paste glistens as it reflects light, flows slightly when the container is tipped, slides 
freely and cleanly off a smooth spatula, and consolidates easily by tapping or jarring 
the container after a trench is formed in the paste with the side of the spatula). Allow 
the sample to stand for another 2 h, preferably overnight, and then recheck for the 
sample for saturation criteria. If the paste is too wet, add known amount of dry soil to 
the paste. Once saturation is attained, weigh the container plus content to get the 
amount of water added. Transfer the paste to a Buchner funnel fitted with highly 
retentive filter paper, and apply a vacuum to collect saturation extract in a test tube. 

4 0.005 M DTPA, 0.01 M CaCI 2 (2 h DTPA test) (Lindsay and Norvell 1978): 10 g 
air-dried soil, screened through a 2 mm sieve, is placed in a 50 mL polypropylene 
centrifuge tube. Add 20 mL of 0.005 M DTPA, 0.01 M CaCI 2 buffered at pH 7.3 
with triethanolamine and shake for 2 h on a reciprocating shaker. Centrifuge 
immediately at 3000 g and filter the supernatant using Whatman No. 42 filter 
paper or its equivalent. 

5 0.5 M Na 2 C0 3 (Jump and Sabey 1 989): 5 g of air-dried soil, screened through a 
2 mm sieve, is shaken on a reciprocating shaker in 20 mL of 0.5 M Na 2 CC>3 
solution at pH 11.3 for 30 min. Filter the extract using Whatman No. 42 filter 
paper or its equivalent. 


7 The soihextractant ratio varies from 1 :2 to 1 :5 and the extraction time from 1 5 min 
to 2 h as given in the above-mentioned references or as summarized by Jump and 
Sabey (1989). 


2 The filtered extracts can be analyzed for Se using a hydride-generating system attached 
to an ICP-AES (Soltanpour et al. 1 996). Filtered extracts to be analyzed for Se can be 
preserved until analysis with either HN0 3 or HCI (pH < 2) to prevent loss of Se from 
solution (through coprecipitation or methylation of Se followed by volatilization). 

All of the above five extractants when tested on soils containing high Se showed high 
correlation between wheat plant Se and Se extracted from soils (Jump and Sabey 1989). 
However, Se extracted in saturated soil pastes and expressed as mg Se L~ ' of extract was 
found to be the best predictor of Se uptake in Se-accumulating plants. Furthermore, the 
results suggest that soil or mine-spoil materials that yield more than 0.1 mg Se L ' in 
saturated extract may produce Se-toxic plants. 

In addition, the AB-DTPA extract has been found to predict Se availability better when Se in 
wheat grain was correlated with Se in the 0-90 cm depth of soil as opposed to the 0-30 cm 
depth (Soltanpour et al. 1982). This was found to be particularly useful to screen soils and 
overburden material for potential toxicity of Se. 

9.4.2 Determination of Selenium 

Selenium in extracting solutions can be accurately determined by hydride generation atomic 
absorption spectrometry (HGAAS), electrothermal, or GFAAS, ICP-AES as well as com- 
bination of chemical methods with colorimetry and fluorometry (APHA 1992). The most 
common method of choice is the continuous HGAAS. For determination of Se at higher 
concentration, the ICP-AES coupled with HG may be preferred, in particular when simul- 
taneous determination of other elements such as As is required (Workman and Soltanpour 
1980). Matrix matching techniques (for example prepare standards in the same matrix as soil 
extracts) and extensive QA/QC procedures should be used to assure the quality of deter- 
mination. For detailed information regarding the HGAAS apparatus and reagents needed for 
determination of Se, refer to APHA (1992) and Huang and Fujii (1996). 


7 The extractants developed have been found to be suitable for predicting the 
availability of Se in Se toxic areas only. Because of rather small quantities of 
available Se in Se-deficient areas, no reliable extractant has yet been developed 
for such soils. Therefore, plant Se and total soil Se will continue to serve as the 
best tools available for testing the Se status of Se-deficient soils. 

2 The term deficiency or deficient in connection with Se has implications in 
livestock and human nutrition only and not in plant nutrition since no known 
yield responses to Se have been found on cultivated crops. 

American Public Health Association (APHA), Wastewater, 18th edn. American Public Health 

American Water Works Association, and Water Association, American Water Works Associa- 

Pollution Control Federation. 1992. Standard tion, and Water Pollution Control Federation, 

Methods for the Examination of Water and Washington, DC. 


Berger, K.C. and Truog, E. 1939. Boron deter- 
mination in soils and plants using the quinalizarin 
reaction. Ind. Eng. Chem. 11: 540-545. 

Bisbjerg, B. and Gissel-Nielsen, G. 1969. The 
uptake of applied selenium In agricultural plants. 
I. The influence of soil type and plant species. 
Plant Soil 31: 287-298. 

Fujii, R., Hatfield, D.B., and Deverel, S.J. 1988. 
Distribution of selenium in soils of agricultural 
fields, western San Joaquin Valley, California. 
Soil Sci. Soc. Am. J. 52: 1274-1283. 

Grigg, J.L. 1953. Determination of the available 
molybdenum of soils. N. Z. J. Sci. Tech. Sect. 
A-34: 405-414. 

Black, C.A., Evans, D.D., White, J.L., Ensmiger, Gupta, S.K. and Stewart, J.W.B. 1978. An auto- 

L.E., and Clark. F.E.. Eds. 1965. Methods of Soil mated procedure for determination of boron in 

Analysis. Port 2. Agronomy 9, ASA, Madison, soils, planls and irrigation waters. Sehweizerische 

WI, 1122 pp. Landwirtschaftliche Eorschung 17: 51-55. 

Cartwright, B., Tiller, K.G., Zarcinas, B.A., and 
Spouncer, L.R. 1983. The chemical assessment of 
the boron status of soils. Aust. J. Soil Res. 21: 

Chao, T.T. and Sanzolone, R.F. 1989. Fraction- 
ation of soil selenium by sequential partial dissol- 
ution. Soil Sci. Soc. Am. J. 53: 385-392. 

Chaudhary, D.R. and Shukla, L.M. 2004. Evalu- 
ation of extiactants for predicting availability of 
boron to mustard in arid soils of India. Commun. 
Soil Sci. Plant Anal. 35: 267-283. 

Davies, B.E. 1980. Applied Soil Trace Elements. 
Wiley-Interscience, Chichester, New York, NY. 

Davis, A.M. 1972a. Selenium accumulation in 
Astragalus species. Agron. J. 64: 751-754. 

Davis, A.M. 1972b. Selenium accumulation in 
a collection of Atriplex species. Agron. J. 64: 

Diaz-Alarcon, J.P., Navarro-Alarcon, M., De la 
Sen-ana, H.L., Asensio-Drima, C, and Lopez- 
Martinez, M.C. 1996. Determination and chem- 
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Chapter 10 
Trace Element Assessment 

W.H. Hendershot, H. Lalande, and D. Reyes 

McGill University 
Sainte Anne de Bellevue, Quebec, (. anada 

J.D. MacDonald 

Agriculture and Agri-Food Canada 
Quebec, Quebec, Canada 


The current literature contains a wide range of extractants that have been used to evaluate 
different fractions of metals and metalloids in soils (Tessier et al. 1979; Ross 1994; Ure 1996; 
Mihaljevic et al. 2003). These techniques fall into two categories: single or sequential 
extractions. Although sequential extractions have gained considerable popularity, they do 
have several drawbacks (Beckett 1989; Lo and Yang 1998; Shiowatana et al. 2001). From an 
analytical point of view, sequential extractions may result in inconsistent results due to 
reprecipitation of the elements of interest from one extractant to the next and errors caused 
by adding the different fractions can lead to values that do not agree with analyses of total 
metals. Perhaps the most important criticism of sequential extractions is that they are not 
really specific for the intended fraction; examples of extractions that do not remove specific 
and identifiable chemical forms are abundant in the literature (Beckett 1989; Mihaljevic 
et al. 2003). 

The approach taken here is to select a series of single extractants that range from weak to 
very strong. Each of the extractants proposed in this chapter selectively dissolves some 
portion of the total element pool in the soil but no attempt is made to relate this to a specific 
type of surface or material. For our purposes it is not really important where the trace 
elements are held on the soil; it is more important that the analysis provides a means of 
predicting or explaining the interactions of the elements with biota or mobility in the soil 
system. In some cases, there is extensive literature that can help to relate the results to 
bioavailability of the elements to specific organisms. 

Some of the metals such as Cd, Cu, Ni, Pb, and Zn have received considerable attention 
over the last 10 years. For these metals there are numerous references that relate the amounts 
of metals extracted by different chemicals to a biological effect (toxicity or uptake). 
However, other metals and metalloids are also significant contaminants in soils affected by 


anthropogenic activity, but these metals have received much less attention (As, Co, Cr, Mo, 
Sb, Se, Tl, etc.)- In these cases the number of references relating selective chemical 
extraction results to biological effects is much less abundant but has been growing in recent 
years (De Gregori et al. 2004). Although in some cases the methods proposed below have not 
been tested for a wide range of metals and metalloids, by providing a series of standard tests 
we hope that more studies will be conducted so that a database of response data 
can be developed. Since the elements forming oxyanions, such as As, Cr, Mo, and Se, 
behave quite differently in soils compared to the cationic metals, some authors prefer to use 
extraction procedures developed for phosphate (Van Herreweghe et al. 2003). However, 
many of the techniques used for the extraction of the metalloid As, for example, were 
originally developed for cationic metals but yield good results nonetheless (Hall et al. 
1996; Mihaljevic et al. 2003). 

Four extraction procedures are proposed here and are presented in the order of increasing 

j A column leaching method using water and 80 [lM CaC^/CaSC^ solution 

2 A weak salt solution using 0.01 M CaCi2 

j A strong chelating agent (0.05 M NH4-ethylenediaminetetraacetic acid [EDTA] par- 
tially neutralized with NHt + ) 

4 A strong acid microwave digestion procedure using HNO3 (USEPA method 3051) 

Recent work in our laboratory has led to the development of a column leaching technique 
that provides a very good simulation of the solubility of trace elements, pH, and ionic 
strength of solutions collected in the field from forest soils in Ontario and Quebec, Canada 
(MacDonald et al. 2004a,b). This method consists of an initial washing of the soil with 
deionized water, followed by an equilibration with very dilute (80 |xM) CaCb and CaSC>4 
solution to simulate the ionic strength observed in forest soils. It has been chosen because 
it provides the extraction procedure best suited to estimate metal mobility under field 

The CaCl2 method is gaining support in Europe and North America as one of the best ways 
of evaluating bioavailability chemically (Houba et al. 1996; Ure 1996; Peijnenburg et al. 
1999; McBride et al. 2003; Walker et al. 2003; Bongers et al. 2004). The method has the 
advantage of being simple to use in the laboratory and the results between laboratories 
are less variable than with some other methods (Quevauviller 1998). This is the same 
solution as is used to measure soil pH in many laboratories. A similar solution but with a 
slightly higher concentration is also recommended in Chapter 11 for use in estimating 
bioavailable Al and Mn. Gray et al. (2003) compared several extraction procedures to the 
"labile pool" as measured by isotope dilution; they found that CaCl2 provided the closest 
comparison to this pool. 

It is well known that metal and metalloids added to soils may become strongly bound to the 
soil particle surfaces (Ross 1994). Whether this is due to specific adsorption or precipitation, 
the elements that become fixed are mostly found on sites that are in contact with the soil 
solution. A strong chelating agent should be able to remove trace elements from a wide 
range of surface adsorption/precipitation sites. Although all of this "fixed" metal would 
not be immediately available, there are studies that show a good correlation between 

EDTA-extractable metal and content in biological tissue (Ure 1996; De Gregori et al. 2004). 
The extraction with 0.05 M EDTA is a good choice for estimating this "potentially 
available" fraction (Quevauviller 1998). 

The choice of digestion methods is wide and the USEPA alone recommends four different 
acid mixtures or procedures (Ming and Ma 1998). Total metal content is only obtained when 
HF is included in the digestion procedure; otherwise silicate minerals are not dissolved. Most 
laboratories prefer to use a method that does not include HF due to the danger of working 
with it; HF causes severe burns to skin or eyes. Trace elements found in the silicates are 
certainly not immediately available and there is a good chance that these trace elements are 
related to minerals found in the parent material rather than added by anthropogenic activity. 
For general laboratory purposes the HNO3 procedure proposed here should provide a very 
good estimate of trace elements in contaminated soils. Although there are several alternate 
methods using HNO3/HCI available (USEPA 1994), an acid mixture without HC1 is 
preferred for inductively coupled plasma-mass spectrometry (ICP-MS) analysis. A prelim- 
inary study prepared by Canada's National Water Research Institute shows very good results 
with this USEPA 3051 method (Alkema and Blum 2001). It is preferred as an appropriate 
method for numerous elements. 

(MACDONALD ET AL. 2004a,b) 

Soil samples collected in the field and brought back to the laboratory for extraction yield 
solutions with significantly higher concentrations than solutions collected by lysimeters in 
the field from the same soil horizon. To obtain soil solutions that are comparable to those 
sampled with lysimeters it is necessary to first remove the relatively soluble material that 
accumulates in a sample following disturbance; this is particularly important here as we are 
using air-dried soil samples. 

The removal of soluble material is done by "washing" the column with deionized water 
until the ionic strength drops to values similar to those found in the field. The column is then 
equilibrated with an artificial soil solution containing Ca, CI, and SO4, the ions most 
common in the soil solutions we have sampled in eastern Canada. 

Initially researchers should monitor changes in pH and electrical conductivity during the 
washing and equilibrium phases so that they can see whether the concentrations are tending 
toward relatively constant values. The procedure described below appears to be suitable for a 
wide range of soils we have tested, but may not work with all soils. 

The suction needed to pull the solution through the soil columns can be generated using two 
very different types of apparatus. The method described below uses a commercially available 
column extraction apparatus, although it is necessary to replace the original syringes that have a 
black rubber seal with ones that have an all polyethylene plunger; the leaching can also be done 
using a multichannel peristaltic pump but it is more difficult to achieve uniform flow rates. 

10.2.1 Materials and Reagents 

1 80 ml_ of 80 (juM CaCI 2 -CaS0 4 for each column for the four leaching days. This is 
prepared by dissolving 0.01 1 8 g of CaCI 2 • 2H 2 plus 0.01 09 g of CaS0 4 in 2 L of 
ultrapure water. 


2 Polyethylene syringes (60 ml_) (HNO3 washed) and high-density polyethylene 
(HDPE) frits that fit the syringes tightly. 

j Vacuum extractor capable of flow rates of 30 ml_ hT 1 and 2-3 ml_ h~ 1 . 

4 10% nitric acid (HNO3) — trace metal grade: dilute 10 mL of concentrated acid to 
1 00 mL in a volumetric flask. 

5 Ultrapure water — usually produced by passing deionized or reverse osmosis 
water through a special system to produce water with an electrical conductivity 
less than 18 |xS cm -1 . 

6 0.45 jjim membrane filters (nylon or polycarbonate). 
10.2.2 Procedure 


1 Extractions should be carried out in an incubator at a temperature between 4°C 
and 6°C. The apparatus for each soil sample consists of three 60 mL syringes that 
are connected together vertically; only the syringe barrels are used for the upper 
two. The upper syringe is used as the reservoir for the solution and is connected to 
the middle syringe, holding the soil, with a tight-fitting stopper. The lowest syringe 
is slowly withdrawn by the vacuum extraction device and the solution is sucked out 
of the upper syringe, through the soil sample and into the bottom syringe. Make sure 
that there are no leaks in the system or the flow rate will be compromised. 

2 Air dry, homogenize, and sieve the soils to 2 mm. Weigh and pack 1 5 g of mineral 
soil (5 g of forest floor) into a 60 mL syringe. Encase the soil between two HDPE 
frits. Insert the upper syringe into the column to hold the solution. 

3 Add 30 mL aliquots of ultrapure water. Apply suction at a rate of 30 mL hT 1 . After 
all the water has passed through the column wait 2 h before starting the next 
leaching. Repeat twice more for a total of 90 mL. 

4 At the end of step 3, wait 2 h before starting the treatment with CaCI 2 -CaS0 4 . 

Leach the columns with 20 mL of 80 (Ji/Vf of CaCI 2 -CaS0 4 at a rate of 2 to 
3 mL h~ 1 every 24 h during 4 days. Make sure that air enters the column at the 
end of each leaching cycle to prevent the columns from becoming anaerobic. 
Collect the leachates in separate acid-washed bottles. 

Measure the pH and EC of each of the leachates. Keep the leachates from the last 3 
days and mix them together to obtain one sample of about 50 mL. Filter solutions 
through 0.45 |xm membrane filters under vacuum, and collect solutions in poly- 
ethylene bottles. Preserve the solution or a subsample of the solution (if part of the 
solution is being kept for other analyses) after filtration by adding 0.2 mL of 1 0% 
HNO3 per 10 mL of solution, and analyze as soon as possible. 

10.2.3 Calculations 

The extraction method is not intended as a quantitative analysis of, for example, the water- 
soluble fraction; however it is appropriate for estimating solid- solution trace element parti- 
tioning or to estimate the concentration of trace metals in water leaching from a site. 

Partitioning coefficients (K^) are calculated as the ratio of total metals (determined through 
hot acid extraction, see Section 10.5) in mg kg -1 over metals in solution as mg LT 1 and have 
the unit L kg -1 : 

Dissolved metal 


7 Care must be taken to assure that all plasticware in contact with final solutions has 
been soaked 24 h in 15% HNO3 and rinsed thoroughly with high-quality deion- 
ized water. Blanks should be carried through the entire extraction procedure to 
assure that solutions are not contaminated by outside sources. 

2 Column methods are prone to variability. Great care must betaken to pack columns 
consistently. We propose adding soil in three steps and compacting the column 
with 1 light taps of a syringe plunger with the seal removed at each step. 

j Work in duplicate, and include blanks and quality control samples in each batch. 

4 This is a fairly time-consuming procedure that takes 5 days to complete. On the 
first day (usually Monday), the three washing solutions are passed through the 
columns and collected — 3 h each washing (1 h to draw the solution through 
and 2 h of equilibration); this makes for a 10 h day. The first CaCI 2 -S0 4 
solution is added to the columns when we leave in the evening of the first 
day, drawn through the columns during the night, and then collected the next 
morning. This leaching with the CaCl2-SC>4 solution is repeated on the evenings 
of days 2-4. 


10.3.1 Materials and Reagents 

7 Centrifuge and 50 ml_ Boston-type polyethylene centrifuge tubes (HNO3 acid 

2 End-over-end shaker (15 rpm). 

3 Calcium chloride, 0.01 M; in a 1 L polyethylene volumetric flask, dissolve 1 .47 g 
of CaCI 2 • 2H 2 in ultrapure water and make to volume. 

4 10% nitric acid (HNO3) — trace metal grade: dilute 10 ml_ of concentrated acid to 
1 00 ml_ in a volumetric flask. 

5 0.45 jjim membrane filters (nylon or polycarbonate). 

10.3.2 Procedure 

/ Work at room temperature. Before taking a subsample, make sure your sample is 
very well homogenized by mixing the sample thoroughly for about a minute. 
Work in triplicates. Take a subsample of each soil to estimate moisture content. 
Include two blank solutions (tube and solution without soil) and two quality 
control samples in each batch of extractions. 

2 Weigh about 2.500 g of soil into a 50 ml_ centrifuge tube and record weight. Add 
25 ml_ of 0.01 M CaCI 2 to each tube, cap and shake on the end-over-end shaker 
for 3 h at 1 5 rpm. 

3 Take a subsample to measure pH and discard (one per triplicate). Centrifuge at 
5000 g for 1 min. Filter, with great care to avoid contamination, through 0.45 |xm 
membrane under low vacuum. Keep the filtrate in a 30 ml_ polyethylene bottle. 
Preserve the solution or a subsample of the solution (if part of the solution is being 
kept for other analyses) after filtration by adding 0.2 ml_ of 1 0% HNO3 per 1 mL 
of solution, and analyze as soon as possible. If dilutions are required, the amount 
of HNO3 should be kept constant. 

10.3.3 Calculations 

M (|xg g- 1 ) = C (|xg L- 1 ) x 0.025 L/(wt. soil g x (1 - mc)) (10.2) 

where M is the metal content, C is the concentration measured, and mc is the moisture 
content expressed as a 2-decimal fraction (i.e., 5% = 0.05). 


1 Great care must be taken to avoid contamination. Polyethylene should be used to 
avoid sorption/desorption of metals to or from the walls of the containers. 
Centrifuge bottles, sample bottles, filtration units must be clean and acid washed 
followed by an acid soaking in 15% HNO3 for 24 h and thoroughly rinsed with 
double-deionized water with a final rinse with ultrapure (or equivalent) water. 

2 The version given here is an adaptation from Quevauviller's method; it is a 
compromise using smaller sample size for routine analysis. The reader is invited 
to read the original reference cited. 

3 Reproducibility is difficult to achieve in this kind of extraction; care should be 
given to each step of the procedure. 


4 As part of the quality control procedure, the analysis of one sample should be 
repeated in each batch of extractions to evaluate the reproducibly of the whole 
experiment. When the value of the quality control sample falls outside 2 standard 
deviations, calculated for all measurements of that sample, the whole batch 
should be reanalyzed. 

10.4.1 Materials and Reagents 

1 Centrifuge and 50 ml_ Boston-type polyethylene centrifuge tubes (in addition 
to the acid-washing procedures described in the comments, the labware 
must be rinsed with EDTA followed by a thorough water rinse before use in 
this experiment). 

2 End-over-end shaker (15 rpm). 

3 NH 4 -EDTA salt solution 0.05 M: EDTA in its ammonium salt form is difficult to 
obtain in a pure form. The following method offers a means of cleaning common 
reagent-grade chemicals. 

4 Ultrapu re water. 

5 0.45 |jim membrane filters (nylon or polycarbonate). 
To purify H A EDTA 

1 Weigh about 100 g H 4 EDTA acid and put in a Teflon beaker. 

2 Add about 1 50 ml_ of 2% HNO3 trace metal grade. 

3 Stir 10 min on magnetic stirrer. 

4 Let settle and decant and discard the supernatant. 

5 Repeat at least three times with the addition of about 1 50 ml_ HNO3, stir, settle, 

6 Rinse with ultrapure water (or equivalent) using the same procedure as above (i.e., 
add about 1 50 ml_ water, stir, settle, decant) at least three times. 

7 Dry the prepared chemical in a warm oven (~40°C) overnight (you might have to 
crush the H 4 EDTA before storing). 

To prepare pure NH4OH 

Trace metal-grade ammonia can be purchased, but it can also be prepared in the laboratory 
using reagent-grade ammonia; you need very clean labware and an efficient fumehood. 


Under the fumehood, in a very clean desiccator, place a beaker with about 100 mL of 
concentrated ACS reagent-grade NH4OH and another Teflon beaker with 100 mL ultrapure 
water. Replace cover and let stand overnight. The next morning you will have pure 1:1 
diluted ammonia in your Teflon beaker. 

To prepare purified ammonium EDTA salt 

In a 2 L volumetric flask containing about 1.8 L ultrapure water, add 29.2 g purified 
H4EDTA. Place on a magnetic stirrer under a fumehood and add about 25 mL of 
purified 1:1 ammonia prepared as described above. Stir. Continue adding NH4OH gradually 
until the H4EDTA completely dissolves (around pH 6). Adjust to pH 7.0 (±0.1) and make to 
volume with ultrapure water. Store in a well stoppered 2 L polyethylene bottle. 

10.4.2 Procedure 

7 Work at room temperature. Before taking a subsample of soil, make sure your 
sample is very well homogenized by mixing thoroughly for about a minute. Work 
in triplicates. Take a subsample of each soil to estimate moisture content. Include 
two blank solutions (tube and solution without soil) within each batch of extrac- 
tion. Weigh about 1 .000 g of soil in a 50 mL centrifuge tubes and record weight. 

2 Add 25 mL of purified 0.05 M NH4-EDTA to each tube, cap and shake on the 
end-over-end shaker for 1 h at 15 rpm. 

j Centrifuge at 5000 g for 10 min, if possible, maintain the temperature of the 
centrifuge at 20°C, filter through a 0.45 jjim membrane, and keep in well-sealed 
polyethylene bottle at 4°C. Dilute with ultrapure water for analysis. Make sure the 
standards used for calibration are in the same matrix as the diluted solution. 

10.4.3 Calculations 

M (|xg g- 1 ) = C (jjig L- 1 ) x 0.025 L/(wt. soil g x (1 - mc)) (10.3) 

where M is the metal content, C is the concentration measured, and mc is the moisture 
content expressed as a 2-decimal fraction (i.e., 5% = 0.05). 


1 EDTA is a powerful extractant that is capable of extracting significant quantities of 
trace elements from high affinity sites on the soil surface. Likewise EDTA will extract 
all elements from the surfaces of plastic and glassware if in contact with the solution. 
Consequently it is very important to preclean labware with purified 0.5 M H 2 EDTA 
followed by a complete water rinse to avoid contamination of the samples. 

2 It is also important to use the same matrix for samples and standards. Do not try to 
acidify the solution before, or while measuring the content of metals as this could 
cause precipitation of the EDTA. 



10.5.1 Materials and Reagents 

1 Specialized microwave digestion system with Teflon liners 

2 Nitric acid (HN0 3 ) — trace metal grade 

3 1 00 ml_ polyethylene volumetric flasks 

4 Ultrapu re water 

10.5.2 Procedure 

7 Follow the safety directions from the microwave system manufacturers. Work 
under a fumehood and wear protective clothing and equipment. 

2 Weigh up to 0.500 g of soil sample. If sample contains high content of organics or 
carbonates, decrease the amount weighed. Organic soils and forest floor horizons 
should be 0.200 g of sample. 

3 Add 1 mL HNO3. If a strong reaction is observed, allow the samples to stand for 
several hours (or over night) before sealing the containers to decrease the possi- 
bility the containers will vent during heating. Close containers and place in the 
microwave system. Follow the manufacturer's recommendations for a heating 
program and maintain a temperature of 185°C for at least 10 min. 

4 After completion of the digestion, let cool and transfer the whole sample to a 
1 00 mL volumetric flask (final acidity 1 0% HNO3). Let settle overnight and decant 
supernatant into a 30 mL polyethylene bottle. 

5 Dilute five times with ultrapure water for analysis on an ICP-MS (final acidity 2%). 
Standards should be prepared in the same matrix. 

10.5.3 Calculations 

M (|xg g _1 ) = C (|xg LT 1 ) x DF x 0.100 L/(wt. soil g x (1 - mc)) (10.4) 

where M is the metal content, C is the concentration measured, mc is the moisture content 
expressed as a 2-decimal fraction (i.e., 5% = 0.05), and DF is the dilution factor. 


Refer to the manufacturer's instructions on the proper use of the microwave digestion system. 
Due to the high pressures that are developed in the reaction vessels, it is important to use a 
microwave digestion system designed specifically for this purpose. In addition to the danger of 
having a vessel explode while being heated, it is also very important to properly cool the 
vessels before trying to open them. Letting them sit for 30 min in an ice bath is recommended. 


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Beckett, P.T.H. 1989. The use of extractants in 
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MacDonald, J.D., Belanger, N., and Hendershot, 
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W.H. 2004b. Column leaching using dry soil repro- 
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McBride, M.B., Mibarger, E.A., Richards, B.K., 
and Steenhuis, T. 2003. Trace metal accumula- 
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Chapter 1 1 

Readily Soluble Aluminum and 

Manganese in Acid Soils 

Y.K. Soon 

Agric. allure and Agri-rood Canada 
Beaverlodge, Alberta, Canada 

N. Belanger 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

W.H. Hendershot 

McGill University 
Sainte Anne de Bellevue, Quebec, Canada 


Approximately 5% of the 3.95 billion ha of acid soils is used for agricultural production 
while 67% supports forests and woodlands (von Uexkiill and Mutert 1995). Plant growth in 
acid soils is usually limited by low pH and/or Al toxicity. The solubility of Al and Mn in 
mineral soils increases rapidly when soil pH drops below a value of 5 so that low pH and 
high soluble Al and Mn concentrations are interrelated. Although the availability of plant 
nutrients such as P and Ca can be limiting at low soil pH (Foy 1984; Asp and Berggren 
1990), Al toxicity is probably the foremost growth-limiting factor in acid soils (Andersson 
1988). Root growth, and consequently water and nutrient uptake, are inhibited when dissolved 
Al attains toxic levels in soil solutions. 

Dissolved Al in acid soil solution is typically comprised of monomeric Al ions (e.g., 
Al 3+ , Al(OH) 2+ , and Al(OH) 2 + ) as well as organically complexed and polynuclear forms 
of Al (e.g., Al 2 (OH) 2 4+ and Al 13 4 (OH) 24 7+ (Akitt et al. 1972)). Polynuclear and organic- 
ally complexed Al species are considered to have little, if any, phytotoxicity (Andersson 
1988; Wright 1989) although there is some contrary evidence for polymeric Al (Bartlett and 
Diego 1972; Hunter and Ross 1991). Ideally, the soil solution in the root zone should be 
analyzed for phytotoxic Al species concentration. For diagnostic purposes, such procedures 


would be too time-consuming. Typically, therefore, the soil is extracted with a dilute neutral 
salt solution that would perturb the ionic equilibrium as little as possible, and for a time 
period sufficient only to bring into solution readily soluble Al (i.e., not associated with the 
solid phase). Included in such extracts would be mainly monomeric and polymeric Al and Al 
complexed by organic ligands of low-molecular weight. 

11.1.1 Aluminum and Manganese Toxicity in Agricultural Soils 

Aluminum concentration in plant tissues cannot be used to confirm Al toxicity since it does 
not accumulate in aboveground plant tissues. Although Mn accumulates in plants somewhat 
in proportion to plant injury in acid soils, its concentration in plant tissues is not a reliable 
indicator of its toxicity (Foy 1984). Therefore, Al and Mn toxicity diagnostic criteria, 
especially Al, have been approached through soil analysis. In a meta-analysis of Al toxicity 
thresholds for crops and forages, Belanger et al. (1999) found that the total dissolved Al 
concentrations associated with negative effects in 10% and 50% of the studies were, 
respectively, 0.003 and 0.02 mM. However, different crops and forages, and even varieties 
within a species, vary in their sensitivity to dissolved Al. 

Soil acidity is usually corrected by liming or adding calcium amendments to the soil. The 
lime requirement (see Chapter 12), i.e., the amount of CaC03 or its equivalent that has to be 
applied to the soil to raise its pH to a certain desired value, usually 6.5, can be determined by 
equilibrating a soil sample with a buffered salt solution and measuring the pH (Shoemaker 
et al. 1961; McLean et al. 1978). Kamprath (1970) suggested that liming can also be based 
on soluble Al extracted from acid soils by a neutral unbuffered salt solution, such as 1 M 
KC1, at least for soil groups such as ultisols and oxisols. In Canada, Hoyt and Nyborg 
(1971a,b, 1972, 1987) showed that crop response on acid soils was closely related to 0.01 M 
and 0.02 M CaCL-soluble Al and Mn. With the exception of alfalfa, yields of the test crops 
were more closely correlated with dilute CaCVextractable Al than soil pH or exchangeable 
Al (1 M KCl-exchangeable) (Webber et al. 1982; Hoyt and Nyborg 1987). There was little 
response of barley (Hordeum vulgare L.), an Al-sensitive crop, to lime when dilute CaCi2- 
extractable Al approached 1 mg kg -1 (Hoyt et al. 1974). Webber et al. (1977) found that the 
amount of lime required to lower 0.02 M CaCl2-extractable Al to 1 mg kg -1 was less than 
the lime requirement to achieve a pH of 6 as determined by the Shoemaker-McLean-Pratt 
(SMP) procedure (Shoemaker et al. 1961). Research studies in Australia, New Zealand, and 
the United States also showed that the Al and Mn extracted by dilute CaCl2 solution are 
suitable diagnostic criteria for Al and Mn toxicities in acid soils (Wright et al. 1988, 1989; 
Close and Powell 1989; Conyers et al. 1991). 

11.1.2 Aluminum and Manganese Toxicity in Forest Soils 

Forest decline since the last 20 years in central Europe and eastern North America has been 
attributed to several environmental stresses such as gaseous pollutant injury and water stress 
(Hinrichsen 1986). As with crop species, it was also shown that increased Al activity in the 
soil solution has adverse effects on tree functions and growth (see review by Cronan and 
Grigal 1995). Forest soils are typically acidic (pH < 5) and thus, the solubility of toxic Al and 
Mn is generally high. Manganese toxicity to trees was not studied as much. However, as for 
crop species, foliage Mn status appears to be a better indicator of solution Mn levels 
compared to Al, but its toxicity is difficult to show due to concomitant high availability of 


Al (Hoyle 1972; Kazda and Zvacek 1989). In solution cultures, Hoyle (1972) found that 
foliage levels of 441 mg Mn kg -1 (solution Mn 0.091 mAf) in yellow birch were optimal for 
growth but levels above 1328 mg Mn kg -1 (solution Mn 0.45 mM) were detrimental. In 
air-polluted European forests, Mn concentrations in soil solutions ranged from 0.018 to 
0.36 mM, depending on the acid load and parent material type (Kazda and Zvacek 1989). 

Joslin and Wolfe (1988) found that dissolved inorganic monomeric Al, total Al, Al 3+ activity 
as well as SrCl 2 -extractable soil Al explained respectively 79%, 74%, 61%, and 61% of 
the variability in root biomass. The SrCl2-extractable Al and inorganic monomeric Al 
concentrations at which significant reductions in root branching and fine root biomass was 
first observed were 10 mg kg -1 and 0.1 mM, respectively (Joslin and Wolfe 1988, 1989). We 
used the data from Joslin and Wolfe (1988) to assess how SrCl2-extractable Al and inorganic 
monomeric Al concentrations are related. Excluding one outlier from the Becket site, 71.2% of 
the variability in inorganic monomeric Al concentrations (y) can be predicted from SrCV 
extractable Al concentrations (x) using the following power function: y = 4.70 a24<k . A soil with 
10 mg kg -1 of SrCl2-extractable Al therefore corresponds to a solution inorganic monomeric 
Al level of about 0.05 mM, which is close to the 0.07 mM toxicity threshold obtained from a 
meta-analysis computed by Belanger (2000). Finally, Joslin and Wolfe (1989) discussed the 
unique response of trees at the Becket site (i.e., substantial root growth despite the relatively high 
foliage Al concentrations and SrC^-extractable soil Al levels) and suggested that most of the 
Al absorbed by trees was organically bound. It is known that the complexation of metal cations 
by organics enhances plant uptake (Arp and Ouimet 1986) but this form is nontoxic to trees 
(Rost-Siebert 1984). Therefore, the SrCh method may not always be a reliable indicator of the 
potential toxicity of Al in soils where organically bound Al dominates. 


Hoyt and Nyborg (1972) reported that when Al and Mn in acid soils were extracted with 2.5 to 
40 mM CaCb solutions, generally better correlations with the yield response of three crops 
were obtained if the extractant was 20 mM CaCl2. A subsequent study showed that a 5 min 
shaking was adequate and gave only slightly lower concentrations of Al than a 1 h shaking and 
twice the Al concentration as 10 mM CaCb (Hoyt and Webber 1974). According to Webber 
et al. (1977), liming is likely not needed for Canadian acid soils when extracted Al is 1 mg kg~ 
or less. Close and Powell (1989) also used this extraction procedure for New Zealand soils. 

11.2.1 Materials and Reagents 

j 0.02 M CaCI 2 : Dissolve 5.88 g of reagent-grade CaCI 2 • 2H 2 in about 250 ml_ of 
deionized water and dilute to 2 L. 

2 50 ml_ centrifuge tubes and rubber stoppers. 

j Whatman No. 42 filter paper or equivalent. 

4 Centrifuge, with rotors accepting 50 ml_ centrifuge tubes. 

5 Reciprocal shaker, 
g Liquid dispenser. 


11.2.2 Procedure 

7 Weigh 10 g of soil (<2mm) into centrifuge tube. 

2 Dispense 20 ml_ of the 0.02 M CaCI 2 reagent into the centrifuge tube and stopper 

3 Shake 5 min on the shaker (1 20 oscillations min~ 1 ). 

4 Remove from shaker and centrifuge at 1 250 gfor 1 min to facilitate rapid filtering. 

5 Filter through a fluted filter paper into receptacle for storing the extract. 


A 0.01 M CaCl2 solution is closer in ionic strength to the soil solution of agricultural soils 
than is a 0.02 M solution and it is also commonly used to assess other soil chemical 
properties such as pH and soluble P (Soon 1990). Therefore, it may be advantageous to 
use 0.01 M instead of 0.02 M CaCl2 solution when those soil properties are also to be 
determined. However, a critical level of 0.01 M CaCl2-soluble Al has not been proposed. 
The above procedure has not been tested for soils with organic matter content much higher 
than 10%. 


Since increased Ca availability alleviates toxic effects of Al on trees (i.e., Al toxicity is mostly 
indirect — it is toxic due to its antagonistic effects on divalent cation uptake (Cronan and Grigal 
1995)), the SrCl2 method is advantageous relative to the commonly used CaCl2 procedure in 
agricultural soils as it allows the quantification of extractable Ca and other cations as well as 
Ca and Mg to Al ratios. Strontium is slightly more efficient at displacing Al than Ca, but the 
difference is only about 5% at this ionic strength (0.01 M). However, no work has been done on 
the relative amounts of Al and Mn exchanged with SrCl2, CaCl2, and BaCl2. Such a study 
would help to clarify the need for using different extractants when the amount of potentially 
toxic elements was under investigation. The SrCl2 method of Joslin and Wolfe (1989) is 
described here after some modifications based on the suggestions of Heisey (1995). 

11.3.1 Materials and Reagents 

/ 0.01 M SrCI 2 : Dissolve 5.332 g of reagent-grade SrCI 2 • 6H 2 in about 250 ml_ of 
distilled/deionized water and make to volume in a 2 L volumetric flask. 

2 50 ml_ centrifuge tubes and screw caps. 

j Ultracentrifuge accepting 50 ml_ tubes. 

4 End-over-end shaker. 

5 Pipette and liquid dispenser. 


11.3.2 Procedure 

1 Weigh 10 g of soil (dried and <2mm) into centrifuge tube. 

2 Add 20 ml_ of 0.01 M SrCI 2 to the centrifuge tube. 

3 Shake for 60 min at 1 5 oscillations min -1 . 

4 Remove from shaker and centrifuge for 30 min at 7000 g. 

^ Pipette off the supernatant and retain in container for analysis. 


An end-over-end shaker is used here because it is more efficient in wetting and mixing forest 
soils with a high litter/organic matter content and using a low soikextractant ratio. The low 
soikextractant ratio results in a thick suspension that typically requires a high-speed centri- 
fuge to separate. 


Aluminum in the extracts can be measured by atomic absorption (Webber 1974), inductively 
coupled plasma atomic emission spectroscopy (ICP-AES) (Carr et al. 1991), or by color- 
imetry (Hoyt and Webber 1974; Carr et al. 1991). Atomic absorption or ICP-AES will give 
results for total dissolved Al whereas colorimetry should yield results for Al "reactive" 
with the chromogen. What will be included as "reactive" Al depends on the equilibration 
time allowed for color development. Shorter reaction times should yield mainly labile 
(monomeric) Al whereas longer reaction times would include the determination of poly- 
meric and complexed Al. Grigg and Morrison (1982) showed that the pyrocatechol violet 
(PCV) method was superior to the Aluminon (aurintricarboxylic acid triammonium salt) 
method in precision, and automating the procedure resulted in further improvement in 
its precision. The pyrocatechol method was also recommended by Conyers et al. (1991). 
The method below is an adaptation of Wilson and Sergeant (1963). The procedure is simple 
and reliable. 

11.4.1 Reagents 

7 0.1 % (w/v) PCV. Keep in a dark glass bottle. 

2 0.1% (w/v) o-phenanthroline (OP). Store in a polyethylene bottle. 

3 10% (w/v) hydroxylamine hydrochloride (HH). Keep in a polyethylene bottle. 

4 10% (w/v) ammonium acetate (buffered at pH 6.2 using acetic acid). Store in a 
polyethylene bottle. 

5 Aluminum working standard solution: from a stock standard solution contain- 
ing 1 g Al L -1 , prepare a working standard containing 100 mg Al L~ 1 in CaCI 2 


solution of the same molarity as the soil extractant. By further dilution, prepare 
5 standards over the range of 0.1 to 2.5 mg Al L in CaCb solution of the same 
molarity. For forest soils, standards should be prepared in 0.01 M SrCI 2 - 

11.4.2 Procedure 

/ Pipette 2 ml_ of extract or standard solution into 1 6 mm x 1 25 mm culture tubes. 
The tubes should be prewashed with 0.1 M HCI. Sample solutions should contain 
no more than 5 |xg Al. 

2 Add sequentially 0.5 ml_ each of PCV, OP, and HH, gently swirling the contents of 
the tube after each addition. In a batch of samples, each reagent should be added 
to all samples before adding the next reagent. 

j Add 6 ml_ of the buffer solution, stopper and invert the tube three times and allow 
to stand for 1 h. 

4 Measure absorbance at 580 nm with a spectrophotometer using 1 cm cuvette. Plot 
the absorbance values against juug Al. The jxg Al value read off the calibration 
curve gives extracted Al level in mg kg -1 soil. If dilution of the extract is required, 
multiply by the dilution factor. 


The extracts should be analyzed with minimum delay. If delays are inevitable, acidify the 
samples slightly to prevent polymerization of Al monomers. The PCV powder and the 
prepared solution should be kept in the dark in tightly sealed containers. Interference by 
iron is diminished by the OP and HH reagents. Color development is maximal and stable 
between 1 and 2 h, after which the color gradually declines. Reagent blank values are 
determined using the soil extractant. It is advisable to use freshly prepared PCV solution. 
The other reagents are stable for at least 4 weeks when stored at room temperature. 

The analysis as described should include monomeric and polymeric Al and weakly com- 
plexed Al. Kerven et al. (1989) described a PCV procedure with a reaction time of 60 s to 
measure only monomeric Al. For forest surface soils, which typically have much higher 
organic matter content than agricultural soils, the difference between measuring total 
dissolved Al and monomeric inorganic Al should be more critical. Also much calibration 
of extractable soil Al with crop response has been done using dissolved total Al (Hoyt and 
Webber 1974; Hoyt and Nyborg 1987). An autoanalyzer PCV method that uses ion-exchange 
to separate inorganic monomeric Al from organically complexed Al has been described by 
McAvoy et al. (1992). 


Manganese in the soil extract is determined by atomic absorption spectrometry using an 
oxidizing air-acetylene flame. ICP-AES analysis would also be convenient, especially if Al 
is to be determined. 


Akitt, J.W., Greenwood, N.N., Khandelwal, B.K., 
and Lester, G.D. 1972. 27 A1 nuclear magnetic 
resonance studies of the hydrolysis and polymer- 
ization of the hexa-aquo-aluminium (III) cation. 
/. Chem. Soc, Dalton Trans: 604-610. 

Andersson, M. 1988. Toxicity and tolerance of 
aluminium in vascular plants: a literature review. 
Water Air Soil Pollut. 39: 439^162. 

Arp, P. and Ouimet, R. 1986. Uptake of Al, Ca, 
and P in black spruce seedlings: effect of organic 
versus inorganic Al in nutrient solutions. Water 
Air Soil Pollut. 31: 367-375. 

Asp, H. and Berggren, D. 1990. Phosphate and 
calcium uptake in beech (Fagus sylvatica L.) in 
the presence of aluminium and natural fulvic 
acids. Physiol. Plant 80: 307-314. 

Cronan, C.S. and Grigal, D.F. 1995. Use of 
calcium-aluminum ratios as indicators of stress in 
forest ecosystems. /. Environ. Qual. 24: 209-226. 

Foy, CD. 1984. Physiological effects of hydro- 
gen, aluminum and manganese toxicities in acid 
soil. In F. Adams, Ed. Soil Acidity and Liming, 
2nd edn. ASA-CSSA-SSSA, Madison, WI, 
pp. 57-97. 

Grigg, J.L. and Morrison, J.D. 1982. An auto- 
matic colorimetric determination of aluminium 
in soil extracts using catechol violet. Commun. 
Soil Sci. Plant Anal. 13: 351-361. 

Heisey, R.M. 1995. Growth trends and nutritional 
status of sugar maple stands on the Appalachian 
plateau of Pennsylvania, USA. Water Air Soil 
Pollut. 82: 675-693. 

Bartlett, R.J. and Diego, D.C. 1972. Toxicity of 
hydroxy aluminum in relation to pH and phos- 
phorus. Soil Sci. 114: 194-200. 

Belanger, N. 2000. Investigating the long-term 
influence of atmospheric deposition and forest 
disturbance on soil chemistry and cation nutrient 
supplies in a forested ecosystem of southern 
Quebec, PhD thesis, Department of Natural 
Resource Science, McGill University, Montreal, 
QC, Canada, 164 pp. 

Belanger, N., Fyles, H., and Hendershot, W.H. 
1999. Chemistry, bioaccumulation and toxicity 
of aluminum in the terrestrial environment — 
PSL2 assessment of aluminum salts. A report pre- 
pared for the Commercial Chemicals Evaluation 
Branch, Environment Canada, ON, Canada, 78 pp. 

Carr, S J., Ritchie, G.S.P., and Porter, W.M. 1991. 
A soil test for aluminium toxicity in acidic sub- 
soils of yellow earths in Western Australia. Aust. 
J. Agric. Res. 42: 875-892. 

Close, E.A. and Powell, H.KJ. 1989. Rapidly 
extracted (0.02 M CaCl 2 ) 'reactive' aluminium 
as a measure of aluminium toxicity in soils. 
Aust. J. Soil. Res. 27: 663-672. 

Conyers, M.K., Poile, G.J., and Cullis, B.R. 1991. 
Lime responses by barley as related to available 
soil aluminium and manganese. Aust. J. Agric. 
Res. 42: 379-390. 

Hinrichsen, D. 1986. Multiple pollutants and 
forest decline. AMBIO 15: 258-265. 

Hoyle, M.C. 1972. Mangan 
birch (Behtla alleghanien. 
Plant Soil 36: 229-232. 

,e toxicity in Yellow 
Butten) seedlings. 

Hoyt, P.B. and Nyborg, M. 1971a. Toxic metals 
in acid soil. I. Estimation of plant-available 
aluminum. Soil Sci. Soc. Am. Proc. 35: 236-240. 

Hoyt, P.B. and Nyborg, M. 1971b. Toxic metals 
in acid soil. II. Estimation of plant-available man- 
ganese. Soil Sci. Soc. Am. Proc. 35: 241-244. 

Hoyt, P.B. and Nyborg, M. 1972. Use of dilute 
calcium chloride for the extraction of plant- 
available aluminum and manganese from acid 
soil. Can. J. Soil Sci. 52: 163-167. 

Hoyt, P.B. and Nyborg, M. 1987. Field calibra- 
tion of liming responses of four crops using soil 
pH, Al and Mn. Plant Soil 102: 21-25. 

Hoyt, P.B., Nyborg, M., and Penney, D. 1974. 
Farming Acid Soils in Alberta and Northeastern 
British Columbia, Publication 1521, Agriculture 
Canada, Ottawa, ON. 

Hoyt, P.B. and Webber, M.D. 1974. Rapid meas- 
urement of plant-available aluminum and manga- 
nese in acid Canadian soils. Can. J. Soil Sci. 54: 


Hunter, D. and Ross, D.S. 1991. Evidence for a 
phytotoxic hydroxyl-aluminum polymer in orga- 
nic soil horizons. Science 251: 1056-1058. 

Joslin, J.D. and Wolfe, M.H. 1988. Responses of 
red spruce seedlings to changes in soil aluminum 
in six amended forest soil horizons. Can. J. Forest 
Res. 18: 1614-1623. 

Joslin, J.D. and Wolfe, M.H. 1989. Aluminum 
effects on northern red oak seedling growth in 
six forest soil horizons. Soil Sci. Soc. Am. J. 53: 

Kamprath, E.J. 1970. Exchangeable aluminum as 
a criteria for liming leached mineral soils. Soil 
Sci. Soc. Am. Proc. 35: 252-254. 

Kazda, M. and Zvacek, L. 1989. Aluminum and 
manganese and their relation to calcium in soil 
solution and needles in three Norway spruce 
{Picea abies L. Karst) stands of Upper Austria. 
Plant Soil 114: 257-267. 

Kerven, G.L., Edwards, D.G., Asher, C.J., 
Hallman, P.S., and Kokot, S. 1989. Aluminium 
determination in soil solution. II. Short-term col- 
orimetric procedures for the measurement of 
inorganic monomeric aluminium in the presence 
of organic acid ligands. Aust. J. Soil Res. 27: 

McAvoy, D.C., Santore, R.C., Shosa, J.D., and 
Driscoll, C.T. 1992. Comparison between pyro- 
catechol violet and 8-hydroxquinoline procedures 
for determining aluminum fractions. Soil Sci. Soc. 
Am. J. 56: 449-455. 

McLean, E.O., Eckert, D.J., Reddy, G.Y., and 
Trieweiler, J.F. 1978. An improved SMP soil 
lime requirement method for incorporating 
double buffer and quick-test features. Commun. 
Soil Sci. Plant Anal. 8: 667-675. 

Rost-Siebert, K. 1984. Aluminum toxicity in 
seedlings of Norway spruce {Picea abies Karst) 
and beech (Fagus sylvatica L.). In F. Andersson 
and J.M. Kelly, Eds. Workshop on Aluminum 
Toxicity to Trees. Sveriges Lantbruks Universitet, 
Uppsala, Sweden. 

Shoemaker, H.E., McLean, E.O., and Pratt, P.F. 
1961. Buffer method for determining lime 
requirement of soils with appreciable amounts of 
extractable aluminum. Soil Sci. Soc. Am. Proc. 

25: 274-277. 

Soon, Y.K. 1990. Comparison of parameters of 
soil phosphate availability for the northwestern 
Canadian prairie. Can. J. Soil Sci. 70: 227-237. 

von Uexkiill, H.R. and Mutert, E. 1995. Global 
extent and economic impact of acid soil. In RA. 
Date, et al., Eds. Plant-Soil Interactions at Low 
pH: Principles and Management. Proceedings of 
the Third International Symposium on Plant-Soil 
Interactions at Low pH, 12-16 September 1993, 
Brisbane, Australia. Kluwer Academic Pub- 
lishers, Dordrecht, the Netherlands, pp. 5-19. 

Webber, M.D. 1974. Atomic absorptio 

ments of Al in plant digests and neutral salt 

extracts of soils. Can. J. Soil Sci. 54: 81-87. 

Webber, M.D., Hoyt, P.B., and Corneau, D. 1982. 
Soluble Al, exchangeable Al, base saturation and 
pH in relation to barley yield on Canadian acid 
soils. Can. J. Soil Sci. 62: 397^05. 

Webber, M.D., Hoyt, P.B., Nyborg, M., and 
Corneau, D. 1977. A comparison of lime require- 
ment methods for acid Canadian soils. Can. J. Soil 
Sci. 57: 361-370. 

Wilson, A.H. and Sergeant, G.A. 1963. The colori- 
metric determination of aluminium in minerals by 
pyrocatechol violet. Analyst 88: 109-112. 

Wright, R.J. 1989. Soil aluminum toxicity and 
plant growth. Commun. Soil Sci. Plant Anal. 20: 

Wright, R.J., Baligar, V.C., and Ahlrichs, J.L. 
1989. The influence of extractable and soil solu- 
tion aluminum on root growth of wheat seedlings. 
Soil Sci. 148: 293-302. 

Wright, R.J., Baligar, V.C., and Wright, S.F. 
1988. Estimation of plant available manganese 
in acidic subsoil horizons. Commun. Soil Sci. 
Plant Anal. 19: 643-662. 


Chapter 1 2 
Lime Requirement 

N. Ziadi 

Agri( allure and Agri-rood Canada 
Quebec, Quebec, Canada 

T. Sen Tran 

Institute of Research and Development in Agroenvironment 
Quebec, Quebec, Canada 


The soil pH indicates the amount of acidity present in the soil solution and is one of the most 
commonly measured soil properties. It is considered as a standard and routine soil analysis. 
Soil pH affects the solubility and availability of many elements as well as microbial activity 
(Curtin et al. 1984; Marschner 1995). An acid soil commonly has concentrations of Al or Mn 
that are high enough to be toxic to some plants. The target soil pH, which represents the soil pH 
value associated with optimum plant growth, varies with crop species and can be influenced by 
soil type. In general, a soil pH of 6.0 to 7.0 is ideal for most agronomic crops such as corn (Zea 
mays L.), soybean (Glycine max L. Merr.), and wheat (Triticum aestivum). However, a lower 
target pH may be acceptable for other plants such as potato (Solatium tuberosum L.) or 
blueberry (Vaccinium spp.). Liming acid soils to maintain an appropriate pH for plants is, 
therefore, an essential practice for soil and crop management in many areas. 

There are two components of soil acidity that are used in determining lime application: 
active acidity and exchangeable (reserve) acidity. Active acidity is the concentration of H + 
ions in the soil solution phase and indicates whether or not liming is required to reduce soil 
acidity. The exchangeable acidity refers to the amounts of H + ions present on exchange sites 
of clay and organic matter fractions of the soils and affects the amount of lime needed to 
achieve the target soil pH. The greater the exchangeable (reserve) acidity, the more the soil is 
said to be buffered against change in pH and the greater the lime requirement (LR). 

Lime requirement is defined as the amount of agricultural limestone (CaCCh), or any other 
basic material, required to increase soil pH from acidic conditions to a target level that is 
optimum for the desired use of the soil. The nature of soil acidity, along with soil physical 
and chemical properties (mainly soil texture and organic matter content), affects the LR. 
The test used such as soil-lime incubations, soil-base titrations, or soil-buffer equilibrations 
can also affect the recommendation for lime (Aitken 1990; Conyers et al. 2000; Alatas et al. 


2005). Accurate methods to assess the amount of liming materials are essential, and different 
LR tests should be used in different geographical areas based both on research and practical 
experience. The selection of one specific technique to determine LR must also be taken into 
consideration some practical aspects such as the time available to conduct the test, the 
required equipment and supplies, the cost, etc. Many techniques and methods have been 
developed and successfully used worldwide to measure LR and are reported in previous 
studies (McLean 1982; van Lierop 1990). The majority of these methods are based on the 
following principles (Sims 1996): (i) the measured LR should reflect the amount of liming 
material needed to reach the target pH when the lime is applied under field conditions; 
(ii) LR test should accurately measure all forms of acidity (dissociated and undissociated) 
present in a soil; (iii) LR test should be calibrated in the geographic area where the test will 
be used; and (iv) LR test should be calibrated to determine conversion factors between 
limestone and the other liming materials used. 

To estimate the amount of lime required to correct soil acidity and attain a desired soil 
pH, different procedures can be used through field or laboratory studies. Soil-lime 
incubations, soil-base titrations, and soil-buffer equilibrations (Viscarra Rossel and 
McBratney 2003; Machacha 2004; Liu et al. 2005) are the most commonly used methods. 
Estimation of LR based on field studies, however, remains the most accurate means to 
determine LR for a soil, and especially to evaluate new liming materials. Although these 
methods are time consuming and expensive, they are the foundation for the more rapid 
and inexpensive procedures. In routine soil testing laboratories in North America, the 
Adam-Evans (A-E) buffer (Adams and Evans 1962) and the Shoemaker-McLean-Pratt 
(SMP) (Shoemaker et al. 1961) procedures are the most commonly used methods. In 
Canada for example, Nova Scotia and Newfoundland currently use the A-E procedure 
while New Brunswick, Prince Edward Island, Ontario, British Columbia, and Quebec 
use the SMP method. Webber et al. (1977) recommend the SMP method for Canadian 
acid soils. Tran and van Lierop (1982) and van Lierop (1983) also found the method 
to be suitable for acid mineral and organic soils in Quebec. Recently, Warman 
et al. (2000) recommended the replacement of the A-E method with the SMP method 
in Nova Scotia and Newfoundland. For these reasons, only the SMP method is described 
in this chapter. 


12.2.1 Principles 

The SMP method was developed in 1961 from a soil-lime (CaCC^) incubation study using 
14 acidic soils from Ohio (Shoemaker et al. 1961). The accuracy of this procedure relies on 
its calibration of decreasing soil-buffer pH values with increasing LR rates. Originally, this 
procedure was particularly well adapted for determining the LR of soils needing 
LR >4.5 Mg ha~', and with pH values <5.8 and organic matter contents <100 g kg -1 
(McLean 1982). van Lierop (1990) improved the accuracy of the SMP single-buffer method 
at low LR values and proposed the amount of lime required to attain target values of 5.5, 6.0, 
6.5, and 7.0 (Table 12.1). This improvement is obtained by fitting curvilinear instead 
of linear equations to the relationships between soil-buffer pH and incubation LR values 
and is based on a number of LR studies (McLean 1982; Soon and Bates 1986; Tran and 
van Lierop 1993). 


TABLE 12.1 Relationships between Soil SMP-Buffer pH and Lime Requirement Values 
to Achieve pH 5.5, 6.0, 6.5, and 7.0 of Mineral Soils 

Quantity of liming material (Mg ha 1 ) required to reach desired pH 































Source: From van Lierop, W., in R.L. Westerman (Ed.), So/7 Testing and Plant Analysis, 
2nd ed., SSSA, Madison, Wisconsin, 1990, 73-126. 

Lime requirement in Mg CaC0 3 for a furrow layer of 20 cm depth of soil. 

12.2.2 Materials and Reagents 

1 pH meter 

2 Disposable plastic beakers 

3 Automatic pipette 

4 Glass stirring rods 

5 Mechanical shaker 

6 Standard buffers, pH 7.0 and 4.0 

7 SMP buffer solution 

a 0.1 M HCI, 4.0 M NaOH, and 4.0 M HCI solutions 


The SMP buffer solution can be prepared as follows: 

a. Weigh and place in a 10 L bottle the following chemicals: 

• 18 gp-nitrophenol (N0 2 C 6 H 4 OH); 

• 30 g potassium chromate (K 2 Cr0 4 ); 

• 531 g calcium chloride dihydrate (CaCI 2 -2H 2 0). 

b. Add approximately 5 L of distilled water. Shake vigorously as the water is added, 
and continue shaking for a few minutes to prevent formation of a crust over the salts. 

c. Dissolve 20 g of calcium acetate [(CH3COO) 2 Ca • H 2 0] in a separate flask containing 
about 1 L of distilled water. 

d. Add solution from step (c) to that from step (b) and continue shaking for about 2 or 3 h. 

e. Add 100 ml_ of dilute triethanolamine (TEA) solution: TEA (N(CH 2 OH) 3 ) is very 
viscous and difficult to pipette accurately. It is recommended that a dilute TEA 
solution be prepared by diluting 250 ml_ (or 280.1 5 g) of TEA to 1 L with distilled 
water and mix well. 

f. Shake the mixture periodically until it is completely dissolved. This takes about 
6 to 8 h. 

g. Dilute to a final volume of 10 L with distilled water. 

h. Adjust pH to 7.5 + 0.02 by titrating with either 4 M NaOH or 4 M HCI as required. 

i. Filter through fiberglass sheet or cotton mat if necessary. 

j. Verify buffer capacity of prepared SMP buffer by titrating 20 ml_ from pH 7.5 to 
5.5 with 0.1 M HCI. This should take 0.28 ± 0.005 cmol (+) HCI/pH unit. 

The 10 L SMP prepared solution can be used for approximately 500 soil samples. 

12.2.3 Procedure 

7 Measure 10 mL or weigh 10 g air-dried, screened (<2 mm) soil samples in 
appropriate beakers. 

2 Add 1 mL of distilled water and stir with glass rod and repeat stirring periodically 
during the next 30 min. 

3 Measure the soil pH in the beaker (soil + H 2 0) and rinse electrodes with a 
minimum of distilled water. 

4 If the soil pH (H 2 0) is less than the desired pH, add 20 mL of SMP buffer to 
the soil-water mixture (soil-water-buffer ratio is 1:1:2 by volume) and stir with 
glass rod. 


5 Place soil-water-buffer samples on a mechanical shaker for 1 5 min at about 200 
oscillations min -1 . Remove samples from shaker and let stand for 15 min. The 
times of shaking and standing are very important and should be respected. Sims 
(1 996) proposed 30 min of shaking and 30 min of standing. 

6 Adjust the pH meter to read 7.5 with SMP buffer. 

j Stir sample thoroughly and read the soil-water-buffer to nearest 0.01 pH unit. 
Record as soil-buffer pH. 

g Select the amount of lime required to bring the soil to the pH you choose to lime 
the soil, based on soil-buffer pH relationships used in local recommendations 
(e.g., CRAAQ 2003; OMAFRA 2003). 

9 As the SMP buffer solution can affect the accuracy of the glass electrode after 
approximately 200 buffer-pH determinations, it is strongly recommended to 
regenerate the electrodes by appropriate procedure. The combined glass elec- 
trode can be regenerated by immersing it into a plastic beaker containing a 
solution of 10% ammonium hydrogen fluoride (NH 4 F • HF) for 1 min. Since 
the NH 4 F-HF is a hazardous compound, appropriate protection should be 
respected according to its Material Safety Data Sheet. After etching, dip elec- 
trode into 1:1 H2O-HCI solution to remove silicate. Rinse the electrode 
thoroughly with distilled water and immerse in hot 3 M KCI solution (50°C) for 
5 h. The electrolytes in the electrode (saturated KCI or calomel) must be replaced 
if necessary. 


For a LR greater than about 7 Mg limestone ha -1 , it is recommended to divide the rate into 
two or more applications to avoid local overtiming (Brunelle and Vanasse 2004). This is 
important as a liming recommendation assumes that the material is homogeneously incorpo- 
rated into the plow-layer, a precept that is difficult to achieve in practice. When surface 
applying liming material, without significant incorporation (i.e., without tillage), the rate 
should be reduced to about a third. Where some tillage is practiced, but not to the typical 
plow-layer depth used in the calibration of the test, then the liming rate should be reduced 
proportionately (van Lierop 1989). 

Adams, F. and Evans, C.E. 1962. A rapid method and lime requirement in some acidic Queensland 

for measuring lime requirement of Red- Yellow soils. Aust. ./. Soil Res. 30: 119-130. 
Podzolic soils. Soil Sci. Soc. Am. Proc. 26: 

355-357. Alatas, J., Tasdilas, CD., and Sgouras, J. 2005. 
Comparison of two methods of lime requirement 

Aitken, R.L. 1990. Relationship between extract- determination. Comm , Sci Plant Anal. 36: 

able Al, selected soil properties, pH buffer capacity 183-190. 


Brunelle, A. and Vanasse, A. 2004. Le chaulage Sims, J.T. 1996. Lime requirement. In D.L. 
des sols. Centre dereference en agriculture etagroa- Sparks et al., Eds. Methods of Soils Analysis. 
limentairedu Quebec, Quebec, QC, Canada, 41 pp. Part 3. SSSA, Madison, WI, pp. 491-515. 

Conyers, M.K., Helyar, K.L., and Poile, G.J. 2000. 
pH buffering: the chemical response of acidic 
soils to added alkali. Soil Sci. 165: 560-566. 

CRAAQ. 2003. Guide de reference en fertilisa- 
tion, Ire edition. Centre de reference en agricul- 
ture et agroalimentaire du Quebec (CRAAQ). 
Quebec, QC, Canada, 294 pp. 

Curtin, D., Rostad, H.P.W., and Huang, P.M. 
1984. Soil acidity in relation to soil properties 
and lime requirement. Can. J. Soil Sci. 64: 

Liu, M., Kissel, D.E., Cabrera, M.L., and 
Vendrell, P.F. 2005. Soil lime requirement by 
direct titration with a single addition of calcium 
hydroxide. Soil Sci. Soc. Am. J. 68: 522-530. 

Machacha, S. 2004. Comparison of laboratory pH 
buffer methods for predicting lime requirement 
(LR) of acidic soils of Eastern Botswana. 
Commun. Soil Sci. Plant Anal. 35: 2675-2687. 

Marschner, H. 1995. Nutrient availability in soils. 

In Mineral Nutrition of Higher Plants, 2nd cd. 
Academic Press, London, UK, pp. 483-505. 

McLean, E.O. 1982. Soil pH and lime require- 
ment. In A.L. Page, R.H. Miller, and D.R. 
Keeney, Eds. Methods of Soil Analysis, Part 2, 
Agronomy 9. SSSA, Madison, WI, pp. 199-224. 

OMAFRA. 2003. Agronomy Guide for Field 
Crops. Publication 811. Ministry of Agriculture, 
Food and Rural Affairs, Toronto, ON, Canada, 
348 pp. 

Shoemaker, H.E., McLean, E.O., and Pratt, P.F. 
1961. Buffer methods for determining lime 
requirement of soils with appreciable amounts of 
extractable aluminium. Soil Sci. Soc. Am. Proc. 

25: 274-277. 

Soon, Y.K. and Bates, T.E. 1986. Determination 
of the lime requirement for acid soils in Ontario 
using the SMP buffer methods. Can. J. Soil Sci. 
66: 373-376. 

Tran, T.S. and van Lierop, W. 1982. Lime 
requirement determination for attaining pH 5.5 
and 6.0 of coarse-textured soils using buffer-pH 
methods. Soil Sci. Soc. Am. J. 46: 1008-1014. 

Tran, T.S. and van Lierop, W. 1993. Lime 
requirement. In M.R. Carter, Ed. Soil Sampling 
and Methods of Analysis. Canadian Society of 
Soil Science, Lewis Publishers, CRC Press, 
Boca Raton, FL, pp. 109-113. 

van Lierop, W. 1983. Lime requirement deter- 
mination of acid organic soils using buffer-pH 
methods. Can. J. Soil Sci. 63: 411^123. 

van Lierop, W. 1989. Effect of assumptions on 
accuracy of analytical results and liming recom- 
mendations when testing a volume or weight of 
soil. Commun. Soil Sci. Plant Anal. 30: 121-137. 

van Lierop, W. 1990. Soil pH and lime requirement. 
In R.L. Westerman, Ed. Soil Testing and Plant An- 
alysis, 2nd edn. SSSA, Madison, WI, pp. 73-126. 

Viscarra Rossel, R.A. and McBratney, A.B. 2003. 
Modelling the kinetics of buffer reactions for 
rapid field predictions of lime requirements. 
Geoderma 114: 49-63. 

Warman, P.R., Walsh, I.Y., and Rodd, A.V. 2000. 
Field testing a lime requirement test for Atlantic 
Canada, and effect of soil pH on nutrient uptake. 
Commun. Soil Sci. Plant Anal. 31: 2163-2169. 

Webber, M.D., Hoyt, P.B., Nyborg, M., and 
Corneau, D. 1977. A comparison of lime require- 
ment methods for acid Canadian soils. Can. J. Soil 
Sci. 57: 361-370. 


Chapter 1 3 

Ion Supply Rates Using 

Ion-Exchange Resins 

P. Qian and J.J. Schoenau 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

N. Ziadi 

Ay it ullure ,»];/ Agri-rood C.iihuL: 
Quebec, Quebec, Canada 


Use of ion-exchange resins to measure nutrient availability in soils was reported as early as 
1951 (Pratt 1951) and 1955 (Amer et al. 1955). Since then, many journal articles have been 
published on the use of ion-exchange resins in agricultural and environmental soil 
research, mostly focusing on measuring nutrient availability in soil. Anion-exchange 
resin extraction as a method to assess P availability in soil has been described earlier by 
Olsen and Sommers (1982) and Kuo (1996). The principle of resin membrane extraction is 
also briefly described and commented on by Havlin et al. (2005). A review of application 
of ion-exchange resins in agricultural and environmental research has been provided by 
Qian and Schoenau (2002a). 

Synthetic ion-exchange resins are solid organic polymers with an electrostatic charge that is 
neutralized by a selected counterion of opposite charge, and hence they function in a manner 
analogous to charged soil colloids. The strongly acidic cation-exchange (sulfonic acid 
functional group) resins and strongly basic anion-exchange (tertiary ammonium functional 
group) resins are chosen for use as a sink to extract nutrient ions in soils and other media. 
When ion-exchange resins are equilibrated with a solution containing a mixture of ions, 
proportions adsorbed by resin will not be the same as ionic proportions in the bulk solution, 
because of preferential selectivity by the resins for various ions. Generally speaking, cations 
and anions with the lowest affinity to the resin are best for use as counterions. There are two 
forms of ion-exchange resins that are commercially available. One is bead form and the other 
is membrane form. The resin beads are normally retained in a sealed nylon bag, while the 


resins in membrane form should be cut into the desired size of strips (Qian et al. 1992) 
before use. Both resin beads and membranes have evolved from initial usage in batch 
systems where beads or membranes are mixed with a certain amount of soil and water, and 
then shaken as a suspension for a fixed time period (Amer et al. 1955; Martin and Sparks 
1983; Turrion et al. 1999) to diffusion-sensitive systems where ion-exchange resins are 
placed in direct contact with soil for extended periods (Skogley 1992; Ziadi et al. 1999; 
Qian and Schoenau 2002b). When ion-exchange resins are used in the diffusion-sensitive 
systems, it is not easy to place them in the soil in situ, especially for resin membrane 
strips. To overcome the difficulty, resin capsule (made by sealing the resin bead inside a 
porous shell to form a compact rigid sphere capsule) and PRS probe (made by encapsu- 
lating the membrane in a plastic frame to create a probe) are commercially available 
from UNIBEST (Bozeman, Montana) and Western Ag Innovations (Saskatoon, SK), 

In batch systems, the resins are in aqueous suspension with soil. During extraction, the resins 
adsorb nutrient ions from soil solution via surface adsorption, and the resins maintain ion 
concentrations at a low level to facilitate continued nutrient ion desorption from the soil until 
equilibrium is reached (Sparks 1987). In diffusion-sensitive procedures, resins are placed in 
direct contact with soil, which provides a measure that includes both the rates of release of 
ions from different soil surfaces as well as their diffusion rates through bulk soil. The system 
integrates both chemical and biological transformations as well as diffusion to a sink into the 
measure of nutrient availability, which accounts for the kinetics of nutrient release and 
transport (Curtin et al. 1987; Abrams and Jarrel 1992). With its nature of action similar to a 
plant root in its extraction of nutrient ions in soils, this method is able to account for factors 
affecting nutrient uptake by plant roots (Qian and Schoenau 1996). The theoretical verifica- 
tion for the procedures has been documented previously (Yang et al. 1991a,b; Yang and 
Skogley 1992). 

With diffusion-sensitive systems, we can easily measure the nutrient supply rate (NSR). 
The NSR is defined as the amount of nutrient ion adsorbed per unit surface area of resin 
membrane over the time of duration of direct contact with soil. It can be expressed as |jig 
(or |xmol) per cm 2 for the time of direct contact (i.e., 24 h). There is no direct calibration 
between supply rate data and soil nutrient concentrations determined by conventional 
s as they are different measurements. Using ion-exchange resin membrane in 
t with soil to assess nutrient availability is an alternative approach to traditional 
chemical extractions in that it provides a measure of nutrient ion flux, and is useful 
in mimicking and tracking the dynamic behavior of ion supply to plant roots in 
soil (Qian and Schoenau 2002a). It can be considered a unique multiple element assess- 
ment that is universal in its application to soils from different regions and of different 

Current efforts in assessing nutrient ion supply rate in soil have focused on direct contact of 
resin with soil either in the laboratory or in the field (Qian and Schoenau 2002a). The 
embodiment of resin membranes into probes facilitates the use of ion-exchange resins 
in situ in the field or fresh bulk soil samples in the laboratory. A so-called "sandwich test" 
for laboratory testing can be used to measure NSR in soil, which only requires a few 
grams of soils, and is suitable for soil samples that have been ground and dried in pre- 
paration for other types of analysis. The "sandwich" test for laboratory testing is described 
in this chapter. 



13.2.1 Principle 

The "sandwich" test was developed to use a minimum amount of processed (air-dried and 
ground) soil to achieve a measurement of NSR. The basic principle is to allow the resin 
membrane to adsorb nutrient ions from soils by directly contacting it with the soil in a moist 
condition for 24 h. 

13.2.2 Materials and Reagents 

7 Resin membrane: supplied from Western Ag Innovation Inc. (Saskatoon, SK). 
Other sources are BDH (Poole, England) and Ionics (Watertown, Massachusetts). 
The membrane sheets are cut into squares of about 8 cm 2 each to ensure the 
square is of a size that just fits inside the vial cap. 

2 Snapcap vial lids (7 dram). 

3 Snapcap vials with lids (7 dram). 

4 Parafilm laboratory film. 

5 Analytical balance. 
5 Shaker. 

7 Pipette (1 or 2 ml_) and tips (or dropper). 

g 0.5 M NaHC0 3 solution: dissolve approximately 42 g of NaHC0 3 in deionized 
water and make to volume in a 1 L volumetric flask. 

g 0.5 M HCI: mix 42 ml_ of concentrated HCI with deionized water and make to 
volume in a 1 L volumetric flask. 

13.2.3 Procedure 


The resin membranes must be cleaned and regenerated before each use. It is very important 
that used membrane strips are not contaminated with ions of interest before making 

Before use, cation-exchange resin membranes must be cleaned/regenerated by 
soaking in 0.5 M HCI twice, for 1 h each time, with 3 ml_ of HCI per 1 cm 2 of 
membrane strip. This will put the cation-exchange membrane exchange sites into 
the proton (H + ) form as the counterion for exchange. The mixture should be 
stirred or agitated every 15 min or if possible, shaken continuously at slow 


speed on a rotary-bench or side-to-side shaker. When the counterions are not 
protons, the cleaning process should be repeated as many as four times. 

Clean brand new or regenerate used anion-exchange membranes by soaking in 
0.5 M NaHC0 3 solution four times, for 2 h each time, with 3 ml_ of NaHC0 3 
solution per 1 cm 2 of membrane strip. The solution should also be stirred on a 
regular basis or slowly shaken. This will put the anion-exchange membrane 
exchange sites into the bicarbonate (HC0 3 ~) form. 

Rinse cleaned or regenerated membrane strips with deionized water, and keep 
them in deionized water before use. 

/ Place subsamples of air-dried soil <2 mm into two Snapcap vial lids, filling the 
lids with soil up to the edges to ensure good contact between the complete surface 
of the membrane and the soil. 

2 Place the Snapcap vial lids with the soil sample on an analytical balance. Add 
deionized water until the soil in each lid is close to saturated or at field capacity. If 
adding water just to field capacity, the field capacity of the soil should be estimated 
in advance to determine how much water is required for the weight of soil used. 

3 Place a cation- or anion-exchange membrane strip onto the surface of the soil in one 
Snapcap vial lid, and then cover with the other Snapcap vial lid, making a "sand- 
wich," with the membrane sandwiched between the two lids containing soil. 
Normally a cation-exchange membrane sandwich and an anion-exchange mem- 
brane sandwich would be made if measurement of all cations and anions is desired. 

4 Seal the "sandwich" with Parafilm laboratory film to avoid loss of soil moisture 
during the extraction. 

5 Extraction time is normally set at a period of 24 h, similar to the burial time of 
membranes used in a commercial laboratory. 


7 Add 20 ml_ of 0.5 M HCI to the Snapcap vial (7 dram). 

2 Remove the Parafilm from the "sandwich," separate the lids and pick out the 
membrane strip with plastic tweezers and then wash with deionized water until 
all soil particles are removed from the membrane surface. It is important that all 
soil particles are removed to avoid any soil entering into the eluent (HCI). 

j Place the washed membrane strips into the vials with 0.5 M HCI. Cap the vials with 
lids and then shake the vials containing the membranes in a shaker at 200 rpm for 1 
h. The cation-exchange membrane and anion-exchange membrane strip from the 
same sample of soil can both be placed into the same 20 ml_ of HCI eluent as 
protons will elute the cations and CI - will elute the anions. The eluent should 
completely cover the membrane strips during shaking to ensure complete elution. 


Ion Measurement 

Nutrient ion concentrations in HC1 can be measured with various instruments commonly 
used in a soil analytical chemistry laboratory, including manual or automated colorimetry, 
ion chromatography, atomic absorption-flame emission (AA-FE) spectrometry, or induct- 
ively coupled plasma (ICP) spectrometry. 

13.2.4 Calculation 

NSR can be calculated as 

NSR=(CxV)/S (13.1) 

where C is the concentration of an adsorbed cation or anion (|jig mLT 1 ) in HC1 eluent, V is 
the volume of eluent (mL), and S is the surface area of membrane strip (cm 2 ). 

Example: a "sandwich" was prepared with an 8 cm 2 anion-exchange membrane. After 
24 h the resin membrane was removed, washed, and placed in 20 mL of 0.5 M HC1. The 
concentration in 0.5 M HC1 was 10 |uug N0 3 -N mL~' as measured by colorimetry. The value 
of NSR is reported as: (10 (xg mL~' x 20 mL)/8 cm" 2 = 25 |xg cm~ 2 . 


1 Ion-exchange resin membranes are a very sensitive measure of nutrient supply. 
Thus, maintaining consistent and uniform contact between soil and membrane is 
an essential condition to achieve reproducible results. If there is incomplete 
contact between the membrane and the soil, the area of membrane surface that 
can actually adsorb ions from soil is different than that assumed in the calculation 
of supply rate. 

2 The tests should be under the same moisture and temperature conditions. Mois- 
ture and temperature have significant effects on ion diffusion and mineraliza- 
tion/solubilization in soil. 

3 The "sandwich" test requires only a small amount of air-dried soil (about 4.5 g per 
7 dram vial lid or 9 g per "sandwich"). As such, there is no need to prepare a large 
amount of sample. 

Abrams, M.M. and Jarrel, W.M. 1992. Bioavail- anion exchange resin adsorption and "~P equili 

ability index for phosphorus using ion exchange bration. Plant Soil 6: 391-408. 
resin impregnated membranes. Soil Sci. Soc. Am. 

J. 56: 1532-1537. Curtin, D., Syers, J.K., and Smillie, G.W. 1987. 
The importance of exchangeable cations and 

Amer, F., Bouldin, D.R., Black, C.A., and Duke, resin-sink characteristics in the release of soil 

F.R. 1955. Characterization of soil phosphorus by phosphorus. J. Soil Sci. 38: 711-716. 


Havlin, J.L., Beaton, J.D., Tisdale, S.L., and 
Nelson, W.L. 2005. Soil fertility evaluation. In 
Soil Fertility and Fertilizers: An Introduction to 
Nutrient Management, 7th ed. Prentice-Hall, 
New Jersey, NJ, pp. 298-361. 

Kuo, S. 1996. Phosphorus. In Sparks, D.L., Ed. 
Methods of Soil Analysis, 3rd edn. Part 3. SSSA, 
Madison, WI, pp. 869-920. 

Martin, H.W. and Sparks, D.L. 1983. Kinetics of 
non-exchangeable potassium release from two 
Coastal Plain soils. Soil Sci. Soc. Am. J. 47: 

Olsen, S.R. and Sommers, L.E. 1982. Phosphorus. 
In Page, A.L. et al., Eds. Methods of Soil Analysis, 
2nd edn. Part 2. Agronomy Monograph No. 9. 
ASA and SSSA, Madison, WI, pp. 403-430. 

Pratt, P.F. 1951. Potassium removal from Iowa 
soils by greenhouse and laboratory procedures. 
Soil Sci. 72: 107-118. 

Qian, P. and Schoenau, J.J. 1996. Ion exchange 
resin membrane (IERM): a new approach for 
in situ measurement of nutrient availability in 
soil. Plant Nutr. Pert. Sci. 2: 322-330. 

Qian, P., Schoenau, J.J., and Huang, W. 1992. Use 
of ion exchange membranes in routine soil testing. 
Commun. Soil Sci. Plant Anal. 23: 1791-1804. 

Skogley, E.O. 1992. The universal bioavailability 
environment/soil test UNIBEST. Commun. Soil 
Sci. Plant Anal. 23: 2225-2246. 

Sparks, D.L. 1987. Potassium dynamics in soils. 
Adv. Soil Sci. 6: 1-63. 

Turrion, M.B., Gallardo, J.F., and Gonzales, M.I. 
1999. Extraction of soil-available phosphate, 
nitrate, and sulphate ions using ion exchange 
membranes and determination by ion exchange 
chromatography. Commun. Soil Sci. Plant Anal. 
30: 1137-1152. 

Yang, J.E. and Skogley, E.O. 1992. Diffusion 
kinetics of nutrient accumulation by mix-bed 
ion-exchange resin. Soil Sci. Soc. Am. J. 56: 

Yang, J.E., Skogley, E.O., Georgitis, S.J., 
Schaff, B.E., and Ferguson, A.H. 1991a. The 
phytoavailability soil test: development and 
verification of theory. Soil Sci. Soc. Am. J. 55: 

Qian, P. and Schoenau, J.J. 2002a. Practical appli- 
cations of ion exchange resins in agricultural and 
environmental soil research. Can. J. Soil Sci. 
82: 9-21. 

Yang, J.E., Skogley, E.O., and Schaff, B.E. 
1991b. Nutrient flux to mix-bed ion-exchange 
resin: temperature effects. Soil Sci. Soc. Am. J. 
55: 762-767. 

Qian, P. and Schoenau, J.J. 2002b. Availability Ziadi, N., Simard, R.R., Allard, G., and Lafond, J. 

of nitrogen in solid manure amendments 1999. Field evaluation of anion exchange mem- 

with different C/N ratios. Can. J. Soil Sci. 82: brane as a N soil testing method for grasslands. 

219-225. Can. J. Soil Sci. 79: 281-294. 

Chapter 14 
Environmental Soil 
Phosphorus Indices 

Andrew N. Sharpley 

University of Arkansas 
Fayetteville, Arkansas, United States 

Peter J.A. Kleinman 

U.S. Department of Agriculture 

University Park, Pennsylvania, United Stales 

Jennifer L. Weld 

The Pennsylvania State University 
University Park, Pennsylvania, United States 


The loss of phosphorus (P) in agricultural runoff and its input to freshwater bodies is known 
to accelerate eutrophication (Carpenter et al. 1998; U.S. Geological Survey 1999; Sharpley 
2000). As eutrophication of surface water impairs its use for recreation, drinking, and 
commercial fishing, several strategies have been put in place to minimize impairment by 
reducing the potential for P loss from agricultural operations (Gibson et al. 2000; U.S. 
Environmental Protection Agency 2004). 

Key components of remedial strategies to decrease P loss from agriculture are the determin- 
ation of soil P levels that are above those required for optimum crop growth, due to the 
continual application of P (Sims et al. 1998; Simard et al. 2000; Daverede et al. 2003) and 
the identification of critical source areas where there is a high risk of P loss due to the 
coincidence of runoff and erosion with high soil P levels (Sharpley et al. 2001, 2003; 
Coale et al. 2002). Traditional soil P tests to estimate for crop P availability have been 
used as surrogate estimates of runoff P enrichment by soil P (Sharpley et al. 1996). Because 
soil P tests were developed to work on certain soil types (e.g., Mehlich-3 and Bray-1 for 
acidic soils and Olsen for calcareous, alkaline soils) and do not mimic soil P release to 
runoff water, efforts have been made to establish environmental soil P tests (Sibbesen and 
Sharpley 1997; Torbert et al. 2002). The more prominent of these environmental tests 
include water-extractable soil P and P sorption saturation. 


Considerable field-based research has provided data to support the use of water-extractable 
soil P as an environmental test, which is independent of soil type, to assess the potential for 
soil to enrich runoff with dissolved P (Pote et al. 1996; McDowell and Sharpley 2001). The 
extraction of soil with water more closely mimics the interaction between surface soil and 
rainfall and the subsequent release of P to runoff water than do acidic or basic soil test P 
extractants. Andraski and Bundy (2003), Andraski et al. (2003), Daverede et al. (2003), 
Hooda et al. (2000), Pote et al. (1999a,b), and Torbert et al. (2002) all reported water- 
extractable soil P to be closely related to runoff-dissolved P for both grassed and cropped 
plots, at a similar or greater level of significance than Bray-1 and Mehlich-3-extractable soil P 
(Vadas et al. 2005). Increasingly, investigators are using water-extractable P in lieu of runoff 
data in laboratory studies aimed at comparing environmental and agronomic effects (e.g., Stout 
et al. 1998). 

Estimation of P sorption saturation is based on the premise that the saturation of P sorbing 
sites for a soil determines P release (intensity factor) as well as the level of soil P (capacity 
factor) (Breeuwsma and Silva 1992; Kleinman and Sharpley 2002). For example, soils of 
similar soil test P may have differing capacities to release P to runoff, based on the fact that P 
would be bound more tightly to clay than sandy soils (Sharpley and Tunney 2000). Phos- 
phorus sorption saturation can also represent the capacity of a soil to sequester further P 
addition and thereby enrich runoff P (Schoumans et al. 1987; Lookman et al. 1996). For 
example, the addition of P to a soil with a high P sorption saturation will enrich runoff P more 
than if P was added to a soil with a low P sorption saturation, independent of soil test P level 
(Sharpley 1995; Leinweber et al. 1997). Traditional techniques to estimate soil P sorption 
saturation have relied upon methods that are not commonly performed by soil testing 
laboratories, such as acid ammonium oxalate extraction in the dark (e.g., Schoumans and 
Breeuwsma 1997) and P sorption isotherms (e.g., Sharpley 1995). Recent research has shown 
soil P sorption saturation in acidic soils can be reliably estimated from Mehlich-3-extractable 
Al and Fe (primary components of P sorption) and P (Beauchemin and Simard 1999; 
Kleinman and Sharpley 2002; Nair and Graetz 2002). 

Soil P sorption has also been used to estimate the potential of a soil to sequester proposed 
additions of P. In specific cases, a detailed assessment of the P sorption capacity of a soil is a 
planning requirement of proposed land applications of biosolids, in order to determine the 
potential for P leaching through a soil profile (U.S. Environmental Protection Agency 1993; 
Bastian 1995). Traditionally, P sorption isotherms are constructed using batch equilibrations 
of soil with P added in a supporting solution, usually as KH2PO4 in a 0.01 M CaCb matrix 
for 24 to 40 h (Syers et al. 1973; Nair et al. 1984). Equations such as the Langmuir, 
Freundlich, and Tempkin models have been used to describe the relationship between the 
amount of P sorbed to the P in solution at equilibrium and to calculate P sorption maximum, 
binding energy, and equilibrium P concentrations for a given soil (Berkheiser et al. 1980; 
Nair et al. 1984). This chapter will discuss the Langmuir approach only. 

While P sorption isotherms can provide a large amount of soil-specific information that is 
useful to agronomic and environmental characterization of P sorption capacity, they are too 
time-consuming, complicated, and expensive for routine use by soil testing laboratories 
(Sharpley et al. 1994). To overcome these limitations, Bache and Williams (1971) suggested 
a single equilibration using a high concentration of P (single-point isotherm), from which a P 
sorption index (PSI) was calculated, to rapidly determine soil P sorption capacity. They 
found that PSI was closely correlated with P sorption maxima determined by the full sorption 
isotherm for 42 acid and calcareous soils from Scotland (r = 0.97; P> 0.001) (Bache 
and Williams 1971). Other researchers have subsequently found PSI to be correlated with 


soil P sorption maxima of soils varying widely in chemical and physical properties 
(Sharpley et al. 1984; Mozaffari and Sims 1994; Simard et al. 1994). 

Finally, most states in the United States have now adopted a P indexing approach as part 
of P-based nutrient management planning requirements, so that areas at greatest risk of 
P loss can be targeted for remediation or more restrictive management (Sharpley et al. 
2003). The P indexing approach is based on the knowledge that most P loss from agricultural 
watersheds (>75% annually), occurs from small, defined areas of a watershed (<20% 
land area) (Smith et al. 1991; Schoumans and Breeuwsma 1997; Pionke et al. 2000). The P 
index ranks these critical source areas by identifying where high P source potential (i.e., soil P 
and the rate, method, timing, and type of P added as fertilizer or manure) coincides with 
high transport potential (i.e., surface runoff, leaching, erosion, and proximity to a stream) 
(Lemunyon and Gilbert 1993). The P index is one of the more successful approaches that 
addresses P source, management, and transport in a holistic way by attempting to combine 
important P loss variables into a practical program that assesses specific field's potential for P 
loss (Gburek et al. 2000; Sharpley et al. 2003). Use of the P index helps farmers, consultants, 
extension agents, and livestock producers identify (i) agricultural areas or practices that have 
the greatest potential to accelerate eutrophication and (ii) management options available to 
land users that will allow them flexibility in developing remedial strategies. 

This chapter details the methods used to estimate water-extractable soil P, P sorption 
saturation, P sorption capacity, and indexing P loss potential for a given site. For all these 
chemical methods and preparation of reagents used, the use of standard laboratory protective 
clothing and eye covering is recommended. 


The extraction of soil P with water provides a rapid and simple means of determining the 
amount of soil P that can be released from soil to runoff water. The method assumes that 
extraction with water replicates the reaction between soil and runoff water and is thus, 
independent of soil type. The following method is a variation of the method described by 
Olsen and Sommers (1982) for determination of water-soluble P in soils. In summary, P 
extracted from a soil sample after it has been shaken with water for a specific period of 
time is measured spectrophotometrically by the colorimetric molybdate-ascorbic 
acid method (Murphy and Riley 1962). Alternatively, filtrates can be analyzed by induct- 
ively coupled plasma-atomic emission spectrometry (ICP-AES), which will measure total 
dissolved P. 

14.2.1 Materials and Reagents 

1 Centrifuge tubes (40 ml_) with screw caps. 

2 End-over-end shaker. 

3 Centrifuge. 

4 Filtration apparatus (0.45 jjim pore diameter membrane filter or Whatman No. 42). 

5 Photometer: Spectrophotometer with infrared phototube for use at 880 nm and 
providing a light path of at least 2.5 cm, preferably a 5 cm path length cell. 

For light path lengths of 0.5, 1 .0, and 5.0 cm, the P ranges are 0.3-2.0, 0.15-1 .30, 
and 0.01-0.25 mg L~ 1 , respectively. 

6 Acid-washed glassware and plastic bottles: Graduated cylinders (5 to 100 mL), 
volumetric flasks (1 00, 500, and 1 000 mL), storage bottles, pipets, dropper bottles, 
and test tubes or flasks for reading sample absorbance. The spectrophotometer 
should be calibrated daily by using factory standard procedures for the laboratory 

j Balances used to weigh reagents and samples are calibrated according to 
factory specifications and routinely cleaned to ensure proper and accurate 
working order. 

g Distilled water. 

9 A series of P standards (0, 0.25, 0.5, 0.75, and 1.00 mg P L~ 1 as KH 2 P0 4 ) is 
prepared fresh on the day of analysis. 

1Q Reagents for ascorbic acid technique for P determination. 

a. 2.5 M H2SO4: Slowly add 70 mL of concentrated H2SO4 to approximately 
400 mL of distilled water in a 500 mL volumetric flask. After the solution has 
cooled, dilute to 500 mL with distilled water, mix, and transfer to a plastic 
bottle for storage. Store in refrigerator until used. 

b. Ammonium molybdate solution: Dissolve 20 g of (NH 4 ) 6 M0 7 024 • 4H 2 in 
500 mL of distilled water. Store in a plastic bottle at 4°C until used. 

c. Ascorbic acid, 0.1 M: Dissolve 1 .76 g of ascorbic acid in 1 00 mL of distilled 
water. The solution is stable for about a week if stored in an opaque plastic 
bottle at 4°C until used. 

d. Potassium antimonyl tartrate solution: Using a 500 mL volumetric flask, 
dissolve 1.3715 g of K(SbO)C 4 H 4 6 • 1/2H 2 in approximately 400 mL of 
distilled water, and dilute to volume. Store in a dark, glass-stoppered bottle at 
4°C until used. 

e. Combined reagent: When making the combined reagent, all reagents must 
be allowed to reach room temperature before they are mixed, and they 
must be mixed in the following order. To make 100 mL of the combined 

Transfer 50 mL of 2.5 M H 2 S0 4 to a plastic bottle. 

Add 15 mL of ammonium molybdate solution to the bottle and mix. 
. Add 30 mL of ascorbic acid solution to the bottle and mix. 
. Add 5 mL of potassium antimonyl tartrate solution to the bottle and mix. 

If turbidity has formed in the combined reagent, shake and let stand for a few 
minutes until turbidity disappears before proceeding. Store in an opaque 
plastic bottle. The combined reagent is stable for less than 8 h, so it must be 
freshly prepared for each run. 


g. Stock phosphate solution: Using a 1 000 ml_ volumetric flask, dissolve 21 9.5 mg 
anhydrous KH2PO4 in distilled water and dilute to 1000 ml_ volume; 1 ml_ 
contains 50 jxg of P. 

h. Standard P solutions: Prepare a series of at least six standard P solutions within 
the desired P range by diluting stock phosphate solution with distilled water. 

14.2.2 Procedure 

7 Weigh out 2 g of air-dried soil into a 40 ml_ centrifuge tube. Conduct in duplicate. 

2 Add 20 ml_ of distilled water and shake at 10 rpm end-over-end for 1 h. 

3 Centrifuge at about 3000 g for 1 min. 

4 Filter the solution through a Whatman No. 42 filter paper or 0.45 |jim membrane 
filter if paper filtrates are not clear. 

5 Measure P by ICP-AES or by the ascorbic acid technique (see Section 1 4.2.1 ). 

g Pipette 20 ml_ of water extraction filtrate into a 25 ml_ volumetric flask and add 
5 ml_ of combined Murphy and Riley color reagent. 

7 If the P concentration of the extract is greater than the highest standard, a smaller 
sample aliquot is required. Add revised sample aliquot to volumetric flask, make 
up to 20 mL with distilled water, and add Murphy and Riley reagent. 

q Measure absorbance (880 nm) and determine concentration from standard curve 
prepared each day. 

14.2.3 Calculations 

7 Water-extractable soil P (mg P kg soil -1 ) 

= [Concentration of P in extract, mg L~ 1 ] x [volume of extractant, 
L/mass of soil, kg] 

2 Minimum detection limit is 0.02 mg kg -1 . 

3 There is no upper limit of detection, as extracts from soils with large amounts of 
P can be diluted. 


Air-dried soils can be stored at room temperature in whirl-pack or closed plastic containers, 
to avoid contamination. Water extracts of soils should be kept at 4°C until P is measured, 
preferably within 2 days of extraction. A large amount of soil common to the users' area and 
similar to that being analyzed should be air-dried and archived. The water-extractable soil P 
concentration of the archived sample is run each day to ensure day-to-day analytical 
reproducibility. Any deviations form this standard value should be addressed immediately. 



Phosphorus sorption saturation provides insight into a soil's ability to release P to solution as 
well as its remaining capacity to sorb added P and is defined as follows: 

P sat = S ° rbedP . (14.2) 

P sorption capacity 

In the method described below, sorbed P is represented by Mehlich-3-extractable soil P and P 
sorption capacity by Mehlich-3-extractable Al and Fe. Notably, in estimating P sorption 
saturation from Mehlich data, this study does not include a, the proportion of Mehlich-3 Al 
and Fe that contribute to P sorption capacity. Use of a in the literature has been primarily 
associated with P sorption saturation calculated from acid ammonium oxalate data (e.g., van der 
Zee and van Riemsdijk 1988). Given soil-specific variations in sorption mechanisms affecting 
P sorption capacity as well as variability in methods used to estimate P sorption, there is little 
justification for the continued use of this value unless it is measured (Hooda et al. 2000). 

14.3.1 Materials and Reagents 

7 Centrifuge tubes (40 mL) with screw caps. 
2 End -over-end shaker. 
j Centrifuge. 

4 Filtration apparatus (0.45 |jim pore diameter membrane filter or Whatman No. 42). 

5 Mehlich-3 solution as 0.2 M CH 3 COOH, 0.25 M NH4NO3, 0.015 /W NH 4 F, 
0.013 M HN0 3/ and 0.001 M EDTA (see Chapter 7 for more detail). Store in 
refrigerator until used. 

6 Acid-washed glassware and plastic bottles. 

14.3.2 Procedure 

1 Weigh out 2.5 g of air-dried soil into a 40 mL centrifuge tube. Conduct in duplicate. 

2 Add 25 mL of Mehlich-3 reagent and shake at 10 rpm for 5 min. 

3 Filter the solution through a Whatman No. 42 filter paper or 0.45 |jim membrane 
filter if paper filtrates are not clear. 

4 Measure P, Al, and Fe by ICP-AES. Represented as P M 3, AI M 3, and Fe M 3, respectively. 

14.3.3 Calculations 

/ In all cases, molar concentrations of extracted elements (mmol kg~ 1 ) were used to 
determine P sat . 

2 For acid soils (pH < 7.0): 



Soil P sorption saturation is increasingly used as an environmental indicator of soil P 
availability to runoff and can be easily calculated from data that is readily available through 
soil testing laboratories and national databases. Several studies show that Mehlich-3 data 
can be effectively used to estimate P sat for a wide range of acidic and alkaline soils. As most 
soil testing laboratories currently conducting Mehlich-3 extraction employ ICPs, analytes 
required to estimate P sat (Pm3, A1 M 3, and Fe M 3) are measured simultaneously. However, 
P estimated by ICP is often greater than by colorimetric methods due to ICP measuring 
near total (inorganic + organic) dissolved P. Care must be taken in building databases or 
comparing studies, which have used different methods of determining P in filtrates. 


Estimates of P sorption vary with soil/solution ratio, ionic strength and cation species of the 
supporting electrolyte, time of equilibration, range of initial P concentrations, volume of soil 
suspension to headspace volume in the equilibration tube, rate and type of shaking, and type 
and extent of solid/solution separation after equilibration (Nair et al. 1984). Even though a 
similar basic procedure is used to measure P sorption, there is considerable variation in the 
above parameters, which makes comparison of results among studies difficult. Thus, Nair et al. 
(1984) proposed a standard P adsorption procedure that would produce consistent results over a 
wide range of soils. This procedure was evaluated, revised, tested among laboratories, and was 
eventually proposed as a standardized P adsorption procedure and is detailed below. 

14.4.1 Materials and Reagents 

7 Centrifuge tubes (40 ml_) with screw caps. 

2 End-over-end shaker. 

3 Centrifuge. 

4 Filtration apparatus (0.45 jjim porediameter membrane filter or Whatman No.42). 

5 Photometer: Spectrophotometer with infrared phototube for use at 880 nm and 
providing a light path of at least 2.5 cm, preferably a 5 cm path length cell. For 
light path lengths of 0.5, 1 .0, and 5.0 cm, the P ranges are 0.3-2.0, 0.1 5-1 .30, and 
0.01-0.25 mg L -1 , respectively. 

g Acid-washed glassware and plastic bottles: Graduated cylinders (5 to 100 ml_), 
volumetric flasks (100, 500, and 1000 mL), storage bottles, pipets, dropper bottles, 
and test tubes or flasks for reading sample absorbance. The spectrophotometer should 
be calibrated daily using factory standard procedures for the laboratory machine. 

7 Balances used to weigh reagents and samples are calibrated according to factory 
specifications and routinely cleaned to ensure proper and accurate working order. 


g Support or equilibrating solution is 0.01 M CaCb- Store in refrigerator until used. 

9 Inorganic P solution of 50 mg L" 1 as KH 2 P0 4 in 0.01 M CaCI 2 . Store in refriger- 
ator until used. 

14.4.2 Procedure 

/ Weigh out 1 g of air-dried soil into a 40 ml_ centrifuge tube. Conduct in duplicate. 

2 Add 0, 1 , 2, 5, 1 0, 1 5, and 20 mL of stock P solution (50 mg L" 1 ) and make up to a 
final volume of 25 mL with distilled water and shake at 1 rpm end-over-end for 
24 h. This gives equilibrating P concentrations of 0, 50, 1 00, 250, 500, 750, and 
1000 mg P kg soil" 1 or 0, 2, 4, 10, 20, 30, and 40 mg P L -1 , respectively. The 
range of P concentrations used can be adjusted as needed to ensure the upper 
concentration represents a distinct curvature of the plotted P sorption isotherm. 

3 Centrifuge at 3000 g for 1 min. 

4 Filter the solution through a Whatman No. 42 filter paper or 0.45 |jim membrane 
filter if paper filtrates are not clear. 

5 Measure P by ICP-AES or by the ascorbic acid technique (see Section 14.2.1). 

g Pipette 5 mL of water extraction filtrate into a 25 mL volumetric flask and add 
5 mL of combined Murphy and Riley color reagent and make up to 25 mL with 
distilled water. 

7 Adjust sample aliquot as required and make up to a final volume of 25 mL after 
addition of Murphy and Riley reagent. 

g Measure absorbance (880 nm) and determine concentration from standard curve 
prepared each day. 

14.4.3 Calculation of P Sorption Isotherm 

7 The amount of P sorbed by soil (S, mg P kg soih 1 ) is calculated as the difference 
between added P and P remaining in solution after the 24 h equilibration. Several 
methods exist for the determination of the amount of P originally sorbed by soil 
(S ) such as the least squares fit model, oxalate-extractable P, and anion- 
membrane exchangeable P (Nair et al. 1998). 

2 The Langmuir sorption isotherm is plotted as equilibrium solution P concentration 
(C, mg P L _1 ) against P sorbed (S) as shown in Figure 14.1a. 

j Using the Langmuir sorption equation below, P sorption maximum 
(Smax/ m 8 P kg soil" 1 ) ar| d binding energy of P to soil (k, L mg P~ 1 ) can be 


" 300 



^^- * 

I 200 

; s^^ 

1 100 

J Equilibrium P concentration, EPC 





| 0.04 


^^ r2 = 0.99 


^^ S max = 370 mg P kg- 1 
^y^^ /c=0.011 Lmg P- 1 

^ EPC =0.73mg PL-1 

FIGURE 14.1. Representation of Langmuir P sorption isotherm (a) and linear (b) plot from which P 
sorption maximum, binding energy, and equilibrium P concentration are calculated. 

where S = S' + S Q , the total amount of P sorbed (mg P kg soil -1 ); 5', P sorbed 
by soil (mg P kg soih 1 ); S Q , P originally sorbed (previously sorbed P) (mg P kg 
soiT 1 ); C, equilibrium solution P concentration after 24 h shaking (mg P L _1 ); 
Smax, P sorption maximum (mg P kg soih 1 ); and k, a constant relating the binding 
energy of P to soil (L mg P -1 )- 

^ P sorption maximum, S max , is calculated as the reciprocal of the slope of the plot 
C/S vs. C (Figure 14.1a). 

5 Binding energy, k, is calculated as the slope/intercept of the same plot 
(Figure 14.1b). 

6 The equilibrium P concentration (EPC , mg P L _1 ), defined as the solution P 
concentration supported by a soil sample at which no net sorption or desorption 
occurs, is calculated as the intercept of the isotherm curve on the x-axis (see 
Figure 14.1). 


The procedure to determine PSI using a single-point isotherm approach, described below, is 
based on Bache and Williams (1971). 


14.5.1 Materials and Reagents 

7 Centrifuge tubes (40 ml_) with screw caps. 

2 End -over-end shaker. 

3 Centrifuge. 

4 Filtration apparatus (0.45 |jim pore diameter membrane filter or Whatman No. 42). 

5 Photometer: Spectrophotometer with infrared phototube for use at 880 nm and 
providing a light path of at least 2.5 cm, preferably a 5 cm path length cell. For 
light path lengths of 0.5, 1.0, and 5.0 cm, the P ranges are 0.3-2.0, 0.15-1.30, 
and 0.01-0.25 mg L~ 1 , respectively. 

6 Acid-washed glassware and plastic bottles: Graduated cylinders (5 to 100 mL), 
volumetric flasks (100, 500, and 1000 mL), storage bottles, pipets, dropper bottles, 
and test tubes or flasks for reading sample absorbance. The spectrophotometer should 
be calibrated daily using factory standard procedures for the laboratory machine. 

y Balances used to weigh reagents and samples are calibrated according to factory 
specifications and routinely cleaned to ensure proper and accurate working order. 

8 Inorganic P solution of 75 mg L~ 1 as KH 2 P0 4 in 0.01 M CaCI 2 . Store in 
refrigerator until used. 

14.5.2 Procedure 

/ Weigh out 1 g of air-dried soil into a 40 mL centrifuge tube. Conduct in duplicate. 

2 Add 20 mL of the 75 mg P L _1 sorption solution to the centrifuge tube. This 
provides a single addition of 1 .5 g P kg soih 1 and a solutiomsoil ratio of 20:1 . 

j Shake at 1 rpm end-over-end for 1 8 h. 

4 Centrifuge at 3000 g for 10 min. 

5 Filter the solution through a Whatman No. 42 filter paper or 0.45 ^m membrane 
filter if paper filtrates are not clear. 

5 Measure P by ICP-AES or by the ascorbic acid technique (see Section 1 4.2.1 ). 

y Pipette 5 mL of water extraction filtrate into a 25 mL volumetric flask and add 5 mLof 
combi ned Mu rphy and Riley color reagent and make up to 25 mL with distil led water. 

g Adjust sample aliquot as required and make up to a final volume of 25 mL after 
addition of Murphy and Riley reagent. 

9 Measure absorbance (880 nm) and determine concentration from standard curve 
prepared each day. 


14.5.3 Calculation of P Sorption Index 

7 The PSI is calculated using the quotient S/log C, where S is the amount of P sorbed 
(mg P kg -1 ) and Cis solution P concentration (mg L" 1 )- 

2 Others have shown that expressing PSI directly as the amount of P sorbed 
(mg P kg" 1 ) is acceptable (Sims 2000). 


Site vulnerability to P loss in runoff is assessed with the P index by selecting rating values for 
a variety of source and transport factors. Although procedures and formats of P indices vary 
regionally, generally the first step in the process is to collect farm information such as farm 
maps, soil test reports, manure analysis, crop rotations, and manure handling and application 
information. The second step is to determine erosion rates, runoff class, and distance to 
receiving water is often needed. A site visit and evaluation is critical to properly evaluate 
field boundaries, areas of runoff and erosion contributions, and options for improved nutrient 
management and best management practices. The following procedure outlines the sources 
of information and calculations for Pennsylvania's P index. English units are most com- 
monly used in P indices, to be consistent with the units used by field practitioners. Factors 
are given to convert English to metric or SI units. 

The screening tool reduces potential time and workload associated with P index evaluations, 
by identifying fields at greatest risk to P loss using one or more readily available P index 
factors. In the Pennsylvania P index, the screening tool is Part A of the P index and uses soil 
test P (Mehlich-3 ppm P) and distance from the bottom edge of a field to a receiving body of 
water (contributing distance) (Table 14.1). 

14.6.1 Procedure 

If a soil test P level for a field is either greater than 200 ppm P or if the bottom edge of the 
field is closer than 150 ft. (50 m) to a receiving body, then the field is determined to have a 
potentially high risk of P loss. To determine the risk of P loss, additional field factors must be 
evaluated using Part B of the P index (Table 14.2). 

If the soil test P level for the field is less than 200 ppm P and the bottom edge of the 
field is more than 150 ft. (50 m) from a receiving body, then the field does not have a 
potentially high risk for P loss and N-based nutrient management recommendations can 
be followed. 

TABLE 14.1 The P Indexing Approach Using a Modified Version of Pennsylvania's Index 
Version of 8/2002, as an Example Part A — Screening Tool 
Evaluation category 

Soil test P— Mehlich-3 P >200 ppm (|jLg g" 1 ) If yes to either factor 

Contributing distance <1 50 ft. (50 m) then proceed to Part B 


TABLE 14.2 The Transport Factors Included in Part B of the Pennsylvania P Index Version of 8/2002; Part B— Transport Factors 


Risk levels 


Soil Erosion 

Risk value = Annual soil loss = to ns/ac re/yea r a 

Runoff Potential 

Very Low 




Very High 

Subsurface Drainage 


Patterned 13 

Leaching Potential 



Contributing Distance 

>500 ft. (>150 m) 

500 to 350 ft. 

(100 to 150 m) 


350 to 250 ft. 

(75 to 100 m) 


150 to 250 ft. 

(50 to 75 m) 


<1 50 ft. (<50m) 

Transport Sum = Erosion + Runoff Potential + Subsurface Drainage + Leaching Potential + Contributing Distance 

Modified Connectivity 

Riparian buffer 

Applies to distances <I50 ft. (<50 m) 


Grassed waterway 

Applies to distances >150 it. (>50 m) 

Transport Factor = Transport Sum X Modified Connectivity/22 

The transport value is divided by 22 (i.e. the highest value obtainable) in order to normalize site transport to a value of 1 , where full 
transport potential is realized. 

Caution: Many states in the United States have a state-specific P index. Although the principles of most P index tools are similar, 
individual factors or weightings of those factors vary among states. If available, review your own state's P index. For more 
specific information on the various indices adopted by states see Sharpley et al. (2003). 

a 1 ton/acre/year is equivalent to 2.24 Mg ha -1 year -1 . 
b Or a rapidly permeable soil near a stream. 


14.6.2 Warning 

Phosphorus indices vary with respect to factors evaluated, coefficients assigned to field 
conditions and management scenarios, and calculations used to determine P index values. 
Additionally, P indices are subject to change and modification to reflect current research and 
policy. In order to ensure the P index is being used and interpreted properly, current 
regionally approved versions must be obtained and regional training and certification 
requirements must be met by those specialists using the P index. The information that 
follows is based on Pennsylvania's P index. 

14.6.3 Materials 

1 Soil erosion: Soil erosion rate can be calculated using RUSLE 1 .06 c (Renard et al. 

2 Runoff potential: Using the predominate soil type in a field (50% or greater of the 
field area) and county specific tables, which can be provided by USDA-NRCS 
staff, the index surface runoff class can be determined for each evaluated field. 
The following describes the USDA-NRCS method for determining index surface 
runoff class. 

3 Subsurface drainage: Using farm information, determine if there is artificial 
drainage in the field or if the field is near a stream and has rapidly permeable 
soils. "Random" drainage is a single or a few tile lines in a field and "patterned" 
drainage is when most or the entire field is drained with a fill-patterned drainage 
system. Rapidly permeable soils must occur within 1 50 ft. (50 m) of a stream and 
be classified as such by USDA-NRCS. 

4 Contributing distance: Determine the contributing distance of each field to be 
evaluated to receiving water. The distance is measured from the lower edge of the 
field closest to the receiving water and can be determined using farm maps or by 
field measurements. 

5 Modified connectivity: Accounts for if and where buffers, grassed waterways, 
ditches, and pipe outlets are present. 

• If the field is within 150 ft. (50 m) of water and a riparian buffer is present, select 
the appropriate modified connectivity factor (i.e., reduces transport value). All 
buffers must be designed and maintained to meet USDA-NRCS standards. 

• If a field is more than 1 50 ft. (50 m) from water but a direct connection such as a 
pipe or ditch from field to receiving water is present, select appropriate modi- 
fied connectivity factor (i.e., increases transport value). 

• If a field has a grassed waterway or has no qualifying management practices, 
then a default coefficient of 1.0 is used (i.e., the transport value is neither 
increased nor is it decreased). 


14.6.4 Calculations 

7 Transport sum: Sum the actual soil loss rate (tons/acre/year) with the coefficients 
for runoff potential, subsurface drainage, and contributing distance. Enter the sum 
into the transport sum risk value column for each field. 

2 Transport factor: Multiply the transport sum by the modified connectivity coeffi- 
cient and divide the product by 22. Twenty-two is the maximum transport sum 
value and dividing by this value allows the transport factor to vary generally 
between and 1. One is the value at which the full (100%) field transport 
potential is reached. Any other value would represent a relative percentage of 
the field's full transport potential. The transport factor only exceeds 1 when 
erosion losses are exceptionally high. Enter the product into the transport factor 
risk value column for each field. 

14.6.5 Materials 

/ Soil test P: Current soil test reports. 

2 Fertilizer and manure rate: Farm records or a nutrient management plan indicating 
the amount of P, in pounds of P20s/acre, to be applied to each field. 

j Loss rating for fertilizer and manure application: Farm records or a nutrient 
management plan indicating the methods and timing used to apply P to each field. 

4 Manure P availability: Farm records or a nutrient management plan indicating the 
manure types, manure groups, or other organic P sources to be applied to each 
field to be evaluated (Table 14.3). 

14.6.6 Calculating the P Index Value 

/ Enter all of the transport factors (Part B) and sums of management factors (Part C) 
into the worksheet below. 

2 Multiply Part B by Part C and then the product by 2. The factor of 2 normalizes the 
final index rating to 100. This is your final P index rating. 

j Look up the associated interpretation and management guidance in Table 14.4. 

Part B Part C P Index Interpretation of 

Field transport risk management risk B x C X 2 the P index 

Example 0.55 92 101 Very high 


TABLE 14.3 Phosphorus Loss Potential due to Source and Site 

Management Factors 

in the P Index; Part C — Source and Site Factors 

Risk Levels 



Very Low 




Very High 

Soil test P risk 3 

Risk Value = So 

Test P(ppm asMehlich-3 P) x 0.20 
e = Soil test P (lbs P 2 5 /acre) x 0.05 


ppm x 0.20 
^Cyacre x 0.05 

Loss rating for P 

application method 

and timing 

Placed with planter 

or injected more 
than 2" (5 cm) deep 


Incorporated <1 
week after 


Incorporated >1 

incorporated >1 

following application 

in spring-summer 


Incorporated >1 

following application 


Surface applied on 
covered soil 


Fertilizer P risk 3 

Risk Value 
Risk Value 

Fertilizer P Application Rate x 
lbs P-,0 5 /acre x 

Loss Rating for 

P Application 

Manure P 

Organic Phosphorus Source Availability Coefficients 

Manure P risk 3 

Risk Value = Man 
Risk Value - 

jre P Application Rate x Loss Rating for P Application x P Availability Coefficient = 
lbs P.CWacre x x 

Total of Management Risk Factors 

Sum of management factors = 

Caution: Many states in the United States have a state-specific P index. Although the principles of most P index tools are similar, 
factors or weightings of those factors vary among states. If available, review your own state's P index. For more specific i 
on the various indices adopted by states see Sharpley et al. (2003). 

a Conversion factor: 10 lbs P 2 5 /acre is equivalent to 4.89 kg P ha -1 . 



TABLE 14.4 General Interpretations and Management Guidance for the P Index 

ral interpretatio 


If current farming practices are 
maintained, there is a low risk of 
adverse impacts on surface waters 

Chance for adverse impacts on 
surface waters exists, and some 
remediation should be taken to 

N-based appl 

N-based applications 


irface waters, 
res and P 
management plan are needed to 

Very high Adver: 

e P loss 
5 impact on s 

must be 
P loss 

and P management plar 
nplemented to minimizi 


limited to 
il of P 

Caution: Many states in the United States have a state-specific P index. Although the principles of 
most P index tools are similar, individual factors or weightings of those factors vary 
among states. If available, review your own region's P index. For more specific 
information on the various indices adopted by states see Sharpley et al. (2003). 

Andraski, T.W. and Bundy, L.G. 2003. Relation- 
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Beauchemin, S. and Simard, R.R. 1999. Soil 
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Berkheiser, V.E., Street, J.J., Rao, P.S.C., and 
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Chapter 1 5 

Electrical Conductivity 

and Soluble Ions 

Jim J. Miller 

Agrii allure and Agri-rood Canada 
Lethbridge, Alberta, Canada 

Denis Curtin 

New Zealand Institute for Crop and Food Research 
Christchurch, New Zealand 


Saline soil is defined as one containing sufficient soluble salts to adversely affect the growth 
of most crop plants (Soil Science Society of America 2001). Soil salinization is a widespread 
limitation to agricultural production in dryland and irrigated soils throughout the world. Soil 
salinity reduces crop growth because depression of the osmotic potential of the soil solution 
limits water uptake by the plant (Corwin and Lesch 2003). Salinity may also cause specific 
ion toxicity or nutrient imbalances, and soil permeability and tilth may deteriorate if 
excessive amounts of Na accumulate on the soil's cation-exchange complex. 

Soil salinity is generally measured by the electrical conductivity (EC) of a soil extract. A soil 
is considered saline if the EC of the saturation extract exceeds 4 dS m _1 at 25°C (Soil 
Science Society of America 2001). The main ions comprising soluble salts are cations Na, 
Ca, Mg; and anions SO4, and CI. Minor amounts of K, HCO3, CO3, and NO3 may also be 
present. Soil sodicity is the accumulation of exchangeable Na, determined by measuring the 
exchangeable sodium percentage (ESP); or more commonly, estimated from the sodium 
adsorption ratio (SAR) of a soil-water extract. If the SAR of the saturation extract exceeds 
13, the soil is considered sodic (Soil Science Society of America 2001). A more detailed 
classification scheme for sodic soils based on physical behavior (clay dispersibility), sodium, 
and salinity levels, has been proposed by Sumner et al. (1998). 

Soil salinity or EC may be measured on the bulk soil (EC a ), in the saturation paste extract 
(EC e ), in water extracts at soil: water ratios of 1:1 to 1:5 (ECi : i, ECi : 2, EChs), or directly on 
soil water extracted from the soil in the field (EC W ) (Corwin and Lesch 2003). The EC a or 
apparent EC has become one of the most reliable and frequently used measurements 
to characterize the spatial distribution of soil salinity at field scales. Field methods used to 


measure EC a include Wenner array or four-electrode, electromagnetic (EM) induction, and 
time domain reflectometry (TDR) (Rhoades and Oster 1986; Rhoades 1990, 1992). The EM 
induction method is the most popular of these three methods because measurements can be 
taken quickly over large areas, the large volume of soil measured reduces local-scale 
variability, and measurements are possible in relatively dry or stony soils because no contact 
is necessary between the soil and EM sensor (Hendrickx et al. 1992). The EM38 meter, and 
to a lesser extent, the EM31 meter (Geonics Ltd., Mississauga, Ontario) are most commonly 
used in soil investigations. The EM38 can measure EC a to a depth of 1.2 m in the vertical 
mode and to 0.6 m in the horizontal mode. Mobile systems have been developed in 
conjunction with global positioning systems (GPS) to allow rapid salinity mapping of 
large fields (Rhoades 1992; Cannon et al. 1994). The EC a readings from the EM38 meter 
are easily converted to EC e values for different soil temperature, texture, and moisture 
conditions (Rhoades and Corwin 1981; Corwin and Rhoades 1982; McKenzie et al. 1989). 

The EC of aqueous extracts of soil has traditionally been defined in terms of the EC of the 
saturated soil paste extract (EC e ) (U.S. Salinity Laboratory Staff 1954). Since the EC and 
concentration of solutes are affected by the soikwater ratio (Reitemeier 1946), this needs to 
be standardized to allow for consistent universal interpretation across soil texture classes. 
Exceptions include sandy soils, organic soils, and soils containing gypsum (Robbins and 
Wiegand 1990). Since it is impractical to routinely extract soil water at typical field-water 
contents, soil solution extracts must be made at higher than normal water contents. The 
saturated soil paste approximates the lowest soikwater ratio at which sufficient extract can be 
routinely removed for analysis of major salinity constituents. The saturated paste method 
relates more closely to the water holding capacity of the soil than do extracts at a fixed 
soikwater ratio. The water content of a saturation paste is about twice that at field capacity 
for most soils (Robbins and Wiegand 1990). Crop tolerance to salinity has traditionally been 
expressed in terms of EC e . 

Because the saturated paste method requires time and skill, laboratories are increasingly 
using fixed soil: water ratios (e.g., 1:1, 1:2, 1:5) when measuring soil EC and solute 
concentrations. However, cation exchange and mineral dissolution as the soikwater ratio 
widens (Reitemeier 1946) may lead to overestimation of EC and changes in solute composi- 
tion. This is especially the case in samples containing gypsum, since Ca and SO4 concen- 
trations remain near-constant over a range of soikwater ratios while the concentration of the 
other ions decreases with dilution (Robbins and Wiegand 1990). Nevertheless, studies have 
shown good correlations between EC, Mg, K, and CI in 1:2 extracts versus saturation paste 
extracts (Sonneveld and Van den Ende 1971); between EC, Na, Ca + Mg, and CI in 1:1 and 
1:2 extracts versus saturation paste extracts (Hogg and Henry 1984); and between EC, 
soluble cations (Na, Ca, Mg, K) and anions (SO4, CI) in 1:1 extracts versus saturation 
paste extracts (Pittman et al. 2004). In an analysis of soluble salt data from 87 laboratories 
in the United States, average residual standard deviation (RSD) was lowest for saturation 
paste extracts (13.4%), followed by 1:1 extracts (24.2%), and then 1:2 extracts (32.5%) 
(Wolf et al. 1996). Ninety percent of results for the 1:1 extracts were within +2 standard 
deviations of the mean value (acceptable laboratory performance) compared with 87% of the 
saturation paste extracts, and 84% of the 1:2 extracts. 

Measurement of EC (EC W ) and solutes in the soil water extracted at field-water content is 
theoretically the best measure of salinity because it indicates the actual salinity level 
experienced by the plant root (Corwin and Lesch 2003). However, EC W has not been widely 
used because it varies as soil-water content changes over time and so it is not a single-valued 
parameter (Rhoades 1978), and the methods for obtaining soil solutions are too laborious and 


costly to be practical (Rhoades et al. 1999). Soil solutions can be obtained from disturbed 
samples by displacement, compaction, centrifugation, molecular adsorption, and vacuum or 
pressure extraction methods (Rhoades and Oster 1986). Soil solutions from undisturbed 
samples can be obtained using various suction-type samplers and salinity sensors (Corwin 
and Lesch 2003). Kohut and Dudas (1994) reported considerable variation between the 
properties of saturation paste extracts and immiscibly displaced solutions, with the saturation 
paste extract having lower EC values, cation concentrations (Na, Mg, K), and anion 

This chapter will focus mainly on laboratory methods used to measure EC of saturation paste 
extracts and extracts at fixed soihwater ratios, and on methods available to analyze soluble 
cations and anions in these extracts. 

15.2.1 Saturation Extract (Janzen 1993; Rhoades 1996) 

Determine moisture content or weight of water in air-dry soil samples to be used. 
Weigh a subsample (30-50 g) of air-dry soil, oven-dry at 105°C, reweigh it, and 
determine weight of water in air-dry soil. 

Weigh from 200 to 400 g of air-dry soil with known moisture content into a 
container with lid. Record the total weight of container and the soil sample. (The 
weight of soil used will depend on the volume of extract required. In general, 
approximately one-third of the water added is recovered in the saturation extract.) 

Add deionized water while mixing to saturate the soil sample. At saturation, the soil 
paste glistens, flows slightly when the container is tipped, slides cleanly from the 
spatula, and readily consolidates afteratrench isformed upon jarringthe container. 

Allow the sample to stand for at least 4 h and check to ensure saturation criteria 
are still met. If free water has accumulated on the surface, add a weighed amount 
of soil and remix. If the soil has stiffened or does not glisten, add distilled water 
and mix thoroughly. 

Weigh the container with contents. Record the increase in weight, which corres- 
ponds to the amount of water added. (Alternatively, the amount of water added 
can be determined volumetrically by dispensing water from a burette.) Calculate 
the saturation percentage (SP) as follows: 

(weight of water added + weight of water in sample) 

SP = - 5 ^^ — ^x100 (15.1) 

oven-dry weight of soil 

Allow the paste to stand long enough to establish equilibrium between the soil 
minerals and the water (at least 4 h, but preferably overnight). If a pH measure- 
ment is needed, the samples are then thoroughly mixed and their pH measured 
with an electrode and pH meter. The pH of the saturation paste is generally 
more meaningful than the pH of the saturation paste extract (Robbins and 
Wiegand 1990). 


7 Transfer the wet soil to a Buchner funnel fitted with highly retentive filter paper. 
Apply vacuum and collect extract until air passes through the filter. Turbid filtrates 
should be refiltered. 

g Store extracts at 4°C until analyzed for EC and soluble cations and anions. 


If possible, organic soils should be extracted without prior drying, which affects the SP. 
Organic soils may require an overnight saturation period and a second addition of water to 
achieve a definite saturation endpoint. For fine-textured soils, sufficient water should be 
added immediately to the soil sample with minimal mixing to bring the sample close to 
saturation. Do not over-wet coarse-textured soils. Free water on the soil surface after 
standing indicates oversaturation of coarse-textured soils. 

15.2.2 Fixed Ratio Extracts (Janzen 1993; Rhoades 1996) 

1 Weigh appropriate amount of air-dry soil into a flask, add sufficient deionized 
water to achieve desired extraction ratio, and shake for 1 h. 

2 Filter the suspension using highly retentive filter paper and store filtrate at 4°C 
before analysis. 

The 1:1 and 1:2 soil: water extraction ratios are preferred over the 1:5 ratio. However, the 
1:5 ratio is commonly used in Australian salinity work (Rengasamy et al. 1984; Sumner 
et al. 1998). 


15.3.1 Electrical Conductivity (EC e , EC 1:1 , EC 1:2 , EC 1:5 ) 

The total solute concentration in the various extracts is normally estimated by measuring EC. 
Although the relationship between conductivity and salt concentration varies somewhat de- 
pending on solution ionic composition, EC provides a rapid and reasonably accurate estimate of 
solute concentration. The procedure below is for modern EC meters that provide automatic 
temperature compensation, automatically adjust cell constant internal to the meter, and readout 
EC directly in |xmho cm~' or similar units. For older EC meters that do not have these three 
features, refer to Rhoades (1996) or American Public Health Association (1998). 


Make up standard 0.0 10A4KCI solution to automatically adjust eel I constant internal 
to the meter. Dissolve 0.7456 g of reagent-grade anhydrous KCI and make up to 1 L 
using pure water (EC < 0.001 dS rrr 1 ). This solution has an EC of 1 .41 3 dS m~ 1 at 
25°Cand issuitableformostsolutionswhenthecell constant is between 1 and2. Use 
stronger or weaker KCI solutions to determine other cell constants. 


Calibrate conductivity meter using standard KCI solution to automatically 
adjust cell constant internal to the meter. Rinse probe three times with 0.01 
M KCI. Adjust temperature of a fourth portion to 25.0°C + 0.1 °C. Adjust tempera- 
ture compensation dial to 0.0191 C _1 . With probe in standard KCI solution, 
adjust meter to read 1 41 3 |j,mho citt 1 or 1 .41 3 dS m~ 1 . 

Read conductivity of extracts u 
ofdSirr 1 . 

ing EC probe and meter. Report results in SI units 


Use an EC meter capable of measuring EC with an error not exceeding 1% or 1 |xmho cm -1 
or 0.001 dS m _1 . The basic unit of EC is mho cm -1 , and is too large for most natural waters 
(Bohn et al. 1979). A more convenient unit is mmho cm -1 . Units in the older literature, or 
when dealing with low salinity waters, have also been reported as pimho cm -1 . The 
SI unit of conductivity is Siemens per meter (S m~'), but results are generally reported 
as dS m _1 . Water with an EC of 0.0002 mho cm -1 has an EC of 0.2 mmho cm -1 , 
200 ixmho cm" 1 , 0.020 S m" 1 , or 0.2 dS m" 1 . 

15.3.2 Soluble Ion Concentrations — Overview and Comparison 
of Methods 

Various methods are available to analyze soluble cations and anions in soil-water e 
(Table 15.1). Most laboratories have used flame-atomic absorption spectroscopy (FL-AAS) 
to analyze soluble cations, colorimetric methods on an autoanalyzer to determine CI and 
SO4, and the titrimetric method to analyze HCO3 and CO3. 

FL-AAS is the preferred instrument for analyzing soluble cations where cost is a major 
limitation, number of samples will not be large, and extremely low detection limits are not 
required (Wright and Stuczynski 1996). Ion chromatography (IC) has been mostly used to 
analyze SO4 and CI in aqueous systems (American Public Health Association 1998) and soil 
extracts (Nieto and Frankenberger 1985a). Although IC can also determine soluble cations in 
soils (Basta and Tabatabai 1985; Nieto and Frankenberger 1985b), it is seldom used for 
cation analysis. 

TABLE 15.1 Methods That Could Be Used to Measure Concentrations of Soluble Cations 
and Anions in Saturation Paste and Fixed Ratio Extracts 









FL-AAS, flame-atomic absorption spectroscopy; IC, i 
Dupled plasma-atomic emission spectroscopy. 

chromatography; ICP-AES, inductively 


Inductively coupled plasma-atomic emission spectroscopy (ICP-AES) has been increasingly 
used to analyze soluble Na, K, Ca, and Mg in soil extracts (Soltanpour et al. 1996; Wright and 
Stuczynski 1996) and waters (Vitale et al. 1991). In addition, ICP-AES can be used to determine 
nonmetals such as S and CI in aqueous extracts (Richter et al. 1999). The advantages of ICP- 
AES are the plasma flux is extremely stable compared to conventional flames with FL-AAS, 
lower detection limits are possible for certain elements, and it has simultaneous multielement 
capability where 15 to 20 metals in a water sample can be measured in a 2 min period (Vitale 
et al. 1991). Disadvantages with ICP-AES are high initial cost, high operating costs (gases, 
power, consumables) (Wright and Stuczynski 1996), and possible severe matrix interferences 
from high concentrations of total dissolved solids, Na, Ca, Fe, and Al (Vitale et al. 1991). 

Soluble Cations 

Sodium has been most commonly analyzed using flame emission photometry at 589 nm, and 
K using flame photometry at 766.5 nm (Robbins and Wiegand 1990; Helmke and Sparks 
1996). Pretreatment involves filtering out any solid particles. Calcium has been traditionally 
analyzed using AAS at 422.7 and 285.2 nm, respectively (Robbins and Wiegand 1990; 
Suarez 1996). Elements that form stable oxysalts (Al, Be, P, Si, Ti, V, Zr) can interfere with 
Ca and Mg analyses, but these can be removed by adding 0.1% to 1.0% lanthanum or 
strontium chloride to the samples. 

Soluble Anions 

Chloride in soil extracts is most commonly analyzed using potentiometric titration 
with AgN03, direct potentiometric analysis using a solid-state selective ion-electrode, 
automated colorimetric analysis (mercury thiocyanate method) on the autoanalyzer, or by 
IC (Frankenberger et al. 1996). The mercury thiocyanate method is widely used to determine 
CI, but there is a trend toward IC and ICP-AES methods to avoid working with, and 
disposing of, Hg and cyanate. Gravimetry, turbidimetry, titrimetry, and colorimetry are the 
most common methods to analyze SO4 in soil extracts; but the most sensitive and accurate 
methods for soil extract analyses are the methylene blue (MB) colorimetric and IC methods 
(Tabatabai 1996). In addition, the automated methylthymol blue method on the autoanalyzer 
is commonly used to measure SO4 in aqueous systems (American Public Health Association 
1998). This method can directly measure SO4 in water, unlike the MB colorimetric method, 
which requires reduction of SO4 to H2S. However, similar to CI, some laboratories are 
increasingly using IC and ICP-AES to measure SO4 to avoid working with, and disposing 
of, thymol. Carbonate and bicarbonate ions are most commonly determined by titrating 
(titrimetric method) samples to an endpoint of pH 8.4 using phenolphthalein (CO3) and then 
to pH 4.7 using methyl orange (HCO3) (U.S. Salinity Laboratory Staff 1954). Alternatively, 
a pH probe and meter (electrometric method) can be used to determine the endpoints. 


15.4.1 Electrical Conductivity 

Salt tolerance data for crops have been developed relating crop yield to EC e . Data have been 
compiled for 69 herbaceous crops based on controlled tests in the United States and India 
(Maas 1990) for soils where chloride salts predominate. Salt tolerance data have also been 
compiled by Ayers and Westcott (1985). Crops grown on gypsiferous soils, such as found 
in the Canadian Prairies, will tolerate an EC e of about 2 dS m _1 higher than those listed in 
Maas's table. In Canada, salt tolerance data based on field tests at specific locations have been 


TABLE 15.2 Crop Response to Salinity Measured as Electrical Conductivity 

(EC) of the Saturation Extract 
EC (dS m~ 1 at 25°C) Crop response 

0-2 Almost negligible effects 

2-4 Yields of very sensitive crops restricted 

4-8 Yields of most crops restricted 

8-1 6 Only tolerant crops yield satisfactorily 

>16 Only very tolerant crops yield satisfactorily 

Source: Adapted from Bernstein, L, Ann. Rev. Phytopathol., 13, 295, 1975. 

reported by Holm (1983) and McKenzie (1988). More recently, research at Canada's salt 
tolerance testing facility (Steppuhn and Wall 1999) reported salt tolerance data for spring-sown 
wheats (Steppuhn and Wall 1997), as well as for canola, field pea, dry bean, and durum wheat 
crops (Steppuhn et al. 2001). General salinity effects are presented in Table 15.2. 

15.4.2 Expressing Results of Soluble Ion Analyses 

Soluble salt data are generally expressed in units such as meq L _1 (mmol c L _1 ), mg L _1 , 
or mmol L~ ' . If the results are to be expressed on a mass basis (e.g., mg of Ca per kg of soil), then 
the mass of air-dry soil, the mass of water added, and water already in the soil need to be known. 

15.4.3 Ion Activities and Saturation Index Values 

Soil solution data are generally reported as ion concentrations. However, it may sometimes 
be desirable to express the results as ion activities or thermodynamically effective concen- 
tration. The activity of an element, rather than its concentration, may be more closely related 
to plant response (Adams 1966) and general chemical reactivity (Freeze and Cherry 1979). 
Ion activity is the product of the ion concentration and the activity coefficient. There is an 
inverse relationship between the activity coefficient and ionic strength of the soil solution. 
As salinity or ionic strength of the aqueous solution increases, the activity coefficient 
decreases, resulting in a lower ion activity that can participate in chemical reactions. 
Increasing salinity also increases the solubilities of minerals via the ionic strength effect. 
Ion activities can be estimated from various geochemical models, and some ion activities 
(e.g., CI, K) can be directly measured in solution extracts using ion-selective electrodes. 
Saturation index (SI) values for minerals can also be estimated from geochemical models by 
dividing the ion activity product of the solution species composing the mineral of interest by 
the solubility product constant (K sp ) of the mineral. SI values <0 indicate undersaturation or 
dissolution with respect to the mineral, SI = indicates saturation or equilibrium between the 
solution and solid phase, and SI > indicates supersaturation or precipitation of the mineral. 
However, SI values for evaporate minerals from saline soils were found to be poor predictors 
of minerals formed in evaporated soil solutions (Kohut and Dudas 1994). 

15.4.4 Sodium Adsorption Ratio 

The SAR, a useful index of the sodicity or relative sodium status of soil solutions, and 
aqueous extracts, or water in equilibrium with soil, is calculated as follows: 

where cation concentrations are in mmol L _1 . 


Soils with SAR values greater than 13 are considered to be sodic (Soil Science Society of 
America 2001), although other critical values have been proposed (Bennett 1988; Sumner 
et al. 1998). Equation 15.2 is often referred to as the practical SAR (SAR p ), whereas 
theoretical SAR (SAR,) values are calculated using the same equation but with free ion 
activities instead of concentrations (Kohut and Dudas 1994). Since exchangeable cations are 
difficult to measure in saline soils because of errors arising from anion exclusion or 
dissolution of slightly soluble minerals, the SAR of soil aqueous extracts has become the 
principal tool for diagnosing sodic soils (Bohn et al. 1979; Jurinak 1990). 

15.4.5 Exchangeable Sodium Percentage 

ESP can be estimated from SAR based on the linear equation: 

where K g is the Gapon selectivity coefficient. The value of K g has traditionally been taken as 
0.015 (mmol L -1 )" 05 (U.S. Salinity Laboratory Staff 1954), though K g can vary depending 
on soil organic matter content and pH (Curtin et al. 1995). In general, the affinity of soils for 
Na decreases as the contribution of organic matter to the cation-exchange capacity increases. 

15.4.6 Potassium Adsorption Ratio 

The potassium adsorption ratio (PAR) is calculated by substituting K for Na in Equation 15.2. 
Excessive K concentrations may interfere with crop uptake of other nutrients, decrease soil 
hydraulic conductivity and permeability, and increase soil erodibility (Hao and Chang 2003). 
Potassium concentrations are high in livestock manures, and K may become the dominant 
soluble cation in manured soils (Pratt 1984). Pratt (1984) reported that the long-term hazard of 
the use of manures on well-leached irrigated lands was more from K than from Na accumu- 
lation. Critical PAR values to define soils with excessive K remain to be determined. 

15.4.7 Critical Calcium Ratio 

A number of studies have shown that crop yield in a salt-affected soil is strongly influenced by 
the ratio of Ca to that of other cations in the soil solution (Howard and Adams 1 965 ; Carter et al. 
1979; Janzen and Chang 1987; Janzen 1993). Yield reductions are typically observed when the 
ratio of Ca:total cations is below approximately 0.10. This ratio can fall below the critical 
value in sodic soils (Carter et al. 1 979) and in saline, gypsiferous soils where Ca concentrations 
are low because of the poor solubility of CaSCU • 2H2O (gypsum) (Curtin et al. 1993). 

Adams, F. 1966. Calcium deficiency as a causal and Wastewater, 20th edn. APHA, Washington, 
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Ayers, R.S. and Westcott, D.W. 1985. Water 
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Standard Methods for the Examination of Water age Paper 29 (Rev. 1). FAO, Rome, Italy. 


Basta, N.T. and Tabatabai, M. 1985. Determin- 
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Bennett, D.R. 1988. Soil chemical criteria for 
irrigation suitability classification of Brown 
Solonetzic soils. Can. J. Soil Sci. 68: 703-714. 

Bernstein, L. 1975. Effects of salinity and sodi- 
city on plant growth. Ann. Rev. Phytopathol. 13: 

Bohn, H., McNeal, B., and O'Connor, G. 1979. 
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Cannon, M.E., McKenzie, R.C., and Lachapelle, 
G. 1994. Soil salinit) mapping with electromag- 
netic induction and satellite-based navigation 
methods. Can. J. Soil Sci. 74: 335-343. 

Carter, M.R., Webster, G.R., and Cairns, R.R. 
1979. Calcium deficiency in some Solonetzic 
soils of Alberta. J. Soil Sci. 30: 161-174. 

Corwin, D.I. and Rhoades, J.D. 1982. An 
improved technique for determining soil electrical 
conductivity-depth relations from aboveground 
electromagnetic measurements. Soil Sci. Soc. Am. 

Corwin, D.L. and Lesch, S.M. 2003. Application 
of soil electrical conductivity to precision agricul- 
ture: theory, principles, and guidelines. Agron. J. 
95: 455-471. 

Curtin, D., Selles, F., and Steppuhn, H. 1995. 
Sodium-calcium exchange selectivity as influ- 
enced by soil properties and method of determin- 
ation. Soil Sci. 159: 176-184. 

Curtin, D., Steppuhn, H., and Selles, F. 1993. 
Plant responses to sulfate and chloride salinity: 
growth and ionic relations. Soil Sci. Soc. Am. J. 
57: 1304-1310. 

Frankenberger, W.T. Jr., Tabatabai, M.A., 
Adriano, D.C., and Doner, H.E. 1996. Bromine, 
chlorine, and fluorine. In D.L. Sparks et al., Eds. 
Methods of Soil Analysis, Part 3 — Chemical 
Methods. SSSA Book Series No. 5, SSSA and 
ASA, Madison, WI, pp. 833-868. 

Freeze, RA. and Cherry, JA. 1979. Ground 
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Hao, X. and Chang, C. 2003. Does long-tt 


clay loam soil 

Agric. Ecos) 

'.. 94: 

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id southern Alberta? 

Helmke, PA. and Sparks, D.L. 1996. Lithium, 
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Part 3— Chemical Methods. SSSA Book Series 
No. 5, SSSA and ASA, Madison, WI, pp. 551-574. 

Hendrickx, J.M.H., Baerends, B., Raza, Z.I., 
Sadig, M., and Akram, M. 1992. Soil salinity 
assessment by electromagnetic induction of irri- 
gated land. Soil Sci. Soc. Am. J. 56: 1933-1941. 

Hogg, T.J. and Henry, J.L. 1984. Comparison of 
1:1 and 1:2 suspensions and extracts with the 
saturation extract in estimating salinity in Sas- 
katchewan soils. Can. J. Soil Sci. 64: 699-704. 

Holm, H.M. 1983. Soil Salinity, a Study in Crop 
Tolerances and Cropping Practices. Saskatch- 
ewan Agriculture Publication No. 25M/3/83. 
Plant Industry Branch, Regina, SK, Canada. 

Howard, D.D. and Adams, F. 1965. Calcium 
requirement for penetration of subsoils by primary 
cotton roots. Soil Sci. Soc. Am. Proc. 29: 558-562. 

Janzen, H.H. 1993. Soluble salts. In M.R. Carter, 
Ed. Soil Sampling and Methods of Analysis. 
Lewis Publishers, Boca Raton, FL, pp. 161-166. 

Janzen, H.H. and Chang, C. 1987. Cation nutri- 
tion of barley as influenced by soil solution 
composition in a saline soil. Can. J. Soil Sci. 67: 

Jurinak, J.J. 1990. The chemistry of salt-affected 
soils and waters. In K.K. Tanji, Ed. Agricultural 

Salinity Assessment and Management. ASCE, 
New York, NY, pp. 42-63. 

Kohut, C.K. and Dudas, M.J. 1994. Comparison 
of immiscibly displaced soil solutions and satur- 
ated paste extracts from saline soils. Can. J. Soil 
Sci. 74: 409^119. 

Maas, E. V. 1990. Crop salt tolerances. In K.K. Tanji, 
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McKenzie, R.C. 1988. Tolerance of plants to soil 
salinity. Soil and Water Program, 1987. Pamphlet 


88-10. Alberta Special Crops and Horticultural Methods. SSSA Book Series No. 5, SSSA and 
Research Centre, Brooks, AB, Canada. ASA, Madison, WI, pp. 417-436. 

McKenzie, R.C., Chomistek, W., and Clark, N.F. 
1989. Conversion of electromagnetic inductance 
readings to saturated paste extract values in soil 
for different temperature, texture, and moisture 
conditions. Can. J. Soil Sci. 69: 25-32. 

Nieto, K.F. and Frankenberger, W.T. 1985a. 
Single ion chromatography. I. Analysis of inor- 
ganic anions in soils. Soil Sci. Soc. Am. J. 49: 

Nieto, K.F. and Frankenberger, W.T. 1985b. 
Single ion chromatography. II. Analysis of ammo- 
nium, alkali metals, and alkaline earth cations in 
soils. Soil Sci. Soc. Am. ./. 49: 592-596. 

Pittman, J.J., Kress, M.W., and Zhang, H. 2004. 
Comparison of two soil salinity extraction methods. 
Available at: 
Conf/zhang_31.pdf (last verified March, 2006). 

Pratt, P.F. 1984. Salinity, sodium, and potassium 
in an irrigated soil treated with bovine manure. 
Soil Sci. Soc. Am. J. 48: 823-828. 

Reitemeier, R.F. 1946. Effect of 
on dissolved and exchangeable io 
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Rengasamy, P., Greene, R.B.S., Ford, G.W., and 
Mehanni, A.H. 1984. Identification of dispersive 
behaviour and the management of red-brown 
Earths. Aust. J. Soil Res. 22: 413-431. 

Rhoades, I.D. 1978. Monitoring soil salinity: a 
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K.D. Schmidt, Eds. Establishment of Water Quality 
Monitoring Programs, Vol. 2. American Water Re- 
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Rhoades, J.D. 1990. Determining soil salinity Soil 35: 505-516. 
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Commun. Soil Sci. Plant Anal. 21: 1887-1926. 

Rhoades, J.D., Chanduvi, F., and Lesch, S. 1999. 
Soil salinity assessment: methods and interpret- 
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FAO Irrigation and Drainage Paper No. 57. 
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Rhoades, J.D. and Corwin, D.L. 1981. Determin- 
ing soil electrical conductivity-depth relations 
using an inductive electromagnetic soil conduct- 
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Rhoades, J.D. and Oster, J.D. 1986. Solute con- 
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2nd ed. Agronomy Monograph 9, ASA and 
SSSA, Madison, WI, pp. 985-1006. 

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P. 1999. In New Applications for Nonmetals 
Determination by ICP-AES. American Labora- 
tory, Shelton, CT, pp. 170-171. 

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and laboratory measurements. In K.K. Tanji, Ed. 

Agricultural Salinity Assessment and Manage- 
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2001. Comparing canola, field pea, dry bean, and 
durum wheat crops grown in saline media. Crop 
Sci. 41: 1827-1833. 

Steppuhn, H. and Wall, K.G. 1997. Grain yields 
from spring-sown Canadian wheats grown in 
saline rooting media. Can. J. Plant Sci. 77: 63-68. 

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WI, pp. 65-90. 



Section Editors: Y.K. Soon and W.H. Hendershot 


Chapter 1 6 

Soil Reaction and 

Exchangeable Acidity 

W.H. Hendershot and H. Lalande 

McGill University 
Sainte Anne de Bellevue, Quebec, Canada 

M. Duquette 


Montreal, Quebec, Canada 


Soil pH is one of the most common and important measurements in standard soil analyses. 
Many soil chemical and biological reactions are controlled by the pH of the soil solution in 
equilibrium with the soil particle surfaces. 

Soil pH is measured in an aqueous matrix such as water or a dilute salt solution. Soil pH 
measured in water is the pH closest to the pH of soil solution in the field (this is true for soils 
with low electrical conductivity and for soils that are not fertilized), but is dependent on the 
degree of dilution (the soil to solution ratio). Measuring soil pH in a matrix of 0.01 M CaCl2, 
as opposed to water, has certain advantages, but the addition of the salt does lower the pH by 
about 0.5 pH units compared to soil pH in water (Schofield and Taylor 1955; Courchesne 
et al. 1995). In soil correlation work, the use of pH in CaCl2 is preferred because the 
measurement will be less dependent on the recent fertilizer history. Other methods for soil 
pH measurement, such as pH in 1 M KC1, are presented elsewhere (Peech 1965); these 
methods are not commonly used in Canada for routine analysis and are not included in 
this chapter. 


When measuring soil pH in water, the main concern is that an increase in the amount of 
water added will cause an increase in pH; it is therefore important to keep the ratio constant 
and as low as possible. However, the supernatant solution must be sufficient to immerse the 


electrode properly without causing too much stress when inserting the tip of the 
electrode into the soil and to allow the porous pin on the electrode to remain in the solution 
above the soil. 

16.2.1 Materials and Reagents 

/ pH meter: an appropriate instrument provided with two calibration points should 
be used. 

2 Combined electrode: since the volume of soil is generally limited and the soil to 
solution ratio kept as low as possible, a combination electrode is a valuable asset. 

j 30 ml_ long form beakers (Pyrex or disposable plastic): beakers that have a narrow 
shape help to immerse the electrode in the supernatant without introducing the tip 
into the soil. 

4 Stirrers: disposable plastic stirrers or glass rods can be used. 

16.2.2 Procedure 

/ Weigh 10 g of air-dried mineral soil (<2 mm) into a beaker and add 20 ml_ of 
double deionized (d.d.) water. For organic samples, use 2 g of soil in 20 ml_ of d.d. 
water. Record the soil to solution ratio used. Include duplicate quality control 
samples in each batch. 

2 Stir the suspension intermittently for 30 min. 

3 Let stand for about 1 h. 

4 Immerse the electrode into the clear supernatant and record the pH once the 
reading is constant. Note: Both the glass membrane and the porous salt bridge 
must be immersed. 


Soil samples containing high amounts of organic matter tend to form a thick dry paste when 
the ratio is kept the same as for mineral samples; therefore, a decreased ratio of sample to 
water must be accepted (1:5 or 1:10). 

Two pH standards should be used to calibrate the pH meter; they must be chosen in accordance 
with the pH range expected for the soils analyzed (pH 4.0 and 7.0 or pH 7.0 and 10.0). 

A large amount of a soil similar to the samples being analyzed should be kept as an indicator 
of the variability of pH results over time; duplicate subsamples of this quality control (QC) 
sample should be run with each batch of samples measured. Failure of the QC to fall within 
acceptable limits means that the whole batch should be reanalyzed. 

16.3 SOIL pH IN 0.01 M CaCl 2 

Standard measurement of soil pH in CaCl2 is probably the most commonly used method to 
characterize soil pH. As mentioned by Peech (1965), Davey and Conyers (1988), and 
Conyers and Davey (1988), the use of CaCi2 has some advantages for pH measurement: 
(1) the pH is not affected within a range of the soil to solution ratios used, (2) the pH is 
almost independent of the soluble salt concentration for nonsaline soils, (3) this method is a 
fairly good approximation of the field pH for agricultural soils, (4) because the suspension 
remains flocculated, errors due to the liquid junction potential are minimized, (5) no 
significant differences in soil pH determination are observed for moist or air-dried soil, 
and (6) one year of storage of air-dried soil does not affect the pH. 

16.3.1 Material and Reagents 

7 pH meter: an appropriate instrument provided with two calibration points should 
be used. 

2 Combined electrode: since the volume of soil is generally limited and the soil 
to solution ratio kept to a minimum, a combination electrode is a valuable asset. 

3 30 ml_ long form beakers (Pyrex or disposable plastic): beakers that have a narrow 
shape help to immerse the electrode in the supernatant without introducing the tip 
of the electrode in the soil thus avoiding breakage. 

4 Stirrers: disposable plastic stirrers or glass rods can be used. 

5 Calcium chloride, 0.01 M: dissolve 2.940 g of calcium chloride dihydrate 
(CaCI 2 • 2H 2 0) with d.d. water in a 2 L volumetric flask. The electrical conduct- 
ivity of the CaCb solution must be between 2.24 and 2.40 mS citT 1 at 25°C. 

16.3.2 Procedure 

1 Weigh 1 g of air-dried mineral soil (<2 mm) or 2 g of organic soil into a 30 ml_ 
beaker and add 20 ml_ of 0.01 M CaCI 2 . Note the soil to solution ratio used. 
Include duplicate quality control samples in each batch. 

2 Stir the suspension intermittently for 30 min. 

3 Let stand for about 1 h. 

4 Immerse a combination electrode into the clear supernatant and record the pH 
once the reading is constant. Note: Both the glass membrane and the porous salt 
bridge must be immersed. 


The pH and electrical conductivity of the CaCl2 should be fairly constant, i.e., pH 
in the range of 5.5-6.5 and the electrical conductivity around 2.3 mS cm~' at 25°C. If the 
pH is outside this range, it should be adjusted with HC1 or Ca(OH) 2 solution. If the electrical 
conductivity is not within the acceptable range, a new solution must be prepared. 



In addition to bases (e.g., Ca, Mg, K, Na) there is also an amount of acidity that can be 
displaced from the exchange complex of a soil. The amount of this acidity is largely a 
function of soil pH and the exchange capacity. In most soils the exchangeable acidity will 
be composed of (i) exchangeable H + , (ii) exchangeable Al as either Al 3+ or partially 
neutralized Al-OH compounds such as A10H 2+ or Al(OH) 2 + , and (iii) weak organic acids. 

When a soil is limed, the exchangeable acidity is neutralized as the pH rises. Hence, 
exchangeable acidity is one measure of the amount of lime that will be needed to correct 
soil pH. 

The method of Thomas (1982) used 1 M KC1 as the displacing salt solution, whereas the 
Expert Panel on Soil (2003) proposes 0.1 M BaCl2. Since the method proposed in this 
manual for measuring exchangeable cations uses 0.1 M BaCi2, it seems more appropri- 
ate to use the same salt solution for measuring exchangeable acidity. Due to the lower 
concentration of the BaCb solution, the amounts of some cations are lower than when the 
extraction is done with KC1; however, Jonsson et al. (2002) have determined regression 
equations that could be used to estimate the difference between the two extraction 

16.4.1 Materials and Reagents 

1 50 ml_ centrifuge tubes, a centrifuge capable of generating 5000 g and an end- 
over-end shaker (15 rpm). 

2 Replacing solution, barium chloride 0.1 M: dissolve 24.43 g of BaCI 2 -2H 2 with 
distilled deionized (d.d.) water and make to volume in a 1 L volumetric flask. 

3 Aluminum complexing solution, 1 M sodium fluoride: dissolve 41 .99 g of NaF in 
about 900 ml_ of d.d. water in a 1 L beaker and then titrate to the phenolphthalein 
endpoint with sodium hydroxide (NaOH). Transfer to a 1 L volumetric flask and 
make to volume. 

4 Sodium hydroxide (NaOH), approximately 0.05 M, standardized. 

5 Phenolphthalein solution: dissolve 1 g of phenolphthalein in 1 00 ml_ of ethanol. 

16.4.2 Procedures 

7 Weigh a 2.5 g sample of mineral soil or 2.0 g of organic soil into a 50 ml_ 
centrifuge tube, add 30 ml of 1 M BaCb solution, and shake for 1 h. Centrifuge 
at 5000 gfor 10 min. Transfer supernatant liquid to a 100 ml_ volumetric flask. 
Repeat by adding 30 ml_ aliquots of BaCb solution, shaking, centrifuging, and 
decanting two more times, collecting all the supernatant in the same 100 ml_ 
volumetric. Make up to volume with BaCb solution and mix. Filter the extract 
(Whatman No. 42 or equivalent) into a plastic bottle and store in a refrigerator 
prior to analysis. 


2 To obtain exchangeable acidity, pipette 25 mL of the extract into a 100 mL 
polyethylene beaker, add 4 or 5 drops of phenolphthalein, and titrate with 
0.05 M NaOH to the first permanent pink endpoint; record the volume 
of NaOH used as VA. (Note: A deep pink is too far.) Titrate a blank (25 mL of 
BaCb solution) to the endpoint and record the amount VB. Centimoles 
of BaCb-extracted acidity per kg of soil (cmol(+) kg -1 ) are calculated as 
shown below. 

3 To determine exchangeable H + acidity, pipette 25 mL of the extract into a 1 00 mL 
polyethylene beaker, then add 2.5 mL of 1 M NaF, and titrate with 0.05 M NaOH 
to the first permanent pink endpoint (Va). Repeat with a blank sample of BaCb (Vb). 

16.4.3 Calculation 

g sample x Vs 

cmol(+) kg -1 H+ acidity is calculated using the same equation replacing VA by Va and 
VB by Vb, where VA or Va are the volumes of titrant used for the determination of 
exchangeable acidity and H + , Vs is the volume of extract titrated and V is the total volume 
of extract collected, M(NaOH) is the concentration of the titrant, and g sample is the mass of 
soil extracted. 


1 The procedure has been written using a pH indicator solution, which is our 
preference for manual titrations. However, if an automated titrator is used, the 
endpoint should be set at pH 7.8. 

2 Exchangeable cations and exchangeable acidity (including H + ) can all be deter- 
mined on the extracts obtained by this multiple washing procedure; this is the 
procedure recommended by the Expert Panel on Soil (2003). Although this 
extraction procedure is somewhat more complicated than the 0.1 M BaCb 
method proposed in Chapter 18 (Section 18.2), it should give similar results. 

Conyers, M.K. and Davey, B.G. 1988. Observa- Davey, B.G. and Conyers, M.K. 1988. Deter- 

tions on some routine methods for soil pH deter- mining the pH of acid soils. Soil Sci. 146: 141-150. 
mination. Soil Sci. 145: 29-36. 

Expert Panel on Soil 2003. Manual on Methods 

Courchesne, F., Savoie, S., and Dufresne, A. and Criteria for Harmonized Sampling, Assess- 

1995. Effects of air-drying on the measurement ment. Monitoring and Analysis of the Effects 

of soil pH in acidic forest soils of Quebec, of Air Pollution on Forests. Part Ilia. Sampling 

Canada. Soil Sci. 160: 56-68. and Analysis of Soil. International co-operative 


programme on assessment and monitoring of air Part 2. American Society of Agronomy, Madison, 
pollution effects on forests, (www.icp-forests. WI, pp. 914-926. 
org/pdf/ manual3a.pdf, verified February 9, 2005) 

Schofield, R.K. and Taylor, A.W. 1955. The 
Jonsson, U., Rosengren, U., Nihlgard, B., and measurement of soil pH. Soil Sci. Soc. Am. Proc. 
Thelin, G. 2002. A comparative study of two 19: 164-167. 
methods for determination of pH, exchangeable 

base cations and aluminium. Commun. Soil Sci. Thomas, G.W. 1982. Exchangeable cations. In 
Plant Anal. 33: 3809-3824. A.L. Page et al., Eds., Methods of Soil Analysis. 

2nd ed. American Society of Agronomy, Madison, 
Peech, M. 1965. Hydrogen-ion activity. In WI, pp. 159-166. 
C.A. Black et al., Eds., Methods of Soil Analysis. 


Chapter 1 7 

Collection and Characterization 

of Soil Solutions 

J.D. MacDonald 

n 1 1 LI I If/ Agri-I f I Mi, 

Quebec, Quebec, Canada 

N. Belanger 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 

S. Sauve and F. Courchesne 

University of Montreal 
Montreal, Quebec, Canada 

W.H. Hendershot 

McGill University 
Sainlt Anne de Bellevue, Quebec, Canada 


The soil solution plays a dominant role in the uptake of nutrients by plants, has direct impacts 
on other soil-living organisms, and under certain conditions it is the vector for the migration 
of dissolved and suspended materials through soils. We are defining the soil solution as the 
liquid phase of the soil present in the field. This definition precludes all methods in which 
salt solutions, or water, are added to a soil sample in the laboratory to simulate soil solutions 
as these procedures may be more accurately defined as soil extractions. 

Studies of atmospheric deposition on soils and watersheds often incorporate monitoring of 
soil solution chemistry. In many cases lysimeters are used to collect soil solution on a regular 
basis to assess the ability of soils to absorb atmospherically deposited material and to control 
the release of nutrients and contaminants to ground and surface waters. Studies of macro- 
it availability in forest soils may also use lysimeters as a means of measuring available 
. Examples of these types of studies are frequently found in the literature (Haines 
et al. 1982; Beier et al. 1992; Foster et al. 1992; Hendershot et al. 1992; MacDonald et al. 
2003; Belanger et al. 2004). 


Field soil solution collectors fall into two main categories: zero-tension lysi 
tension lysimeters. Both forms of lysimeters consist of an apparatus that is inserted into 
the soil column and collects water either moving through or held within soil capillaries. 
Recently, tension lysimetry has been further refined to include microlysimeters that can be 
used in the study of microenvironments in the soil. The development of microlysimeters 
shows potential for the study of the heterogeneity of soil solution chemistry at the microscale 
that is known to occur in soils, yet is poorly understood. 

Methods of obtaining soil solution from fresh soil in the laboratory or directly in the field 
have been developed over the years (Heinrichs et al. 1995; Lawrence and David 1996). 
These techniques typically result in soil solutions with much higher concentrations of many 
ions, including dissolved organic matter, than those collected using field lysimeters (Ludwig 
et al. 1999) and some inconsistencies are observed in results depending on soil moisture 
content (Jones and Edwards 1993). Nonetheless, laboratory methods for sampling soil 
solutions from freshly sampled soils, e.g., centrifugation, miscible displacement, and syringe 
pressure, are useful tools for investigating plant nutrition and worthy of examination 
(Smethurst 2000). 

We include a centrifugation method in this chapter; however, due to concerns raised about 
the influence of the force of extraction used to extract these solutions, we also propose a 
simple method developed by Ross and Bartlett (1990) that extracts solutions by applying 
pressure to moist soils packed in syringes in the field. Like the proposed centrifugation 
method, the syringe compression method is rapid and simple, and provides an alternative 
approach in cases where there is concern that centrifugation may overestimate concentra- 
tions of certain elements in solutions. 

It is not always possible to carry out field studies. To approximate soil solution chemistry in 
the laboratory using bulk soil samples, we suggest a weak CaCL; (0.01 M) shake and 
centrifuge extraction (Quevauviller 1998) due to its simplicity and because it is an approach 
that is easily standardized. We also propose a column leaching extraction method that has 
been developed for bulk soil samples and has been shown to provide results similar to those 
obtained from zero-tension lysimeters (MacDonald et al. 2004a,b). 

In this chapter, we provide five different procedures to separate the solution phase from the 
solid phase of soils. For in situ studies we propose zero-tension and tension lysimeters, as 
well as microlysimeters. To acquire solutions from fresh soil samples in the laboratory, we 
suggest centrifugation and syringe pressure techniques. The preferred methods of laboratory 
approximations of soil solutions, the column leaching method and weak CaCb (0.01 M) 
extraction, can be found in Chapter 10. The applications, advantages, and disadvantages of 
the different methods of acquiring the soil solution from soils and the solution extractions 
described in Chapter 10 are summarized in Table 17.1. 


Zero-tension lysimeters collect water moving through the soil profile only when the soil 
moisture content is greater than the field capacity. Several different designs of zero-tension 
lysimeters have been proposed. These include simple plates inserted in the soil, models with 
pierced plates installed in funnels, fiberglass wick-type collectors, and funnels filled with 
quartz sand. Our preferred design uses a plastic funnel filled with 2 mm quartz sand. Quartz 
sand is relatively unreactive and when the lysimeter is installed with a good contact between 


TABLE 17.1 Advantages, Disadvantages, and Potential Applications of Different Sample Methods and Sampler Types Proposed in the Present Chapter and 
Chapter 1 
Type Advantages Disadvantages Potential applications 


Samples can be collected from 
column over many years 
Simple system with low 

n except u 


Does not colle< 

Requires long equilibration (many months 

to a year) period before samples are 


Requires long equilibration (many months 
to a year) period before samples are 

Variable results with soil moisture 

Not efficient in surface organic horizons 

High maintenance, fragile devices that are 
difficult to maintain under field conditions 
Requires equilibration period 
Very small (sub ml_) solution volumes, 
hence need adapted analytical procedures 
and systems 

Samples can be extracted rapidly from fresh Lack of reproducibility of results as solution 
moist soils concentrations are dependent on moisture 

Reasonable alternative when lysimetry is content of soils and time of sampling 

not available or feasible Small solution volumes and potential 

overestimations of certain anions and 
dissolved organic carbon (DOC) 

Tension lysimeters Samples can be collected from s 
column over many years 

Microlysimeters Samples 

Sample solutions that move through the soil 
with only gravitational potential 
Long-term solution chemistry monitoring 
Mobility of elements and fine particles in 

Sample solutions held at potentials greater 
than gravitational potential 
Long-term soil solution monitoring 
Availability of elements in soils 

n be collected a 

Very high spatial resolution 
Samples can be repeatedly collected from 
the same location 

Sample solutions in soil microenvironment, 
e.g., in the rhizosphere, the outer layer of 
peds, or in the coatings lining macropores 

Microscale heterogeneity of soil solutions, 
nutrient uptake, biogeochemical processes 

Provides a single point in time extraction of 
capillary solutions present in soils 
Provides estimates of nutrient availability in 


TABLE 1 7.1 (continued) Advantages, Disadvantages, and Potential Applications of Different Sample Methods and Sampler Types Proposed ii 
Chapter and Chapter 1 



Potential applications 

Column leaching 

0.01 M CaCI 2 : 

Samples can be extracted rapidly from 

Minimum disturbance of the soil during 

solution acquisition 
Reasonable alternative when lysimetry is 

not available or feasible 

Soil samples can be collected and extracted 

relatively quickly 
Solution samples can be replicated from the 

same bulk soil sample 

Lack of reproducibility of results as solution 
concentrations are dependent on moisture 
content of soils and time of sampling 

May yield small amounts of solution during 
dry periods, especially in coarse soil 
horizons (e.g., sandy loam) with low 
organic matter content 

Estimates of field partitioning, but the 
method does not reproduce true field 

Does not provide information on ni 
availability, N, or P 

Provides a single point in time extraction of 
capillary solutions present in soils 
Provides estimates of nutrient availability in 

May avoid overestimation of certain 
elements associated with the force of 
extraction used in centrifugation 

Estimation of field partitioning coefficients 
of divalent metals 
Adsorption-desorption studies 

Solution samples can be replicated from the Sol 
same bulk soil sample sc 

Easily standardized methodology 

e often very different from field 
ollected from the same soil at 

Cannot measure Ca 2+ and CI 


about the chemistry of soil 
-desorption and bioavailability 


the quartz sand and the soil column above it, the soil-lysimeter interface has a large c 
area. Capillary flow is not as likely to be interrupted and consequently the quartz sand design 
will tend to collect water under circumstances where the plate lysimeters and pierced plate 
lysimeters do not. The material is inexpensive relative to commercially available lysimeters. 
It is better to avoid using the fiberglass wick collectors since the fiberglass is more reactive 
than the quartz sand, and wick samplers have been suggested to cause modifications to soil 
solution chemistry (Goyne et al. 2000; Brahy and Delvaux 2001). 

17.2.1 Materials 

7 180 mm diameter polyethylene or polypropylene funnels 

2 2 mm quartz sand 

j 75 mm diameter (3") or 50 mm (2") acrylonitrile-butadiene-styrene (ABS) drain 
pipe about 2 m long 

4 ABS end cap, and flexible cap for ABS drain pipe 

5 19 mm (3/4") plastic hose pipe fittings, one L-shaped and one straight 

g 19 mm (3/4") clear plastic hose with 3 mm (1/8") thick wall and about 70 cm 

j Epoxy two component cement and ABS solvent glue 

8 Nylon screening 25 x 50 mm 

9 5% (v/v) HCI 

17.2.2 Preparation (Figure 17.1) 

7 The funnels are prepared by gluing the L pipe fitting into the bottom with epoxy. 

2 Cut the ABS drain pipe so that once it is buried it extends above the ground by at 
least 45 cm (to avoid rain-splash), or in areas of snow, at least the depth of normal 
snow cover. Attach the hard plastic ABS end cap to one end with either epoxy or 
ABS glue. Clearly label (paint or engrave) each pipe near the top. 

3 Drill a hole in the ABS pipe and insert the straight hose fitting by coating with 
epoxy and hammering into place. The distance of the hose fitting from the base of 
the pipe is dependent on the desired volume of the lysimeter reservoir. For 75 mm 
ABS and a reservoir volume of 1 L, the hose fitting is installed at 23 cm from the 
bottom, whereas for 2 L, the distance is 45 cm. With 50 mm ABS, the distance for 
a 1 L reservoir volume is 51 cm. In shallow or rocky soil the 75 mm ABS is easier 
to install. 

4 Acid wash the ABS pipe, the funnel, the clear plastic tube, and enough quartz 
sand to fill the funnel with 5% HCI. Rinse with deionized water until the electrical 
conductivity (EC) of wash water is equal or close to that of deionized water. 


Install fitting 23 cm from 
pipe base for 1 L solution 
n 3" ABS-tubing. 

Flexible cap 

ABS drain pipe 

FIGURE 17.1. Schematic of zero-tension lysimeter constructed from ABS-tubing using a silic 
sand filled funnel as a solution collection device. 

Place the quartz sand and the funnels with the clear tubing attached into separate 
plastic bags for transport to the field. The ABS tubes should have plastic taped over 
the hose fitting and the flexible plastic cap placed on the upper end. 

17.2.3 Installation Procedure 

Cordon off the location of the lysimeter station taking care to avoid walking on the 
area or contaminating it with soil or other debris. 

Dig a pit downslope, if the site is not flat, approximately 1 m 2 to a depth greater 
than that at which the lowest lysimeter is to be installed. Separate the surface 
layers from the underlying soil so that they can be replaced in such a way that the 
site is disturbed as little as possible at the end of the installation. 

Starting with the deepest lysimeter, dig a tunnel into the side of the pit under the 
delineated area. Use a spare funnel to make sure the tunnel is cut to the correct 
size to avoid contaminating the acid-washed funnels. 

Insert the nylon screen into the L fitting, fill the funnel with the quartz sand, and 
carefully slide it into place making sure that it makes good contact with the soil 
above it. Press it into place, pack rocks under the L fitting at the bottom, and 
backfill carefully all around the under surface of the funnel. Ensure that the soil 
has been well packed around the lysimeter such that contact between the silica 
sand and the soil column is solidly maintained. Attach the clear tube to the hose 
fitting in the ABS tube. 


5 Several lysimeters at different depths can be installed in the same pit; however, it 
must be ensured that funnels are placed such that each has an undisturbed cone of 
soil above it (often requiring a larger pit than anticipated). When all the lysimeters 
have been installed in the face of the pit, make sure that the clear tube slopes 
downward from the funnel to the ABS tube. Record the placement of the 
collectors. Carefully refill the pit. 


Porous cup tension lysimeters are inserted into the soil such that the porous surface is in 
contact with the capillaries of the soil column. When a vacuum is applied to the porous cup, 
solution is drawn out of the capillaries into the lysimeter reservoir. Tension lysimeters 
extract soil solutions that are maintained within the micropores of the soil and consequently 
may be immobile. The solutions that they extract have been observed to differ significantly 
from zero-tension lysimeters (Haines et al. 1982; Hendershot and Courchesne 1991). 

Various types of tension lysimeters are available, differing in the type of porous cup that is 
inserted into the soil. The most commonly used tension lysimeters observed in the 
literature are the ceramic cup lysimeters that are installed from the surface. Recently, porous 
poly(tetrafluoroethene) or Teflon* cups have been developed for tension lysimeters to avoid 
the impact that the exchange capacity of ceramic cups can have on solution chemistry 
(Swenson 1997; Russell et al. 2004). We have used ceramic cups in the past; and we feel 
that after adequate stabilization periods in the soil, the ceramic cups are representative of 
macroelements in soil solution. However, the new Teflon-treated cups appear to be less 
reactive and are therefore a more reliable method to extract solutions under tension. 

17.3.1 Preparation 

The lysimeters should be cleaned following the manufacturer's recommendation or using the 
following procedure. Place the lysimeters in a container with 5% HC1 and draw the solution 
through the porous cup and into the lysimeters using suction. Repeat this procedure three 
times and ensure that the PVC shaft above the porous cup is also effectively acid washed. 
Rinse with deionized water until the EC is close or equal to that of deionized water and is 
constant (may take up to 10 washings). When clean, place the lysimeters in clean plastic bags 
ready to go into the field. 

17.3.2 Installation Procedure 

7 Cordon off the location of the lysimeter station and take care to avoid walking on 
the area or contaminating it with soil or other debris. 

2 a Surface installation: place a plastic sheet with a hole the same diameter as the 
lysimeters on the soil surface to trap soil as it is excavated. Using an auger the 
same size as the lysimeters, dig a hole to the required depth. 

i. Install the lysimeters and refill the hole around the lysimeter shaft with 

soil from the same soil horizon in which the lysimeters are installed. 

Carefully reconstruct the soil horizons above the lysimeters until the hole 
is filled. 


ii. Ensure that the soil is tightly sealed around the lysimeter shaft so that prefer- 
ential flow does not occur. In soils where good soil-to-lysimeter contact is 
difficult to establish, a slurry can be prepared using soil taken from the same 
depth as that of the lysimeter. A small amount of slurry is poured into the auger 
hole before installation of the lysimeter. 

2/5 Pit installation: dig a pit approximately 1 m 2 to a depth greater than that at which 
the lowest lysimeter is to be installed. 

i. Separate the surface layers from the underlying soil so that they can be 
replaced in such a way that the site is disturbed as little as possible at the 
end of the installation. 

ii. Starting with the deepest lysimeter, dig a tunnel into the side of the pit under 
the delineated area equal in diameter to the porous cup. Carefully insert the 
porous cup ensuring good contact with the tunnel walls. Repeat for all 
lysimeter depths. 

iii. Connect the vacuum and sample tubes. Record the position of the lysimeters 
and carefully refill the pit and replace the surface layers. 

j Apply a vacuum of 30-60 kPa to the lysimeter. It is recommended that a constant 
vacuum be maintained in the lysimeter. Constant vacuum systems will provide 
cumulative samples over periods between sample collections; however, systems 
that maintain a constant vacuum between sampling periods are expensive. It is 
also possible to use discontinuous systems and apply a vacuum for a period of 
several days before sample collection. It should be noted that discontinuous 
vacuum systems will provide samples that are representative of the short time 
period over which the vacuum is maintained. 


Soil solutions can be extracted from lysimeter reservoirs using handheld vacuum pumps or 
peristaltic pumps ensuring that solutions are not cross-contaminated during collection. 

7 Lysimeters should be completely emptied each time they are sampled. Record the 
total volume of solution removed from the lysimeter. 

2 Solutions should be transferred immediately to coolers and maintained at 4°C in 
the dark for transport to the laboratory. 

j Once solutions are in the laboratory, set aside a small subsample of soil 
solutions (1 0-20 mL) and filter the rest of the solution using low vacuum through 
0.4 (jum polycarbonate filters. Solutions intended for analysis of elements that could 
be modified through contact with the air (nitrogen species for example) should be 
sealed in polycarbonate vials immediately after filtration, leaving little to no air 
space. A subsample for metal analysis should be acidified (0.2% HNO3 v/v); trace 
metal-grade acid should be used if trace elements are to be analyzed. 

4 Filtration will modify solution pH, therefore take the pH and EC of unfiltered 
subsamples of solutions immediately at room temperature. 



7 Solutions should be drawn from the lysimeter reservoirs on a regular sampling 
schedule. Typically, lysimeter monitoring is carried out on a weekly, biweekly, or 
monthly schedule. Solutions that remain in the reservoir for long time periods may 
be modified, due to decomposition of dissolved organic carbon or the dissolution 
of suspended colloidal materials. Furthermore, it should be noted that lysimeter 
solutions, once separated from the soil, do not preserve in situ gas partial 
pressures and their associated chemistry. 

2 The installation of lysimeters causes significant disturbance to the soil. Ensure that 
the lysimeters have stabilized before beginning a sampling regime. After installa- 
tion, the pH and EC of lysimeter solutions should be monitored. Solutions cannot 
be considered representative of the soil chemistry until the pH and EC of the 
solution have stabilized. Stabilization periods for lysimeters can be long (6 months 
to 1 year). The pH and EC are good indicators of the stabilization point of soil 
solutions, but the initial data produced from lysimeters should be examined to 
ensure that stabilization of all elements of interest has occurred, particularly for 
nitrogen species. 


The investigation of the microscale heterogeneity of soil materials, in particular the spatial 
variability in the liquid phase, requires a lysimeter system that is adapted to the character- 
istic small scale of the soil environment of interest. Gottlein et al. (1996) described a system 
for microscale lysimetry that allowed the monitoring of soil solution at a high spatial 
resolution to study gradients in concentrations of elements in the root-soil interface. The 
lysimeter unit consists of a 1 mm diameter ceramic cell with 1 |jim pore size attached to 
1.59 mm capillary tubing and connected to a vacuum device to extract the solution from 
the soil matrix. At a suction of 35 kPa, these cylindrical cups can sample solution in the 
volume of soil extending to a distance of > 1 cm from their surface (Gottlein et al. 1996) and 
sample volumes range from 50 to 300 jjlL collected on a weekly basis at a suction of 40 kPa. 
Other microlysimeter designs have been proposed, but the cylindrical microlysimeters 
developed by Gottlein et al. (1996) are presented in this chapter because their design has 
been the most widely tested. 

17.5.1 Materials 

7 Ceramic capillaries with porosity of about 48%, 1 mm wide, and a suggested 
maximum pore size of 1 |xm. 

2 Polyetheretherketone (PEEK) tubing 1 .59 mm (1/16") wide, 50 mm long with an 
inside diameter (ID) of 0.75 mm; this tubing, used for high-pressure liquid 
chromatography (HPLC), is widely available (see Section 17.5.4). 

3 Epoxy, two component cement. 

4 PEEK tubing with an ID of 0.25 mm. 

5 HPLC fitting to couple microlysimeters with 0.25 mm ID tubing. 

6 Vacuum pump. 

7 Vacuum chamber made of PVC with a Plexiglas cover with a connector to attach 
the vacuum pump (see Figure 1 7.2). 

g Sampling vials 2 ml_ in volume with caps. 

g Vial rack. 

1q Plexiglas plate, rigid and about 20 mm thick, or rhizotron made of transparent 
Plexiglas plates. 

7 7 Stainless steel rod with the same dimensions as the individual microlysimeters 
(1 .59 mm wide x 50 mm long). 

17.5.2 Construction and Preparation (Figure 17.2) 

7 Cut the ceramic capillary into 12 mm long segments. 

2 Seal the tip (exterior end) of the ceramic capillary by melting over a Bunsen 
burner to obtain a microceramic cup 10 mm long with a glass tip. 

3 Cut the 0.75 mm ID PEEK tubing into 50 mm lengths. 

4 Insert the 1 mm long ceramic cup 5 mm into the 0.75 mm ID PEEK tube. 

5 Glue the ceramic cup to the PEEK tube using a two-part cement to complete 
assembly of the microlysimeter (Figure 1 7.2). 

g Clean the microlysimeters by drawing 5% HCI through the porous cup and into 
the tubing using suction. Repeat three times and then rinse with deionized water 
until the EC is close or equal to that of deionized water and is constant (may take 
up to 10 washings). 

j Fix a 0.25 mm ID PEEK tubing of the appropriate length to each of the micro- 

q Construct a vacuum chamber made of transparent Plexiglas and connected to a 
vacuum pump, as in Figure 17.2. 

g Install the vial rack and vials with caps in the vacuum chamber. Pierce holes in the 

1q Connect the tubing fixed to the microlysimeters to the vial through the hole 
pierced in the cap to avoid contamination and limit evaporation. 

7 7 When clean, place all equipment in clean plastic bags ready to go into the field. 


Polyetheretherketone (PEEK) 

capillary 1.59 mm (1/16") 

50 mm out to collection device 

Long nails for fixation 

FIGURE 17.2. Schematic of microlysimeter suction device, support plate, and sample collection 
chamber for solutions from microlysimeters for microlysimeter installation. (From 
Gottlein, A., Hell, U., and Blasek, R., Ceoderma, 69, 1 47, 1 996. With permission.) 

17.5.3 Installation Procedure (Figure 17.3) 

7 Determine the location where the microlysimeters are to be installed in the soil, 
either on the face of a natural profile or in soil materials contained in a rhizotron. 

At that point, make a hole in the soil having the din 
using the stainless steel rod. 

n of the microlysimeters 


Grid of microlysimeters 

FIGURE 1 7.3. Installation of microlysimeters. (From Dieffenbach, A., Gottlein, A., and Matzner, E., 
Plant Soil, 192, 57, 1997. With permission.) 

3 Remove the rod from the channel and insert the microlysimeter in the soil to the 
desired depth. Determine the exact position of the tip of the suction cup. 

4 Use the Plexiglas plate, with holes the size of the microlysimeters (or one of the 
faces of the rhizotron), to support individual microlysimeters and to ensure their 
precise and constant position in the soil (Figure 17.3). 

5 Apply a vacuum of 30-40 kPa to the microlysimeter. It is recommended that a 
constant vacuum be maintained in the lysimeter. It is also possible to use 
discontinuous systems and apply a vacuum for short time periods. 

6 Like any lysimeter, microlysimeters should be allowed to equilibrate with the 
surrounding soil and the pH and EC of solutions should be monitored. Once pH 
and EC are stable, data from the microlysimeters can be considered to be 
representative of soil solution chemistry. 

y Solutions should be transferred immediately to coolers and maintained at 4°C in 
the dark for transport to the laboratory. 


Microlysimeters solution volumes are small and can easily be contaminated, so 
the selection of tubing and container types is crucial to limit the adsorption of 
major ions, trace metals, or organic acids to surfaces during sampling and storage. 
Nylon or Teflon is recommended to reduce the sorption of trace metals whereas 
glass materials are suggested for dissolved organic substances. 

The solution volumes collected with microlysimeters are in the range 50-300 |xL. 
Therefore, analytical methods adapted to very small solution volumes are needed; 
for example, capillary electrophoresis (CE) (Gottlein and Blasek 1996) and 
other methods based on high-resolution inductively coupled plasma-mass 


spectrometry (Puschenreiter et al. 2005) have been used to analyze major anions 
and cations in very small sample volumes. 


A variety of methods to obtain soil solutions in the laboratory from freshly sampled soils 
have been proposed. These methods include low- and high-speed centrifugation (Gillman 
1976; Reynolds 1984) displacement methods with miscible (Adams 1974; Wolt and Graveel 
1986) and immiscible liquids (Kinniburgh and Miles 1983) and positive air pressure in 
sealed cylinders (Lawrence and David 1996). These methods have been compared and 
generally produce similar results (Adams et al. 1980; Wolt and Graveel 1986; Elkhatib 
et al. 1987). In all cases, the key to obtaining minimally altered results is the processing of the 
sample shortly after collection. Centrifugation is recognized as a rapid and simple method. 
The method that we propose is the classic Davies and Davies (1963) method outlined in the 
previous edition of this book with the exception that we propose the use of high-density 
polyethylene (HDPE) frits to contain the soil in the syringe as opposed to glass wool. 

Although centrifugation is probably the most commonly used method to separate the soil 
solution from the solid phase in the laboratory, Ross and Bartlett (1990), when comparing 
high-speed centrifugation with miscible displacement and syringe compression on forest 
floor and Bhf horizons, came to the conclusion that high-speed centrifugation should be 
avoided as it yields high H + and F concentrations as well as occasionally high CT~, S0 4 2 ~, 
and N0 3 ~ levels. The miscible displacement method, though it yielded large amounts of 
solution, was tedious and time-consuming. Since increased processing time inevitably results 
in increased alteration of soil solutions, we feel that the simple and relatively rapid syringe 
pressure technique is a good alternative for extracting solutions from moist soils. The syringe 
technique yielded solutions with similar chemistry to that of the miscible displacement 
method and the precision of analyses on duplicated samples was as good, or better, than 
the displacement or centrifugation methods. 

17.6.1 Centrifugation (Davies and Davies 1963) 
Material and Equipment 

1 The centrifuge apparatus is a 60 ml_ syringe that has been cut to 55 mm and is 
used to contain the fresh moist soil sample and a solution cup that can be made by 
cutting the top of a 50 ml_ HDPE centrifuge tube (see Figure 1 7.4). 

2 Centrifuge with horizontal rotors and 50 mL centrifuge shields or adaptors, 
preferably with refrigeration. 

3 HDPE frits, 27 mm in diameter. 

4 Small solution bottles (HDPE). 

5 Parafilm. 

g 0.4 |j,m polycarbonate membrane filters. 


Cut 60 ml HDPE syringe 

W- n 

~ Soil solution repository 

FIGURE 17.4. Schematic of device used to collect soil solutions during separation with a 
centrifuge. (From Soon, Y.K. and Warren, C.J., in M.R. Carter (Ed.), So/7 Sampling 
and Methods of Analysis, Lewis Publishers, CRC Press, Boca Raton, Florida, 1 993. 
With permission.) 

1 All plasticware in contact with soil samples and solutions should be acid washed 
(5% HCI) and rinsed with deionized water until the EC of the rinse water is close 
or equal to that of deionized water and is constant. If trace elements are of 
interest, plasticware should be prepared according to procedures outlined in 
Chapter 10. 

2 Insert an HDPE frit into the base of the modified 60 ml_ syringe. 

3 Place about 25 g of moist soil in the soil container (1 g if the soil is organic) and 
cover with parafilm to avoid evaporation during the centrifugation procedure. 
A subsample of each soil may be kept to determine the moisture. 

4 Place the solution collecting cup under the syringe containing the soil in the 
centrifuge shield. 

5 Centrifuge at a relative centrifugal force (RCF) of 1 500 g at the bottom of the soil 
column for 30 min. 

5 Set aside a portion of the solutions for analysis of pH and EC. Transfer the 
rest of the solution to clean storage bottles. Solutions may be further 
filtered using low vacuum through 0.4 jjim polycarbonate filters before storage 
and analysis. A subsample for metal analysis should be acidified (0.2% HN0 3 
v/v); trace metal-grade acid should be used if trace elements are to be 

7 Replicate all samples and include blanks. 


17.6.2 Syringe Pressure Method (Ross and Bartlett 1990) 

1 60 ml_ polyethylene syringes 

2 HDPE frits, 27 mm in diameter 

3 Deionized H 2 

4 Compression apparatus (see Figure 1 7.5) 

5 0.4 |jim polycarbonate membrane filters 

7 Wash HDPE frits with deionized H 2 0. 

2 Fit the HDPE frits into the bottom of the syringes. 

3 Pack fresh soil samples (ideally within 12 h of sampling) into the polyethylene 

4 Initiate pressure in the compression apparatus. Discard the first 5-10 drops. 
Reapply pressure for 15 min and collect remaining solution. 

5 Set aside a portion of the solutions for analysis of pH and EC. Transfer the rest 
of the solution to clean storage bottles. Solutions may be further filtered using 
low vacuum through 0.4 jjim polycarbonate filters before storage and analysis. 

FIGURE 17.5. The compression d 
tesy of Don Ross). 

e for the syringe pressu 


A subsample for metal analysis should be acidified (0.2% HNO3 v/v); trace 
metal-grade acid should be used if trace elements are to be analyzed. 

6 Replicate all samples and include blanks. 


/ In both methods, soil solutions should be separated from the soils as rapidly as 
possible after sampling. Soil samples should be kept cool (4°C in the dark but not 
frozen) before solutions are extracted. The time taken to separate the soil solution 
from the soil solid phase after the disturbance of taking the soil out of its natural 
environment is important in reducing sampling artifacts (Qian and Wolt 1990; Ross 
and Bartlett 1990). 

2 The force of extraction during centrifugation can be calculated as the RCF: 

nr . r (Iirnfr ,„ .,, 

where n is the number of revolutions per second, r the distance from the center of 
rotation in centimeters, and g is 981 cm s~ 2 . The RCF is related to the size of 
pores (assumed to be capillary pores) drained by the centrifugal force. For 
example, pores of 1 jjum diameter are drained at an RCF of roughly 1000 g 
(Edmunds and Bath 1976; Soon and Warren 1993). The force of extraction used 
in the syringe pressure method should also be measured and recorded to ensure 
comparable and consistent results. 

Both methods will produce low volumes of solution (1-3 ml_) and may require 
several replicates bulked together to produce enough solution for a range of 
solution analyses. Bulked solutions should also be replicated to provide a clear 
idea of the reproducibility of the procedure (i.e., if three extracted solutions are 
bulked together to produce a 5-1 ml_ sample; six solutions should be extracted to 
produce a replicate). 

Adams, F. 1974. Soil solutions. In E.W. Carson, oethene) porous cup soil water samplers. Environ. 

Ed. The Plant Root and Its Environment. Univer- Sci. Techno!. 26: 2005-201 1. 

sity Press of Virginia, Charlottesville, VA, 

pp. 44i_482. Belanger, N., Cote B., Fyles, J.W., Courchesne, 

F., and Hendershot, W.H. 2004. Forest regrowth 

Adams, F., Burmester, C, Hue, N.V., and Long, as the controlling factor of soil nutrient availabil- 

F.L. 1980. A comparison of column displacement ity 75 years after fire in a deciduous forest of 

and centrifuge methods for obtaining soil solu- Southern Quebec. Plant Soil 262: 363-372. 
tions. Soil Sci. Soc. Am. J. 44: 733-735. 

Brahy, V. and Delvaux, B. 2001. Comments on 

Beier, C, Hansen, K., Gundersen, P., Andersen, Artifacts caused by collection of soil solution 

B.R., and Rasmussen, L. 1992. Long-term with passive capillary samplers. Soil Sci. Soc. 

field comparison of ceramic and poly(tetrafluor- Am. J. 65: 1571-1572. 


Davies, B.E. and Davies, R.I. 1963. A simple 
centrifugation method for obtaining small sam- 
ples of soil solution. Nature 198: 216-217. 

Dieffenbach, A., Gottlein, A., and Matzner, E. 1997. 
In-situ soil solution chemistry in an acid forest soil 
as influenced by growing roots of Norway spruce 
(Picea abies [L.] Karst.). Plant Soil 192: 57-61. 

Elkhatib, E.A., Hern, J.L., and Staley, T.E. 1987. 
A rapid centrifugation method for obtaining soil 
solution. Soil Sci. Soc. Am. J. 51: 578-583. 

Foster, N.W., Mitchell, M.J., Morrison, I.K., and 
Shepard, J.P. 1992. Cycling of acid and base 
cations in deciduous stands of Huntington Forest, 
New York, and Turkey Lakes, Ontario. Can. 
J. Forest Res. 22: 167-174. 

Gillman, G.P. 1976. A centrifuge method for 
obtaining soil solution. CSIRO Division of Soils, 
Report No. 16, Adelaide, Australia. 

Gottlein, A. and Blasek, R. 1996. Analysis of 
small volumes of soil solution by capillary elec- 
trophoresis. Soil Sci. 161: 705-715. 

Gottlein, A., Hell, U., and Blasek, R. 1996. A 
system for microscale tensiometry and lysimetry. 
Geoderma 69: 147-156. 

Goyne, K.W., Day, R.L., and Chorover, C. 2000. 
Artifacts caused by collection of soil solution 
with passive capillary samplers. Soil Sci. Soc. 
Am. J. 64: 1330-1336. 

Haines, B.L., Waide, J.B., and Todd, R.L. 1982. 
Soil solution nutrient concentrations sampled with 
tension and zero-tension lysimeters: report of 
discrepancies. Soil Sci. Soc. Am. ./. 46: 547-555. 

Heinrichs, H, Bottcher, G., Brumsack, H., 
and Pohlman, M. 1995. Squeezed soil-pore 
solutes — a comparison to lysimeter samples and 
percolation experiments. Water Air Soil Poll. 89: 

Hendershot, W.H., Mendes, L., Lalande, H., 
Courchesne, F., and Savoie, S. 1992. Soil and 
stream water chemistry during spring snowmelt. 
Nord.Hydrol. 23: 13-26. 

Jones, D.L. and Edwards, A.C. 1993. Effect of 
moisture content and preparation technique on the 
composition of soil solution obtained by centrifu- 
gation. Commun. Soil Sci. Plant Anal. 24: 

Edmunds, W.M. and Bath, A.H. 1976. Centrifuge 171-186. 
l and chemical analysis of interstitial 
5. Environ. Sci. Technol. 10: 467^172. 

Kinniburgh, D.G. and Miles, D.L. 1983. Extrac- 
tion and chemical analysis of interstitial water 
from soils and rocks. Environ. Sci. Technol. 17: 

Lawrence, G.B. and David, M.B. 1996. Chemical 
evaluation of soil-solution in acid forest soil. Soil 
Sci. 161: 298-313. 

Ludwig, B., Meiwes, K.J., Khanna, P., Gehlen, R., 
Fortmann, H., and Hildebrand, E.E. 1999. 
Comparison of different laboratory methods 
with lysimetry for soil solution composition — 
experimental and model results. /. Plant Nutr. 
Soil Sci. 162: 343-351. 

MacDonald, J.D., Belanger, N., and Hendershot, 
W.H. 2004a. Column leaching using dry soil 
reproduces solid-solution partitioning observed 
in zero-tension lysimeters. 2. Trace metals. Soil 
Sed. Contam. 13: 361-374. 

MacDonald, J.D., Belanger, N., and Hendershot, 
W.H. 2004b. Column leaching using dry soil 
reproduces solid-solution partitioning observed in 
zero-tension lysimeters. 1. Method Development. 
Soil Sed. Contam. 13: 375-390. 

MacDonald, J.D., Johnson, D., Taillon, K., Hale, B., 
and Hendershot, W.H. 2003. Modeling the effect 
of trace metals emissions on boreal forest soils. 
/. Human Ecol. Risk Assess. 9: 123-141. 

Puschenreiter, M., Wenzel, W.W., Wieshammer, G., 
Fitz, W.J., Wieczorek, S., Kanitsar, K., and 
Kollensperger, G. 2005. Novel micro-suction-cup 
design for sampling soil solution at defined dis- 
tances from roots. ./. Plant Nutr. Soil Sci. 168: 

Hendershot, W.H. and Courchesne, F. 
Comparison of soil solution chemistry i 
tension and ceramic cup tension lysii 
/. Soil Sci. 42: 577-583. 

1991. Qian, P. and Wolt, J.D. 1990. Effects of 
zero drying and time of incubation on the compos- 
leters. jtion of displaced soil solution. Soil Sci. 149: 



Quevauviller, P.H. 1998. Operationally defined Smethurst, PJ. 2000. Soil solution and other soil 

extraction procedures for soil and sediment analyses as indicators of nutrient supply: a review, 

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17: 289-298. 

Soon, Y.K. and Warren, C.J. 1993. Soil solution. 

Reynolds, B. 1984. A simple method for the InM.R. Carter, Ed. Soil Sampling and Methods of 

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365-381. 602-605. 


Chapter 18 

Ion Exchange and 

Exchangeable Cations 

W.H. Hendershot and H. Lalande 

McGill University 
Sainte Anne de Bellevue, Quebec, Canada 

M. Duquette 


Montreal, Quebec, Canada 


Soils possess electrostatic charge as a result of atomic substitution in the lattices of soil 
minerals (permanent charge) and because of hydrolysis reactions on broken edges of the 
lattices and the surfaces of oxides, hydroxides, hydrous oxides, and organic matter (pH- 
dependent charge). These charges attract counterions (exchangeable ions) and form the 
exchange complex. The principle of the methods used to measure exchangeable ions is to 
saturate the exchange complex with some ion that forces the exchangeable ions already 
present on the charged surfaces into solution (law of mass action). Exchange capacity can 
then be calculated as the sum of the individual cations displaced from the soil (summation 
method); or the ion used to saturate the exchange complex, termed the index ion, can be 
displaced with a concentrated solution of a different salt and the exchange capacity calcu- 
lated as the amount of the index ion displaced (displacement method). 

The cation-exchange capacity (CEC) is a measure of the amount of ions that can be 
adsorbed, in an exchangeable fashion, on the negative charge sites of the soil (Bache 
1976). The results are commonly expressed in centimoles of positive charge per kilogram 
of soil (cmol(+) kg -1 ). Anion-exchange capacity (AEC) is expressed in terms of negative 
charge (cmol( — ) kg -1 ). In most Canadian soils, CEC is much greater than AEC; as a result, 
in most routine soil analysis, only CEC and exchangeable cations are measured. 

The measurement of CEC is complicated by (1) errors due to the dissolution of soluble salts, 
CaC03, and gypsum (CaSCU • H2O); (2) specific adsorption of K and NH4 in the interlayer 
position in vermiculites and micas (including illite or hydrous mica); and (3) the specific 
adsorption of trivalent cations such as Al 3+ or Fe 3+ on the surface of soil particles. 

In general, the errors can be reduced by using a method of CEC determination that employs 
reagents of similar concentration and pH to those of the soil to be analyzed. For this reason a 
method buffered at pH 7.0 or 8.2 using relatively high concentrations of saturating and 
extracting solutions will decrease errors due to dissolution of CaCC>3 and gypsum in soils 
from arid regions (Thomas 1982). In acidic soils, solutions buffered at pH 7.0 or 8.2 are less 
effective in replacing trivalent cations and an unbuffered method will provide a better 
estimate of the CEC and exchangeable cations. 

Methods using a solution at a buffered pH are commonly used with agricultural soils 
providing a measurement that is independent of recent fertilization and liming practices. 
For forest soils and other low pH soils, it is often preferable to measure CEC at the pH of the 
soil (see Section 18.2), thus providing a more accurate measure of exchangeable cations and 
CEC under field conditions. 

Soils containing appreciable amounts of amorphous materials (e.g. podzols, some brunisols, 
and soils containing volcanic ash) will show order of magnitude changes in CEC and AEC as 
a result of acidification or liming. The method for measuring pH-dependent CEC and AEC 
(see Section 18.3) is provided for those who wish to study the variation in charge properties 
as a function of pH. The method provides more useful information than does the potentio- 
metric titration method. Although both can be used to give an estimate of the point of zero 
charge (PZC), the pH-dependent CEC and AEC method also provides a measure of the 
absolute amount of exchange capacity at any pH. 


The BaCh method provides a rapid means of determining the exchangeable cations 
and the "effective" CEC of a wide range of soil types. In this method CEC is calculated 
as the sum of exchangeable cations (Ca, Mg, K, Na, Al, Fe, and Mn). The method is 
particularly applicable in forestry or studies of environmental problems related to soils 
where information on the CEC at the pH of the soil in the field is of prime importance. In 
soils with large amounts of pH-dependent cation-exchange sites, the value measured at pH 7 
will be considerably higher than that measured by this method. Problems may arise if this 
method is used with saline soils containing very high levels of SO4 since BaSC>4 will 

This method has been compared to other methods of determining the CEC at the soil pH and 
provides comparable results (Hendershot and Duquette 1986; Ngewoh et al. 1989). Barium is 
a good flocculant and is able to displace trivalent cations. The relatively low ionic strength of 
the equilibrating solution causes a smaller change in pH than do more concentrated salt 
solutions. This method is simple and rapid; however, it is recommended that exchangeable 
iron and manganese be measured since they may be more abundant in some acidic soils than 
other commonly considered cations such as potassium and sodium. 

The Expert Panel on Soil (2003) proposes an alternative method that involves three succes- 
sive additions of 0.1 M BaC^. The soihsolution ratio of 1:60 and the successive shaking 
and decanting steps result in higher measured values of exchangeable cation. However, the 
more complicated procedure is less suitable for routine laboratory analysis. Since the method 
proposed in this chapter for measuring exchangeable cations uses 0.1 M BaC^, it seems 
more appropriate to use the same salt solution for measuring exchangeable acidity. 


Jonsson et al. (2002) have determined regression equations that could be used t 
the difference between the two extraction procedures. 

The results of this method are dependent on the soil:solution ratio used, with higher values of 
exchangeable cations obtained with smaller amounts of soil. The suggested weights of soil 
are a reasonable compromise. We have decreased the maximum amount of soil to be used 
from 3.0 to 1.5 g compared to the previously published methodology (Hendershot et al. 
1993). If results are to be compared over time, or between sites, it is important that standard 
weights of sample be used. 

18.2.1 Materials and Reagents 

7 Centrifuge tubes (50 ml_) with screw caps and low-speed centrifuge. 

3 Barium chloride, 0.1 M: dissolve 24.43 g of BaCI 2 -2H 2 with double deionized 
(d.d.) water and make to volume in a 1 L volumetric flask. 

4 Standards of Ca, Mg, K, Na, Al, Fe, and Mn are prepared using atomic absorption 
reagent-grade liquid standards of 1000 mg L~ 1 . The matrix in the standards must 
correspond to the BaCI 2 concentration of the analyzed sample (diluted or non- 
diluted matrix). 

5 Lanthanum solution, 100 mg L : dissolve 53.5 g of LaCl3-7H 2 in a 200 ml_ 
volumetric flask and make to volume (for analysis by atomic absorption spectro- 
photometry [AAS]). 

6 Cesium solution, 100 g L~ 1 : dissolve 25.2 g CsCI in a 200 ml_ volumetric flask 
and make to volume (for analysis by AAS). 

18.2.2 Procedure 

1 Weigh out about 0.5 g of air-dry (<2 mm) organic soil or fine-textured soil to 1 .5 g 
of coarse-textured soil into a 50 ml_ centrifuge tube. Record the exact weight of 
soil used to the nearest 0.001 g. Include blanks, duplicates, and quality control 

2 Add 30.0 ml_ of 0.1 M BaCI 2 to each tube and shake slowly on an end-over-end 
shaker (15 rpm) for 2 h. 

3 Centrifuge (1 5 min, 700 g) and filter the supernatant (SN) with Whatman No. 41 
filter paper. 

4 Analyze the following cations in the SN solution with an AAS or any other 
suitable instrument: Ca, Mg, K, Na, Al, Fe, and Mn. Dilution (10- or 100-fold) is 
usually required for Ca, K, and Mg. The addition of 0.1 ml_ of La solution 
and 0.1 mL of Cs solution to a 10 mL aliquot of diluted extract is required for 
the determination of Ca, Mg, and K by AAS. (For detailed instructions on this 
and other aspects of analysis refer to the manual for your AAS.) Preservation 


of samples by acidifying to 0.2% HNO3 will prevent the loss of metals, such as 
Fe and Al. 

5 If desired, the pH of the equilibrating solution can be measured on a separate 
aliquot of the BaCI 2 solution before filtering. Leakage of K from the KCI salt bridge 
of the pH electrode is significant and therefore the same aliquot cannot be used 
for K analysis and pH measurement. 

18.2.3 Calculations 

7 Exchangeable cations 

M+ cmol(+) kg -1 = C cmol(+) L~ 1 x (0.03 L/wt.soil g) x 1000 g kg~ 1 x DF 


where M + is the concentration of an adsorbed cation, cmol(+) kg~ 1 , C is the 
concentration of the same cation measured in the BaCb extract (cmol(+) L~ 1 ), and 
DF is the dilution factor, if applicable. 

2 Effective CEC 

Effective CEC cmol(+) kg" 1 = SM+cmol(+) kg" 1 (1 8.2) 

3 Percent base saturation 

% BS = (SCa + Mg + Na + K/Effective CEC) x 1 00 (1 8.3) 


7 A large amount of a soil similar to the samples being analyzed should be kept as an 
indicator of the variability of results over time; duplicate subsamples of this quality 
control (QC) sample should be run with each batch of samples measured. Failure of 
the QC to fall within acceptable limits means that the whole batch should be 
reanalyzed. Analysis of QC samples is also useful to verify that samples analyzed 
by different people in the same laboratory are comparable, and that results do not 
change from one year to another or from one batch of chemicals to another. 

2 For the sake of simplicity AAS standards are usually made up by diluting 
1000 mg L~ 1 concentrate to lower concentration values suitable for the range of 
the instrument being used. Calibrate the machine using the corresponding 
cmol(+) L~ 1 value; the conversion values are as follows: 



In the literature, the method of Fey and LeRoux (1976) is often cited in research on pH- 
dependent CEC and AEC. The method is time-consuming because of the multiple saturation 
and pH adjustment steps. An alternative is to add different amounts of acid or base to the 
soil suspensions and measure the resulting pH. This method is preferred because there are 
fewer steps, and therefore it is faster with less chance of errors due to contamination or loss of 
soil. The only disadvantage with the modified procedure is that it is more difficult to obtain an 
even distribution of pH values than with the method of Fey and LeRoux, but this can be 
corrected by rerunning the analysis and adjusting the amounts of HNO3 or Ca(OH) 2 added. 

18.3.1 Materials and Reagents 

1 Centrifuge tubes (50 ml_) with screw caps and low-speed centrifuge. 

2 Vortex centrifuge tube mixer and end-over-end shaker. 

3 Calcium nitrate, 0.05 Mr. dissolve 23.62 g of calcium nitrate tetrahydrate 
(Ca(N0 3 ) 2 -4H 2 0) with d.d. water in a 2 L volumetric flask. 

4 Nitric acid, 0.1 M: dilute 6.3 ml_ of concentrated nitric acid (HNO3) with d.d. 
water in a 1 L volumetric flask. 

5 Calcium hydroxide, 0.05 M: dissolve 3.70 g of calcium hydroxide (Ca(OH) 2 ) 
with d.d. water in a 1 L volumetric flask, and filter through a Whatman No. 41 filter 
(a prefiltration step can be done using a glass microfiber filter [Whatman GF/C]). 

6 Calcium nitrate, 0.005 M. dilute 200 ml_ of 0.05 M Ca(N0 3 ) 2 solution with d.d. 
water in a 2 L volumetric flask. 

j Potassium chloride, 1 .0 M: dissolve 149.12 g of potassium chloride (KCI) with d.d. 
water in a 2 L volumetric flask. 

g Lanthanum solution, 100 mg L~ 1 : dissolve 53.5 g of LaCl3-7H 2 in a 200 ml_ 
volumetric flask and make to volume (for analysis by AAS). 

9 Cesium solution, 100 g L~ 1 : dissolve 25.2 g CsCI in a 200 ml_ volumetric flask 
and make to volume (for analysis by AAS). 

18.3.2 Procedure 

1 Weigh 20 empty 50 mL centrifuge tubes to the nearest 0.001 g (one set of 20 tubes 
for each soil sample to be analyzed). 

2 Add 1 .0 g subsamples of air-dry <2 mm soil to each tube and record the weight 
of the tube plus soil to the nearest 0.001 g. The analysis is done in duplicate for 
each targeted pH and corresponds to one pair of quality control samples per 
batch. If moist soil is used, start by weighing out four additional samples into 
small beakers and air-dry to determine the weight of moist soil equivalent to 1 gof 
air-dried soil. 


j Add 25 ml_ 0.05 M Ca(NC>3)2 solution, cap the tubes, and shake for 1 h using an 
end-over-end shaker (15 rpm). 

4 Centrifuge (1 min, 700 g) and discard SN by decantation. Be careful to avoid loss 
of soil during decantation. 

5 Add a new 25 ml_ aliquot of 0.05 M Ca(NC>3)2 solution to each tube. Then 
add 0, 0.25, 0.5, 1.0, or 2.5 ml_ of 0.1 M HN0 3 to tubes in duplicate, and 
finally add 0.25, 0.5, 1 .0, or 2.5 ml_ of 0.05 M Ca(OH) 2 to the remaining tubes in 
duplicate. Add 1 .0 ml_ of 0.1 M HN0 3 or 0.05 M Ca(OH) 2 to the quality control 
sample. A vortex mixer is useful to resuspend the soil after addition of the solution. 

6 Cap and shake overnight on an end-over-end shaker. 

7 Centrifuge (10 min, 700 g) and discard SN. 

8 Resuspend the soil in 25 ml_ of 0.005 M Ca(N0 3 ) 2 , centrifuge (10 min, 700 g), 
and discard SN. 

9 Repeat step 8, but measure pH in a separate aliquot of the SN and keep the 
remaining SN for the analysis of Ca and N0 3 (after 100-fold dilution with d.d. 
water). Weigh tubes plus the soil and the interstitial soil solution. 

/0 Add 25 ml_ of 1 .0 M KCI, shake for 1 h, and centrifuge (1 min, 700 g). 

7 7 Keep this SN for determination of displaced Ca and N0 3 . Dilute this KCI extract 
10-fold with d.d. water. 

72 Measure Ca by AAS in the 1 0-fold diluted KCI extract (saved in step 1 1 ) and in the 
0.005 M Ca(NO B ) 2 equilibration solution (saved in step 9). The addition of 0.1 ml_ 
of La solution and 0.1 mL of Cs solution to a 10 mL aliquot of diluted extract is 
required for the determination of Ca by AAS. (For detailed instructions on this and 
other aspects of analysis refer to the AAS manual.) 

73 Measure N0 3 in the undiluted KCI extract (saved in step 11) and in the diluted 
0.005 M Ca(NO B ) 2 equilibration solution (saved in step 9). 

18.3.3 Calculations 

7 Residual Ca and N0 3 

a. Volume of interstitial solution 

Subtract the weight of the empty tube with the soil (step 2) from weight 
measured in step 9 to calculate weight of residual 0.005 M Ca(N0 3 ) 2 solution 
(Vol res ). Assume 1 g equals 1 mL. 

b. Residual amount of Ca and NO B (Ca res and N0 3 res ): 

Ca res (mol) = Vol res (mL) x Ca so , (mM) x 0.001 (L mL" 1 ) x DF (18.4) 


NO B res (mol) = Vol res (itiL) x N0 3 so | (itiM) x 0.001 (L mL~ 1 ) x DF (1 8.5) 

where Ca so | and N03 S0 | are the measured concentrations of calcium and 
nitrate in the 0.005 M Ca(N0 3 ) 2 wash solution saved in step 9 (units in mM) 
and DF is the dilution factor if applicable. 

Total amount of calcium and nitrate (Ca t N0 3 1 ) in the KCI extract (including the 

Ca t (mmol) = Ca KC i (mM) x 25 (mL) x 0.001 (L mL" 1 ) x DF 

N0 3 1 (mmol) = N0 3 kg (mM) x 25 (mL) x 0.001 (L mL" 1 ) x DF 

where Ca K ci and NO3 K ci are the calcium and nitrate concentrations (mM ) in the 
KCI extract saved in step 1 1; and DF is the dilution factor if applicable. 

Calculation of the CEC and AEC: 

CEC cmol(+) kg~ 1 = (Ca t - Ca res ) (mmol) x 0.2 (cmol(+) mmor 1 ) 

x1000(gkg- 1 )/wt.soil (g) (18.7) 

AEC cmol(-) kg" 1 = (NO3 1 - NO3 res) (mmol) x 0.1 (cmol(-) mmol" 1 ) 

x1000 (g kg^/wtsoil (g) (18.8) 

Plot CEC and AEC as a function of final equilibrium pH measure in step 9 of 
Section 18.3.2. 



AT pH 7.0 (LAVKULICH 1981) 

The method described here was developed by Lavkulich (1981) for standard analysis of a 
wide range of soil types. It involves fewer steps than some other similar methods such as that 
of McKeague (1978). Problems with this approach to measuring exchangeable cations and 
CEC have been discussed extensively in the literature (Chapman 1965; Bache 1976; Rhoades 
1982; Thomas 1982) but we agree with the conclusion of Thomas (1982) that "there is no 
evidence at the present time that cations other than NH4+ give results that are less arbitrary 
than those obtained using NH4+." 

Errors due to the dissolution of CaC0 3 and gypsum will result in an excess of Ca 2+ being 
extracted by NH4+ and a decrease in the amount of NH4+ retained due to competition 
between Ca 2+ and NH4+ during equilibration in the saturating step. In soils containing these 
minerals, exchangeable Ca will be too high and total CEC too low. The former problem can 
not easily be corrected (Thomas 1982); however, more accurate measurement of CEC in 
this type of soil can be obtained by using the method described by Rhoades (1982). 

Fixation of K + and NFi4 + in phyllosilicates can result in either an over- or underestimation 
of exchangeable K + when NH4 4 " is used as an extractant depending on whether the NH4+ 
moves through the interlayer positions replacing the K + or whether it causes the collapse of 
the edges preventing further exchange. 


Compared to the other methods presented in this chapter, this method uses a larger sample 
size, which helps to decrease the sample to sample variability. Another advantage of this 
procedure is that there are no decantation steps that can cause the loss of sample, particularly 
in the case of organic soils. 

The method described below can be used to measure either exchangeable cations and CEC or 
just exchangeable cations. In the latter case, the sum of exchangeable cations (including Al) 
could be used as an estimate of CEC. Due to the high pH of the extracting solution, the 
amount of Al measured will usually be lower than that displaced by BaCl2 or KC1. 

18.4.1 Materials and Reagents 

7 Centrifuge tubes: 100 ml_ centrifuge tubes and stoppers. 

2 Reciprocal shaker. 

j Buchner funnels (55 mm diameter) and 500 mL filtering flasks connected to low- 
pressure vacuum line. 

4 Ammonium acetate, 1 M: dissolve 77.08 g of NH 4 OAc with d.d. water and make 
to volume in a 1 L volumetric flask. Adjust pH to 7.0 with ammonium hydroxide 
or acetic acid. 

5 Isopropanol. 

6 Potassium chloride, 1 M: dissolve 74.6 g of KCI with d.d. water and make to 
volume in a 1 L volumetric flask. 

7 Standard ammonium solution, 200 mg L" 1 N: dissolve 0.238 g of (NH 4 ) 2 S0 4 
(dried for 3-4 h at 40°C) in about 1 00 mL of d.d. water and then dilute to volume 
in a 250 mL volumetric flask. Prepare diluted standards of 10, 20, 40, and 
80 mg L~ 1 from the 200 mg L~ 1 stock. 

g Prepare Ca, Mg, K, and Na standards using 1 M NH 4 OAc as the matrix. 

18.4.2 Procedures 

For Exchangeable Cations 

/ a. For samples low in organic matter: weigh out 10.000 g of soil into a 100 mL 
centrifuge tube. 

b. For samples high in organic matter: weigh out 5.000 or 2.000 g. 

c. Prepare a blank and include a quality control sample. 

2 Add 40 mL of 1 M NH 4 OAc to the centrifuge tube. Stopper the tube and shake for 
5 min on a reciprocal shaker (115 rpm). Remove tubes from shaker, agitate to 
rinse down soil adhering to the sides of the tube, and let stand overnight. 


3 Shake tube again for 1 5 min. Prepare Buchner funnels with Whatman No. 42 filter 
paper and place them above 500 ml_ filtering flasks. 

4 Transfer contents of the tube to the funnel with suction applied. Rinse the tube and 
the stopper with 1 M NH 4 OAc from a wash bottle. 

5 Wash the soil in the Buchner funnel with four 30 ml_ portions of 1 M NH 4 OAc. 
Let each portion drain completely before adding the next, but do not allow the soil 
to become dry or cracked. 

g Transfer the leachate to a 250 ml_ volumetric flask; rinse the filtering flask with 
1 M NH 4 OAc and make up to volume with 1 M NH 4 OAc. Mix well and save a 
portion of the extract for analysis of Al, Ca, Mg, K, and Na. Keep samples 
refrigerated prior to analysis. 

For Total-Exchange Capacity (CEC) 

7 Replace the funnels containing the ammonium-saturated soil onto the filtering 
flasks. To remove the residual NH 4 OAc from the soil, wash the soil in the Buchner 
funnel with three 40 ml_ portions of isopropanol, again letting each portion drain 
completely before adding the next (turn off the suction after the last washing 
before the soil dries out). Discard the isopropanol washings and rinse the flask 
well with tap water followed by d.d. water. 

2 Replace the funnels onto the flasks and leach the soil with four 50 ml_ portions 
of 1 M KCI, again letting each portion drain completely before adding the 
next. Transfer the leachate to a 250 ml_ volumetric flask. Rinse the filtering 
flask into the volumetric flask with d.d. water and make up to volume with d.d. 
water. Mix well and save a portion of the extract for analysis of NH 4 by auto 

18.4.3 Calculations 

7 Exchangeable cations: 

M+ cmol(+) kg" 1 = C cmol(+) L~ 1 x (0.25 L/wt soil g) x 1000 g kg" 1 (18.9) 

where M + is the concentration of adsorbed cation, cmol(+) kg~ 1 ; and C is the 
concentration of cation in the NH 4 OAc extract (cmol(+) L~ 1 ). 

Note: see Section 18.2.4 for conversion of mg L~ 1 to cmol(+) L" 1 . 

2 CEC: 

CEC cmol(+) kg" 1 = (mg L~ 1 N x (1 cmol(+)/140 mg)) 

x (0.25 L/wt.soil g) x 1000 g kg~ 1 (18.10) 


Bache, B.W. 1976. The n 

exchange capacity of soils. J. Sci. Food Agric. 27: 


Chapman, H.D. 1965. Cation exchange capacity. 
In C.A. Black et al., Eds., Methods of Soil Analy- 
sis. Agronomy 9, American Society of Agron- 
omy, Madison, WI, pp. 891-901. 

Expert Panel on Soil. 2003. Manual on Methods 
and Criteria for I larmonizcd Sampling, Assessment, 
Monitoring and 1; \e t Kir Pa, 

Pa lit i >ling and Analysis 

of Soil. International co-operative programme on 
assessment and monitoring of air pollution effects 
on forests, ( 
pdf, verified February 9, 2005) 

Fey, M.V. and LeRoux, J. 1976. Electric charges 
on sesquioxidic soil clays. Soil Sci. Soc. Am. J. 40: 

Hendershot, W.H. and Duquette, M. 1986. A sim- 
ple barium chloride method for determining cat- 
ion exchange capacity and exchangeable cations. 
Soil Sci. Soc. Am. J. 50: 605-608. 

Hendershot, W.H., Lalande, H., and Duquette, M. 
1993. Ion exchange and exchangeable cations. In 
M.R. Carter, Ed., Soil Sampling and Methods of 
Analysis. Lewis Publishers, Boca Raton, FL, 
pp. 167-175. 

Jonsson, U., Rosengren, U., Nihlgard, B., and 
Thelin, G. 2002. A comparative study of two 
methods for determination of pH, exchangeable 
base cations and aluminium. Commun. Soil Sci. 
Plant Anal. 33: 3809-3824. 

Lavkulich, L.M. 1981. Methods Manual, Ped- 
ology Laboratory. Department of Soil Science, 
University of British Columbia, Vancouver, British 
Columbia, Canada. 

McKeague, JA. 1978. Manual on Soil Sampling 
and Methods of Analysis, 2nd ed. Canadian 
Society of Soil Science, AAFC, Ottawa, Ontario, 

Ngewoh, Z.S., Taylor, R.W., and Shuford, J.W. 
1989. Exchangeable cations and CEC determin- 
ations of some highly weathered soils. Commun. 
Soil Sci. Plant Anal. 20: 1833-1855. 

Rhoades, J.D. 1982. Cation exchange capacity. In 
A.L. Page et al., Eds., Methods of Soil Analysis. 
Agronomy 9, 2nd ed. American Society of 
Agronomy, Madison, WI, pp. 149-157. 

Thomas, G.W. 1982. Exchangeable cations. In 
A.L. Page et al., Eds., Methods of Soil Analysis. 
Agronomy 9, 2nd ed. American Society of 
Agronomy, Madison, WI, pp. 159-165. 


Chapter 19 
Nonexchangeable Ammonium 

Y.K. Soon 

Ai;rii allure jnd Agri-rood Canada 
Beaverlodge, Alberta, Canada 

B.C. Liang 

Environment Canada 
Gatineau, Quebec, Canada 


It has been known since the early part of the twentieth century that some types of soils have 
the ability to bind ammonium (to certain types of clay minerals, predominantly vermiculite 
and mica types) such that it is not readily recovered by extraction with dilute acid or alkali 
(McBeth 1917). This form of ammonium is referred to as fixed or nonexchangeable 
ammonium (NEA). Barshad (1951) proposed that fixed ammonium should be defined as 
ammonium that is not displaceable with prolonged extraction or leaching of soil with 
potassium salt solution. The proportion of soil N as NEA usually does not exceed 10% in 
surface soils, but it can increase with depth of soil to over 50% in some subsoil horizons 
(Hinman 1964; Bremner 1965). Sources of NEA in the soil include (i) NH 4 + produced by 
mineralization of organic matter, and added through ammoniacal-N fertilizer material, and 
(ii) indigenous or native fixed ammonium found in parent rock materials. There is consid- 
erable interest in quantifying the NEA pool because the amount in the soil through the 
rooting depth can be considerable, and its availability to plants and microorganisms has been 
demonstrated in many studies (Kudeyarov 1981; Scherer 1993; Green et al. 1994; Scherer 
and Werner 1996; Soon 1998). Soderland and Svensson (1976) estimated that there is as 
much fixed NH 4 + -N as there is plant biomass N in the global soil-plant system. The NEA 
pool in the soil has been found to be a slow-release reservoir of available ammonium when 
the exchangeable NH 4 + levels become depleted (Drury and Beauchamp 1991). Ammonium 
fixation and release must be characterized and quantified especially in soils with a high 
ammonium fixation capacity (i.e., soils with a high vermiculite or mica content) in order to 
efficiently manage N use in soils for agronomic and environmental reasons. 

Several procedures have been developed for the determination of NEA (Young and Aldag 
1982); however, the most widely accepted method is that of Silva and Bremner (1966). 
Bremner et al. (1967) evaluated several methods and found that all except the Silva and 
Bremner method have defects: (i) the pretreatments used to eliminate interference by organic 


N compounds were either inefficient, or led to gain or loss of NEA, and (ii) the procedures 
used to release NEA were not quantitative, or led to formation of NH4-N from organic N 
compounds. According to Keeney and Nelson (1982), the Silva-Bremner method enjoys 
widespread use because of its apparent lack of defects; however, they cautioned that "there 
is no way of establishing that the KOBr-HF method is accurate . . .". Bremner et al. (1967) 
mentioned two possible problems associated with the determination of NEA: intercalated 
organic materials containing N that are released by the HF treatment and the presence of 
metal ammonium phosphates that are not soluble in KOBr or KC1 but soluble in HF. 
Although either one will result in an overestimation of NEA, under normal conditions the 
contribution of either one would be remote or very slight. The Silva-Bremner procedure 
involves and comprises three basic steps: (i) removal of exchangeable NH 4 + cations, 
(ii) oxidation and removal of organic matter including organic N, and (iii) extraction of 
NEA with HF and HC1, and determination of the released NH 4 + . A slightly and a substan- 
tially modified version of the method will be described below. 

Zhang and Scherer (1998) proposed a simplified version (method A) of the Silva-Bremner 
method, which reduced the time involved and the amount of reagents used. A more 
substantial modification that eliminated entirely the HF extraction step (method B) was 
proposed by Nieder et al. (1996) and Liang et al. (1999): here, NEA in the soil residue left 
from the KOBr and KC1 extractions is determined directly by dry combustion in an 
automated N-analyzer. This is a major advancement for the procedure because it eliminates 
the hazardous HF extraction step and the subsequent disposal of the HF, saving time in the 
process by reducing the number of steps in the procedure. Nitrogen isotope ratios can also be 
very conveniently determined when the N-analyzer is connected by a continuous flow 
linkage to a 15 N/ 14 N isotope ratio mass spectrometer. 


The procedure described below is an adaptation of the Silva and Bremner (1966) method by 
Zhang and Scherer (1998). In this variation of the method, organic matter in the sample is 
oxidized in a centrifuge tube immersed in a boiling water bath (instead of a beaker heated 
with a hot plate) and subsequent extraction steps are carried out without having to transfer 
the residual soil to a centrifuge tube. Zhang and Scherer (1998) also found that heating in a 
microwave oven (1150 watt) at 50% of full power for 10 min gave similar NEA values to 
heating in a boiling water bath. Use of microwave ovens of different power would likely 
require adjustments by trial and error. The method using a boiling water bath is described. 

19.2.1 Materials and Reagents 

7 50 ml_ polypropylene or polyethylene centrifuge tubes with lined screw caps. 

2 Potassium hydroxide (KOH) solution, 2 M: Dissolve 1 12.2 g of KOH in approxi- 
mately 600 ml_ of distilled deionized water and, after cooling, dilute to 1 L volume. 

3 Potassium hypobromite (KOBr) solution, prepared immediately before use: Add 
6 ml_ of Br to 200 ml of 2 M KOH solution. Add the Br slowly (approximately 
0.5 ml_ min -1 ) with constant stirring, keeping the KOH solution cool in an 
ice-bath during the addition. 


4 0.5 M Potassium chloride (KCI) solution: Dissolve 149 g of KCI in 600 ml_ of 
deionized water and make up to 4 L. 

5 Hydrofluoric acid-hydrochloric acid solution (approximately 5 M HF-1 M HCI): 
With a 1 L measuring cylinder, transfer 1 .5 L of deionized water to a 2 L graduated 
polypropylene or polyethylene conical flask. Add slowly, with continuous stirring, 
1 67 ml_ of cone. HCI (specific gravity 1 .1 9) followed by 325 ml_ of approximately 
52% HF (approximately 31 M). Dilute with deionized water up to the 2 L mark 
and mix well. 

g Boiling water bath. 

j Reciprocal shaker. 

19.2.2 Extraction Procedure 

1 Weigh 0.5 g of finely ground soil (<60 mesh) in a 50 ml_ centrifuge tube. 
Record the weight of tube and soil. Add 1 ml_ of KOBr solution, and screw the 
cap on. Invert the centrifuge tube several times to mix up the contents, loosen 
screw cap, and leave it standing for 2 h. In the mean time, heat up the 
water bath. 

2 Place the centrifuge tubes in a rack and then immerse the rack in a boiling water- 
bath so that the water level in the water bath exceeds the level of the KOBr 
solution in the centrifuge tube. Once the solution in the centrifuge tubes starts to 
boil, allow it to continue boiling for 1 min. 

3 Remove the tubes and allow the contents to cool and settle. If necessary, centri- 
fuge at 1 000 g for 5 min. 

4 Decant and discard the clear supernatant solution. 

5 Add 30 ml_ of 0.5 M KCI, suspend the soil by shaking for 5 min, and centrifuge at 
1000 gfor 5 min. Decant the clear supernatant solution. 

5 Repeat step 5 two more times. 

7 Weigh the centrifuge tube and soil. The increase over the initial weight (in step 1) 
is taken to represent the volume of KCI retained by the soil. This liquid volume has 
to be added to the acid reagent volume added in step 8 when calculating mg kg~' 

g Add 10 ml_ of 5 M HF-1 M HCI working solution and shake for 24 h on a 
reciprocal shaker at 120 cycles min -1 . If the sample contains carbonates, allow 
the evolved C0 2 to escape before starting the overnight shaking. 

9 Centrifuge at 1000 gfor 5 min. Decant the clear supernatant solution into a plastic 
vial for subsequent NH 4 -N determination. 


19.2.3 Determination of Extracted NH 4 -N 

The original Silva-Bremner method determines nonexchangeable NH4-N in the acid extrac- 
tant by steam distillation and subsequent titrimetry. However, colorimetric determination 
using the development of indophenol has been used by Doram and Evans (1983) and Soon 
(1998). A manual procedure is outlined below, which is easily adaptable for automated 
analysis. The autoanalyzer method outlined in Chapter 6 for exchangeable NH4-N determin- 
ation or the procedure described by Kempers and Zweers (1986) can be readily adapted for 
analysis of NEA. 


Unless specified otherwise, all reagents used must be of analytical grade. 

1 Trisodium citrate solution: Dissolve 20.0 g of Na 3 C 6 H 8 7 -2H 2 in 700 ml_ 
of deionized water. Dissolve 10.0 g of NaOH in deionized water and dilute to 
700 mL. Combine the citrate and NaOH solution (reagent A). 

2 Salicylate-nitroprusside reagent: Dissolve 18.0 g of sodium salicylate 
(HOC 6 H 4 C02Na, 2-hydroxybenzoic acid, sodium salt) in 250 mL of water. 
Dissolve 0.20 g of sodium nitroprusside (Na 2 Fe(CN) 5 NO • 2H 2 0) in 250 mL of 
water. Combine the two reagents and store in a brown bottle (reagent B). 

j Alkaline hypochlorite solution: Dissolve 1 .5 g of NaOH in 50 mL of deionized 
water, add 8 mL of sodium hypochlorite (5%-5.25% NaOCI), and dilute to 100 
mL (reagent C). Prepare fresh as needed. 

4 Ammonium standard solution: 0, 5, 10, 15, 20, and 25 |xg NH 4 -N mL -1 in 5 M 
HF-1 M HCI prepared by dilution of 1000 mg N L~ 1 stock solution. 

Pipette 0.2 mL of the HF-HCI soil extract or ammonium standard solutions 
(containing up to 5 |xg NH 4 -N) into a 1 6 mm x 1 25 mm culture tube. 

Add 7 mL of reagent A and mix immediately. 

Add 2 mL of reagent B and mix immediately. 

Add 0.5 mL of reagent C and mix immediately. 

Immediately cover with a dark colored plastic sheet and leave for 60 min for color 
to develop. 

Measure absorbance of standard and test solutions at 660 nm using 1 cm cuvette. 

The concentrations of NH 4 -N in the test solutions are read off the calibration 
curve, either manually or by the processor in the spectrophotometer. 

g Results can be calculated as follows: 

mg NEA kg T soil = (jug NH 4 -N per ml_ extract x (10 + increase 
in weight in step 7 of Section 19.2.2) 
x F/weight of soil (19.1) 

where F is the dilution factor if dilution of the extract is required, and weight is 
measured in grams. 

7 A final solution pH of about 13 will result in maximum color development. The 
advantages of salicylate as a substitute for phenol are increased sensitivity, lower 
toxicity, and increased stability (Kempers and Zweers 1986). Sodium citrate was 
found to be a better complexing agent for removing interfering elements than 
either EDTA or potassium sodium tartrate (Willis et al. 1993). 

2 If the automated procedure is to be used, the following steps should be taken 
to minimize the slow corrosion of glass elements of the analytical cartridge. 
An analytical cartridge with dialyzer is used to further dilute the fluoride concent- 
ration in the test solution. The wash solution used need not contain hydrofluoric 
acid, and this does not influence the baseline: its acidity is maintained using 6 M HCI. 


This major modification of the Silva and Bremner (1966) method was proposed by Nieder 
et al. (1996). However, the procedure gained greater prominence only after more extensive 
testing and validation by Liang et al. (1999). The method follows the Silva-Bremner method 
from the oxidation and removal of organic materials through the removal of exchangeable 
NH 4 + cations. It is assumed that any NH 4 + not removed from the soil at this stage would 
be nonexchangeable or fixed. This N fraction is then determined by dry (Dumas) combustion 
of the sample using an automated N-analyzer. Liang et al. (1999) showed that dry combus- 
tion recovered 100% of fixed NH 4 + and gave results similar to those obtained using the 
full Silva-Bremner method. The coefficient of variation for 17 soils was 6.4% for the full 
Silva-Bremner method and 2.0% for the modified version. 

19.3.1 Materials and Reagents 

7 50 ml_ polypropylene or polyethylene centrifuge tubes with lined screw caps. 

2 Potassium hydroxide (KOH) solution, 2 M: Dissolve 1 12.2 g of KOH in approxi- 
mately 600 ml_ of distilled deionized water and, after cooling, dilute to 1 L volume. 

3 Potassium hypobromite (KOBr) solution, prepared immediately before use: Add 
6 mL of Br to 200 mL of 2 M KOH solution. Add the Br slowly (approximately 
0.5 mL min~ 1 ) with constant stirring, keeping the KOH cool in an ice-bath during 
the addition. 


4 0.5 M Potassium chloride (KCI) solution: Dissolve 149 g KCI in 600 ml_ of 
deionized water and make up to 4 L. 

5 Boiling water bath. 
5 Reciprocal shaker. 

19.3.2 Procedure 

1 Weigh 0.5 g of finely ground soil (100 mesh) into a centrifuge tube. Record the 
weight of tube and soil. Add 1 ml_ of KOBr solution and screw the cap on. Invert 
centrifuge tube several times to mix up contents, loosen screw cap, and allow to 
stand for 2 h. In the mean time, heat the water bath. 

2 Place the centrifuge tubes in a rack and then immerse the rack in a boiling water 
bath so that the water level in the water bath exceeds the level of the KOBr 
solution in the centrifuge tube. Once the solution in the centrifuge tubes starts to 
boil, allow it to continue boiling for 1 min. 

3 Remove the tubes and allow the contents to cool and settle. Centrifuge at 1 000 g 
for 5 min if necessary. 

4 Decant and discard the clear supernatant solution. 

5 Add 30 ml_ of 0.5 M KCI, suspend the soil by shaking for 5 min, centrifuge at 
1 000 g for 5 min. Decant the clear supernatant solution. 

g Repeat step 5 two more times. 

y Dry residue overnight in a drying oven at 105°C or freeze-dry the residue. 

g Weigh dry residue. (Let this weight be Y g.) 

19.3.3 Determination of NEA in the Residue 

Follow the procedure for the elemental N-analyzer that is to be used. Samples (normally 
50-100 mg) are weighed in tin foil sample cups which are loaded into autosamplers. 
Combustion of the samples with oxygen at 1030°C converts NEA to N2 and NO., gases 
(with other combustion products). These gases are routed to a reduction furnace containing 
heated Cu, which removes excess oxygen and converts NO A to N2, which is separated by gas 
chromatography and the concentration of N2 is measured using a thermal conductivity 
detector (for more on dry combustion for N determination see Chapter 22). The analyzer 
is calibrated with certified standards. 

Since the determination is done on a sample that is free of organic matter, a c 
needed to convert the analytical result to a whole soil basis. Suppose that (i) the ii 
determines NEA to be Z% of residue, and (ii) the weight of the dry residue (in grams) as 
determined in step 8 is Y. Multiply Z% by 10 to convert percent to milligram per gram or 
gram per kilogram basis. Since the weight of the original whole soil samples is 0.5 g, the 
corrected value of NEA in the original soil (in milligram per kilogram) = Z x 10 x 7/0.5. 



The method has several advantages of the original Silva and Bremner (1966) method. The 
use of an automated N-analyzer for the determination of NEA in the soil residue 
(i) eliminates the use and handling of hydrofluoric acid, (ii) increases the precision of the 
method, and (iii) simplifies and simultaneously allows the determination of N isotopic ratios 
by linking an isotopic ratio mass spectrometer to the elemental analyzer. It also shortens the 
time required for analysis. 

Barshad, I. 1951. Cation exchange in soils: I. Kempers, A.J. and Zweers, A. 1986. Ammonium 

Ammonium fixation and its relation to potassium determination in soil extracts by the salicylate 

fixation and to determination of ammonium method. Commun. Soil Sci. Plant Anal. 17: 

exchange capacity. Soil Sci. 72: 361-371. 715-723. 

Bremner, J.M. 1965. Inorganic forms of nitrogen. 
In C.A. Black et al., Eds., Methods of Soil Analy- 
sis, Part 2 — Chemical and Microbiological 
Properties. American Society of Agronomy, 
Madison, WI, pp. 1179-1238. 

Bremner, J.M., Nelson, D.W., and Silva, J.A. 
1967. Comparison and evaluation of methods of 
determining fixed ammonium in soils. Soil Sci. 
Soc. Am. Proc. 31:466-472. 

Doram, D.R. and Evans, L.J. 1983. Native fixed 
ammonium and fixation of added ammonium in 
relation to clay mineralogy in some Ontario soils. 
Can. J. Soil Sci. 63: 631-639. 

Drury, C.F. and Beauchamp, E.G. 1991. Ammo- 
nium fixation, release, nitrification and immobil- 
ization in high and low fixing soils. Soil Sci. Soc. 
Am. J. 55: 125-129. 

Green, C.J., Blackmer, A.M., and Yang, N.C. 
1994. Release of fixed ammonium during 
nitrification in soils. Soil Sci. Soc. Am. J. 58: 

Hinman, W.C. 1964. Fixed a 

Saskatchewan soils. Can. J. Soil Sci. 44: 151-157. 

Keeney, D.R. and Nelson, D.W. 1982. Nitro- 
gen — inorganic forms. In A.L. Page et al., Eds., 
Methods of Soil Analysis, Part 2 — Chemical and 
Microbiological Properties, 2nd edn. American 
Society of Agronomy, Soil Science Society 
America, Madison, WI, pp. 643-698. 

Kudeyarov, V.N. 1981. Mobility of fixed 
ammonium in soil. Ecol. Bull. (Stockholm) 33: 

Liang, B.C., MacKenzie, A.F., and Gregorich, 
E.G. 1999. Measurement of fixed ammonium 
and nitrogen isotope ratios using dry combustion. 
Soil Sci. Soc. Am. J. 63: 1667-1669. 

McBeth, I.G 1917. Fixation of ammonia in soils. 
J. Agr. Res. 9: 141-155. 

Nieder, R., Neugebauer, E., Willenbockel, A., 
Kersebaum, K.C., and Richter, J. 1996. Nitrogen 
transformation in arable soils of north-west 
Germany during the cereal growing season. Biol. 
Fert. Soils. 22: 179-183. 

Scherer, H.W. 1993. Dynamics and availability of 
the non-exchangeable NH4-N — a review. Eur. 
J. Agron. 2: 149-160. 

Scherer, H.W. and Werner, W. 1996. Significance 
of soil micro-organisms for the mobilization of 
non-exchangeable ammonium. Biol. Fert. Soils. 
22: 248-251. 

Silva, J.A. and Bremner, J.M. 1966. Determin- 
ation and isotope-ratio analysis of different 
forms of nitrogen in soils. 5. Fixed ammonium. 
Soil Sci. Soc. Am. Proc. 30: 586-594. 

Soderland, R. and Svensson, B.H. 1976. The 
global nitrogen cycle. In B.H. Svensson and 
R. Soderland, Eds., Nitrogen, Phosphorus, and 
Sulphur— Global Cycles. SCOPE Report 7. 


Soon, Y.K. 1998. Nitrogen cycling involving Young, J.L. and Aldag, R.W. 1982. Inorganic 

non-exchangeable ammonium in a gray luvisol. forms of nitrogen in soil. In F.J. Stevenson, Ed., 

Biol. Fert. Soils. 27: 425^-29. Nitrogen in Agricultural Soils. American Society 

of Agronomy, Madison, WI, pp. 43-66. 
Willis, R.B., Schwab, G.J., and Gentry, C.E. 

1993. Elimination of interferences in the colori- Zhang, Y. and Scherer, H.W. 1998. Simplifi- 

metric analysis of ammonium in water and cation of the standard method for the determin- 

soil extracts. Commun. Soil Sci. Plant Anal. 24: ation of nonexchangeable NH 4 -N in soil. 

1009-1019. Z. Pflanenerndhr. Bodenk. 161: 101-103. 


Chapter 20 

Tee Boon Goh 

University of Manitoba 
Winnipeg, Manitoba, Canada 

A.R. Mermut 

University of Saskatchewan 

Saskatoon, Saskatchewan Canada 


Inorganic carbon occurs in soils commonly as the carbonate minerals calcite (CaCC^), 
dolomite (CaMg(C03) 2 ), and magnesian calcites (Cai_^Mg v C03). Other less common 
forms are aragonite (CaCOs) and siderite (FeCOs). Carbonate in soils can be of primary 
(inherited from parent material) or secondary (pedogenic) origin. Secondary carbonates are 
usually aggregates of silt- and clay-sized calcite crystals that are easily identified in grain 
mounts. Larger crystals of calcite or dolomite are of primary origin (Doner and Lynn 1989). 
Once routinely reported by sedimentologists, the qualitative and quantitative determination, 
especially of Ca and Mg carbonates, is useful in studies of soil genesis and classification, and 
micronutrient and phosphorus sorption. Furthermore, soil carbonates affect root and water 
movement, soil pH (Nelson 1982), and the nature of the exchange complex (St. Arnaud and 
Herbillon 1973). The variability in topsoil carbonate content due to incorporation of subsoil 
calcite and dolomite has been used successfully to explain differences in crop yield in eroded 
landscapes (Papiernik et al. 2005). 

A variety of methods can be used for the determination of calcite, dolomite, and magnesian 
calcite in soils. Chemical determinations of carbonates include the use of empirical standard 
curves relating pH to known carbonate content as well as the measurement of CO2 evolved 
when treated with acid. These permit a measurement of inorganic C from carbonates in soil. 
Most procedures express the carbonate content as the calcium carbonate equivalent. Further 
analysis of the Ca and Mg content provides a means of estimating the kind of inorganic 
carbonate in soil. The largest source of error is in apportioning the cations between the 
carbonate minerals and the soluble cations from the exchange complex of the soil. 

In instances where the carbonates are of primary origin, and hence consist of larger crystals, it 
may first be useful to separate them by density fractionation techniques (Jackson 1985; Laird 
and Dowdy 1994) before further attempting to distinguish between calcite and dolomite. 



The analysis is suitable for rapid and routine analysis of large numbers of samples. A known 
quantity of acetic acid is consumed by reaction with carbonates, and the final pH following 
complete dissolution of CaCCh is recorded for each sample. Calcium carbonate content is 
determined empirically from a standard curve relating pH to weight of CaC03 according to 
the equation 

pH = K + n log [CaC0 3 /(T - CaC0 3 )] (20.1) 

where K and n are constants and T is the total amount of CaC03 that could be completely 
neutralized by the quantity of acetic acid used. 

20.2.1 Reagents and Equipment 

7 Calcite standard: Pure calcite such as Iceland spar calcite ground to <270 mesh in 
size is suitable. 

2 Acetic acid, 0.4 M: Dilute 400 ml_ of 1 M CH 3 COOH to the mark in a 1 L 
volumetric flask with deionized distilled water. 

j pH meter: A digital pH meter is recommended. 

4 Ultrasonic probe: A suitable model with a probe that can be inserted into a 50 ml_ 
centrifuge tube. 

20.2.2 Procedure 

Standard Curve 

7 Weigh accurately, Iceland spar calcite, ranging from 5 to 500 mg into separate 
50 ml_ polypropylene centrifuge tubes. 

2 Add 25 ml_ of 0.4 M acetic acid, which is sufficient to exactly neutralize all the 
CaCC>3 in the largest sample of the standard (500 mg CaC0 3 ), according to the 

CaC0 3 + 2CH3COOH -» Ca 2+ + 2CH 3 COCr + H 2 + C0 2 

3 Shake tubes intermittently for 8 h. At approximately hourly intervals, swirl the 
contents for a few minutes to allow for adequate mixing and degassing. Allow 
tubes to stand overnight with caps loosened to allow escape of CO2. 

4 A final degassing is carried out for approximately 30 s using an ultrasonic probe at 
low setting to prevent excessive splashing. 

5 Centrifuge and record pH of the supernatant to two decimal places after 4 min. 


g Plot standard curve of pH versus log [CaC03/(T — CaCC>3)]. Note: T is the weight 
of CaCC>3 (mg) used to exactly neutralize the volume of acetic acid used and will 
vary if either the volume or concentration of acetic acid is changed. 

Calcium Carbonate Content of Soil Samples 

1 Weigh accurately, up to 2 g soil (<100 mesh size) containing up to 400 mg 
CaC0 3 . Reduce the soil sample weight if the carbonate content exceeds 20%. 

2 Repeat steps (2) through (5) as for the standard curve above. 
20.2.3 Calculations 

From the pH value recorded, determine the value of log [CaCC>3/(T - CaCC>3)] using the 
standard curve and calculate the weight of CaCC>3 (mg) in the soil sample. The total 
carbonate content so determined is expressed as percent calcium carbonate equivalent. 

% CaC0 3 equivalent = m g CaC ° 3 x 100 (2 0.2) 

mg sample 


If dolomite is present in the soil sample, increased reaction times may be required for the 
dissolution to go to completion. The accuracy of results is influenced by (i) proton consumption 
by soil constituents, (ii) acid-generating hydrolysis reactions during mineral decomposition, 
(iii) high PCO2, (iv) volatilization of acetic acid, and (v) errors in pH determination. These 
can be minimized by standard additions of Ca 2+ , from a solution of CaCl2, to all samples and 
standards; grinding of samples to increase reactivity of sand-sized carbonates and reduction 
of reaction time between acetic acid and other minerals; use of covers to reduce loss of acetic 
acid; degassing CO2; and reduction of suspension effects in pH reading (Loeppert et al. 1984). 


A preweighed soil sample containing carbonates is reacted with acid. The resultant loss in 
weight from CO2 released is used to calculate the calcium carbonate content. Calcite and 
dolomite cannot be accurately distinguished, but a fair estimate of the proportion of dolomite 
in the sample can be obtained by checking the weight loss with time. 

20.3.1 Reagents 

7 Hydrochloric acid (HCI), 4 M. 

2 Hydrochloric acid (HCI)-ferrous chloride (FeCI 2 • 4H 2 0) reagent: Dissolve 3 g of 
FeCI 2 • 4H 2 per 1 00 ml_ of 4 M HCI immediately before use. 

20.3.2 Procedure 

7 Weigh a stoppered, 50 ml_ Erlenmeyer flask containing 10 ml_ of the HCI — FeCb 

2 Transfer a 1-10 g soil sample containing between 100 and 300 mg of carbonate 
to the flask gradually to avoid excessive frothing. 

j After effervescence has subsided, replace the stopper loosely and allow the 
carbonate to decompose further in the mixture for about 30 min with occasional 
swirling to displace any accumulated C0 2 . Replace the stopper and weigh the 
flask with its contents. 

4 Repeat step (3) until the change in weight of the flask and its contents is no more 
than 2-3 mg. The reaction is usually complete within 2 h. 

20.3.3 Calculations 

Weight of CO2 lost from carbonates = difference in initial and final weights of (flask 
+ stopper contents) 


When dolomite is present, it is considerably less reactive to cold HC1. Therefore, if the 
weight is observed to decrease markedly after 30 min, some dolomite is present. The use of 
acid containing FeCh as an antioxidant eliminates errors caused by oxidizing interferences 
due to MnC>2 in soil. The accuracy of this method depends upon the accuracy of weighing 
and the degree to which CO2 retained in solution is compensated for by loss of water vapor. 


The loss in weight of a soil sample is measured accurately after reaction between carbonates 
in the soil and acid. In this method, the loss of water vapor evolved with CO2 is eliminated by 
a trap containing anhy drone. The addition of a CO2 trap to the apparatus is an alternative to 
the method, by measuring the gain rather than the loss in weight, and provides a check 
against any leaks in the connections to the glassware. With several units in operation, the 
method is quite rapid and accurate. 

20.4.1 Apparatus 

The apparatus is assembled as depicted in Figure 20.1 for the weight loss method. A 
polyethylene drying tube packed with Ascarite II R to trap CO2 can also be attached to the 
end of the gas train after stopcock D in the weight gain method. 

20.4.2 Reagents 

/ Hydrochloric acid (HCI), 6 M. 

2 HCI-ferrous chloride (FeCI 2 • 4H 2 0) reagent: Dissolve 3 g of FeCI 2 • 4H 2 per 
1 00 ml_ of 6 M HCI immediately before use. 

A. Glass wool plugs 

B. Anhydrone, Mg(CI0 4 ) 2 

C. Vial containing 6 moi L~ 1 HCI 

D. Stopcock 

E. Stopcock 

F. 125 mL Erlenmeyer flask 

G. Stopper 
H. U-tube 

I . Calcium chloride tube (shortened) 
J . Glass tube 

FIGURE 20.1. Apparatus for accurate quantitative determination of calcium carbonate equiva- 
lent. (From Raad, A. A., in J. A. McKeague (Ed.), Manual on Soil Sampling and 
Methods of Analysis, 2nd edn. Canadian Society of Soil Science, AAFC, Ottawa, 
Ontario, Canada, 1978.) 

3 Anhydrone (Mg(CI04)2), drying agent. 

4 Ascarite ll R : 20-30 mesh, optional. 

20.4.3 Procedure 

Weight Loss Method 

1 Weigh a 1-10 g sample of oven-dry soil (<100 mesh) containing <1 g CaC03 

equivalent in a 125 mL Erlenmeyer flask. 

Wash down the sides of the flask with 1 mL of deio 

ized distilled water. 

Transfer 7 mL of HCI-FeCI 2 reagent into vial C (Figure 20.1), and place the vial 
upright in the flask without spilling any acid. 

Moisten stopper G with glycerin, sprinkle it with a small amount of 180 mesh 
abrasive to overcome slipperiness, and assemble the apparatus as in Figure 20.1 
without connecting the U-tube to stopcock E. Close stopcocks D and E. 

Place the apparatus beside the balance and allow the temperature in the flask to 
equilibrate with that of the air in the balance. 

Using tongs, place the apparatus on the weighing pan, open stopcock D, and 
record weight to the nearest 0.1 mg. Close stopcock D immediately. Weigh again 
after 1 min to ensure that weight has stabilized. 


7 Open stopcock D and shake the flask to upset vial C, thus allowing the acid to 
react with the soil. 

q After 10 min, attach the U-tube H to the apparatus, open stopcock E, and apply 
gentle suction at stopcock D at a rate of 5-1 bubbles per second at tube J to 
sweep out CO2 with dry air. Shake the flask at 10 min intervals. 

g Stop the suction when the reaction is complete (usually 30 min; 1 h if dolomite is 
present). Close stopcocks D and E. Disconnect the U-tube H. Wait for 1 h and 
weigh the apparatus and its contents with stopcock D open. Check the weight 
after 10 min. 

1q Calculate as follows: 

weightg -Final weight, g)^^ {2Q4) 

Sample weight, g 

Weight Gain Method 

7 Weigh drying tube containing Ascarite ll R to the nearest 0.1 mg. Attach to the 
apparatus depicted in Figure 20.1 at stopcock D. 

2 Proceed as described in the weight loss method but apply suction at the end 
of the polyethylene drying tube so that the gas train passes through the 
C0 2 trap. 

j Disconnect from the CO2 trap and weigh drying tube and its contents. 

4 Calculate as follows: 

, „ „„ , (Final weight, g — Initial weight, g) 

% CaCOs equivalent = s_L5 6 - 5 x 227.3 (20.5) 

Sample weight, g 


The results obtained by the two methods should agree within the limits of weighing error. 
Larger discrepancies may indicate leaks in the connections of the apparatus. 


The citrate buffer method described by Raad (1978) is presented here. Calcite and dolomite 
are selectively dissolved in a citrate buffer solution, Ca and Mg in solution are determined, 
and the calcite content of the sample is calculated. It is assumed that dolomite has a Ca:Mg 
molar ratio of 1:1 and the only sources of Ca and Mg in the solution are calcite and 
dolomite (i.e., no magnesian calcite is present), and exchangeable calcium and magnesium 
have been removed first or otherwise accounted for (Hesse 1971). The portion of dolomite 


dissolved in the citrate buffer is calculated from the Mg in solution; an equivalent a 
of Ca is assigned to it; and the remaining Ca determines the calcite content of the sample. 
The total dolomite content of the sample is obtained by the difference between the total 
carbonate content previously determined in another subsample and the portion of carbon- 
ate from calcite. As a check of accuracy, the dolomite content of the sample can also be 
calculated from the Mg in solution. The method is useful if clay-sized dolomite is present 
in the sample. 

20.5.1 Reagents 

7 Citrate buffer: Dissolve 64 g citric acid (CeHsOz) in 1 L of deionized water. Titrate 
to pH 5.85 with concentrated NH 4 OH. 

2 Sodium chloride-ethanol: Dissolve 58.5 g NaCI in 30% (v/v) ethanol and bring to 
1 L with deionized water. 

3 Sodium dithionite. 

20.5.2 Procedure 

1 Weigh 50-500 mg oven-dry soil ground to pass a 100 mesh sieve into a 50 ml_ 
centrifuge tube. 

2 Wash twice with NaCI-ethanol solution and discard washing. 

j Add 25 ml_ of citrate buffer solution, and heat in a water bath at 80°C. Add 
approximately 0.5 g of sodium dithionite and continue heating with stirring for 
about 15 min. 

4 Centrifuge and collect supernatant in a 250 mL volumetric flask. Wash the residue 
once with 25 mL of citrate buffer, centrifuge, and collect the supernatant. Make to 
volume with deionized water. 

5 Determine Ca and Mg in solution by atomic absorption spectroscopy usi 
standards made up in the same concentrations of citrate buffer and dithioni 
Standards and sample solutions should contain 1 mg La mL~ 1 to minim 
interference effects. 

6 The total carbonate content is determined in another subsample by other quan- 
titative methods (Section 20.2). 

20.5.3 Calculations 

The molecular formula of calcite is CaCC>3, and the molecular formula of dolomite is 
CaMg(C03) 2 . If total citrate-soluble Ca = X mmol, and citrate-soluble dolomite Mg = Y 
mmol, then citrate-soluble dolomite-Ca = Y mmol, and the mmol calcite-Ca = X — Y. 
Since 1 mmol of Ca is contained in 1 mmol of calcite, and since 1 mmol calcite weighs 
100 mg, then: 

% calcite in sample 


(X - Y) mmol 100 mg 
mg sample mmol 

The dolomite content of the sample can be calculated if total carbonate in sample = Z mmol. 
Since total carbonate = calcite carbonate + dolomite carbonate, the mmol dolomite- 
CO3 = Z — (X — Y), and since 2 mmol of carbonate is contained in 1 mmol of dolomite, 
then, the mmol dolomite = \ (mmol dolomite-C03) = \\Z — (X — F)], and since 1 mmol 
dolomite weighs 184 mg, then 

i [Z - (X - Y)] mmol 184 mg 

% dolomite in sample = ^ x - x 100 (20.7) 

mg sample mmol 

Alternatively, the dolomite content can be calculated by the mmol dolomite-Mg = Y: 

% dolomite in sample = Y m ™\ x ™^ x 100 (20.8) 

mg sample mmol 

The % CaCC>3 equivalent of the dolomite present 

= [ Z-(X-F)]mmol x 100mg xi()0 
mg sample mmol 


The dolomite content calculated from dolomite-C03 should agree with that calculated from 
the Mg in solution unless some source of citrate-soluble Mg other than dolomite, or 
magnesian calcite is present. If magnesian calcite is present, the calcite content of the sample 
will be underestimated. Therefore, identification and quantification of magnesian calcite 
(e.g., St. Arnaud et al. 1993) should be conducted for more specialized research. 

Allison, L.E. and Moodie, CD. 1965. Carbonate. Hesse, P.R. 1971. A Textbook of Soil Chem- 

In C.A. Black et al., Eds., Methods of Soil Ana- ical Analysis. Chemical Publishing Co., Inc. 

lysis, Part 2 — Chi •• cal ndMici biolo ical Pro- New York, NY. 
perties 1st ed. American Society for Agronomy, 

Madison, WI, pp. 1379-1396. lackson, M.L. 1985. Soil Chemical Analysis- 
Advanced Course, 2nd edn. M.L. lackson, 

Doner, H.E. and Lynn, W.C. 1989. Carbonate, Madison, WI. 
halide, sulfate and sulfide minerals. In J.B. 

Dixon and S.B. Weed, Eds., Minerals in Soil Laird, D.A. and Dowdy, R.H. 1994. Preconcen- 

Enviroiunents, 2nd ed. Soil Science Society of tration techniques in soil mineralogical analyses. 

America, Madison, WI, pp. 279-330. In I.E. Amonette and L.W. Zelazny, Eds., 


Quantitative Methods in Soil Mineralogy. Soil 
Science Society of America, Madison, WI, 
pp. 236-266. 

Loeppert, R.H., Hallmark, C.T., and Koshy, M.M. 
1984. Routine procedure for rapid determina- 
tion of soil carbonates. Soil Sci. Soc. Am. J. 
48: 1030-1033. 

Nelson, R.E. 1982. Carbonate and gypsum. In 
A.L. Page et al., Eds., Methods of Soil Analysis, 
Part 2 — Chemical and Microbiological Proper- 
ties, 2nd edn. American Society of Agronomy, 
Madison, WI, pp. 81-197. 

Papiernik, S.K., Lindstrom, M.J., Schumacher, J.A., 
Farenhorst, A., Stephens, K.D., Schumacher, T.E., 
and Lobb, DA. 2005. Variations in soil properties 
and crop yield across an eroded prairie landscape. 
/. Soil Water Conserv. 60: 47-54. 

Peterson, G.W., Chesters, G., and Lee, G.B. 1966. 
Quantitative determination of calcite and dolo- 
mite in soils. J. Soil Sci. 17: 328-338. 

Raad, A.A. 1978. Carbonates. In J.A. McKeague, 
Ed., Manual on Soil Sampling and Methods of 
Analysis, 2nd ed. Canadian Society of Soil 
Science, AAFC, Ottawa, ON, Canada, pp. 86-98. 

St. Arnaud, RJ. and Herbillon, A.J. 1973. Occur- 
rence and genesis of secondary magnesium- 
bearing calcites in soils. Geoderma 9: 279-298. 

St. Arnaud, R.J., Mermut, A.R., and Goh, Tee 
Boon. 1993. Identification and measurement of 
carbonate minerals. In M.R. Carter, Ed., Soil 
Sampling and Methods of Analysis. Lewis Pub 
lishers, Boca Raton, FL, pp. 737-754. 

USDA Soil Conservation Service. 1967. Calcium 
carbonate. In Soil survey investigations report 
No. 1. Soil Survey Laboratory Methods and 
Procedures for Collecting Soil Samples. U.S. 
Government Printing Office, Washington DC, 
pp. 28-30. 



Chapter 21 
Total and Organic Carbon 

J.O. Skjemstad and J. A. Baldock 

Commonwealth Scientific aiul industrial Research Organize 
Glen Osmond, South Australia, Australia 


Carbon in soils exists in both organic and inorganic forms. Carbonate, in a variety of forms, 
makes up the inorganic component of total carbon (TC), whereas a range of organic moieties 
make up the organic carbon (OC) component. The terms OC or organic matter associated 
with soil have been defined in various ways. Stevenson (1994) and Baldock and Nelson 
(1998) defined OC as the total of all organic materials existing within and on soil, whereas 
Oades (1988) excluded charcoal and charred materials and MacCarthy et al. (1990) excluded 
nondecayed plant and animal tissues, their partial decomposition products, and the living soil 
biomass. In reality, however, the methods used to determine both TC and OC do not 
discriminate between the various fractions described above, and consequently, the all 
encompassing definition of OC used by Stevenson (1994) and Baldock and Nelson (1998) 
is also used in this chapter. 

There are a number of approaches available for the determination of TC and OC in soils. 
These are broadly based on either the chemical or thermal oxidation of soil OC. Chemical 
or wet oxidation is followed by the measurement of expelled CO2 (Snyder and Trofymow 
1984) or the consumption of oxidant required to quantitatively oxidize the OC (Walkley and 
Black 1934). Under acidic conditions, any chemical or wet oxidation methods that measure 
expelled CO2 will also include carbonate C and will be a measure of TC. In dry combustion 
methods, samples are heated to high temperatures, usually exceeding 1000°C in the 
presence of excess O2. Under these conditions, all C present in OC and carbonate is 
quantitatively converted to CO2. Liberated CO2 may be determined gravimetrically (Allison 
et al. 1965), volumetrically (Rayment and Higginson 1992), titrimetrically (Snyder and 
Trofymow 1984), or spectrometrically (Merry and Spouncer 1988). If thermal oxidation 
(dry combustion) at temperatures exceeding 1100°C is used, all carbon in the sample 
including carbonates will be determined (Giovannini et al. 1975). For both chemical and 
thermal oxidation methods where CO2 is measured, a correction for carbonate can be made 
from a separate carbonate measurement or the carbonate may be removed with acid before 
carbon analysis. 


A comparison of several titrimetric and gravimetric methods was made by Kalembasa and 
Jenkinson (1973) and showed that dry combustion methods were the more precise. Loss on 
ignition at various temperatures has also been used as a simple predictor of soil organic 
matter in soil types where clay contents are low (Ball 1964; Lowther et al. 1980) but these 
methods are not recommended if alternative methods are available. 


Currently, there are a large number of instruments available commercially for the determin- 
ation of TC. Some instruments will simultaneously determine C and one or more of the 
following additional elements: N, H, and S. For some instruments, an induction furnace is 
used to heat the sample rather than the more commonly available resistance furnace. These 
methods rely on the addition of Fe chips and catalysts to the sample to produce high 
temperatures of up to 1400°C (Rayment and Higginson 1992) or the use of a quartz enclosed 
graphite crucible to heat the sample externally (Allison et al. 1965). 

For all instruments, combustion is usually carried out at high temperatures (>1000°C) and in 
a stream of O2. This is to ensure that all C species are quantitatively converted to CO2. At 
lower temperatures, combustion may not be complete, resulting in the generation of some 
CO or incomplete decomposition of carbonate species. Unless the CO is converted to CO2, 
losses will be recorded due to the inability of detectors tuned to CO2 to detect CO. This can 
be overcome by mixing a catalyst such as V2O5 (Morris and Schnitzer 1967) with the 
sample, or by the inclusion of a catalytic conversion furnace (usually CuO) in the train 
prior to detection. For some instruments, samples are loaded into Sn or Al cups and the 
sample and cup are ignited. The exothermic reaction that takes place as the Sn or Al cups 
ignite increases the combustion temperature significantly, even though the furnace may 
operate at 1000°C or less. 

Carbonates will be decomposed at elevated temperatures (500°C-1000°C) to also produce 
CO2; and so for OC measurements, carbonates must be either removed prior to combustion 
or a correction must be made. Some authors have suggested using low combustion temper- 
atures to confine the carbon measured to only that contained within organic materials. 
However, this requires temperatures <720°C, since dolomite starts to decompose above 
this temperature. At temperatures <720°C, not only is the production of CO an issue, but 
also a substantial amount of charcoal can be generated and may not fully oxidize if the 
combustion duration is short. 

Carbonate is removed through the addition of acid before analysis. Sulfurous acid (H2SO3) is 
the only suitable acid because its reducing properties minimize the oxidation of OC during 
the process (Piper 1944). This can be accomplished in two ways: either by destruction of the 
carbonate within the combustion vessel or before subsampling for OC analysis. The former 
approach is preferable because it minimizes sample handling, but the combustion vessel 
needs to be large enough in relation to the sample size to enable the addition of excess acid 
and the potential effervescence that may result. Three methods are described here; addition 
of acid directly to the combustion vessels at two different scales and a method for removing 
carbonate before subsampling. 

A range of modern instruments with several levels of automation are commercially available 
for dry combustion methods. The two most common detection systems are based on infrared 
or thermal conductivity measurement. All instruments are provided with comprehensive 


instructions on the setting up, standardization, and use of the equipment for routine analysis. 
For the remainder of this section, we will not describe TC and OC analysis based around one 
or more specific instruments but will discuss the issues that arise at different parts of the 
carbon analysis train and how these may be dealt with. 

21.2.1 Removal of Carbonate — Large Combustion Vessel 

This method is directed toward instruments such as those manufactured by Laboratory Equip- 
ment Corporation (LECO), which utilize large ceramic combustion boats with a 5-6 mL 
capacity. These boats are porous and so acid treatment cannot be performed directly within 
the boat, since some leakage will occur and OC rendered soluble by the treatment will be lost. 
The sample is therefore weighed into a commercially available Ni liner placed within the 
combustion boat. 


6% (w/v) H 2 S0 3 solution. 

7 Weigh 0.1-1.0 g of soil (<0.15 mm) into a Ni liner placed within a ceramic 
combustion boat, and place on a hot plate. Larger samples are not recommended 
unless the samples are known to contain only very small concentrations of 
carbonate and low concentrations of OC. 

2 Moisten samples with a little distilled or deionized water and add 1 .0 mL of 6% 
H2SO3 to each boat and allow to stand. Meanwhile, turn on the heating plate and 
set the temperature to ensure that the samples do not exceed 70°C. Because of the 
insulating properties of the ceramic boat and the generally large loss of heat from 
the hot plate between samples, the hot plate temperature may need to be set as high 

3 When the samples have stopped reacting, add a further 1 .0 mL of 6% H2SO3. It is 
important that the samples are not allowed to dry until the treatment is completed, 
since this will lead to deterioration in the Ni liner and ultimately leakage. Some 
evaporation will be necessary, however, to allow the addition of sufficient acid to 
complete the carbonate removal. 

4 When addition of acid no longer promotes a reaction, allow the samples to dry. 

5 Analyze samples using an OC determinator as described by the manufacturer, but 
ensure that the initial weight of the sample (prior to 6% H2SO3 treatment) is 
entered into the calculation and not the final weight after treatment. 

14 mL of 6% H2SO3 is required to neutralize 1.0 g of CaCC»3 ■ H2SO3 solutions will slowly 
deteriorate with time, losing SO2; and more acid may be needed depending on the age of the 
H2SO3 solution. 


21.2.2 Removal of Carbonate — Small Combustion Vessel 

This method is directed toward instruments such as those manufactured by Carlo Erba, 
which utilize small samples placed in metal combustion cups. For pretreatment for carbonate, 
Ag cups are recommended instead of the normal Sn or Al cups, since Ag has a much greater 
resistance to 6% H2SO3. The reaction is carried out in a small Al block that has been 
drilled out to hold the capsules with a reasonably tight fit similar to that described by Verardo 
et al. (1990). 


6% (w/v) H2SO3 solution. 

/ Place Ag cups into a small Al block. Weigh up to 20 mgof the sample (<0.02 mm) 
into the cups and place the small block onto a heating plate. 

2 Add 10 |xL distilled or deionized water to each sample. This slows the initial 
reaction when 6% H2SO3 is added and minimizes the risk of losing sample due to 
a strong effervescence. Add 1 (xL of 6% H2SO3 to each sample and allow to 
stand. Meanwhile, turn on the heating plate and set temperature to 70°C. 

j When reaction has ceased, add another 1 |xL of acid. It is important that the 
samples are not allowed to dry until the treatment is complete, because this will 
lead to deterioration in the Ag cups and they may crumble during the balling 
process. Some evaporation will be necessary, however, to allow the addition of 
sufficient acid to complete the carbonate removal. 

4 When addition of acid no longer promotes a reaction, allow the samples to 
dry and analyze samples using an OC determinator as described by the manu- 
facturer ensuring that the initial weight of the untreated soil is used in the 

21.2.3 Removal of Carbonate Prior to Subsampling 


6% (w/v) H2SO3 solution. 


7 Weigh 1 .0-2.0 g of sample into a preweighed test tube or beaker and place in a 
digestion block or on a hot plate. 

2 Add 1 .0 ml_ of 6% H 2 S0 3 and allow the reaction to subside. Meanwhile, turn on 
the block/hot plate and set temperature to 70°C. 

3 Continue to add 6% H 2 S0 3 in 1.0 ml_ aliquots until further addition no longer 
yields a reaction and allow samples to dry. 


4 When dry, remove samples from block, place in a desiccator or cabinet with silica 
gel, and allow to cool. Weigh tube or beaker, and quantitatively remove treated 
sample. Maintain samples in an oven dry state (105°C) under desiccation for OC 

5 Take 1 g of soil and place in a preweighed beaker or silica dish. Dry for 24 h at 
105°C in an oven. Cool in a desiccator and reweigh. 


This pretreatment modifies the sample in two ways. First, because the sample is heated, the 
water content of the sample is changed; and second, because SC>3 2 ~ has more mass than 
C03 2 ~, the mass of the sample will increase. These changes need to be taken into consid- 
eration when calculating the OC content of the sample. 

j Oven dry factor (ODF) is calculated as 

ODF = (weight air dry soil)/(weight oven dry soil) (21 .1 ) 

2 Soil OC content (105°C) g kg -1 = (OC content of treated sample g kg" 1 ) 

x (weight of sample post treatment at 105°C)/ 
(weight of sample taken for treatment/ODF) 

21.2.4 Correction for Carbonate 

For analyzers that rely on a continuous flow of exhaust gases for the determination of CO2, 
and the temperature of the combustion is high (>1000°C), a correction for carbonate content 
can be made mathematically. Some analyzers, however, rely on the collection of a given 
volume of exhaust gases that limits the time over which the sample is heated to high 
temperature. This is common in instruments that analyze for C and N simultaneously. 
Under these conditions, carbonates may not be fully decomposed and the TC may be 
underestimated. This can often be overcome by taking much smaller samples, but such 
analyzers need to be tested to determine whether a TC measurement in the presence of 
carbonate is quantitative. If not, then a simple correction is not quantitative and carbonates 
will need to be removed before analysis as described by one of the procedures given above. 

For those analyzers that quantitatively determine TC, the following correction can be made: 

OC g kg" 1 = TC g kg" 1 - 0.12 x CaC0 3 g kg" 1 (21.3) 

21.2.5 Standards 

Preignition of combustion vessels may be necessary to eliminate contamination. For metal 
cups, this can be achieved at 550°C and for ceramic boats at 1000°C for 16 h. 

A wide range of materials can be used as OC standards for these instruments. These 
include CaC03, EDTA, sucrose, glucose, potassium hydrogen phthalate, and urea. 


Standard materials of given C content can also be purchased from companies such as LECO. 
It is recommended, but not essential, that the standards used exhibit similar combustion 
characteristics to the samples. This becomes an issue when highly organic standards or 
samples are used. For continuously monitoring instruments, the linear operating range of the 
detector may be exceeded if the standards or samples "flash" and an intense pulse of CO2 
passes through the detector within a short time. This can be overcome by reducing the rate of 
combustion by covering the sample with a layer of ignited sand. The sand used should be < 1 mm 
and ignited at >1000°C over at least 24 h with frequent stirring of the sand to ensure any 
C present is totally combusted to CO2. 

The linear operating range of the detector can be determined by analyzing different 
weights of standards and samples over a range that encompasses expected TC and 
OC contents. 


With these methods, dichromate (Cr207 2 ~) solution in combination with sulfuric acid 
(H2SO4) is used to oxidize OC to CO2. The orange dichromate is reduced to the green 
Cr 3+ form according to the following equation: 

2H 2 Cr 2 7 + 6H2SO4 + 3C -> 2Cr 2 (S0 4 ) 3 + 3C0 2 + 8H 2 (21 .4) 

The oxidation state (Baldock et al. 2004) of the C in the organic matter can influence the 
consumption of oxidant. Molecules with a high H/C ratio, such as lipids, give higher 
recoveries than molecules with high O/C ratio (Skjemstad 1992). Because soil organic 
matter is highly diverse in chemistry, these two effects tend to cancel one another when 
the whole soil is considered; however, variations in oxidation state of OC with increasing 
extent of decomposition have been documented (Baldock et al. 2004). This issue, therefore, 
may be more serious if specific soil OC fractions are being considered. 

If consumption of oxidant is to be used, the analysis can be performed with heating (Heanes 
1984) or without external heating (Walkley and Black 1934). The consumption of oxidant 
can be determined either by titration using an indicator or platinum-calomel electrode or 
colorimetrically. If only the heat of reaction is used with no applied external heating, as in 
the case of the Walkley and Black (1934) method, then a 75%-80% recovery of carbon is 
usually obtained and a conversion factor of 1.3 is commonly used to equate the OC value to 
the thermal oxidation (dry combustion) methods. This factor will vary among soil types and 
with depth and must be applied with caution. 

Any material that can be oxidized by the dichromate will be measured as OC. Chlorides are 
quantitatively oxidized to free chlorine by chromic acid. Thus, where consumption of 
dichromate is used to determine OC, the presence of CI can result in erroneously high OC 
contents. This methodological error can be corrected when the CI content of the sample is 
known. Four CI ions have the same reducing power as 1 C atom (4C1 = 20 = 1C) and hence 
11.83 g of CI is equivalent to 1 g of C. Several workers have suggested that the addition of 
Ag 2 S04 can suppress CI interference (Walkley 1947). Heanes (1984), however, demon- 
strated that the addition of Ag 2 S04 either as a solid or in solution with the H2SO4 was 
ineffective. For saline soils therefore, it is recommended that the correction for measured CI 
be used rather than additions of Ag2S04. 


In this chapter, we detail two methods. One based on the Schollenberger (1945) method uses 
external heating and titration with an indicator. The alternative method uses external heating 
and colorimetric determination of Cr m (Heanes 1984). Reduced forms of Fe and Mn may 
also interfere with these methods. These interferences are rare but can be overcome by the 
procedures described in Jackson (1958). The use of steel or iron mills should also be avoided 
since these can act as a source of Fe metal, which is readily oxidized under the conditions of 
the reaction. 

21.3.1 Dichromate Redox Colorimetric Method (Heanes 1984) 

The dichromate redox colorimetric method utilizes the formation of the green Cr m species 
resulting from the reduction of the orange dichromate (Cr VI ) species. The amount of 
dichromate consumed is determined against a set of standards and measured on a spectro- 
meter in the visual range. Carbonates are not determined by this procedure but CI will 
interfere. Because the dichromate solution is not used as the primary standard in 
this method, we describe here the use of the more soluble Na2Cr207 salt rather than 
the K salt. 


Na dichromate solution: dissolve 50 g of Na 2 Cr 2 07 in distilled or deionized 
water and dilute to 1 L. 

Sulfuric acid: 98% cone. H 2 S0 4 . 

Standards: dissolve 1 .376 g of glucose monohydrate in distilled or deionized water 
and dilute to 250 ml_. A small crystal of HgCI 2 can be added to preserve the 
standard against microbial decomposition. 1 .0 ml_ of this solution = 2.0 mg of OC. 

Prepare standards by adding a range of aliquots of the glucose solution to 
borosilicate tubes (25 mm OD) marked at 100 ml_. A convenient range is 1-12 ml_ 
of standard that equates to 2-24 mg of OC. Tubes containing glucose solution and a 
blank are dried in an oven at a temperature not exceeding 60°C. 

Weigh 0.1-2.0 g of air-dried soil (<0.15 mm) containing <20 mg of OC into 
digestion tubes. 

Add 10.0 ml_ of Na 2 Cr 2 7 solution, and while agitating add 20.0 ml_ of 98% 
H 2 S04 cautiously so that the reaction is confined to the bottom of the tube. 
Agitate for a further 30 s before inserting into a preheated (135°C) digestion 
block. Agitate tubes occasionally to ensure all of the soil material is exposed to 
the chromic acid mixture. 

After 45 min, remove tubes from the block and allow to cool. Add 50 ml_ of 
distilled or deionized water to digest and agitate with a thick-walled glass 
capillary tube that has a stream of air passing through it so that the samples are 
thoroughly mixed. After removal from the block, the samples still contain H 2 S0 4 
at a strong enough concentration to cause heating when water is added. 


If the tubes are inverted after the addition of water, enough heat is generated to 
potentially cause hot chromic acid to be lost. Agitation with the assistance of a 
stream of air prevents any losses. When cool, make the tubes up to 100 mL with 
distilled or deionized water and invert to mix using a rubber bung. 

Decant diluted chromic acid mixture into 15 mL centrifuge tubes and centrifuge 
at 2000 rpm for 15 min. Measure the absorbance of the centrifuged samples at 
600 nm in a 10 mm cell. 

Construct a standard curve plotting absorbance at 600 nm against mg C present in the 
standards. Using this curve, estimate the mg C in the unknown samples. 

g C kg _I soil = mg C in digest/weight soil in grams (21.5) 

If the mg C content of samples is <2 or >20, analysis should be repeated with more or less 
weight to bring them within the optimum range of the determination. 

Modification for Saline Soils 

For saline soils, a separate determination of the chloride content of the soil is required and 
expressed as g CI kg~' soil. The OC content of the soil is then corrected for the CI 

g C kg^'soil = apparent g C kg -1 soil - (g CI kg _1 soil/12) (21.6) 

21.3.2 Dichromate Redox Titration Method 

This procedure is similar to the spectroscopic method but utilizes the unreacted dichromate 
(Cr VI ) that remains following the reaction of OC with acid dichromate. Back titration with 
Fe 11 solution is used to determine the remaining dichromate. The procedure described here is 
based on that described by Schollenberger (1945) with the modification by Jackson (1962) 
for the o-phenanthroline indicator. N-phenanthranilic acid (Nelson and Sommers 1982) or 
diphenylamine (Piper 1944) can be substituted. Carbonates do not interfere but CI does and a 
correction must be made if CI levels are high. 


Digestion mixture: dissolve 39.22 g of K 2 Cr 2 7 (dried at 90°C) in 800-900 mL of 
distilled or deionized water in a large glass beaker. Carefully add 1 L of 98% 
H2SO4. As the acid is added, the mixture will become very hot and will boil. 
When cool, make to 2 L with distilled or deionized water. This solution is 0.067 M 
(0.4 N) in dichromate and 9 M in H2SO4 and is the primary standard for the OC 

Ferrous ammonium sulphate: dissolve 157 g of Fe(NH 4 )2(S0 4 )2 • 6H 2 in about 
1 L of distilled or deionized water containing 100 mL of 98% H 2 S0 4 . Make to 2 L 
to give a -0.2 M (-0.2 N) solution. The solution does not store well and must be 
standardized against the dichromate solution at each use. 

Phosphoric acid: 85% H3PO4. 

Indicator solution: dissolve 3.00 g of o-phenanthroline monohydrate (Ferroin) 
and 1 .40 g of FeS0 4 • 7H 2 in distilled or deionized water and dilute to 200 ml_. 
Alternatively, dissolve 0.1 g of /V-phenanthranilic acid and 0.1 g of Na 2 C0 3 in 
100 ml_ of distilled or deionized water or dissolve 0.5 g of diphenyl- 
amine in 100 ml_ of 98% H 2 S0 4 containing 20 ml_ of distilled or deionized 

1 Weigh samples (<0.1 5 mm) up to 1 .0 g that contain between 1 and 1 mg of OC 
into 100 mL digestion tubes. Add 15 mL of digestion mixture and place on 
digestion block preheated to 1 50°C. 

2 After 45 min, remove samples from the block and allow to cool before quantita- 
tively transferring solution and sample to a titration vessel with approximately 
50 mL of distilled or deionized water. Add 5 mL of 85% H3PO4 and four drops of 
indicator. The H3PCU eliminates interference from Fe'". 

3 Titrate with Fe(NH 4 )2(S0 4 )2 solution to a color change from green to reddish 
brown for the o-phenanthroline, dark violet-green to light green for the 
/V-phenanthranilic acid, and violet-blue to green for the diphenylamine. 

4 Two unheated blanks are also titrated to standardize the Fe(NH 4 ) 2 (S0 4 )2 solution. 

7 Calculate the molarity (normality) of the Fe(NH 4 ) 2 (S0 4 ) 2 solution as 

Molarity of Fe(NH 4 ) 2 (S0 4 ) 2 solution = (1 5 x 0.4)/ 7^ (21 .7) 

where T y is the titer of the Fe(NH 4 ) 2 (S0 4 ) 2 solution in mL. 

2 Calculate the OC concentration in the sample as 

gOC kg- 1 soil = (6- T 2 )xMx3/W (21.8) 

where B and T 2 are titers in mL of heated blank and sample, respectively, M is the 
molarity of the Fe(NH 4 ) 2 (S0 4 ) 2 solution, and W is the weight of sample in grams. 


With this method, the sample is oxidized with a H 2 S0 4 -dichromate mixture and the evolved 
C0 2 is captured in NaOH solution and determined by titration using either an indicator or pH 
meter. This approach is more complex than the redox approach but most of the interferences 
encountered with the redox methods are eliminated. A further advantage of this method is 
that the trapped C0 2 can also be used to determine isotopic composition (Amato 1983). 
Various vessels have been used to contain the reaction and collect the C0 2 . Snyder and 


Trofymow (1984) used tubes with screw caps, Amato (1983) used tubes with subaseals, and 
Dalai (1979) used McCartney bottles. 

Because of the acidic conditions under which the reaction progresses, any carbonates present 
will also be quantitatively converted to CO2 and determined. If carbonates are present, OC is 
determined by either first removing the carbonates in the reaction vessel or correcting for 
their presence. Carbonate content can either be determined using the same reaction vessel or 
can be determined by another suitable method of analysis. 

The method as described by Snyder and Trofymow (1984) can handle solid or liquid samples 
and uses a temperature of 120°C. If only solid samples are processed, the digestion can be 
performed at a higher temperature, provided it remains below the boiling point of the acid 
mixture. If boiling occurs, the vessels may leak or even break. For the determination of OC 
only in the presence of carbonates, an acid pretreatment is required or the TC can be 
corrected for carbonate as outlined in Section 21.2.4. 

21.4.1 Preparation of Reaction Tubes 

Standard culture tubes capped with screw caps containing a conical polyseal are modified 
with three indentations near the top capable of supporting an inserted glass vial (15 x 45 mm). 
Alternatively, a glass rod bent at one end can be inserted into the tube so that the bend in the 
rod supports the vial at an appropriate distance above the reaction mixture (Amato 1983). 
Amato (1983) also suggests the use of regular digestion tubes sealed with subaseals. Either 
approach is satisfactory. 

21.4.2 Reagents 

7 Pretreatment acid mixture (for elimination of carbonates): dilute 57 mL of 98% 
H2SO4 in 600 mL of distilled or deionized water and add 92 g of FeS0 4 • 7H 2 0. 
Dissolve and make to 1 L to give approximately 1 M H 2 S0 4 containing 5% 

2 Digestion mixture: these are kept separate and only combined in the reaction tube: 
(a) K 2 Cr 2 7 and (b) a mixture of three parts 98% H 2 S0 4 and two parts 85% H3PO4. 

j C0 2 absorption solution: dissolve 16.0 g of NaOH and bring to 200 mL with 
distilled or deionized water to give a ~2 M solution. This should be kept in an 
airtight flask or under a C0 2 trap. 

4 Indicator solution: dissolve 0.4 g of thymolphthalein in 100 mL in a mixture of 1 :1 
ethanokdistilled or deionized water. 

5 Barium chloride solution: dissolve 41 .66 g of BaCI 2 (48.86 g of BaCI 2 ■ H 2 0) in 
distilled or deionized water and make to 200 mL to give a ~1 M solution. 

6 Titrant: use exactly 1 .000 M HCI. 

The following reagents are for use with the two endpoint titration procedure in combination 
with a pH meter or autotitrator. 


7 Tris standard solution: dissolve 2.8000 g of Tris (hydroxy-methyl)-amino-methane 
(MW= 121 .14) in distilled or deionized water and make to 100 ml_. 

8 -0.5 M HCI: dilute 100 mL of cone. HCI to 2 L with distilled or deionized water. 

21.4.3 Oxidation Procedure 

Soil sample (ground to <0.15 mm) weights are limited to 2.0 g. Liquid samples up to 5 mL 
can be digested without pretreatment; larger samples must be evaporated to <5 mL in the 
digestion tube at 100°C. When liquid samples are processed, the temperature of the digest 
must be limited to 120°C. 

For samples containing up to 10% carbonates, 3 mL of pretreatment acid is added per gram 
of soil. The pretreatment is done in the digestion tube by shaking them uncapped for 60 min 
on a reciprocal or orbital shaker set at slow speed. The water added with the acid limits the 
digestion temperature to 120°C. 

1 Place samples into the bottom of the digestion tubes with a long spatula and then 
pretreat to remove carbonates. 

2 Approximately 1 g of K 2 Cr 2 7 is added using a long glass funnel. Add 25 mL of 
the digestion acid mixture and quickly insert the CO2 trap (vial containing 

3 Tightly cap the tubes and place in a digestion block preheated to 1 50°C (1 20°C for 
wet samples) for 2 h. 

4 Remove the tubes from the block and after 1 2 h, remove and titrate contents of the 
C0 2 trap. 


The amount of NaOH in the trap limits the amount of CO2 that can be absorbed. Using 1 mL 
of 2 M NaOH in a 6 mL capacity vial allows the titration to be made directly in the vial. 

1 mL of 2 M NaOH will trap 12 mg of CO2-C, but absorption efficiency drops before this 
maximum is approached. 

21.4.5 Titration Procedure 

Carbonic acid trapped in the NaOH can be titrated by the direct, two endpoint method, or by 
back titration. 

Back Titration Procedure 

7 Add 2 mL of 1 M BaCI 2 solution to the NaOH to precipitate BaC0 3 . 

2 Add approximately five drops of the thymolphthalein indicator solution and titrate 
the NaOH with 1.000 M HCI using a microburette accurate to 0.001 mL. Four 
blanks per 40 tube digestion batch should be included. 



The OC content of the soil or plant material is calculated as 

g OC kg^'sample = (mL HC1 blank - mL HC1 sample) x 6/weight sample (21.9) 
since 2 mol of OH are equivalent to 1 mol (12 g) of C and molarity of the acid is 1.00. 
Two Endpoint Titration Procedure 

/ Pipette accurately 20.0 mL of Tris standard into a titration vessel and titrate with 
~0.5 M acid to pH 4.7. Perform standardization of acid at least three times. 

2 Three blanks of the NaOH solution and NaOH traps are then titrated against 
standardized 0.5 M HCI as follows. 

j Titrate each solution slowly using standardized 0.5 M HCI to pH 8.3 and note 
volume (7]). Continue titration to pH 3.8 and note volume (T 2 ). 

If using an auto burette, the speed of the titrations and endpoints will need to be optimized 
for the burette and strength of acid to ensure the endpoints at pH 8.3 and 3.8 are not overshot. 

Molarity of HCl(M H ci) = 0.23114 x 20/mean titre HCI 
g C kg -1 sample = (Ti — T 2 ) x M H ci x 6/weight sample 

Allison, L.E., Bollen, W.B., and Moodie, CD. Ball, D.F. 1964. Loss-on-ignition as 

1965. Total carbon. In C.A. Black et al. Eds., of organic matter and organic carbon in non- 

Mctlwds of Soil Analysis, Part 2 — Chemical and calcareous soils. J. Soil Sci. 15: 84-92. 

Microbiological Pro/terries. American Society of 

Agronomy, Madison, WI, pp. 1346-1366. Dalai, R.C. 1979. Simple procedure for the 

determination of total carbon and its radioactiv- 
Amato, M. 1983. Determination of carbon 12 C ity in soils and plant materials. Analyst 101: 
and 14 C in plant and soil. Soil Biol. Biochem. 151-154. 

Giovannini, G., Poggio, G., and Sequi, P. 1975. 
Baldock, J.A., Masiello, C.A., Gelinas, Y., and Use of an automatic CHN analyser to determine 
Hedges, J.I. 2004. Cycling and composition of organic and inorganic carbon in soils. Commun. 
organic matter in terrestrial and marine ecosys- Soil Sci. Plant Anal. 6: 39^-9. 
terns. Mar. Chem. 92: 39-64. 

Heanes, D.L. 1984. Determination of total 
Baldock, J.A. and Nelson, P.N. 1998. Soil organic organic-C in soils by an improved chromic acid 
matter. In M. Sumner, Ed., Handbook of Soil digestion and spectrophotometric procedure. 
Science. CRC Press, Boca Raton, FL, pp. B25-B84. Commun. Soil Sci. Plant Anal. 15: 1191-1213. 


Jackson, M.L. 1958. Soil Chemical Analyst 
Prentice Hall, Englewood Cliffs, NJ. 

Piper, C.S. 1944. Soil and Plant Analysis. The 
University of Adelaide, Adelaide, SA, Australia. 

Rayment, G.E. and Higginson, F.R. 1992. Austra- 
lian Laboratory Handbook of Soil and Water 
i Vl< Inkata Piess, Melbourne, 

Kalembasa, S.J. and Jenkinson, D.S. 1973. A Victoria, Australia. 

comparative study of titrimetric and gravimetric 

methods for the determination of organic carbon in Schollenberger, C.J. 1945. Determination of soil 

soil. ./. Sci. FoodAgric. 24: 1085-1090. organic matter. Soil Sci. 59: 53-56. 

Lowther, J.R., Smethurst, P.J., Carlyle, J.C., and Skjemstad, J.O. 1992. Genesis of podzols on 

Nambiar, E.K.S. 1980. Methods for determining coastal dunes in southern Queensland. III. The 

organic carbon in podsolic sands. Commun. Soil role of aluminium-organic complexes in profile 

Sci. Plant Anal. 21: 457-470. development. Aust. J. Soil Res. 30: 645-665. 

MacCarthy, P., Malcolm, R.L., Clapp, C.E., and 
Bloom, P.R. 1990. An introduction to soil humic 
substances. In P. McCarthy et al. Eds., Humic Sub- 
stances in Crop and Soil Science: Selected Read- 
ings. Soil Science Society America, Madison, WI, 
pp. 1-12. 

Merry, R.H. and Spouncer, L.R. 1988. The meas- 
urement of carbon in soils using a microprocessor- 
controlled resistance furnace. Commun. Soil Sci. 
Plant Anal. 19:707-720. 

Morris, G.F. and Schnitzer, M. 1967. Rapid 
determination of carbon in organic matter by 
dry-combustion. Can. J. Soil Sci. 47: 143-144. 

Nelson, D.W. and Sommers, L.E. 1982. Total 
carbon, organic carbon, and organic matter. In 
A.L. Page et al. Eds., Methods of Soil Analysis, 

Part2 ' . ,, ' dM rob logical Properties, 
2nd edn. American Society of Agronomy, Soil Sci- 
ence Society America, Madison, WI, pp. 539-579. 

Oades, J.M. 1988. The retention of organic n 
in soils. Biogeochemistry 5: 35-70. 

Snyder, J.D. and Trofymow, J. A. 1984. A rapid 
accurate wet oxidation diffusion procedure for 
determining organic and inorganic carbon in 
plant and soil samples. Commun. Soil Sci. Plant 
Anal. 15: 587-597. 

Stevenson, F.J. 1994. Humus Chemistry. Genesis, 
Composition, Reactions. 2nd edn. John Wiley & 
Sons, New York, NY. 

Verardo, D.J., Froelich, P.N., and Mclntyre, A. 
1 990. Determination of organic carbon and nitrogen 
in marine sediments using the Carlo Erba NA- 
1500 analyser. Deep-Sea Res. 37: 157-165. 

Walkley, A. 1947. A critical examination of a rapid 
method for determining organic carbon in soils — 
effect of variations in digestion conditions and of 
inorganic soil constituents. Soil Sci. 63: 251-264. 

Walkley, A. and Black, I. A. 1934. An examin- 
ation of the Degtjareff method for determining 
soil organic matter, and a proposed modification 
of the chromic acid titration method. Soil Sci. 37: 



Chapter 22 
Total Nitrogen 

P.M. Rutherford, W.B. McGill, and J.M. Arocena 

University of Northern British Columbia 
Prince George, British Columbia, Canada 

C.T. Figueiredo 

University of Alberta 
Edmonton, Alberta, Canada 


Total soil N includes all forms of inorganic and organic soil N. Inorganic N includes soluble 
forms (e.g., N02~ and N03~), exchangeable NH4+, and clay-fixed nonexchangeable NH4+. 
Organic N content includes numerous identifiable and nonidentifiable forms (Stevenson 
1986) and can be determined by the difference between total soil N and inorganic soil N 
content. Total N analyses may be divided into two main types: (i) wet digestion (e.g., 
Kjeldahl method) or (ii) dry combustion (e.g., Dumas method). Wet digestion techniques 
involve conversion of organic and inorganic N to NH 4 + in acid and its subsequent meas- 
urement. Salts (e.g., K2SO4) and catalysts (e.g., Cu) are usually added to increase digestion 
temperatures and accelerate oxidation of organic matter (Bremner 1996). The dry combus- 
tion method normally involves an initial oxidation step followed by passage of the gases 
through a reduction furnace to reduce NO A to N2. The quantity of N2 is usually determined 
using a thermal conductivity detector. Near-infrared reflectance spectrometry has recently 
been used for the determination of total soil N (Chang and Laird 2002), but the method will 
not be described here. 

The Dumas method is becoming increasingly common due to greater availability and 
simplicity of modern automated instruments, which can determine C, H, N, or S on the 
same sample, and O with a simple modification. Modern systems are available in various 
configurations and have improved accuracy and precision for total N determination over 
earlier models (Bremner 1996). Dry combustion instruments for total N may be connected 
in-line with an isotope ratio mass spectrometer for simultaneous analyses of 15 N (Minagawa 
et al. 1984; Marshall and White way 1985; Kirsten and Jansson 1986). Bellomonte et al. 
(1987) concluded that the automated Dumas procedure was comparable to Kjeldahl analysis 
for heterogeneous substrates. They reported, using a commercial Dumas system for total N 
analysis, coefficients of variation of 0.79% for cereal flour and 1.08% for meat. Reports 
since then confirm total N in plant materials, feeds, excreta, carcasses, and other agricultural 


materials determined by dry combustion to be comparable to or slightly greater than by 
Kjeldahl digestion (Matejovic 1995; Etheridge et al. 1998; Schindler and Knighton 1999; 
Marco et al. 2002). Results are slightly more variable with soils. While results for total soil N 
have been reported as comparable for Kjeldahl and dry combustion methods in some studies 
(Artiola 1990; Yeomans and Bremner 1991), total N determined by dry combustion was 
found to be slightly lower but proportional to Kjeldahl determination by Kowalenko (2001). 
Others have found total soil N by dry combustion to be slightly greater than conventional 
Kjeldahl digestion (McGeehan and Naylor 1988; Vittori Antisari and Sequi 1988). High 
NC>3~ concentrations in sample materials contribute to lower total N by the Kjeldahl method 
if pretreatments to include NC>3~ (see below) are not used (Matejovic 1995; Watson and 
Galliher 2001); however, NC>3~ alone may not account for lower Kjeldahl values (Simonne 
et al. 1998). 

Kjeldahl procedures are still widely used for total N determination. Although some fixed or 
intercalary NHt + is normally included in Kjeldahl digestion, it may not be quantitatively 
extracted from soils with a high proportion of their N constituted as fixed NH/t + . In such 
cases, an HF-HC1 pretreatment, as described by Bremner (1996), may be necessary to free 
intercalary NH4+. Corti et al. (1999) reported a method to measure fixed NHzt + using a 
Kjeldahl digestion followed by distillation, and a digestion of the residue with 5 M HF: 1 M 
HC1 and a second distillation to quantify strongly fixed NH4+. Vittori Antisari and Sequi 
(1988) reported that both dry combustion and microwave digestion with HF-HC1, H3BO3, 
and H2O2 followed by micro-Kjeldahl distillation were effective at including fixed NH4+ in 
total N analysis. 

Given the time involved in Kjeldahl analyses, efforts have been made to speed up digestions. 
One such example is the peroxy method, which replaces K2SO4 and metal catalyst with 
peroxymonosulfuric acid (H2SO5), and involves carbonizing the sample in H2SO4 before 
adding the peroxy reagent (viz. H2SO4 + H2O2). It is 25 times faster than conventional 
Kjeldahl procedures and fully recovers the N of a variety of plant materials and one of the 
most refractory organic compounds, nicotinic acid (Hach et al. 1985). To enhance safety and 
improve speed, Hach et al. (1987) developed a system using a Vigreux fractionation head to 
simplify the addition of the peroxy reagent, and to maintain constant residual H2O2 in the 
digestion solution. A procedure for soils described by Christianson and Holt (1989) takes 
only 38 min; however, N recovery from six soils ranged from 89% to 98% compared to 
Kjeldahl digestion. Further investigation on soil materials would be warranted. Mason et al. 
(1999) used microwave heating to accelerate digestion using only H2SO4 and CUSO4 for 
twofold reduction in time for soil samples. Their system can perform six digests simul- 
taneously. Although microwave-assisted digestion has been widely adopted for sample 
preparation for metal analyses (Smith and Arsenault 1996), it has not received wide 
application for digestion of soils for total N analysis. 

The total time required for Kjeldahl determination of N can be reduced considerably by using 
automated colorimetric analysis of NH4+. The most common method uses the Berthelot or 
indophenol reaction, which is specific for ammonia, and is well-documented (Searle 1984). 
The method has similar results and precision as the distillation procedure (Schuman et al. 
1973). Mason et al. (1999) used the method following microwave-assisted digestion of 
soil. A manual version of the Berthelot reaction has been used to quantify NH 4 + from 
Kjeldahl digestion of soils (Wang and Oien 1986). 

The presence of N03~ can be a concern because unmodified Kjeldahl digestion recovers 
some, but not all, N03~, thereby precluding the addition of N03~ from a separate analysis to 


the Kjeldahl total N values (Goh 1972; Wikoff and Moraghan 1985; Bremner 1996). Hence, 
methods were developed to include NC>3~ in Kjeldahl digestions. Pruden et al. (1985) 
proposed pretreatment of soil with Zn and CrK(S04) 2 to reduce NC>3~ to NH 4 + before 
proceeding with normal Kjeldahl digestion. Dalai et al. (1984) proposed adding sodium 
thiosulfate to the digestion mixture to reduce NC>3~ or NC"2~. The method requires no 
pretreatment and is satisfactory with wet or dry samples. DuPreez and Bate (1989) reported 
that phenyl acetate added to dry samples resulted in quantitative recovery of NC>3~ or NC>2~. 
Phenyl acetate reacts with NC>3 _ or N02~ under acidic conditions to form nitrophenolic 
compounds from which N is fully recovered. The procedure requires no additional reductant 
or pretreatment but is suitable for dry samples only. 

Selection of the most suitable combination of variables for the Kjeldahl method must be 
based on local requirements and facilities. Digestion options include H2SO4 with or 
without H2O2, heating mantles or digestion blocks, macro- or semimicro digestion, inclu- 
sion or omission of NCh" plus NC>3~. Subsequent measurement of NH4+ may use the 
Berthelot reaction, NH.4+ electrode, diffusion in digestion tubes or Conway dishes, steam 
distillation directly from digestion tubes or from standard taper flasks, macro- or semi- 
micro distillation, titration with an indicator or using automated titrators. Several 
semiautomatic to fully automatic distillation systems are now commercially available 
and allow for very rapid analysis. Digestion in tubes using block heaters with the tem- 
perature controlled electronically is now common. Several modern infrared digestion 
systems are available and have much faster heat-up and cool-down times than traditional 
aluminum block systems. 

The choice here has been to present details of modern methods that could be widely used, to 
introduce special purpose methods by way of comments, and to provide citations for older or 
classical methods. Micro-Kjeldahl digestion procedures are given with and without steps to 
include N02~ and N03~. Usually NCh - plus NO3" is a negligible component of soil total N, 
and procedure given in Section 22.2 should be satisfactory. Macro-Kjeldahl procedures are 
little used nowadays because of the cost and disposal of chemicals used in the digestion, and 
the high precision of micro-Kjeldahl and Dumas procedures. 




This method is appropriate for total N determination on samples of surface soil horizons in 
which the N03~ and N02~ contents are negligible. If used with samples containing 
significant amounts of NC"3~ or N02~, the results will be higher than for the fixed NH 4 + 
plus organic N content alone, but lower than for total N including NC>3~ and N02~. This 
method is not recommended for analysis of total N in soil samples from 15 N tracer studies 
because of the significant influence of highly labeled N02~ or NC"3~ on 15 N analyses. The 
method outlined in Section 22.3 is recommended for such samples. 

22.2.1 Materials and Reagents: Digestion 

1 Heating block with digestion tubes, timer, and temperature controller (see Section 
22.2.5). The block must be capable of maintaining a temperature of 360°C for up 
to 5 h. Blocks holding 40 tubes (20 mm OD x 350 mm long) calibrated to hold 
0.1 L are commonly used for micro-Kjeldahl digestions. 


2 Air condenser designed to fit over the digestion tubes in the block (see Section 

3 Concentrated (1 8 M) H 2 S0 4 . 

4 Low N content K 2 S0 4 , CuS0 4 : mixed in mass ratio of 8.8:1 (K 2 S0 4 : 
CuS0 4 • 5H 2 0). Approximately 3.5 g of mix is required per sample. 

5 Hengar granules, both selenized and nonselenized. 
22.2.2 Materials and Reagents: Distillation and Titration 

7 Micro-Kjeldahl steam distillation apparatus (Figure 22.1). See Section 22.2.5. 

2 Steam distillation flasks: 0.5 L round bottom, with 19/38 standard taper ground 
glass joint. 

3 NaOH: 10 M and 0.1 M, prepared in C0 2 -free deionized water. 

Round bottom 

0.6 cm Thick 
plywood blocks 

FIGURE 22.1. Steam distill; 


4 Boric acid (2% w/v) plus indicator: place 80 g of boric acid (H3BO3) powder into 
a 0.25 L beaker. Add -20-40 ml_ of H 2 and mix with a glass rod to wet all the 
H3BO3. Pour into ~3 L of H 2 in a 4 L flask and stir with an electric stir rod. Once 
wet, the H3BO3 dissolves readily. Add 80 ml_ of mixed indicator prepared as 
follows: 0.099 g bromocresol green and 0.066 g methyl red dissolved in 100 ml_ 
ethanol. Add 0.1 M NaOH cautiously until the solution turns reddish-purple 
(pH 4.8-5.0). Make up to 4 L with deionized H 2 and mix thoroughly. 

5 Graduated beakers: 100 mL. 

5 Burette: 1 mL graduated at 0.02 or 0.01 mL intervals. A magnetic stirrer is desirable. 

7 H 2 S0 4 : 0.01 M (standardized). 

22.2.3 Procedure: Digestion, Distillation, and Titration 

7 Place sample, containing about 1 mg N, in a dry digestion tube. This will usually 
vary from 0.25 to 2.0 g. 

2 Add 2 mL deionized H 2 (3 mL if using 2 g soil) and swirl to wet all the soil. 

3 To each tube add 3.5 g of K 2 S0 4 : CuS0 4 , mix. 

4 Add one selenized and one nonselenized Hengar granule. 

5 Add 10 mL concentrated H 2 S0 4 . 

5 Place the digestion tubes into the digestion block. 

j Program the block to raise the temperature to 220°C and maintain it there for 1 .5 h. 
Digestion will start and water will be removed during this time. 

g After 1.5 h of digestion at 220°C, put the air condensers onto the digestion tubes in 
the block. 

9 Program the block to raise the temperature to 360°C and maintain it there for 3.5 h. 

10 After digestion is complete, cool the samples overnight in the block or on a 
fiberglass pad. 

7 7 Remove the air condenser and rinse with water. 

72 Slowly and with swirling, add 25 mL deionized water to each cooled digestion 
tube. Vortex the sample to dissolve salts that may have solidified during cooling. If 
all the material does not enter into suspension, warm gently until it does. Transfer 
the sample quantitatively, with three washes of deionized water, to a 0.5 L round- 
bottom distillation flask. 

73 With the condensers on, connect the distillation flask to the steam distillation 
apparatus; secure with a clamp. 


74 Open the steam supply to the distillation head to allow steam into the tubing. Be 
sure the drain line is already open so that steam can exit to the drain. 

75 Place a 1 00 ml_ graduated beaker with 5 ml_ of 2% H3BO3 under the condenser so 
that the tip of the condenser is immersed in the H3BO3. 

76 Very slowly add an excess (usually 30 ml_; see Section 22.2.5) of 10 M NaOH 
through the distillation head. Do not completely empty the NaOH reservoir, 
otherwise NH 3 may be lost through the stopcock. 

77 Close the pinch clamp, or stopcock, going to the steam drain; this directs steam 
into the distillation flask. The steam generation rate should be such that the 
distillate is collected at about 6 ml_ min~ 1 . Collect 40 ml_ of distillate. 

7g Open the pinch clamp to the steam drain and remove the distillation flask 
then close the pinch clamp to the distillation head. This sequence is important 
to prevent steam burns and drawback of fluid from the distillation flask into the 
steam line. 

79 Wash the tip of the condenser into the beaker. 

20 Titrate the distillate with 0.01 M H2SO4. The color change at the endpoint is from 
green to pink (pH w 5.4). 

22.2.4 Calculations 

One mL of 1 M H 2 S0 4 is equivalent to 28.01 mg of N. 

, (mL sample - mL blank) x M x 28.01 

Total N, g kg~' = (22.1) 

oven-dry mass of soil sample (g) 

where M is the molarity of standard H2SO4, mL sample is the volume of standard H2SO4 
used during titration of sample, and mL blank is the volume of standard H2SO4 used during 
titration of blank. 

The blank is included from the digestion step onward; it contains all materials excluding a 
soil sample. 


7 Soil samples should be dried (usually air-drying) and ground to pass a 100 mesh 
(1 50 |xm) sieve. 

2 Heating blocks and tubes supplied by Tecator or Technicon have been found 
satisfactory for Kjeldahl digestions in our laboratory. The air condenser described 
by Panasiuk and Redshaw (1 977) can be constructed by a competent glassblower. 
Equivalent devices are available from Tecator. A variety of sophisticated digestion 
systems are also available from VELP, Gernhart, and other manufacturers (e.g., 
Fisher, VWR, Cole Parmer, or other suppliers). Some of these units use infrared 


digestion to greatly decrease heating and cooling times over traditional aluminum 
block systems. 

The K 2 S0 4 : CuS0 4 mix can be prepared in the laboratory, obtained as a loose mix 
or in appropriately sized packages (-3.5 g) from commercial suppliers (e.g., 
Kjeltabs — trademark of Tecator, Inc.). Bulk mixes should use low N materials 
and be kept tightly sealed during storage to avoid absorption of moisture and 

A variety of manual, semiautomatic to fully automatic distillation systems are 
available from a variety of manufacturers (e.g., Labconco, Gerhardt, VELP). 
These systems are quite rapid, often reducing distillation times to ~2 min 
from ~7 to 8 min for the setup shown in Figure 22.1. Distillation systems 
similar to those in Figure 22.1 are described by Bremner (1996) and Bremner 
and Breitenbeck (1983). Commercial systems specifically designed for use with 
digestion tubes are available. We have used two distillation heads, each 
supplied with steam from a 5 L round-bottom boiling flask heated by a 600 W 
heating mantle. Several dozen Hengar granules are placed in each flask. 
Concentrated phosphoric acid (about 2 ml_) is added to each boiling flask to 
absorb NH 3 . 

We have eliminated sample transfer from the digestion tube to a distillation 
flask by doing digestions in a 0.25 L digestion tube designed for a block 
that holds 20, rather than 40, tubes. The distillation head was modified by 
attaching a rubber stopper with a hole through which the standard taper joint of 
the distillation head fits. The 0.25 L digestion tube is attached to the distillation 
unit by fitting the end over the rubber stopper. It is held securely in place with a 
clamp, allowing distillation directly from the digestion tube. The distillation 
system described by Bremner and Breitenbeck (1983) uses tubes designed for 
the 40 tube blocks. 

The NaOH must be added slowly and carefully to avoid violent bubbling that 
would force the liquid into the condenser and contaminate the distillation head. 
The amount of NaOH needed varies with the amount of H2SO4 consumed during 
digestion of the sample. Consumption of H 2 S0 4 varies with the amount of soil 
organic matter and reduced minerals present; e.g., 1 g of C consumes 10 mL of 
H2SO4 (Bremner 1996). 

Distillation can be replaced by autoanalyzer analysis of the digested sample. When 
we use this approach, the digested sample is diluted to 0.1 L followed by an 
autoanalyzer method for colorimetric measurement of NH 4 + (Smith and Scott 1 991 ). 

Use of an automatic titrator can improve consistency and eliminate the need to 
mix an indicator into the boric acid solution. 

The above Kjeldahl digestion method does not quantitatively recover fixed NH 4 + 
in most soils. For total N analysis of soils with a high proportion of their N as 
fixed NH 4 +, an HF-HCI modification as described by Bremner (1996) may be 
necessary to release fixed NH 4 +. 





This is the method of choice for total N analysis of samples of surface soil horizons 
containing an appreciable quantity of N as N0 2 ~ or NC>3~. Because of the significant 
influence of highly labeled N0 2 ~ or NC>3~ on 15 N analyses, this method is recommended 
for analysis of total N in all soil samples from 15 N tracer studies. This method is the same as 
in Section 22.2, except for the addition of a pretreatment to oxidize N0 2 ~ to NC>3~ and then 
to reduce the N0 3 ~ to NH 4 +. 

22.3.1 Materials and Reagents: Pretreatment and Digestion 

1 All items from Section 22.2.1 , plus the following: 

2 Potassium permanganate solution: dissolve 50 g KMn0 4 in 1 L deionized H 2 0; 
store in an amber bottle. 

3 H 2 S0 4 , 9 M: dilute concentrated H 2 S0 4 to twice its volume with deionized H 2 0. 

4 Fe powder; finer than 1 00 mesh sieve. 

5 N-octyl alcohol. 

22.3.2 Materials and Reagents: Distillation and Titration 
All items from Section 22.2.2. 

22.3.3 Procedure: Pretreatment to Reduce N0 2 ~ and N0 3 ~ to NH 4 + 

7 Place sample, containing about 1 mg N, in a dry digestion tube. Usually this is 
0.25-2.0 g of soil. 

2 Add 2 ml_ deionized H 2 (3 ml_ if using 2 g soil) and swirl to wet all the soil. 

3 Add 1 ml_ KMn0 4 and swirl for 30 s. 

4 Hold the digestion tube at a 45° angle and very slowly pipette 2 ml_ dilute H 2 S0 4 . 

5 Allow to stand for 5 min. 

g Add one drop N-octyl alcohol (to control frothing). 

j Add 0.5 g reduced Fe using a scoop, through a dry, long-stemmed funnel or thistle 
funnel tube. 

q Immediately cover the digestion tube with an inverted 25 ml_ beaker or 50 mL 
Erlenmeyer flask inverted to prevent loss of water. 

9 Swirl to bring the Fe into contact with the acid. 

1q Allow to stand (about 15 min) until strong effervescence has ceased. 

77 Place digestion tubes into the digestion block and program it to raise the 
temperature to 100°C and hold it there for 1 h. 

72 Cool the tubes before proceeding to digestion. 

22.3.4 Procedure: Digestion, Distillation, and Titration 

Follow steps 3 to 20 inclusive, of Section 22.2.3. 

22.3.5 Calculations 

Use Equation 22.1, described in Section 22.2.4. 


7 The KMn0 4 oxidizes N0 2 ~ to N0 3 ~, which is reduced to NH 4 + by reduced Fe. 

2 N-octyl alcohol is added to reduce frothing. 

3 Goh (1972) concluded that it was not necessary to include the permanganate 
pretreatment when reduced iron is used as a reductant in the procedure to 
include N0 3 _ . 

4 See Section 22.2.5 for additional important information. 


Several automated Dumas systems are available (Kirsten and Jansson 1986; Tabatabai and 
Bremner 1 99 1 ; Bremner 1 996). Many systems combust the sample in a stream of pure O2 at high 
temperatures, producing NO A and N2. An aliquot of gas is carried by pure He into a reduction 
zone where elemental Cu reduces NO., to N2, which is subsequently measured by a thermal 
conductivity detector. Other systems convert total N to N 2 by fusing the sample in a graphite 
crucible at very high temperatures in a He atmosphere, followed by determination of N 2 by gas 
chromatography (Bremner 1996). Numerous configurations are available depending on what 
other elements (e.g., C, H, S, O) are also to be determined. Since Dumas procedures are quite 
variable and instrument dependent, it is not possible to provide a generic methodology here. 

Compared to Kjeldahl, Dumas techniques have the advantage of requiring less laboratory 
space, provide rapid analysis, require less chemical reactants, do not produce noxious fumes 
or hazardous chemical wastes, and include all forms of N without lengthy pretreatments 
(Bellomonte et al. 1987; Vittori Antisari and Sequi 1988). They are suitable for 15 N tracer 
studies when linked by a continuous flow interface from the nitrogen analyzer to an isotope 
ratio mass spectrometer (Fiedler and Proksch 1975; Minagawa et al. 1984; Marshall and 
Whiteway 1985). For tracer studies, they avoid digestion, distillation, titration, evaporation, 
and subsequent oxidation of NH3 to N 2 . 

Sample variability is a concern with combustion techniques because of the small sample size 
required in some instruments (<50 mg). Schepers et al. (1989) recommended that plant and 


soil samples should be ball milled before combustion analysis. Arnold and Schepers (2004) 
reported on a simple roller-mill grinding procedure as an alternative to ball milling plant and 
soil samples. See Bremner (1996), Kowalenko (2001), Perez et al. (2001), and Wang et al. 
(1993) for further information on grinding and sample preparation. 

We recommend that soil samples be air-dried and passed through a 2 mm (10 mesh) sieve. 
Subsamples are then finely ground using a ball-mill such as the Brinkmann, Mixer Mill, model 
MM2. With the grinder set to its maximum setting, soils take 1.5-2 min to grind to a fine 
powder (<100 mesh). Ensure grinding capsules and balls are well cleaned before attempting 
the next sample. If samples are resinous, grind a small scoop of pure silica sand between 
samples (for «30 s) to help clean the resin from the ball and capsules; vacuum and wipe 
capsules and balls before grinding the next sample. After fine grinding, samples should be 
dried overnight at 60°C-70°C and cooled in desiccators before weighing for analysis. 

Arnold, S.L. and Schepers, J.S. 2004. A simple 
roller-mill grinding procedure for plant and soil 
samples. Commun. Soil Sci. Plant Anal. 35: 


Corti, G., Agnelli, A., and Ugolini, F.C. 1999. A 
modified Kjeldahl procedure for determining 
strongly fixed NH 4 +-N. Eur. J. Soil Sci. 50: 


Artiola, J.F. 1990. Determination of carbon, nitrogen 
and sulfur in soils, sediments and wastes: a compara- 
tive study. Int. J. Environ. Chem. 41: 159-171. 

Bellomonte, G., Costantini, A., and Giammarioli, S. 
1987. Comparison of modified automatic Dumas 
method and the traditional Kjeldahl method for 
nitrogen determination in infant food. J. Assoc. 
Off. Anal. Chem. 70: 227-229. 

Bremner, J.M. 1996. Nitrogen — total. In D.L. 
Sparks et al, Eds u, th ds oj mil Analysis, Part 
3 — Chemical Methods. Soil Science Society of 
America, American Society of Agronomy, Madi- 
son, WI, pp. 1085-1 121. 

Bremner, J.M. and Breitenbeck, GA. 1983. A 
simple method for determination of ammonium 
in semimicro-Kjeldahl analysis of soils and plant 
materials using a block digester. Commun. Soil 
Sci. Plant Anal. 14:905-913. 

Chang, C.W. and Laird, DA. 2002. Near-infrared 
reflectance spectroscopic analysis of soil C and 
N. Soil Sci. 167: 110-116. 

Christianson, C.B. and Holt, L.S. 1989. Rapid 
digestion procedure for the determination of 
total N and nitrogen- 15 content of soils. Soil Sci. 
Soc. Am. J. 53: 1917-1919. 

Dalai, R.C., Sahrawat, K.L., and Myers, R.J.K. 
1984. Inclusion of nitrate and nitrite in the Kjeldahl 

nitrogen determination of soi Is and plant materials 
using sodium thiosulphate. Commun. Soil Sci. 
Plant Anal. 15: 1453-1461. 

DuPreez, D.R. and Bate, G.C. 1989. A simple 
method for the quantitative recovery of nitrate-N 
during Kjeldahl analysis of dry soil and plant sam- 
ples. Commun. Soil Sci. Plant Anal. 20: 345-357. 

Etheridge, R.D., Pesti, G.M., and Foster, E.H. 
1998. A comparison of nitrogen values obtained 
utilizing the Kjeldahl nitrogen and Dumas 
combustion methodologies (Leco CNS 2000) on 
samples typical of an animal nutrition analyt- 
ical laboratory. Anim. Feed Sci. Technol. 73: 

Fiedler, R. and Proksch, G. 1975. The determin- 
ation of nitrogen-15 by emission and mass spec- 
trometry in biochemical analysis: a review. Anal. 
Chim. Acta 78: 1-62. 

Goh, K.M. 1972. Comparison and evaluation of 
methods for including nitrate in the total determin- 
ation of soils. J. Sci. FoodAgric. 23: 275-284. 

Hach, C.C., Bowden, B.K., Kopelove, A.B., and 
Brayton, S.V. 1987. More powerful peroxide 


Kjeldahl digestion method. ./. Assoc. Off. Anal. 
Chem. 70: 783-787. 

Hach, C.C., Brayton, S.V., and Kopelove, A.B. 
1985. A powerful Kjeldahl nitrogen method 
using peroxymonosulfuric acid. J. Agric. Food 
Chem. 33: 1117-1123. 

Kirsten, W.J. and Jansson, K.H. 1986. Rapid and 
automatic determination of nitrogen using flash 
combustion of large samples. Anal. Chem. 58: 

Kowalenko, C.G. 2001. Assessment of LECO 
CNS-2000 analyzer for simultaneously mea- 
suring total carbon, nitrogen and sulfur in soil. 
Commun. Soil Sci. Plant Anal. 32: 2065-2078. 

Marco, A., Rubio, R., Compano, R., and 
Casals, 1. 2002. Comparison of the Kjeldahl method 
and a combustion method for total nitrogen deter- 
n animal feed. Talanta 57: 1019-1026. 

Marshall, R.B. and Whiteway, J.N. 1985. Auto- 
mation of an interface between a nitrogen ana- 
lyzer and an isotope ratio mass spectrometer. 
Analyst 110: 867-871. 

Mason, C.J., Coe, G., Edwards, M., and Riby, 
P.G. 1999. The use of microwaves in the acceler- 
ation of digestion and colour development in the 
determination of total Kjeldahl nitrogen in soil. 
Analyst 124: 1719-1726. 

Matejovic, 1. 1995. Total nitrogen in plant-material 
determined by means of dry combustion — a pos- 
sible alternative to determination by Kjeldahl 
digestion. Commun. Soil Sci. Plant Anal. 26: 

McGeehan, S.L. and Naylor, D.V. 1988. Auto- 
mated instrumental analysis of carbon and nitro- 
gen in plant and soil samples. Commun. Soil Sci. 
Plant Anal. 19: 493-505. 

Minagawa, M., Winter, D.A., and Kaplan, I.R. 
1984. Comparison of Kjeldahl and combustion 
methods for measurement of nitrogen isotope 
ratios in organic matter. Anal. Chem. 56: 

Panasiuk, R. and Redshaw, E.S. 1977. A simple 
apparatus used for effective fume control 
during plant tissue digestion using a heating 
block. Commun. Soil Sci. Plant Anal. 8: 411-416. 

Perez, D.V., de Alcantara, S., Arruda, R.J., and 
Meneghelli, N.D.A. 2001. Comparing two 
methods for soil carbon and nitrogen determin- 
ation using selected Brazilian soils. Commun. Soil 
Sci. Plant Anal. 32: 295-309. 

Pruden, G., Kalembasa, S.J., and Jenkinson, D.S. 
1985. Reduction of nitrate prior to Kjeldahl 
digestion. ./. Sci. Food Agric. 36: 71-73. 

Schepers, J.S., Francis, D.D., and Thompson, 
M.T. 1989. Simultaneous determination of total 
C, total N and 15 N in soil and plant material. 
Commun. Soil Sci. Plant Anal. 20: 949-959. 

Schindler, F.V. and Knighton, R.E. 1999. Sample 
preparation for total nitrogen and N-15 ratio 
analysis by the automated Dumas combustion 
method. Commun. Soil Sci. Plant Anal. 30: 

Schuman, G.E., Stanley, M.A., and Knudsen, D. 
1973. Automated total nitrogen analysis of soil and 
plant samples. Soil Sci. Soc. Am. Proc. 37: 480-481. 

Searle, P.L. 1984. The Bertholet or indophenol 
reaction and its use in the analytical chemistry of 
nitrogen. Analyst 109: 549-568. 

Simonne, E.H., Harris, C.E., and Mills, H.A. 
1998. Does the nitrate fraction account for differ- 
ences between Dumas-N and Kjeldahl-N values 
in vegetable leaves? J. Plant Nutr. 21: 2527- 

Smith, F.E. and Arsenault, E.A. 1996. Microwave 
assisted sample preparation in analytical chemis- 
try. Talanta 43: 1207-1268. 

Smith, K.A. and Scott, A. 1991. Continuous-flow, 
flow-injection, and discrete analysis. In K.A. 
Smith, Ed. Soil Analysis: Modern Instrumental 
Techniques, 2nd ed. Marcel Dekker, New York, 
pp. 183-227. 

Stevenson, F.J. 1986. Cycles of Soil: Carbon, 
Nitrogen, Phosphorus, Sulfur, Micronutrients. 
John Wiley & Sons, New York, NY. 

Tabatabai, M.A. and Bremner, J.M. 1991. Auto- 
mated instruments for determination of total car- 
bon, nitrogen, and sulfur in soils by combustion 
techniques. In K.A. Smith, Ed. Soil Analysis: 
Modern Instrumental Techniques. 2nd cd. Marcel 
Dekker, New York, NY, pp. 261-286. 


Vittori Antisari, L. and Sequi, P. 1988. Comparison 
of total nitrogen by four procedures and sequential 
determination of exchangeable ammonium, 
organic nitrogen and fixed ammonium in soil. Soil 
Sci. Soc. Am. J. 52: 1020-1023. 

Watson, M.E. and Galliher, T.L. 2001. Compari- 
son of Dumas and Kjeldahl methods with auto- 
matic analyzers on agricultural samples under 
routine rapid analysis conditions. Commun. Soil 
Sci. Plant Anal. 32: 2007-2019. 

Wang, D., Snyder, M.C., and Bormann, F.H. 
1993. Potential errors in measuring nitrogen con- 
tent of soils low in nitrogen. Soil Sci. Soc. Am. 
J. 57: 1533-1536. 

Wang, L. and Oien, A. 1986. Determination of 
Kjeldahl nitrogen and exchangeable ammonium 
in soil by the indophenol method. Acta Agric. 
Scand. 36: 60-70. 

Wikoff, L. and Moraghan, J.T. 1985. Recovery of 
soil nitrate by Kjeldahl analysis. Commun. Soil 
Sci. Plant Anal. 16: 923-929. 

Yeomans, J.C. and Bremner, J.M. 1991. Carbon 
and nitrogen analysis of soils by automated com- 
bustion techniques. Commun. Soil Sci. Plant 
Anal. 22: 843-850. 


Chapter 23 

Chemical Characterization 

of Soil Sulfur 

C.G. Kowalenko 

Agrii allure and Agri-rood Canada 
Agassiz, British Columbia, Canada 

M. Grimmett 

Agric ullure ,:nd Agri-rood Canada 
Charlottetown, Prince Edward Island, Canada 


Sulfur is relatively abundant in the terrestrial environment and assumed to be the 15th most 
abundant element (Arnhold and Stoeppler 2004). It is present in the soil in a variety of forms, 
both organic and inorganic, and various valence states (Blanchar 1986), each having 
different chemical, biological, and environmental significance. Various chemical analyses 
have been proposed to measure the various forms of sulfur in soils, for different purposes 
(e.g., soil genesis, plant availability, or environmental assessment). The relative success for 
measuring different forms, however, is limited by available chemical quantification 
methods. The most common basic measurements of soil sulfur forms are total, organic, 
inorganic, and extractable (i.e., plant-available) sulfur. 

23.1.1 Total Sulfur 

Measurement of total sulfur is an important measurement on its own but it is often also 
involved in quantifying specific forms by difference calculation (e.g., total organic S = total 
S minus total inorganic S). Numerous methods have been proposed for determining 
total sulfur, but none are universally accepted (Tabatabai 1982; Blanchar 1986). Almost 
all methods for measuring total soil require two steps: (1) conversion of all sulfur to one form 
and (2) quantification of the resulting form. Methods available for the conversion step 
include ashing (or dry combustion) and wet digestion (Tabatabai 1982; Blanchar 1986). 
Dry ashing includes use of ovens, heating elements, open flames (e.g., fusion), enclosed 
flames (e.g., oxygen flask), or high-temperature combustion using induction or resistance 
furnaces. Wet digestion may be either alkaline or acidic. Numerous sulfur quantification 
methods are available for gases and solutions in oxidized or reduced forms. Gases can be 


quantified directly by infrared, chemiluminescent, coulometric, flame photometric, and other 
methods. Solutions (either absorbed/dissolved gases, liquid digests, or solubilized solids) are 
usually analyzed as sulfide or sulfate by spectrometry (colorimetry, flame emission, atomic 
absorption, etc.), ion-selective electrode, titration, gravimetry, and chromatography. Both or 
either of the conversion and the quantification steps can be automated. X-ray fluorescence 
(Jenkins 1984) measures sulfur in one step. 

Comparisons of methods for determining total sulfur in soils have been conducted (e.g., 
Gerzabek and Schaffer 1986); however, the comparisons have been limited with respect to 
the scope of soils analyzed and the type of methods compared (Tabatabai and Bremner 
1970b; Matrai 1989; Kowalenko 2001). Variable results occurred and no one method could 
clearly be said to give a true estimate of total sulfur (Hogan and Maynard 1984; Kowalenko 
2000). The highest value cannot necessarily be taken as the true value. 

Several studies have found that high-temperature combustion has not resulted in particularly 
satisfactory results for some soil samples, but more recent instrumentation has shown good 
results (Kowalenko 2001). Many combustion units have relatively high detection limits for 
analysis of materials, such as coal, that have substantial concentrations of sulfur. X-ray 
fluorescence requires an adjustment for the organic matter content in the sample (Brown 
and Kanaris-Sotiriou 1969). The success of low-temperature ashing has been variable 
(Tabatabai and Bremner 1970a; Killham and Wainwright 1981). Acid digestion should be 
used with caution, as gaseous losses of sulfur are possible (Randall and Spencer 1980). Dry 
ashing with sodium bicarbonate and silver oxide followed by ion chromatography (IC) 
or hydriodic acid reduction has shown variable results (Tabatabai and Bremner 1970b; 
Tabatabai et al. 1988). A fusion technique proposed for geological samples using sodium 
peroxide followed by IC was not completely satisfactory (Stallings et al. 1988). Hordijk 
et al. (1989) found good agreement for total sulfur measurement in freshwater sediments by 
IC with inductively coupled plasma (ICP) and roentgen fluorescence methods after 
Na 2 C0 3 /KN0 3 fusion. 

23.1.2 Organic Sulfur 

Organic sulfur accounts for most of the sulfur present in the surface horizons of soils (Tabatabai 
1982; Blanchar 1986). A number of methods have been attempted to directly determine organic 
sulfur compounds in soils (Kowalenko 1978), but none have been universally accepted. Meas- 
urement of sulfur-containing amino acids such as methionine, cystine, etc., requires special 
precautions and these compounds do not appear to account for a very large portion of the total 
sulfur present. There has been some success in determining the sulfur content of lipid extracts 
(Chae and Lowe 1981; Chae and Tabatabai 1981), but this fraction also accounts for only a small 
portion of the sulfur in the soil. Sulfur that is present in microorganisms does not constitute a 
specific organic compound and may include inorganic as well as various organic forms. This 
fraction comprises only a small portion of the total sulfur (Strick and Nakas 1984; Chapman 
1987) but may have considerable biological significance. In order to estimate microbial sulfur, 
the method requires the measurement of extractable inorganic sulfate. 

Lowe and DeLong (1963) proposed the determination of carbon-bonded sulfur using a 
digestion with Raney nickel in sodium hydroxide. Although this method was shown to be 
quite specific for carbon-bonded sulfur, and hence most of the organic sulfur in soils, the 
method is not quantitative due to interference problems in soils and soil extracts (Freney et al. 
1970; Scott et al. 1981). The difference between total sulfur and hydriodic acid-reducible 
sulfur appears to provide a better estimate of carbon-bonded sulfur than direct determination 


by Raney nickel digestion. The success of this or other difference approaches to quantifying 
organic sulfur in soils depends on accurate measurements of all fractions involved in the 
calculation. Likewise, determining organic sulfur by subtracting inorganic from total sulfur 
requires accurate measurement of all inorganic forms that are present in the sample. 

23.1.3 Inorganic Sulfur 

Inorganic sulfur is largely in oxidized (sulfate) form in aerobic soils, and in reduced forms 
(sulfide, elemental sulfur, etc.) in anaerobic soils (Tabatabai 1982; Blanchar 1986). No 
single method has been developed to measure total inorganic sulfur that includes reduced 
and oxidized forms. Highly reduced forms of sulfur are not very soluble; therefore, cannot be 
readily extracted for quantification. Methods for directly determining reduced inorganic 
sulfur have been proposed, but not thoroughly evaluated (Barrow 1970; Watkinson et al. 
1987) because these forms are limited in agricultural soils. Zinc-hydrochloric acid distilla- 
tion has been used to measure reduced inorganic sulfur in soils (David et al. 1983; Roberts 
and Bettany 1985), but would not include all inorganic forms (Aspiras et al. 1972). Digestion 
of soil with tin and hydrochloric or phosphoric acid has been proposed for measuring 
sulfide, but this method is not specific to inorganic sulfur (Melville et al. 1971; Pirela and 
Tabatabai 1988). 

Elemental sulfur can occur naturally in some soils such as those that are anaerobic and 
associated with marine or marsh situations, or from aerial depositions (e.g., industrial 
pollution) and fertilizer applications. Measurement of elemental sulfur in soils usually 
requires extraction with an organic solvent (e.g., chloroform, acetone, toluene) followed 
by colorimetry, liquid or gas chromatography, or ICP (Maynard and Addison 1985; Clark 
and Lesage 1989; O'Donnell et al. 1992; Zhao et al. 1996). 

Inorganic sulfate may be present in water in the soil, bound or adsorbed on soil surfaces, as 
relatively insoluble compounds such as gypsum (Nelson 1982), or in association with 
calcium carbonate (Roberts and Bettany 1985). Sulfate is adsorbed on positive charges 
that occur in acidic soils (Tabatabai 1982), although a recent study has shown that sulfate- 
binding mechanisms are complex (Kowalenko 2005). Solution and adsorbed inorganic 
sulfate are assumed to be immediately available for plant uptake. To measure total inorganic 
sulfate in soils, all of these forms would need to be measured. Although there is a good 
theoretical basis for solution and adsorbed pools being present in the soil, there are practical 
limitations in their extraction and subsequent quantification. The choice of the extractant will 
depend on analytical equipment available, form of sulfate (e.g., solution, sorbed) to be 
examined, and type of soil to be analyzed. Numerous solutions have been used for extracting 
combined soluble and adsorbed sulfate including acetates, carbonates, chlorides, phosphates, 
citrates, and oxalates (Beaton et al. 1968; Jones 1986). Most of these studies have focused on 
measurement of plant-available rather than total inorganic sulfate. If only soil solution 
sulfate is to be measured, water would theoretically be sufficient. However, weak calcium 
chloride is often preferred, since it depresses clay and organic matter during extraction 
(Tabatabai 1982). Lithium chloride is also used, since lithium would inhibit microbial 
activity that may mineralize organic sulfur during and after extraction (Tabatabai 1982). 
Adsorbed sulfate (together with solution sulfate) is usually extracted with sodium, potas- 
sium, or preferably, calcium phosphate (Beaton et al. 1968). A concentration of 
500 mg PL -1 is usually adequate to displace sulfate in most soils; however, for soils 
that fix considerable phosphate, 2000 mg P L _1 may be required. Alkaline solutions 
theoretically are effective for extracting adsorbed sulfate, since the adsorption mechanism 
is pH dependent, but would extract additional, highly colored organic materials that cause 


problems for some sulfate quantification methods. Acidic extractants may extract portions 
of gypsum- or carbonate-associated sulfate that may be present in some soils. Buffered 
extractants may result in more consistent results. Preextraction treatment on the sample 
such as air drying will also influence the results (Kowalenko and Lowe 1975; Tabatabai 
1982). Specialized methods are required to measure insoluble sulfate in gypsiferous (Khan 
and Webster 1968; Nelson 1982) or acid-sulfate (Begheijn et al. 1978) soils. It is possible 
that oxidized forms other than sulfate such as thiosulfate, tetrathionate, or sulfite (Nor and 
Tabatabai 1976; Wainwright and Johnson 1980) may be found in soils, but are probably 
present only as intermediates during oxidation or reduction of sulfur. 

There are a number of methods for quantifying sulfate (Patterson and Pappenhagen 1978; 
Tabatabai 1982), but not all are compatible with all soil extract solutions. The method should be 
quantitative, adequately sensitive, free from interferences, and specific for sulfate. Unfortu- 
nately, there does not appear to be a specific, direct colorimetric method to determine sulfate. 
The most frequently used sulfate quantification methods applied to soil extracts include 
precipitation with barium or sulfide analysis after hydriodic acid reduction. There are numer- 
ous variations for barium-precipitation methods, including titrimetric, turbidimetric, gravi- 
metric, and colorimetric methods (Beaton et al. 1968), but all are subject to interferences. The 
hydriodic acid reduction method is quite sensitive and relatively free from interferences, but 
the reduction procedure is time consuming, difficult to automate, and the chemicals are costly. 
The hydriodic acid reagent (Johnson and Nishita 1952; Beaton et al. 1968) reduces both organic 
and inorganic sulfate to sulfide. This method has been used extensively for soil analyses. It is 
quite specific for sulfate (Tabatabai 1982), whether organic or inorganic, therefore, is not 
specific to inorganic sulfate. Various pretreatments have been attempted to make the methods 
specific to inorganic sulfate, but each has distinct limitations (Kowalenko and Lowe 1975). 
Pirela and Tabatabai (1988) have shown that the hydriodic reagent will decompose some 
elemental sulfur, thus, the resulting values should be interpreted accordingly. More recently, 
IC, ICP, and x-ray fluorescence have been used for sulfur analysis of soil extracts (Gibson and 
Giltrap 1979; Tabatabai 1982; Maynard et al. 1987). Although IC is specific for inorganic 
sulfate analysis, it is quite sensitive and not affected significantly by interferences; specialized 
instrumentation and attention to the choice of the extraction salts (concentration and types) are 
required. Inductively coupled plasma spectrophotometry and x-ray fluorescence provide fast 
quantification; however, they include all forms (organic and inorganic) of sulfur. 


Since there is potential for sulfur to be lost by volatilization from hot, acidic solutions, an 
alkaline solution is preferred for digesting soil samples for total sulfur determination. 
A method that uses sodium hypobromite for the digestion developed by Tabatabai and 
Bremner (1970a) is described here. The method is modified from the original by the use 
of a different custom-built reduction/distillation apparatus (Kowalenko 1985) that allows the 
measurement of sulfur as sulfide using bismuth to quantify the resulting sulfide rather than 
methylene blue reaction (Kowalenko and Lowe 1972). The bismuth method combined with 
the modified glassware apparatus has the advantages of short analysis time, versatility for 
types of sulfur analyses, and the equipment requirements are small. The digestion and 
quantification must be done in the same vessel; therefore, an alternate sulfur quantification 
method (e.g., ICP or IC) should not be substituted for the hydriodic acid method without 
it of effectiveness and making appropriate modifications. 


23.2.1 Materials and Reagents 

7 Sodium hypobromite digestion reagent: in a fume hood, add slowly with con- 
stant stirring 3 ml_ of bromine to 100 ml_ 2 M sodium hydroxide. This 
reagent has limited stability, and should be prepared immediately before use 
or at least daily. 

2 Formic acid (90%). 

3 Temperature-controlled digestion block: the block heater (commercially available 
or can be custom-built) must accommodate the sample vessel and be capable of 
maintaining a temperature of 250°C-260°C. 

23.2.2 Apparatus 

The recommended apparatus (Kowalenko 1985) for the sample digestion and subsequent 
sulfur determination by hydriodic acid reduction is custom-built glassware and includes two 
components, the vessel into which the sample is placed (capable of being heated in a block) 
and a part that fits onto this vessel to form an airtight unit that includes the ability to 
dispensing the reducing solution for nitrogen gas flushing (Figure 23.1). The inlet of the gas 
flush is attached to a steady and controllable source of nitrogen gas, and the outlet should 
include an arm with a capillary dropper attached by flexible inert tubing such that the exiting 
gas is bubbled through the sodium hydroxide absorbing solution. The hydriodic acid 
reduction procedure is described in Section 23.3. The apparatus must be supported to 
allow it to be lowered to and lifted from a block to heat the sample vessel at 110°C-115°C 
during the hydriodic acid reduction phase. 

23.2.3 Procedure 

7 Weigh a finely ground (<100 mesh) soil sample directly into the dry sample 
vessel. The sample size (an oven-dry basis is recommended) should be adjusted 
so that the sulfur content is within the range of the calibration standards. For 
example, digest 0.1-0.4 g of Ah horizon of mineral soil or 0.1 g or less of organic 
horizon with a 30 ml_ bismuth sulfide final volume that provides a 0-200 mg S 
range of analysis. 

2 Add 3 ml_ of the sodium hypobromite digestion reagent and thoroughly wet the 
soil sample with the reagent by swirling the tube. After letting the sample stand for 
5 min, evaporate the mixture to dryness in the digestion block at 250°C-260°C, 
then continue heating for an additional 30 min. Remove from heat and allow the 
tube to cool for about 5 min. 

3 Resuspend the digested sample in 1 mL of water by swirling and heating briefly. 
After cooling, add 1 mL of formic acid to eliminate any excess bromine that may 
be present. 

4 Quantify the sulfur content with the hydriodic acid reduction reagent method (as 
described in Section 23.3). 


FIGURE 23.1. Simplified digestion-distillation apparatus for total- or sulfate-S analyses. (From 
Kowalenko, C.G., Commun. SoilSci. Plant Anal., 1 6, 289, 1 985. With permission.) 

23.2.4 Calculations 

Calculate the sulfur content (as mg S kg -1 ) of the sample by taking into consideration the 
oven-dry weight of the sample and using the standard curve produced by the hydriodic acid 
determination of sulfur in the sample (see Section 23.3.3). 


/ The sulfur content of soil samples containing a large amount of organic material 
may be underestimated by this method as described, and sequential digestions, 
and/or longer heating times may be required (Guthrie and Lowe 1984). 

This digestion method can also be used to determine total sulfur content in 
solutions, but the aliquot of the solution should be dried before the digestion is 
conducted (Kowalenko and Lowe 1972). However, an ICP instrument capable of 


sulfur determination could be used as an alternative direct analysis since the 
instrument operates on liquid samples and measures total sulfur. Ion chromato- 
graphy which also works with liquids, measures a specific form of sulfur (e.g., 
sulfate); therefore, would have to be applied to a digested sample to determine 
total sulfur. 


The quantification of sulfate in soil studies has been limited by the lack of an accurate, 
suitable, and direct colorimetric analysis method. As noted earlier, sulfate colorimetric 
methods that have been used are essentially based on precipitation with barium, and, 
hence, all are subject to interference and most lack sufficient sensitivity. The other method 
that has been used widely in soil studies is based on the reduction of sulfate with hydriodic 
acid reagent and sulfur measured on the resulting hydrogen sulfide. More recently, several 
new sulfur analytical methods have been developed and used for soil studies with ICP and IC 
being predominant. Both of these methods require specialized and expensive instrumenta- 
tion. Since their operation is specific to the instrument and defined by the manufacturer, they 
will not be described here. Although instruments for these methods are now widely available, 
there have been few assessments of their effectiveness or comparisons with values using 
traditional methods in soil studies. It is generally accepted that interference is limited for ICP 
measurements, but the sulfur measurement includes all forms rather than sulfate specifically. 
Ion chromatography can be specific for sulfate measurements, but is subject to interference 
by a wide array of anions and cations including those that are a part of traditional digestion 
and extraction solutions. These potential interferences must be addressed in each specific 
case. These influences can be circumvented in a variety of ways and depend on the specific 
instrument that is used. The hydriodic acid method is, therefore, outlined here because it does 
not require specialized instrumentation and procedures, and has a long history of application 
to soils. The method involves hydriodic acid reagent described by Johnson and Nishita 
(1952) used in a modified apparatus (Kowalenko 1985) and bismuth sulfide (Kowalenko and 
Lowe 1972) rather than methylene blue quantification of the sulfur evolved as hydrogen 

23.3.1 Materials and Reagents 

1 Custom-built reduction/distillation apparatus and heating block (Kowalenko 
1985) as outlined in Section 23.2.2. 

2 Nitrogen gas: the gas must be relatively pure and free from sulfides in particular. 
The gas may be purified by bubbling it through a solution containing 5 to 10 g 
mercuric chloride in 1 00 mL of 2% (w/v) potassium permanganate. The flow of 
the nitrogen gas to the reduction/distillation apparatus should be regulated 
to approximately 200 mL min -1 . This can be done by commercially available 
flow meters (e.g., Rotometer [Kowalenko 1985]) or forcing the gas through 
an appropriate length (e.g., 30 cm) of capillary glass tubing (Kowalenko and 
Lowe 1972). 

3 Hydriodic acid reducing reagent: mix 4 volumes (e.g., 400 mL) of hydriodic 
acid (e.g., 57% with 1%-2.5% hypophosphorus acid preservative), 1 volume 
(e.g., 100 mL) of hypophosphorus acid (50%), and 2 volumes (e.g., 200 mL) of 


formic acid (90%) and, while bubbling purified nitrogen gas through it, heat for 
1 min at 1 1 5°C-1 1 7°C. Continue the nitrogen gas flow through the reagent while 
cooling. The heating should be done in a well-ventilated hood; refluxing or a 
special gas-trapping apparatus (Tabatabai 1982) is recommended. Since this 
reagent is not very stable, only sufficient reagent for several days of sample and 
standard analyses should be prepared. Storage in a brown bottle and refrigeration 
will extend its stability. 

4 1 M sodium hydroxide: for absorbing hydrogen sulfide. 

5 Bismuth reagent: dissolve 3.4 g bismuth nitrate pentahydrate in 230 ml_ glacial 
acetic acid. Also, dissolve 30 g gelatin in 500 ml_ water and mix thoroughly. 
Both solutions will require gentle heating for dissolution. Filter the bismuth 
solution if it is not clear. The final bismuth reagent is prepared by combining 
the bismuth and gelatin solutions and diluting to 1 L. This reagent is quite stable at 
room temperature. 

g Sulfate-S standards: prepare a 1 000 mg S L _1 stock solution by dissolving 5.435 g 
dried reagent-grade potassium sulfate and diluting to 1 L. Working standards are 
made by appropriate dilutions. 

7 Spectrophotometer: the instrument should be suitable for measurement at 400 nm 
and capable of accommodating small (e.g., 7.5 ml_) volumes, including provision 
for rinsing the cuvette or analysis chamber. 

23.3.2 Procedure 

1 Weigh a portion of dry, whole soil or pipette an appropriate volume (e.g., 2-5 ml_) 
of the filtered extract into the modified Taylor tube for the hydriodic acid 
reduction/distillation apparatus and evaporate (up to 100°C) the aliquot of the 
extract solution to dryness. The weight of soil or volume of extract solution should 
be adjusted in size such that the amount of sulfur in the apparatus will be within 
the range of calibration standards that is conducted. 

2 Assemble the custom-built reduction/distillation apparatus above the small heat- 
ing block in such a way that the modified Taylor tube can be easily installed on or 
removed from the dispenser portion. This can be accomplished by either having the 
heater in a fixed position and the dispenser portion with the Taylor tube easily 
raised and lowered, or the dispenser plus Taylor tube fixed and the heater on a jack 
assembly. A tube (50 mL test tube or larger, depending on the range of the standard 
curve) containing the sodium hydroxide solution for absorbing the hydrogen 
sulfide should be fixed in a position such that the nitrogen gas from the outlet of 
the apparatus will bubble through several centimeters of the absorbing solution. 
The volume of the absorbing solution is adjusted for the concentration range of 
sulfate to be analyzed. Adjust the nitrogen gas at the appropriate rate and fill the 
burette with reducing reagent. As each distillation is completed and the Taylor tube 
is removed, a watch glass should be placed above the heater to intercept any drops 
of reducing reagent. The entire apparatus should be adequately ventilated. 

j Condition the reduction/distillation apparatus by attaching a modified Taylor tube 
containing a high (e.g., 200 mg S L~ 1 ) sulfate standard to the reduction/distillation 


apparatus, place the apparatus in heating position, adjust the nitrogen gas flow, and 
dispense 4 ml_ of reducing reagent into the attached Taylor tube. The apparatus 
requires conditioning at the beginning of each new session with high sulfate-sulfur 
standard to ensure quantitative initial distillation. Distill until all the sulfate has 
been reduced and transferred into the absorbing solution. The time required for this 
process will vary with the flow rate of the nitrogen and the "dead" volume within 
the apparatus. About 8 to 10 min should be adequate, but calibration under 
specific conditions is recommended (Kowalenko 1985). After distillation of the 
hydrogen sulfide is complete, remove the tube containing the absorption solution 
from the apparatus and check that the distillation process is functioning by 
immediately adding an appropriate volume of bismuth reagent and mix thor- 
oughly. This volume should correspond to the volume of the absorbing solution 
(2:1 absorbing solutiombismuth reagent) depending on the range of the standard 
sulfate-sulfur required. For example, 20 ml_ of absorbing solution is suitable for a 
1 -200 mg S L~ 1 range and 5 ml_ for a 1 -40 mg S L~ 1 range. 

4 After the initial setup and conditioning of the apparatus, digested soil samples, 
whole soil sample or dried soil extract aliquot, and dried standards are distilled 
into appropriate volumes of absorption solution, and bismuth reagent is immedi- 
ately added in preparation for quantitative measurements of the bismuth sulfide 
produced. Measurements are best conducted in batches and adequate standards 
included in each batch. Measure absorbance of the sample and standard solutions 
at 400 nm. 

23.3.3 Calculations 

For measurements of sulfate in a soil sample placed directly into the sample vessel of the 
hydriodic acid analysis apparatus, prepare the standard curve for the bismuth colorimetric 
determination as an appropriate range of a quantity (e.g., mg S) of sulfur distilled in a single 
analysis. Then the concentration of sulfur in the soil sample is calculated as the quantity 
of sulfur relative to the standard curve divided by the weight of the soil (oven-dry basis) in 
the vessel during the analysis. Also, for measurements on solution samples, standardize the 
apparatus on an appropriate range of a quantity of sulfur in a single analysis, but calculate 
the concentration of the sulfur on an oven-dry basis taking into account the aliquot size of 
the analyzed solution that was dried in the apparatus sample vessel and the ratio of the 
analyzed solution to the weight of the soil. 


7 Filter paper has been found to contain variable amounts of sulfate which will be 
leached during the filtration. Washing the filter paper with some of the extractant 
prior to filtration is recommended. 

2 Water has been shown to reduce the efficiency of hydriodic acid reagent 
to reduce sulfate to sulfide (Kowalenko and Lowe 1975); therefore, the standard 
and sample solution aliquot volumes should either be the same throughout 
or all the liquid of the aliquot evaporated to dryness. Although evaporating 
the sample to dryness is time consuming, it does provide an opportunity for 
altering the sensitivity of the analysis (i.e., evaporate a small volume for soils 
with a significant sulfate-sulfur content or a large volume for soils with a low 
sulfate-sulfur content). 


The methylene blue color reaction is subject to interference; therefore, passing the 
nitrogen gas through a pyrogallol-sodium phosphate wash just prior to the hydrogen 
sulfide absorption is recommended (Johnson and Nishita 1952). The bismuth 
sulfide method is much less sensitive to interference; therefore, the pyrogallol- 
sodium phosphate wash can be eliminated (Kowalenko and Lowe 1 972). Use of a 
Taylor tube rather than a condenser to provide refluxing can shorten reduc- 
tion/distillation times from 60 to 10 min (Kowalenko 1985). The apparatus is 
also simpler to fabricate and is versatile for different types of analyses. Although 
the sensitivity of the bismuth reaction is considerably lower than the methylene 
blue reaction, it can be adequately enhanced for most soil studies by decreasing 
the volume into which the hydrogen sulfide is absorbed and/or by increasing the 
size of the original sample being analyzed. However, as the volumes of the 
absorbing solution and bismuth reagent are decreased to increase the sensitivity, 
increased attention must be given to the precision and reproducibility of absorbing 
solution and bismuth reagent volume measurements, particularly relative 
to the standard samples. The spectrophotometer should be capable of accom- 
modating the small sample sizes involved, including appropriate rinsing between 

The original sulfate determination procedure (Johnson and Nishita 1952) recom- 
mended that the nitrogen gas should be purified before use. Currently available 
sources of nitrogen are more uniform and free from impurities; therefore, purifi- 
cation of the gas may be omitted. The purity of the gas for analysis purposes can 
be evaluated by examining blanks. There should also be fairly good control of the 
flow rate of the nitrogen gas with a high enough rate to transfer the hydrogen 
sulfide produced into the receiver solution quickly, but slow enough that the 
sulfide gas can be absorbed by the sodium hydroxide. 

Sources of contamination, such as rubber connectors or lubricants for sealing 
connections, should be considered, particularly in the reduction/distillation 
procedure. A small amount of water is adequate to seal the Taylor tube to the 
rest of the apparatus during the reduction/distillation. 

Hydriodic acid is available in concentrations ranging from 48% to 66% and with 
or without preservative. Although these products contain varying quantities of 
sulfate contamination, the sulfate is removed by heating the mixed reagent. The 
57% hydriodic acid with preservative has been found to be acceptable. If other 
products are used, the proportion of hydriodic acid to the other acids may need 
adjustment and the final reagent tested for effectiveness. Adequate time should be 
allowed for acquisition of hydriodic acid, as stocks are often limited. 

Although the hydriodic acid reduction procedure is not influenced by a wide 
variety of salts, it is recommended that the standards should be similar to the 
sample's matrix (e.g., water for whole soil or the solution used for extraction). For 
the alkaline digested soil, standards should be included through the digestion 
process. This precaution will also provide a check on sulfate or sulfide contam- 
ination that may be present in the extract or digestion solutions. 

The hydriodic acid method, although relatively specific for sulfate, includes both 
inorganic and organic forms. This should not be neglected when the results are 
being interpreted. When this method is used to determine total organic sulfate 


(or total carbon-bonded sulfur by difference) the capability of accurately deter- 
mining total inorganic sulfate or total sulfur must be considered, particularly when 
unusual samples (e.g., subsurface, anaerobic, organic, etc. samples) are being 
examined. The difference value will involve the error or variability associated 
with two analyses rather than one. 

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Kowalenko, C.G. and Lowe, L.E. 1975. Evalu- 
ation of several extraction methods and of a 
closed incubation method for sulfur mineraliza- 
tion. Can. J. Soil Sci. 55: 1-8. 

Roberts, T.L. and Bettany, J.R. 1985. The influ- 
ence of topography on the nature and distribu- 
tion of soil sulfur across a narrow environmental 
gradient. Can. J. Soil Sci. 65: 419^134. 


Scott, N.M., Bick, W., and Anderson, H.A. 1981. 
The measurement of sulfur-containing amino 
acids in some Scottish soils. /. Sci. Food Agric. 

32: 21-24. 

Tabatabai, M.A. and Bremner, J.M. 1970a. 
An alkaline oxidation method for determination of 
total sulfur in soils. Soil Sci. Soc. Am. Proc. 34: 

Stallings, E.A., Candelaria, L.M., and Gladney, 
E.S. 1988. Investigation of a fusion technique for 
the determination of total sulfur in geological 
samples by ion chromatography. Anal. Chem. 
60: 1246-1248. 

Strick, J.E. and Nakas, J.P. 1984. Calibration of a 
microbial sulfur technique for use in forest soils. 
Soil Biol. Biochem. 16: 289-291. 

Tabatabai, M.A. 1982. Sulfur. In A.L. Page, R.H. 
Miller, and D.R. Keeney, Eds. Methods of Soil 
Analysis, Part 2 — Chemical and Microbiological 
Properties, 2nd ed. American Society of Agron- 
omy, Madison, WI, pp. 501-538. 

Tabatabai, M.A., Basta, N.T., and Pirela, 
H.J. 1988. Determination of total sulfur in soils 
and plant materials by ion chromatography. Com- 
mun. Soil Sci. Plant Anal. 19: 1701-1714. 

Tabatabai, M.A. and Bremner, J.M. 1970b. 
Comparison of some methods for determination 
of total sulfur in soils. Soil Sci. Soc. Am. Proc. 34: 


Wainwright, M. and Johnson, J. 1980. Determin- 
ation of sulfite in mineral soils. Plant Soil 54: 

Watkinson, J.L., Lee, A., and Lauren, D.R. 1987. 
Measurement of elemental sulfur in soil and 
sediments: field sampling, sample storage, pre- 
treatment, extraction and analysis by high- 
performance liquid chromatography. Aust. J. Soil 
Res. 25: 167-178. 

Zhao, F.J., Loke, S.Y., Crosland, A.R., and 
McGrath, S.P. 1996. Method to determine elem- 
ental sulfur in soils applied to measure sulfur 
oxidation. Soil Biol. Biochem. 28: 1083-1087. 



Chapter 24 
Total and Organic Phosphorus 

I. P. O'Halloran 

University of Guelph 
Ridgetown, Ontario, Canada 

B.J. Cade-Menun 

Stanford University 

Stanford, California, United States 


This chapter describes several methods used for the determination and characterization of 
total phosphorus (P t ) and organic phosphorus (P ) in soils. 

Determination of the P t in soil requires the solubilization of P through the decomposition 
or destruction of mineral and P containing materials in the soil. Historically, the two most 
widely recognized procedures for the determination of soil P t are the sodium carbonate 
(Na 2 C03) fusion method and the perchloric acid (HCIO4) digestion method (Olsen and 
Sommers 1982). Currently, neither method is widely used in studies involving the determination 
of soil P t . Although the Na2C03 fusion method is considered the most reliable procedure for 
quantitative determination of P t in soils, it is laborious, tedious, and generally unsuitable for the 
analyses of large numbers of samples. Digestion with HCIO4, although more adaptable as a 
routine laboratory procedure, requires the use of fume hoods specifically designed for the HCIO4 
digestion. The potential danger of explosions, due to HCIO4 buildup or reaction of HCIO4 with 
organic materials, has led many institutions and laboratories to discontinue the use of HCIO4 
digestions. Given the greater applicability of the HCIO4 digestion method as a routine laboratory 
procedure for the analysis of a large number of samples, this procedure has been included in the 
current text. The reader is referred to the previous edition of this text (O'Halloran 1993) or Olsen 
and Sommers (1982) for details on the Na2C03 fusion method. Three additional methods for the 
determination of P t in soil are presented in this chapter. One involves the alkaline oxidation of 
the sample using sodium hypobromite (NaOBr)/sodium hydroxide (NaOH) (Dick and 
Tabatabai 1977) and the other two are wet acid digestion procedures using either sulfuric 
acid (H 2 S0 4 )/hydrogen peroxide (H 2 2 )/hydrofluoric acid (HF) (Bowman 1988) or 
H 2 S0 4 /H 2 2 /lithium sulfate (Li 2 S0 4 )/selenium (Se) (Parkinson and Allen 1975). 

Soil P methods can be divided into methods that attempt to measure the total P and 
methods that attempt to characterize both the amount and the forms of P in the soil. 


The first group consists of extraction (Mehta et al. 1954; Bowman and Moir 1993) or ignition 
techniques (Saunders and Williams 1955). In each case, total soil P is not determined 
directly, but rather is calculated as the increase in inorganic P (P ; ) measured after the 
digestion of a soil extract or ignition of a soil sample. The ignition technique is less laborious 
than the extraction techniques, but it is also subject to a greater number of errors. 

Characterization of soil P forms can be done either by sequential fractionation techniques 
that provide operationally defined pools of P or techniques that identify specific groups or 
forms of P materials in soils. Sequential fractionation techniques, such as the modified 
procedure of Hedley et al. (1982) that is presented in Chapter 25 of this manual, characterize 
soil P forms by measuring P t and soluble-reactive P (srP) in each fraction and then assuming 
that the difference between P t and srP is P . However, caution should be used when 
interpreting P results by sequential fractionation: first, because there is the potential for 
P forms to be altered by previous extractants within any sequential fractionation procedure; 
and second, because srP measures only orthophosphate (HPO4 2 , H2P04~), the "P " 
measured by the difference between P t and srP in a solution may also include complex P; 
forms such as pyrophosphate or polyphosphates. 

In the identification of specific P compounds, four groups of soil P compounds have been 
detected in soils. Orthophosphate monoesters are esters of phosphoric acid, with one C moiety 
per P, and include inositol phosphates, sugar phosphates, phosphoproteins, and mononucleo- 
tides. Orthophosphate diesters are also esters of phosphoric acid, but have two C moieties per P. 
These include DNA, RNA, phospholipids, teichoic acid, and aromatic compounds. Phos- 
phonates contain C-P bonds rather than ester linkages, and occur as phosphonic acids and 
phosphonolipids. The final group of soil P compounds is orthophosphate anhydrides. 
Although most of these are complex Pi compounds such as pyrophosphate and polyphosphate, 
this grouping also includes important organic orthophosphate anhydrides such as adenosine 
diphosphate (ADP) and adenosine triphosphate (ATP). More details on soil P compounds can 
be found in Condron et al. (2005). 

A range of techniques is available to examine specific soil P compounds. Most involve 
extraction with a reagent specific to the recovery of a particular P compound, followed by 
analysis using techniques such as 31 P nuclear magnetic resonance (NMR) spectroscopy, 
enzyme hydrolysis, thin-layer chromatography, high-performance liquid chromatography 
(HPLC), and mass spectroscopy. These methods are often laborious, may require complex 
and expensive instrumentation, and may be specific to only one P compound or group of P 
compounds. In addition, the interpretation of results is limited by incomplete extraction or 
poor detection. Presented in this chapter are two common procedures to characterize P 
compounds in soil extracts: 31 P NMR spectroscopy and enzyme hydrolysis. Both allow the 
determination of the relative proportions of a range of P compounds in soil samples. 


Currently, the most popular approaches to soil P t determinations involve either alkali or acid 
oxidation or digestion of the soil samples. Each of the following methods has an advantage 
over the Na 2 C03 fusion method in that they are more adaptable for the routine analysis of a 
greater number of soil samples. As mentioned previously, digestion with HCIO4 requires 
special HCIO4 fume hoods, and care must be taken to avoid explosions. It has been reported 
that the HCIO4 digestion method gives relatively low P t values in highly weathered materials 
and with samples containing apatite inclusions (Syers et al. 1967, 1968, 1969). Each of the 


remaining three methods has been reported to produce similar to slightly greater results to 
those obtained with the HCIO4 digestion procedure (Dick and Tabatabai 1977; Bowman 
1988; Rowland and Grimshaw 1985) without the need for a special fume hood to carry out 
the oxidation or digestion. The H2SO4/H2O2/HF digestion method is relatively fast and is 
ideally suited for small numbers of samples. This procedure, however, requires HF-resistant 
materials, since HF attacks glass. The NaOBr/NaOH and I^SCVl^CVl^SCVSe 
methods are well suited for the analysis of large numbers of samples. 

The reader is also referred to two additional procedures for the determination of soil P t that 
may be of interest. The first involves a fusion method using NaOH that melts at a lower 
temperature than Na2CC>3 (325°C versus 850°C), and therefore allows the use of nickel rather 
than platinum crucibles (Smith and Bain 1982). Although this procedure was initially 
reported to give similar soil P t values as the Na 2 C03 fusion method for 10 Scottish soils 
(Smith and Bain 1982), problems with low recoveries of soil P t in New Zealand soils with 
high organic matter content (soils that have >80% weight loss on ignition) have been 
identified (Taylor 2000). The second method is based on the Thomas et al. (1967) method 
for plant tissue digestion using H2SO4/H2O2 to digest a soil sample in an aluminum block 
digestor. This procedure is presented as the final step (i.e., digestion of soil residue) in the 
sequential fractionation procedure described in Chapter 25. Agbenin and Tiessen (1994) 
found for semiarid tropical soils in Brazil that the H2SO4/H2O2 method gave comparable to 
slightly higher soil P t values than the Na2CC>3 fusion method. Gasparatos and Haidouti 
(2001) studying 15 soils varying in extractable P levels reported that this method gave soil P t 
values that were 95%-105% of those obtained with HCIO4 digestion. 

Regardless of the procedures selected, it is recommended that finely ground soil, 0.15-0.18 mm 
(100-80 mesh), be used to allow for efficient recovery or extraction of P from the soil material, 
and to improve the reproducibility. The moisture content of the soil should be known so as to 
allow expression of P content on an oven-dried basis. Blank samples containing no soil should 
also be included to assess the possibility of P contamination and to serve as a suitable reagent 
blank for the colorimetric determination of P. Inclusion of a reference sample with known P, 
and the use of duplicate samples are also encouraged. 

24.2.1 Perchloric Acid Digestion (Olsen and Sommers 1982) 

As a safety precaution, samples should routinely be predigested in concentrated nitric acid 
(HNO3) before proceeding with the HCIO4 digestion. This method is suitable for the 
determination of soil P t in a large number of samples, although the use of an HCIO4 fume 
hood is essential. The digestion can be carried out using 250 mL Erlenmeyer flasks and a hot 
plate, or by using an aluminum block digestor with 75, 100, or 250 mL digestion tubes (see 
comments on p. 268). The HCIO4 digestion typically recovers 92%-96% as much P as the 
Na2C03 fusion method (Sommers and Nelson 1972; Dick and Tabatabai 1977; Bowman 
1988), although the pretreatment of soil samples with HF can increase the recovery of P 
(Kara et al. 1997). 

Materials and Reagents 

1 HCIO4 fume hood 

2 Hot plate (with 250 mL Erlenmeyer flasks) or aluminum block digestor (with 250 mL 
digestion tubes) 

j HN03, concentrated 

4 HCI0 4/ 60% 

5 Color developing solution (see Section 24.5.1) 


1 Accurately weigh approximately 2.0 g of finely ground soil into a 250 ml_ 
volumetric or Erlenmeyer flask. 

2 Add 20 mL concentrated HNO3 to flask and mix well. Heat (approximately 
130°C) to oxidize the organic matter in the sample. Organic matter oxidation is 
complete when the dark color due to the organic matter in the sample disappears. 

3 Allow the S0N-HNO3 mixture to cool slightly. In the HCIO4 fume hood, add 
30 mL of HCIO4 and digest the sample at the boiling temperature (approximately 
200°C) for 20 min. During this time dense white fumes should appear and the 
insoluble solid material left in the bottom of the flask or digestion tube should 
appear like white sand. If necessary, use a little (less than 2 mL) extra HCIO4 to 
wash down any black particles that have stuck to the sides of the flask or digestion 
tube. Heat for another 10-15 min. 

4 Allow the mixture to cool. With distilled/deionized water transfer the mixture to 
a 250 mL volumetric flask and make to volume with distilled/deionized water. 
Mix thoroughly. 

5 Allow sediment to settle before taking an aliquot for analysis. 

6 Determine P concentration in an aliquot of the clear supernatant as indicated in 
Section 24.5. 

A 40 tube aluminum block digestion system with volumetric 75 or 100 mL digestion tubes 
can also be used in this procedure by using half the amounts of sample, HNO3, and HCIO4 
described above. The digested material is diluted to a final volume of 75 or 100 mL (step 4). 

24.2.2 Sodium Hypobromite/Sodium Hydroxide Alkaline Oxidation 
Method (Dick and Tabatabai 1977) 

This method involves boiling to dryness a mixture of soil and NaOBr-NaOH solution using a 
sand bath or, as modified by Cihacek and Lizotte (1990), an aluminum block digestor. Formic 
acid is added after completion of the NaOBr-NaOH treatment to destroy residual NaOBr 
remaining after oxidation of the sample. Soil P t is then extracted from the sample using 
0.5 M H2SO4. The method permits the digestion of a large number of samples at one time, 
although more manipulation of the sample is required compared to the HCIO4 method. Dick 
and Tabatabai (1977) using a wide range of soils from the United States and Brazil found that 
for soil P, this method removed about 96% as much P as the Na2CC>3 fusion method, and was 
comparable (about 1% higher) to the P determined by HCIO4 digestion. Cihacek and Lizotte 
(1990), using soils from the Great Plains region of the United States, found that this procedure 


removed significantly (about 3%) more P than the HCIO4 digestion. Kara et al. (1997) found 
this method to recover 93%-100% of the P determined by Na 2 C0 3 fusion and 99%-102% of 
the P determined by HCIO4 digestion on soils from Scotland and Turkey. 

Materials and Reagents 

1 Sand bath for which the temperature of the sand can be regulated at 260°C-280°C 
or an aluminum block digestor (see Comment 3 on p. 270). 

j Boiling flask (50 mL) with stoppers or digestion tubes for the aluminum block 

4 Centrifuge and 50 mL centrifuge tubes. 

5 NaOH, 2 M: dissolve 80 g NaOH in a 1 L volumetric flask containing 600 mL 
of distilled/deionized water. Allow to cool and make to volume with 
distilled/deionized water. 

g NaOBr/NaOH solution: prepare this solution in a fume hood by adding 3 mL of 
bromine slowly (0.5 mL min -1 ) and with constant stirring to 100 mL of 2 M 
NaOH. Prepare the NaOBr-NaOH solution immediately before use. 

7 Formic acid (HCOOH), 90%. 

8 H2SO4, 0.5 M: add 28 mL concentrated H 2 S0 4 to 600 mL distilled/deionized 
water in a 1 L volumetric flask. Mix, allow to cool, and make to volume using 
distilled/deionized water. 

9 Color developing solutions (see Section 24.5.1). 


7 Accurately weigh between 0.1 and 0.20 g of finely ground soil into a dry 50 mL 
boiling flask. 

2 Add 3 mL of NaOBr-NaOH solution to the boiling flask, and swirl the flask for a 
few seconds to mix the contents. Allow the flask to stand for 5 min, and then swirl 
the flask again for a few seconds. 

3 Place the flask upright in a sand bath (temperature regulated between 260°C and 
280°C) situated in a fume hood. Heat the flask for 1 0-1 5 min until its contents are 
evaporated to dryness, and continue heating for an additional 30 min. 

4 Remove the flask from the sand bath, cool for about 5 min, add 4 mL of 
distilled/deionized water and 1 mL of 90% HCOOH. Mix the contents, and 
then add 25 mL of 0.5 M H 2 S0 4 . Stopper the flask and mix the contents. 

5 Transfer the mixture to a 50 mL plastic centrifuge tube and centrifuge at 1 5,000 g 
for 1 min. 

Determine P concentration in an aliquot of the clear supernatant as indicated in 

Section 24.5. 

/ It is very important that the NaOBr-NaOH solution be prepared immediately 
before use. Dick and Tabatabai (1977) reported that storing the NaOBr-NaOH at 
4°C for 24 h reduced P t values by 2%-4%. 

2 Sample sizes up to 0.5 g can be analyzed for most soils. However, samples 
containing high amounts of Fe show large decreases in the value of P t when 
sample sizes are increased above 0.2 g. 

3 A sand bath may be prepared by placing 3-4 cm of silica sand on a hot plate; 
however, even temperature regulation across the sand bath can be difficult to 
achieve. Cihacek and Lizotte (1990) found that the use of an aluminum block 
digestor resulted in a more uniform heating of all samples and improved the 
precision of P t determination. 

24.2.3 Sulfuric Acid/Hydrogen Peroxide/Hydrofluoric Acid Digestion 
(Bowman 1988) 

This method involves the digestion of the soil sample by the sequential additions of 
concentrated H2SO4, H2O2, and HF. The precision and accuracy is similar to that of the 
HCIO4 method and gives soil P t values that are approximately 94% of those obtained with 
Na2CC>3 fusion (Bowman 1988). This method is suited for the analysis of a small number of 
samples at one time. The time required for manual additions of the H2O2 and HF is similar to 
the manipulations required for the NaOBr-NaOH method, which makes the procedure 
slightly more labor intensive than the HCIO4 or H2S04/H 2 02/Li 2 S04/Se methods. 

Materials and Reagents 

1 Fluoropolymer beaker (100 ml_) of known weight 

2 Fume hood 

3 Balance or 50 mL volumetric flask 

4 Quantitative fine filter paper (e.g., Whatman No. 42) 

5 H 2 S0 4 , concentrated 

6 H 2 2 , 30% 

7 HF, concentrated (see Comment 1, p. 271 ) 

q Color developing solutions (see Section 24.5.1) 


1 Accurately weigh 0.5 g of finely ground soil into a 1 00 mL fluoropolymer beaker 
of known weight. Use 0.25 g for soil high in organic matter. 


In the fume hood add 5 ml_ (9.2 g) of H2SO4 to the soil and gently swirl to suspend 
solid materials adhering to the bottom of the beaker. 

In the fume hood, slowly add 0.5 mL of H2O2 and mix vigorously to promote the 
oxidation of organic materials. Repeat this step until 3 mL of H 2 2 has been 
added to the beaker. Let the sample sit until the reaction with H2O2 has subsided. 

Add 0.5 mL of HF to the beaker and mix. Repeat this step again so that a total of 
1 mL of HF has been added. 

Place beaker on a preheated hot plate (approximately 150°C) for 10-12 min to 
eliminate excess H2O2. 

Remove beaker, and while sample is still warm, wash down the sides of the beaker 
with 1 0-20 mL of distilled/deionized water. Mix and cool to room temperature. 

Weigh the beaker and its contents and add sufficient distilled/deionized water to 
bring the final contents weight to 55 g (equivalent to 50 mL volume). Alterna- 
tively, the material in the beaker can be quantitatively transferred to a 50 mL 
volumetric flask and made to volume using distilled/deionized water. 

Mix and filter the extract. 

Determine P concentration in an aliquot of the clear filtrate as indicated in 
Section 24.5. 

7 HF acid attacks glass and it is very important that HF-resistant materials such as 
polytetra fluoroethylene (PTFE) are used. An example of such material is Teflon. 
(The use of this trade name is provided for the benefit of the reader and does not 
imply endorsement by the CSSS.) 

2 Excess H 2 2 will interfere with the colorimetric determination of P in the digested 
sample. The formation of a yellow color instead of the blue color normally 
associated with the reduced molybdophosphate complex indicates the presence 
of excess H 2 2 . 

24.2.4 Sulfuric Acid/Hydrogen Peroxide/Lithium Sulfate/Selenium 
Digestion (Parkinson and Allen 1975) 

This method involves the digestion of the soil sample with H2SO4 and H2O2. The addition of 
the salt Li 2 S04 allows for the use of a higher digestion temperature and Se is added as a 
catalyst in the oxidation of the organic material present in the sample. Rowland and 
Grimshaw (1985) studying 103 soils across eight major soil types in Britain found that this 
procedure has a similar accuracy, and on average removed slightly more P (103%) than the 
HCIO4 digestion. The digestion is usually completed after 2-2.5 h and is therefore somewhat 
longer than the other procedures. However, using an aluminum block digestor with either 
75 or 100 mL tubes enables the analysis of a large number of samples at a time, with a 
relatively modest amount of sample manipulation. Samples may also be digested using 
50 mL boiling flasks, although it is critical that the proper soikdigestion mixture ratio is 
maintained to ensure proper digestion of the sample. 


Materials and Reagents 

/ Aluminum digestion block with 100 ml_ volumetric digestion tubes having a 
suitable stopper or sealing device (i.e., silicon stopper). A hot plate is required if 
using 50 ml_ boiling flasks. 

2 Fume hood. 

j Vortex mixer (optional). 

4 Silicon stopper (or other device) for sealing digestion tubes. 

5 H 2 S0 4 , concentrated. 

6 H 2 2/ 30%. 

j Lithium sulfate monohydrate (Li 2 S0 4 • H 2 0). 

g Selenium (Se) powder. 

g Color developing solutions (see Section 24.5.1). 

Digestion solution: the day before sample digestion, mix 175 mL of H 2 2 with 
0.21 g Se powder and 7 g Li 2 S0 4 • H 2 in a suitable container. A plastic, sealable 
bottle is preferred as some pressure may develop in the container. Store this 
solution in a refrigerator overnight; the Se powder should be dissolved by the 
following day. Do not heat the solution to dissolve the Se as this may severely 
decrease the effectiveness of the H 2 2 . This solution is stable for 2-3 weeks. 

Accurately weigh 0.2-0.4 g of finely ground soil into a 100 mL digestion tube. 

To reduce the risk of "bumping" an inert boiling stone, glass, or PTFE bead can be 
added to the digestion tube (see Comment 1 , p. 273). 

In a fume hood add 5 mL of concentrated H 2 S0 4 to the digestion tube and swirl 
(or use vortex mixer) until soil is thoroughly mixed with the acid and turns a dark 
brown or black color. 

Carefully and slowly add 1 mL of the digestion mixture to the digestion tube. The 
sample should react by foaming or spattering and due caution should be exer- 
cised. If there is no apparent reaction with the addition of the digestion mixture, 
gently tap the tube to facilitate the mixing of the digestion solution with the acid- 
soil mixture. Repeat this step three more times to add a total of 4 mL of digestion 

Place the digestion tubes in a cold block digestor and gradually increase the 
temperature over a 1-1.5 h period until a temperature of 360°C is reached and 
maintained for 30 min. There should be evidence of H 2 S0 4 vapors refluxing in 
the tubes. 


7 Remove tubes from the block digestor and allow to cool (5-10 min). Add 
0.5 ml_ H2O2, washing down any soil particles that are stuck to the sides of the 
digestion tube, mix well and replace on block for 30 min. 

q Repeat step 7 until solution is a clear to milky-white color (usually requires two 
0.5 ml_ additions of H 2 2 ) indicating a complete digestion of the soil organic 
matter (SOM). Note, some samples with relatively high iron contents may have a 
yellowish tinge, which will not change with further additions of H 2 C>2. 

g After the final heating for 30 min, remove the tubes from the block digestor and 
allow to cool for 30 min. Slowly add 20-30 ml_ distilled/deionized water and mix 
(vortex) the sample to ensure the residue is easily suspended in the solution. 

7 Add distilled/deionized water until liquid level is slightly below the volumetric 
mark on the tubes. Allow the solution to cool before making to final volume with 
distilled/deionized water. 

1 1 Stopper or seal the tube and thoroughly mix the contents by slowly inverting the 
tubes several times. 

12 Allow the contents to settle before decanting into storage containers. Let the 
samples sit overnight in a refrigerator or filter through quantitative fine filter 
paper before colorimetric analysis. 

73 Determine P concentration in an aliquot of the clear filtrate as indicated in 
Section 24.5. 

Digestions at high temperatures involving soil-acid mixtures can cause bumping 
resulting in the violent and dangerous ejection of materials from the digestion 
tube. The risk of bumping can be reduced through the use of inert boiling stones, 
glass, or PTFE beads that facilitate a consistent, smooth boiling of the acid. This is 
more likely to be a problem with blanks than with tubes containing the sample. 

Alternatively, 210 ml_ of concentrated H 2 S0 4 can be added to the digestion 
solution prepared in step 1, and the addition of concentrated H 2 S0 4 directly to 
the sample (step 3) omitted. A total of 9 ml_ of the digestion mixture would be 
required for each sample, and should be added in careful and incremental 
additions to avoid too vigorous a reaction that may cause loss of material from 
the tube or flask. This digestion mixture should be refrigerated and is stable for 
2-3 weeks. 


Total soil P is not measured directly, but rather as the increase in P; resulting from the 
ignition of a soil sample or digestion of a soil extract. Differences among techniques or soil 
types may reflect a change in the efficiency of the procedure, rather than a true change in the 
amount of P in the soil. The extraction techniques (Anderson 1960; Bowman 1989; 
Bowman and Moir 1993) involve the use of various acid and base treatments with the 


subsequent determination of the P; and P t in the extractants. The two major problems with 
these techniques are the incomplete extraction of soil P and the possible hydrolysis of P by 
the extractants. In general, these techniques tend to give the lower range of soil P values. 
The ignition techniques use either high (550°C, Saunders and Williams 1955) or low (250°C, 
Legg and Black 1955) temperatures to oxidize soil P to P;. Matched ignited and unignited 
samples are then extracted with either weak or strong acids. The difference between P; 
(ignited sample) and P; (unignited sample) is considered P . This technique may result in 
erroneous estimates of P due to incomplete oxidation of P and changes in the solubilities of 
P minerals by ignition at either high or low temperatures, while ignition at higher temper- 
atures may also cause volatilization of P. Each technique has its advantages and disadvan- 
tages, depending on the situation and the purpose of the study in question. As indicated by 
Bowman (1989), the extraction techniques are more suited for comparisons of P levels 
across different soil types, whereas ignition techniques are more suitable for comparisons 
among treatments within a soil type. Due to the errors that may be associated with P 
determinations and since the P is determined by difference, little significance can be given 
to treatments that differ by less than 20 |xg P g _1 soil (Olsen and Sommers 1982). 

As indicated by Condron et al. (1990), several studies have reported good agreement 
between ignition and extraction techniques with ignition methods tending to give higher 
soil P values, although studies have shown higher soil P levels with extraction compared to 
ignition techniques (Condron et al. 1990; Agbenin et al. 1999). In addition, considerable 
differences between the two techniques have been noted for certain soil types. Further 
information regarding comparisons of various methods for the determination of P in soils 
can be obtained by referring to Condron et al. (1990, 2005), Dormaar and Webster (1964), 
Steward and Oades (1972), and Turner et al. (2005). 

For improved accuracy and precision of analysis, it is recommended that the soils used be 
air-dried and finely ground (0.15-0.18 mm; 100-80 mesh). Duplicate soil samples and 
blanks containing no sample should be used in each analysis. There are no certified reference 
materials for total organic P. 

24.3.1 Hydrochloric Acid/Sodium Hydroxide Extraction Method 
(Anderson 1960 as Modified by Condron et al. 1990) 

In this method, soils are sequentially extracted with 0.3 M NaOH, concentrated HC1 (hot and 
then at room temperature), 0.5 M NaOH at room temperature, and 0.5 M NaOH at 90°C. The 
Pi in extracts is determined immediately after extraction and the P t is determined after the 
oxidation of the organic matter with persulfate digestion. 

Materials and Reagents 

/ Heat-resistant polypropylene screw-top centrifuge tubes (50 ml_) with caps. 

2 Water bath at 90°C. 

3 Centrifuge. 

4 Vortex mixer (optional), 
c End -over-end shaker. 

g Oven for NaOH extraction of samples at 90°C. 
j Autoclave. 
q Aluminum foil. 

9 Volumetric flasks (50 and 100 mL). 

10 H2SO4, concentrated. 

77 H 2 S0 4 0.9 M: add 50 mL of concentrated H 2 S0 4 to a 1 L volumetric flask 
containing 600 mL of distilled/deionized water. Mix and make to volume using 
distilled/deionized water. 

12 HCI, concentrated. 

13 NaOH, 0.3 M: dissolve 12 g NaOH in a 1 L volumetric flask containing appro- 
ximately 700 mL of distilled/deionized water. Make to volume with 
distilled/deionized water. 

14 NaOH, 0.5 M: dissolve 20 g NaOH in a 1 L volumetric flask containing appro- 
ximately 700 mL of distilled/deionized water. Make to volume with 
distilled/deionized water. 

75 Ammonium persulfate: (NH 4 ) 2 S 2 08. 

75 Color developing solutions (see Section 24.5.1). 

Extraction Procedure 

7 Weigh 0.5 g of finely ground soil into a 50 mL polypropylene centrifuge tube. 

2 0.3 M NaOH extraction: add 30 mL of 0.3 M NaOH, cap the tube, and shake on 
an end-over-end shaker for 16 h at room temperature. After shaking, centrifuge 
(12,500 g) the soil suspension for 10 min and then carefully decant the super- 
natant into a 100 mL volumetric flask ensuring that the soil residue remains in 
the tube. 

3 Concentrated HCI extraction: to the soil residue in the centrifuge tube add 1 mL 
of concentrated HCI, mix thoroughly, and then place the tube in an 82°C water 
bath for 10 min. Remove the tube from the water bath, add 5 mL concentrated 
HCI, and allow to stand at room temperature for 1 h with regular (approximately 
every 15 min) vortex shaking. Centrifuge (12,500 g) for 10 min, carefully decant 
the supernatant into a 50 mL volumetric flask, and make to volume using 
distilled/deionized water. 

4 Room temperature 0.5 M NaOH extraction: to the soil residue in the centrifuge 
tube add 20 mL 0.5 M NaOH, mix well, and allow to stand for 1 h at room 
temperature with regular (approximately every 1 5 min) vortex shaking. Centrifuge 
(12,500 g) the soil suspension for 10 min, and carefully decant the supernatant 
into the 100 mL volumetric flask containing the previous 0.3 M NaOH extract. 


5 Hot 0.5 M NaOH extraction: to the soil residue in the tube, add 30 ml_ of 0.5 M 
NaOH, shake to suspend the soil in solution. Loosely cover the tubes with an inverted 
50 mL beaker or funnel and place in an 82°C oven for 8 h. Remove tubes from 
the oven, allow to cool, centrifuge (1 2,500 g), and decant the supernatant into the 
100 mL volumetric flask containing the previous two NaOH extracts. Make the 
contents of the 1 00 mL volumetric flask to volume using distilled/deionized water. 

Determination of Pj and Total P in the Extracts 

Dclei ruination of Pj 

7 To determine Pj in the NaOH extract, pipette a suitable aliquot (usually <5 mL) into 
a 50 mL centrifuge tube. Acidify to precipitate organic material by adding 2.0 mL of 
0.9 M H 2 S0 4 and set in a refrigerator for 30 min. Centrifuge at 25,000 gfor 1 min 
at 0°C. Decantthe supernatant into a 50 mL volumetric flask. Using a little acidified 
water, rinse the tube carefully so as not to dislodge any of the precipitated organic 
matter, and add the liquid to the contents of the flask (repeat two or three times). 
Develop color as described in Section 24.5.2 starting with pH adjustment (step 3). 

2 To determine Pj in the HCI extract, pipette a suitable aliquot (usually <5 mL) into 
a 50 mL volumetric flask. Develop color as described in Section 24.5.2 starting 
with pH adjustment (step 3). 

Determination of total P 

/ To colorimetrically determine total P in the extracts, pipette a suitable aliquot 
(usually <2 mL) of solution into a 50 mL volumetric flask. 

2 To the NaOH extract add «0.5 g (NH 4 ) 2 S 2 8 and 10 mL of 0.9 M H 2 S0 4 . 

3 To HCI extract add «0.4 g (NH 4 ) 2 S 2 8 and 10 mL deionized/distilled water. 

4 Cover the mouth of the flask with aluminum foil (double layer for HCI extract) and 
autoclave (60 min for HCI extracts and 90 min for NaOH extracts). 

5 Cool, add approximately 10 mL distilled/deionized water. Develop color as 
described in Section 24.5.2 starting with pH adjustment (step 3). 


Total P in the soil sample is determined as the summation of the total P in the HCI and 
NaOH extracts minus the summation of the Pj in the HCI and NaOH extracts. After 
determining the concentration of P in the digests and extracts and converting each to a soil 
weight basis (e.g., mg P kg~' soil), P is calculated as 

P = (HC1-P, + NaOH-P t ) - (HCl-Pi + NaOH-Pj) (24. 1 ) 

Pi should be determined in the extracts as soon as possible to redut 
P hydrolysis resulting in an underestimation of soil P . 


2 Soils high in humic materials and metals, such as forest soils and wetland soils, 
may lose Pj through the formation of P-metal-organic matter complexes during 
the precipitation of organic matter (Darke and Walbridge 2000), resulting in 
overestimation of P Q concentrations. P concentrations for these types of soils 
should be confirmed with a second method, such as ignition (Section 24.3.3). 

3 Instead of acidifying an aliquot of the NaOH extract to precipitate organic matter, 
Pj can be determined directly, provided a suitable blank is used to correct for 
absorbance by organic matter in solution. To do this, pipette equal aliquots of the 
same sample extract into two separate 50 ml_ volumetric flasks and adjust the pH. 
To one add 8 ml_ of the color developing solution as described in Section 24.5. To 
the other add 8 ml_ of the color developing solution without ascorbic acid added. 
Dilute to volume and measure absorbance. The absorbance of the solution 
without ascorbic acid is subtracted from the absorbance of the solution with 
ascorbic acid. 

4 Total P in the extracts can be determined directly using ICP, although dilution may 
be required if samples are relatively high in organic matter. 

5 Total P in the extracts can also be determined by other digestion techniques such 
as the procedure of Thomas et al. (1967) using H2SO4/H2O2, or the potassium 
persulfate digestion using a hotplate (Bowman 1989). 

24.3.2 Basic EDTA Extraction Method (Bowman and Moir 1993) 

In this procedure, P is extracted from the soil using 0.25 M NaOH and 0.05 M disodium ethylene 
diamine tetraacetic acid (Na2EDTA). This method is simple, faster than either the HCl/NaOH 
extraction method of Anderson (1960) or the ignition method of Saunders and Williams (1955), 
and is equally efficient. Excessive amounts of EDTA in solution can interfere with the colori- 
metric determination of P. Acidification of the extracts to pH < 1 .5 will precipitate both 
extracted SOM and EDTA, and this precipitate can then be removed by centrifugation or 
filtration (Nnadi et al. 1975) allowing for the determination of srP in solution. 

Materials and Reagents 

7 Heat-resistant polypropylene screw-top centrifuge tubes (50 ml_). 

2 Centrifuge. 

3 Incubator or oven set at 85°C. 

4 Quantitative fine filter paper (e.g., Whatman No. 42). 

5 NaOH, 0.5 M: dissolve 1 g NaOH in a 500 ml_ volumetric flask containing 300 mL 
of distilled/deionized water. Make to volume with distilled/deionized water. 

6 Na 2 EDTA, 0.1 M: dissolve 1 8.6 g Na 2 EDTA in a 500 mL volumetric flask containing 
300 mL of distilled/deionized water. Make to volume with distilled/deionized water. 

7 NaOH-EDTA mixture: combine the 0.5 M NaOH and 0.1 M Na 2 EDTA solutions 
(final concentration 0.25 M NaOH + 0.05 M Na 2 EDTA). 

g Ammonium persulfate: (NH 4 ) 2 S208. 

g H 2 S0 4 5.5 M: add 306 ml_ of concentrated H 2 S0 4 to a 1 L volumetric flask 
containing 500 ml_ of distilled/deionized water. Mix, cool, and make to volume 
using distilled/deionized water. 

1Q H2SO4 0.9 M: add 50 mL of concentrated H 2 S0 4 to a 1 L volumetric flask 
containing 600 mL of distilled/deionized water. Mix and make to volume using 
distilled/deionized water. 

7 7 Color developing solutions (see Section 24.5.1 ). 

Extraction Procedure 

7 Weigh 0.5 g of finely ground soil into a heat-resistant 50 mL centrifuge tube. 

2 Add 25 mL of combined NaOH-EDTA solution to the tube, cap tightly, and shake 
briefly to mix. 

j Loosen caps. Place in incubator or oven (preheated to 85°C) for 10 min. 

4 Cap tightly and incubate for 1 h 50 min (2 h incubation total). 

5 Centrifuge at 25,000 g for 10 min. 

5 Filter supernatant and keep the filtrate for analysis. 

Determination of Pj and Total P in the Extract 

7 Determination of P,: pipette <5 mL of extract into a 50 mL centrifuge tube. Acidify 
by adding 0.5 mL 0.9 M H2SO4. Cool in refrigerator for 30 min. Centrifuge at 
25,000 gfor 10 min at 0°C. Decant the supernatant into a 50 mL volumetric flask. 
Using a I ittle acidified water, rinse the tube carefully so as to not dislodge any of the 
precipitated organic matter and add the liquid to the contents of the flask (repeat 
two or three times). Develop color as described in Section 24.5.2 starting with pH 
adjustment (step 3). 

2 Determination of total P: pipette <5 mL extract into a 25 mL volumetric flask. Add 
«0.5 g (NH 4 ) 2 S 2 8 and 10 mL of 0.9 M H 2 S0 4 . Cover the mouth of the flask 
with aluminum foil and autoclave for 90 min. Cool, add approximately 10 mL 
distilled/deionized water, and develop color as described in Section 24.5.2 
starting with pH adjustment (step 3). 

After determining the concentration of P in the digests and extracts and converting each to a 
soil weight basis (e.g., mg P kg -1 soil), P is calculated as 

P = P t (digest or ICP) - P; (extract) (24.2) 



1 The initial 10 min period with loose caps is to minimize gas buildup. 

2 The combination of EDTA and NaOH simultaneously eliminates the formation of 
cationic bridges with SOM and solubilized organic matter (Bowman and Moir 
1993). This eliminates the need for acid pretreatment. 

j Extraction at room temperature for 16 h produced similar P Q concentrations as 
extracting at 85°C for 2 h. 

4 As discussed in Comments, pp. 276 and 277, the removal of organic matter by 
precipitation may produce inaccurate estimations of P concentrations in soils 
high in humic materials and metals. 

5 Pj in the NaOH-EDTA extract can also be determined directly following the same 
procedure as outlined in the third point in Comments, p. 277. If this alternative 
procedure is used, the aliquot size should not exceed 4 ml_, because excessive 
EDTA will retard color development. 

g Total P in the extract can be determined using an ICP, or by other methods such as 
the procedure of Thomas et al. (1967) using H2SO4/H2O2, or potassium per- 
sulfate digestion using a hotplate (Bowman 1989). 

24.3.3 Ignition Method (Saunders and Williams 1955, as Modified by 
Walker and Adams 1958) 

In this method, P is estimated by the difference between 0.5 M IT^SCVextractable P in a soil 
sample ignited at 550°C and an unignited sample. The method is suitable for the determination of 
soil P in a large number of samples. Dormaar and Webster (1964) have indicated that 
significant volatile losses of P may occur at temperatures above 400°C, especially with 
peat soils. 

Materials and Reagents 

7 Muffle furnace and porcelain crucibles for igniting soils at 550°C. 
2 Polypropylene centrifuge tubes (100 mL) with caps or stoppers. 
2 Shaker capable of holding the above tubes. 

4 Centrifuge. 

5 H2SO4, 0.5 M: add 28 mL concentrated H 2 S0 4 to 600 mL distilled/deionized 
water in a 1 L volumetric flask. Allow to cool and make to volume using 
distilled/deionized water. 

g Color developing solutions (see Section 24.5.1). 


/ Weigh 1 .0 g of finely ground soil in a porcelain crucible, and place the crucible in 
a cool muffle furnace. 

2 Slowly raise the temperature of the muffle furnace to 550°C over a period of 
approximately 2 h. Continue to heat the samples at 550°C for 1 h, then remove 
the samples and allow them to cool. 

3 Transfer the ignited soil to a 1 00 ml_ polypropylene centrifuge tube for extraction. 

4 To a separate 1 00 mL polypropylene centrifuge tube, weigh 1 .0 g of unignited soil 
for the extraction of Pj. 

5 Add 50 mL of 0.5 M H 2 S0 4 to both samples, mix well, and allow to sit lightly 
stoppered for a few minutes to relieve pressure from C0 2 released from any 
carbonates that may be present in the soil sample. Tightly stopper the tubes and 
place them on a shaker for 16 h. Blank samples containing only 0.5 M H2SO4 
should also be included. 

g Centrifuge the samples at approximately 1 500 g for 15 min. If the extract is not 
clear, filtration using acid-resistant filter paper may be required. 

7 Determine P concentration in an aliquot of clear supernatant or filtrate as indi- 
cated in Section 24.5. 

After determining the concentration of P in the extracts and 
basis (e.g., mg P kg -1 soil), P is calculated as 

P = P ; (ignited sample) — P ; (unignited sample) 

To prevent volatilization of P from the sample, care must be taken to not allow temperatures 
in the muffle furnace to exceed 550°C when using mineral soils (Sommers et al. 1970; 
Williams et al. 1970). 


There are no direct methods to speciate soil P . Although there have been attempts to 
characterize P directly in soil using solid-state 31 P NMR spectroscopy, results have generally 
been poor due to line broadening from the close association of soil P with paramagnetics 
such as Fe (Cade-Menun 2005). As such, soil P must be extracted before speciation. 
Association with mineral components stabilizes much of the soil P , making it difficult to 
extract, and using strong acid or base extraction introduces the risk of P hydrolysis. Thus, 
the ideal extractant for chemical characterization of soil P should maximize recovery while 
minimizing alteration of chemical structure. Post extraction, the ideal speciation technique 
should allow the quantitative determination of the relative proportions of a range of P 


compounds. Two techniques that best fit these criteria are solution 31 P NMR spectroscopy 
and enzyme hydrolysis. 

24.4.1 NaOH-EDTA Extraction for Solution 31 P NMR Spectroscopy 

From its first use on soil extracts by Newman and Tate (1980), solution 31 P NMR spectroscopy 
has substantially advanced our knowledge of P compounds in soil and other environmental 
samples. Various extractants have been used including 0.5 M NaOH alone or combined with 
either EDTA or the cation-exchange resin Chelex (Bio-Rad Laboratories). (The use of this 
trade name is provided for the benefit of the reader and does not imply endorsement by the 
CSSS.) The choice of extractant will influence both the recovery of P from soil and the 
composition of extracted compounds (Cade-Menun and Preston 1996; Cade-Menun et al. 
2002). The extractant most commonly used at present is a combination of NaOH-EDTA 
based on the Bowman and Moir (1993) extraction procedure for total P , described in 
Section 24.3.2. 

31 P NMR spectroscopy allows the characterization of the relative abundances of both P and 
Pi forms in an extract. Figure 24.1 shows 31 P NMR spectra for standard reference materials 
available from the National Institute of Standards and Testing (NIST) in the United States. 
The top sample is apple leaf reference, and the bottom is the San Joaquin soil reference. Note 
the differences in the relative abundances of P and P; compounds. 

It is beyond the scope of this chapter to fully describe the workings of a 31 P NMR spectrometer, 
and exact analytical procedures will vary with each spectrometer. Please see Cade-Menun 
(2005) for important considerations on conducting a successful 3I P NMR experiment on soil 
extracts. Included here is a protocol to extract soil samples for 31 P NMR spectroscopy. 

Materials and Reagents 

1 Polypropylene screw-top 50 mL centrifuge tubes. 

2 Mechanical shaker for the centrifuge tubes. 

3 Vortex mixer (optional). 

4 Centrifuge. 

5 Freezer. 

g Freeze dryer. 

j NMR spectrometer with broadband probe. Ideally, a 500 MHz (for proton) 
spectrometer and a 10 mm probe (see Cade-Menun 2005). 

8 NaOH, 10 M: dissolve 20 g NaOH in a 50 mL volumetric flask containing 
30 mL of distilled/deionized water. Allow to cool and make to volume with 
distilled/deionized water. 

9 NaOH, 0.5 M: dissolve 10 g NaOH in a 500 mL volumetric flask containing 
300 mL of distilled/deionized water. Allow to cool and make to volume with 
distilled/deionized water. 


NIST reference materials 

Apple leaf 



Orthophosphate 26% 









San Joaquin s 




Orthophosphate 84% 









R— O— P=0 

:e (NMR) spectra of N 1ST apple leaf reference and N 1ST 
San Joaquin reference soil, showing the range of P compounds that can be identified 
using this technique, and their relative abundance in soil and foliar samples. (PL is 
phospholipids; DNA is deoxyribonucleic acid.) The inset for the soil spectrum shows 
the expanded orthophosphate monoester, diester, and pyrophosphate region. 

7 Na 2 EDTA, 0.1 M: dissolve 1 8.6 g Na 2 EDTA in a 500 ml_ volumetric flask contain- 
ing 300 ml_ of distil led/deionized water. Make to volume with distilled/deionized 

7 7 NaOH-EDTA mixture: combine the 0.5 M NaOH and 0.1 M Na 2 EDTA solutions 
(final concentration 0.25 M NaOH + 0.05 M Na 2 EDTA). 

72 Deuterium oxide (D 2 0) suitable for NMR analyses. 


7 Weigh 1-2 g of soil into a 50 ml_ centrifuge tube. Use larger sample if soil is 
known to be low in P t . Use smaller sample if high in Fe or organic matter. 

2 Add 30 mL of combined NaOH-EDTA solution to the tube and cap tightly. 

j Shake at room temperature for 5-16 h. Longer extractions can increase the 
recovery of total P, particularly occluded P forms, but may also increase the risk 
of degradation of P forms such as RNA and phospholipids. 

4 Centrifuge at 1 500 g for 20 min. Decant supernatant into another 50 mL centri- 
fuge tube. If the supernatant contains particulate material, samples should be 
filtered before decanting into the second centrifuge tube. 

5 Remove 1 mL of supernatant. Dilute to 1 mL with distilled/deionized water, and 
analyze for P, Fe, and Mn. 

6 Cap centrifuge tubes containing remainder of supernatant tightly, and freeze for 
16-24 h, until completely frozen. Note: freeze tubes on a slant to maximize 
surface area. 

7 Remove caps from tubes and cover loosely with Parafilm (poke small holes in Parafilm 
to allow air to circulate) or similar material. Place tubes upright in freeze-dryer flasks. 
Lyophilize for 24-48 h according to freeze-dryer instructions, until completely dry. 
Remove tubes from freeze-dryer flask. Cap tightly. Store at room temperature. 

g If using a spectrometer with a 1 mm probe, samples can be redissolved directly 
in the centrifuge tube by adding 1.6 mL of distilled/deionized water, 1 mL of 
D 2 0, and 0.4 mL of 10 M NaOH (to adjust the pH to >12, for maximum peak 
separation). Let stand for 30 min, mixing or vortexing occasionally to dissolve all 
solids. Centrifuge at 1500 g for 20 min. Decant into NMR tube. If using a 
spectrometer with a 5 mm probe, adjust volumes accordingly. 

g See Cade-Menun (2005) for a discussion of suitable spectrometer parameters to 
conduct a successful 31 P NMR experiment on soil extracts. 

24.4.2 Organic Phosphorus Characterization by Enzyme Hydrolysis 

Characterization of P is based on the principle that substrate-specific phosphatase enzymes 
will release Pj from specific P forms. Thus, by adding commercially available phosphatase 
enzymes to soil extracts and colorimetrically analyzing the P ; released, the P forms within the 
extracts can be grouped into P compound categories. The specific classification of P forms will 
depend on the enzymes used in the assay. For example, acid phosphatase or alkaline phos- 
phatase will hydrolyze orthophosphate monoesters in general, while phytase will hydrolyze 
one specific orthophosphate monoester, phytic acid («?yo-inositol hexakisphosphate). 

One aspect of enzyme hydrolysis is that it can be conducted on a number of different soil 
extracts, including water, sodium bicarbonate (NaHC03), NaOH, and HC1, and has been 
used with sequential extraction procedures (e.g., He and Honeycutt 2001). However, solu- 
tions should be adjusted to the suitable pH range for each enzyme before characterization 
with enzyme hydrolysis. 

A number of different protocols exist for enzyme hydrolysis, including the universal buffer 
procedure recently developed by He et al. (2004). The following protocol was adapted from 
Turner et al. (2002, 2003) and Toor et al. (2003). 


Materials and Reagents 

/ Polypropylene screw-top 50 mL centrifuge tubes. 

2 Shaker capable of holding the above tubes. 

3 Calibrated disposable plastic centrifuge tubes (1 5 mL), 5 for each soil sample to be 

4 Incubator or shaking water bath, set at 37°C. 

5 Centrifuge. 

5 0.45 jjum membrane filter and vacuum filtration apparatus. 

7 NaOH, 1 M: dissolve 20 g NaOH in a 500 mL volumetric flask containing 
300 mL of distilled/deionized water. Allow to cool and make to volume with 
distilled/deionized water. 

q NaHC0 3 , 0.5 M pH 8.5: dissolve 21 g NaHC0 3 and 0.25 g of NaOH in a 
500 mL beaker containing 300 mL of distilled/deionized water. Transfer to 
a 500 mL volumetric flask, and make to volume with distilled/deionized 

9 Sodium azide (NaN 3 ), 25 m/Vf: dissolve 0.163 g NaN 3 in a 100 mL volumetric 
flask containing 40 mL of distilled/deionized water. Make to volume with 
distilled/deionized water. 

1q H 2 S0 4 , 3 M: add 83 mL of concentrated H 2 S0 4 to 500 mL volumetric flask 
containing 300 mL of distilled/deionized water. Mix and allow to cool before 
making to final volume with distilled/deionized water. 

11 Tris-HCI buffer, 2 M: dissolve 31.5 g Tris-HCI powder (Polysciences, Inc., 
Warrington, PA) and 0.041 g MgCI 2 • 6H 2 in a 100 mL beaker containing 
60 mL of distilled/deionized water. Adjust to pH 8. Transfer to a 100 mL 
volumetric flask, and make to volume with distilled/deionized water. 

12 Glycine-HCI buffer, 2 M: dissolve 0.041 g MgCI 2 • 6H 2 in a 100 mL beaker 
containing 60 mL of distilled/deionized water. Add 20 mL of 1 M Glycine-HCI 
buffer, 10x concentrate (Polysciences, Inc., Warrington, PA). Check pH, which 
should be 2.5. Transfer to a 100 mL volumetric flask, and make to volume with 
distilled/deionized water. 

73 Enzymes: suitable enzymes can be obtained from a variety of sources. (All the 
following enzymes are available from Sigma Chemicals, St. Louis, MO: trade 
names are mentioned only for the benefit of the reader.) 

a. Alkaline phosphatase (EC 3.1 .3.2), Type V-IIS, from bovine intestinal mucosa, 
activity of preparation 1 unit mL -1 : add 0.1 mL of alkaline phosphatase (2.2 mg 
protein per mL, 2420 units activity per mg protein) to 20 mL of 2 M Tris-HCI 
buffer, pH 8. 


b. Phospholipase C (EC, Type XI, from Bacillus cereus, activity of pre- 
paration 1 unit ml_~ 1 : add 24.94 mg of phospholipase (16.04 units activity 
mg~ 1 solid) and 0.1 ml_ of alkaline phosphatase to 20 ml_ of 2 M Tris-HCI 
buffer, pH 8. See Comment 5, p. 286. 

c. Phosphodiesterase (EC, Type IV, from Crotalus atrox venom, activity 
of preparation 0.03 units mL" 1 : add 20 mg of phosphodiesterase (0.02 units 
activity mg~ 1 solid) and 0.1 ml_ of alkaline phosphatase to 20 mL of 2 M Tris- 
HCI buffer, pH 8. See Comment 5, p. 286. 

d. Phytase (EC, Type myo- inositol hexakisphosphate 3-phosphohydro- 
lase, from Aspergillus ficuum, activity of preparation 1 unit mL~ 1 : add 23 mg 
of phytase (1 .1 units activity mg- 1 solid) to 80 mL of 2 M Glycine-HCI buffer, 
pH 2.5. Centrifuge for 1 min at 1 500 g. 

74 Magnesium chloride (MgCI 2 ), 2 m/VJ: dissolve 0.041 g MgCI 2 • 6H 2 in a 1 00 mL 
volumetric flask containing 60 mL of distilled/deionized water. Make to volume 
with distilled/deionized water. 

75 Color developing solution: see Section 24.5.1 . 


7 Weigh 1 .5 g of soil into a 50 mL centrifuge tube. Blank samples containing no soil 
should also be analyzed. 

2 Add 30 mL of 0.5 M NaHC0 3 . Shake for 30 min. 

3 Centrifuge at approximately 1500 g for 15 min. Filter supernatant through 
0.45 (Jim filters. 

4 Label five 15 mL centrifuge tubes for each extracted soil sample or blank. Four 
tubes should be labeled with the names of the enzymes (one per enzyme), while 
the fifth tube should be labeled "control." Add 1 mL of the NaHC0 3 extract to 
five 15 mL centrifuge tubes for each extracted soil sample. Preacidify by adding 
0.1 mL of 3 M H 2 S0 4 and neutralize by adding 0.12 mL of 1 M NaOH (see 
Comment 2, p. 286. 

5 Add 1 mL of 25 mM NaN 3 to prevent microbial activity. 

g Add 0.25 mL of each enzyme-buffer mixture to the appropriately labeled tube for 
each sample or blank. Add 0.25 mL of the MgCI 2 solution to the controls. Dilute 
to 5 mL with distilled/deionized water. 

j Incubate with shaking at 37°C for 1 6 h (incubator or shaking water bath). 

q Terminate enzyme reaction by adding 1 mL of color developing solution (see 
Section 24.5). Final volume for samples (and standards) is 6 mL. 

g Measure the absorbance after 12 min at 880 nm. Calculate P, concentration in 
solution by comparison to a standard curve. Note that phosphodiesterase and 


phytase cause slight interferences with the molybdate blue reaction. Prepare separate 
calibration curves from orthophosphate standards containing the enzymes. 

/ All buffers contain 2 mM MgCI 2 because Mg 2+ ions are natural activators of 
phosphatase enzymes (Dixon and Webb 1966). 

2 The preacidification and neutralization steps are necessary to remove carbonates 
from bicarbonate extractions, to prevent foaming during subsequent colorimetric 

j "Activity of preparation" refers to the activity in the centrifuge tube with the soil 
extract and other reagents. 

4 Commercial phytase is not purified, and contains other P-hydrolyzing enzymes. 
For a purification procedure, see Hayes et al. (2000). 

5 Alkaline phosphatase was added to the phospholipase and phosphodiesterase 
preparations because phosphodiesterase and phospholipase hydrolyze only one 
ester-P bond on the diester molecule. This leaves an orthophosphate monoester, 
which requires alkaline phosphatase to completely release orthophosphate 
(Turner et al. 2002). 

6 If working with acid extracts, use acid phosphatase (EC rather than 
alkaline phosphatase. 

7 Sodium azide and phosphodiesterase from Crotalus atrox venom are both 
poisons, and should be handled and disposed of accordingly. 


Functional classes of organic P compounds are calculated as 

7 labile monoester P: hydrolyzed by alkaline phosphatase; 

2 phospholipids: the difference between the P released by phospholipase + alkaline 
phosphatase and the P released by alkaline phosphatase alone; 

j nucleic acids: the difference between the P released by phosphodiesterase + 
alkaline phosphatase and the P released by alkaline phosphatase alone; and 

4 inositol hexakisphosphate (phytic acid): the difference between the P released by 
phytase and all other treatments. 


The determination of P in solutions is usually conducted by colorimetric methods or by 
inductively coupled plasma (ICP) spectroscopy. Colorimetric methods for the determination 
of P in solution require that the desired pool of soil P that is extracted from the soil is 


completely converted to orthophosphate, while total P in solution may be analyzed without 
prior digestion by ICP. 

One of the most commonly used methods for the colorimetric determination of orthopho- 
sphate concentration in solutions is the method developed by Murphy and Riley (1962). This 
method uses the blue color developed by a phosphoantimonylmolybdenum complex (Going 
and Eisenreich 1974; Drummond and Maher 1995) reduced by ascorbic acid to estimate the 
concentration of orthophosphate in solution. As the procedure was originally developed 
for seawater, Murphy and Riley (1962) only assessed adherence to Beer's law up to a 
final solution concentration (i.e., solution in which the color has been developed) of 
0.2 mg PL -1 . The procedure is suitable for final solution P concentrations of approximately 
0.8 mg P L _1 (Rodriguez et al. 1994) and subsequent modifications of the strength of the 
antimony (Sb) and ascorbic acid solutions have extended this to a final solution concentra- 
tion of 3 mg P L" 1 (Harwood et al. 1969). This procedure is fairly simple, less susceptible to 
interferences than procedures using SnCl2 as a reductant, and is capable of being used 
manually or adapted to automated systems (Drummond and Maher 1995). A manual method 
as modified by Watanabe and Olsen (1965) is present here, and is suitable for aliquots 
containing between 1 and 40 |xg srP when made to a final volume of 50 mL for color 

24.5.1 Reagents 

1 Ammonium molybdate solution: dissolve 12 g of ammonium molybdate tetrahy- 
drate ((NH 4 ) 6 Mo 7 024 • 4H 2 0) in 250 mL distilled/deionized water. 

2 Potassium antimony tartrate solution: dissolve 0.2908 g of potassium antimony 
tartrate (KSbOC 4 H 4 6 ) in 1 00 mL distilled/deionized water. 

3 H 2 S0 4 2.5 M: to a 1 L volumetric flask containing approximately 600 mL of 
distilled/deionized water slowly add 1 39 mL of concentrated (1 8 M) H 2 S0 4 . Mix 
by swirling the contents of the flask, allow to cool, and make to volume with 
distilled/deionized water. 

4 Reagent A: combine the three solutions above in a 2 L volumetric flask, make to 
volume with distilled/deionized water and mix thoroughly. Store in a Pyrex glass 
bottle in a refrigerator. 

5 Color developing solution: dissolve 1. 056 g of ascorbic acid in200mLof reagent A 
and mix. Prepare this solution daily and do not use if more than 24 h old. 

5 p-Nitrophenol solution: dissolve approximately 0.25 gof p-nitrophenol in 100 mL 
of distilled/deionized water. 

7 NaOH 4 M: in a 1 L volumetric flask dissolve 160 g of NaOH in approximately 
800 mL of distilled/deionized water. Allow to cool, make to volume using 
distilled/deionized water, and mix thoroughly. 

8 H 2 S0 4 0.25 M: slowly add 1 4 mL of concentrated H 2 S0 4 to 1 L volumetric flask 
containing approximately 800 mL of distilled/deionized water, make to volume 
with distilled/deionized water, and mix thoroughly. 


9 Standard P stock solution (100 mg P L~ 1 ): dissolve 0.4394 g of potassium dihy- 
drogen phosphate (KH2PO4) in 1 Lof distilled/deionized water. Prepare a working 
standard (10 mg P L" 1 ) by dilution with distilled/deionized water. 

24.5.2 Procedure 

/ Pipette an aliquot containing 1 -40 jULg of P into a 50 mL volumetric flask contain- 
ing approximately 15 mL distilled/deionized water. 

2 Pipette standard P solutions into a set of volumetric flasks so as to encompass the 
range of P concentrations anticipated in the extracts. To each flask containing a 
standard solution, pipette an aliquot of blank solution equal to the aliquot size of 
the sample. 

j Add 1-2 drops of p-nitrophenol and adjust the pH of the solution to ~5. If the 
sample aliquot has a pH <5, add 4 M NaOH drop wise until the solution turns 
yellow in color and then add 0.25 M H 2 S0 4 until colorless. If the sample aliquot 
has a pH >5, add 0.25 M H 2 S0 4 until colorless. 

4 Add 8 mLofthe color developing solution, make to volume with distilled/deionized 
water, and mix thoroughly. After 1 min read absorbance at either 882 or 71 2 nm 
(if solution is slightly colored due to the presence of organic matter). 

5 Appropriate standards (final solution concentrations of 0-0.8 |xg P ml_~ 1 , or 
0-40 |jLg P in the 50 mL volumetric flask) should be analyzed in the same manner 
as samples, and contain similar amounts of extracting or digestion solutions as 
the samples. 


/ Many versions of the Murphy and Riley (1962) procedure have been published, 
and the reader is cautioned that deviations from proposed methodologies can lead 
to erroneous results. The development of a stable blue color that adheres to Beer's 
law requires the proper adjustment of solution pH, as well as specific ranges of 
Mo, Sb, and ascorbic acid concentrations relative each other, to the amount of P 
in the sample, or both (Harwood et al. 1969; Going and Eisenreich 1974; 
Rodriguez et al. 1994; Drummond and Maher 1995). Any changes to proposed 
methods should be verified using samples and standards of known P content. 

2 The original method described by Watanabe and Olsen (1965) for sodium 
bicarbonate extracts of soil P used 25 mL volumetric flasks, and therefore only 
4 mL of color developing reagent and a sample aliquot containing a maximum of 

j Arsenate (As0 4 ) will also form a blue color with the Murphy and Riley solution. 
Olsen and Sommers (1982) indicate that in most soils the average As concentra- 
tion is 6 mg kg~ , and as such would be a negligible amount compared to typical 
P concentrations in soils. However, if a soil has been contaminated with As, this 
could lead to substantial overestimation of P in the sample. In soils with high As 
contents, Olsen and Sommers (1982) recommend reducing As0 4 to As0 3 by 


adding 5 ml_ of sodium hydrogen sulfite solution (5.2 g of NaHSOs dissolved in 
1 00 mL of 0.5 M H2SO4) to the sample aliquot and either heating the mixture in a 
water bath for 30 min (20 min at 95°C) or letting it stand for 4 h before adjusting 
pH and developing the color. 

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Bowman, R.A. 1988. A rapid method to deter- 
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Bowman, R.A. 1989. A sequential extraction pro- 
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Bowman, R.A. and Moir, J.O. 1993. Basic EDTA 
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Extractants, metals and P relaxation times. 
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Cade-Menun, B.J. and Preston, CM. 1996. A 
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Dormaar, J.F. and Webster, G.R. 1964. Losses 
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Drummond, L. and Maher, W. 1995. Determin- 
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formation of the phosphoantimonylmolybdenum 
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Gasparatos, D. and Haidouti, C. 2001. A compari- 
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Olsen, S.R. and Sommers, L.E. 1982. Phosphorus. 
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Parkinson, J.A. and Allen, S.E. 1975. A wet oxi- 
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Rodriguez, J.B., Self, J.R., and Soltanpour, P.N. 
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by the ascorbic acid-molybdenum blue method. 
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Rowland, A.P. and Grimshaw, H.M. 1985. A wet 
oxidation procedure suitable for total nitrogen and 
phosphoius in soil ( n I Sci. Plant Anal. 

16: 551-560. 

Saunders, W.M. and Williams, E.G. 1955. 
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Harwood, J.E., van Steenderen, R.A., and Kiihn, 
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Hayes, J.E., Richardson, A.E., and Simpson, 
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He, Z., Griffin, T.S., and Honeycutt, C.W. 2004. 
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Hedley, M.J., Stewart, J.W.B., and Chauhan, B.S. 
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and by laboratory incubations. Soil Sci. Soc. Am. 
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Legg, J.O. and Black, C.A. 1955. Determina 
of organic phosphorus in soils. II. Ignition 
method. Soil Sci. Soc. Am. Proc. 19: 139-143. 

Mehta, N.C., Legg, J.O., Goring, C.A.I., and 
Black, C.A. 1954. Determination of organic phos- 
phorus in soils. I. Extraction method. Soil Sci. 
Soc. Am. Proc. 18: 443^49. 

Murphy, J. and Riley, J.P. 1962. A modified single 
solution method for the determination of phosphate 
in natural waters. Anal. Chini. Acta. 27: 31-36. 

Newman, R.H. and Tate, K.R. 1980. Soil phosphorus 
characterization by 31 P nuclear magnetic reson- 
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Nnadi, L.A., Tabatabai, M.A., and Hanaway, J.J. 
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O'Halloran, LP. 1993. Total and organic phos- 
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Sampling and Methods of Analysis. Canadian 
Society of Soil Science. Lewis Publishers, Boca 
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Sommers, L.E., Harris, R.F., Williams, J.D.H., 
Armstrong, D.E., and Syers, J.K. 1970. Determin- 
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Sommers, L.E. and Nelson, D.W. 1972. Deter- 
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Steward, J.H. and Oades, J.M. 1972. The deter- 
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Sci. 23: 38^19. 

Syers, J.K., Williams, J.D.H., Campbell, A.S., 
and Walker, T.W. 1967. The significance of apa- 
tite inclusions in soil phosphorus studies. Soil Sci. 
Soc. Am. Proc. 31: 752-756. 

Syers, J.K., Williams, J.D.H., Tyner, E.H., and 
Walker, T.W. 1969. Primary and secondary origin 
of "nonextractable" soil inorganic phosphorus. 
Soil Sci. Soc. Am. Proc. 33: 635-636. 

Syers, J.K., Williams, J.D.H., and Walker, T.W. 
1968. The determination of total phosphorus in 


a-ials. AT. Z. /. Agric. Res. 11: 

Taylor, M.D. 2000. Determination of total phos- 
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Commun. Soil Sci. Plant Anal. 31: 2665-2670. 

Thomas, R.L., Sheard, R.W., and Moyer, J.R. 
1967. Comparison of conventional and automated 
procedures for nitrogen, phosphorus and potas- 
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digestion. Agron. J. 59: 240-243. 

Toor, G.S., Condron, L.M., Di, H.J., Cameron, 
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Turner, B.L., Cade-Menun, B.J., Condron, L.M., 
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Turner, B.L., Cade-Menun, B.J., and Wester- 
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ition and potential bioavailability in semi-arid 
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Turner, B.L., McKelvie, I.D., and Haygarth, P.M. 
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Walker, T.W. and Adams, A.F.R. 1958. Studies 
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content of parent materials on accumulation of 
carbon, nitrogen, sulfur and organic phosphorus 
in grassland soils. Soil Sci. 85: 307-318. 

Watanabe, F.S. and Olsen, S.R. 1965. Test of an 
ascorbic acid method for determining phosphorus 
in water and NaHC03 extracts from soil. Soil 
Sci. Soc. Am. Proc. 29: 677-678. 

Williams, J.D.H., Syers, J.K., Walker, T.W., and 
Rex, R.W. 1970. A comparison of methods for 
the determination of soil organic phosphorus. Soil 
Sci. 110: 13-18. 



Chapter 25 

Characterization of Available P 

by Sequential Extraction 

H. Tiessen 

nstitute for Global Change Research 
Sao Jose dos Campos, Sao Paulo, Brazil 

J.O. Moir 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 


Phosphate availability is a function of chemical equilibrium-controlled solubility and rate- 
limited processes. Most methods for available P determination attempt to quantify P solubility 
using different extractants, but few relate this to P supply rates that are relevant to plant uptake. 

Soil test methods for P do not measure the quantity of P available to a crop, but extract a portion 
of soil P that is related to plant-available P. This relationship is usually established over 
years of agronomic experimentation and testing of fertilizer responses through regression 
equations. These equations relate plant performance to soil test P levels, or indicate fertilizer 
requirement for optimum crop production. Results obtained with this approach are not 
always transferable between crops or soil types, and different equations are established by 
soil testing services for varying crops and soils. The approach does not work when perennial 
plants or natural ecosystems are examined, because measurable pools are often small, and P 
cycling is the major determinant of P availability. Since any "immediately available" pool 
of P is constantly replenished through dissolution or desorption of "less-available" P, and 
through the mineralization of organic P, "plant-available" P is strongly time-dependent. 


Agronomic tests for available P are designed with several aims; they should: 

1 Be simple enough for routine application. 

2 Extract sufficient P to be easily measurable. 


j Extract P that represents a significant portion of potential plant uptake, so that 
plant supply is represented closely by the quantity measured rather than being 
dependent on P turnover and replenishment of the measured pool. 

4 Not extract significant amounts of P that are not plant available over the growing 

This is achieved with moderately acidic or alkaline solutions which release P associated with 
the soil mineral phase without solubilizing significant amounts of phosphatic minerals. 
Alternatively, or in combination with these pH changes, specific anions are introduced that 
bring P into solution by competing with P sorption sites or by lowering the solubility of 
cations that bind P in the soil. Based on these principles, numerous extraction methods exist, 
all of which have some merits and limitations and are used in various parts of the world, 
where their value relies on long-term correlation studies that establish the relationship 
between extractable P and crop response. An exhaustive review of extraction methods by a 
working group in Spain (Anon. 1982) listed 50 different methods and more than 50 
publications comparing different extracts. 

The most common methods are probably the alkaline bicarbonate method of Olsen et al. 
(1954) and the acid ammonium fluoride extraction (Bray and Kurtz 1945) with various 
modifications. An extraction using lactate (Egner et al. 1960) is popular in Europe. The 
rationale for the use of bicarbonate or lactate for the extraction of available P is that plant 
roots produce CO2 which forms bicarbonate in the soil solution as well as various organic 
acids similar to lactate that may solubilize soil P. It is proposed that these extractants 
somehow simulate the solubilizing action of plant roots and, thus, give a more appropriate 
measure of plant-available P. Chelating extracts (Onken et al. 1980) have been proposed for 
similar reasons. An advantage of chelating extracts is that the same extract can also be used 
for cation soil testing (micronutrients and K). 

The bicarbonate extractant (Olsen et al. 1954) has been used successfully on a wide range of 
acid to alkaline soils. Available P is extracted with a solution of sodium bicarbonate of pH 
8.5 for 30 min. Interference from organic matter dissolved in the solution has frequently been 
eliminated by sorbing the organic matter onto activated acid-washed charcoal (carbon black) 
added to the extract, but it is difficult to obtain P-free charcoal. An alternative was therefore 
developed which eliminates organic interference with polyacrylamide (Banderis et al. 1976). 
If organic matter content in the extract is low (as judged by its yellow coloring) a blank 
correction can be used. When the blue phosphomolybdate complex is measured at a wave- 
length of 712 nm, color interference from the yellow organic matter is negligible. However, 
using color correction with blanks will not work at high organic matter concentrations in the 
extract because the organic matter will precipitate upon acidification during the Murphy and 
Riley (1962) procedure and interfere with P colorimetry. The extraction time of 30 min has 
been designed for rapid routine soil testing. A more complete extraction is obtained by 
extracting for 16 h (Colwell 1963). For all applications that attempt to functionally evaluate 
the bicarbonate-extractable P pool, and that include organic P determination, the more 
complete 16 h extract should be used, because at 30 min the extraction is far from complete. 

The acid ammonium fluoride extraction (Bray and Kurtz 1945) has been widely used on acid 
and neutral soils, and a large database exists. This is a purely chemical test that cannot be 
interpreted in terms of plant function like the bicarbonate or some of the organic acid or 
chelating extracts. Fluoride has been used to extract Al-associated P, but it is not obvious 
what the link to plant availability would be. In addition, Ca phosphates which are of low 


plant availability in high-pH soils would be extracted by the acid and give excessive values 
for available P. The relatively low acid strength and importance of acidity for the extraction 
mechanism make the method unsuitable for calcareous or strongly alkaline soils, which 
would partially neutralize the acidity and eliminate the standard test conditions. However, 
buffered variations of this soil test have been reported to correlate well with bicarbonate- 
extracted P and plant response to P (van Lierop 1988; Soon 1990). 


Since available P is a functional concept rather than a measurable quantity, no simple direct 
measurements are available. Plant-available P is that P taken up by a plant during a specific 
period, such as a cropping season, year, or growth cycle. Since the plant obtains P from the 
soil solution through its roots or root symbionts, available P is composed of solution P plus P 
that enters the solution during the period used to define availability. Phosphorus may enter 
the solution by desorption or dissolution of inorganic P (Pj) associated with the soil's solid 
phase, or by the mineralization of organic P (P ). In some dystrophic rain forests, P may not 
even cycle through the soil, but can be taken up directly from plant litter. 

It is difficult to resolve whether desorption or dissolution replenishes solution P from P ; forms. 
In one case the solubility product of the least-soluble P compound, and in the other, the 
saturation of sorbent surfaces would determine the P supply at equilibrium. Countless 
publications have fitted theoretical equations to the reverse of these reactions — precipitation 
and adsorption. Empirical data usually fit either process to some degree (Syers and Curtin 1 989 ). 
There is an increasing realization, though, that solid-phase P is not static, and that sorption- 
desorption and precipitation-dissolution equilibria change with time due to secondary processes 
(Parfitt et al. 1989) such as recrystallization (Barrow 1983) or solid-state diffusion (Willett et al. 
1988). A measurement of available Pj therefore needs to consider both the amounts and rates 
of release of P from the solid phase. Very few appropriate methods have been published. 
Among the approaches taken are repeated water extracts and sorption-desorption isotherms 
(Fox and Kamprath 1970; Bache and Williams 1971), possibly at elevated temperatures to 
substitute for impractically long reaction times (Barrow and Shaw 1975). 

A simple and more realistic approach is the use of anion-exchange resin, a sink for solution 
Pi. The resin offsets the equilibrium between dissolved and soluble P;, and "exchangeable" 
P ; as well as some of the more soluble precipitated P forms will enter the depleted solution 
and be absorbed by the resin. The P sorbed by the resin is subsequently measured. Several 
different methods have been developed and tested, using different anionic forms, ratios of 
soikwatenresin, times and methods of shaking, and enclosure in bags or mixing through the 
suspension (Sibbesen 1977, 1978; Barrow and Shaw 1977). By far the simplest method uses 
polyester- or Teflon-based anion-exchange membranes, which can be cut into strips and used 
repeatedly and easily (Saggar et al. 1990; Schoenau and Huang 1991). These ion-exchange 
membranes have also been used in situ, inserted or buried in soil where they integrate 
processes of nutrient release and diffusion (to the membrane) over time (Qian and Schoenau 
1997). When choosing resin membranes, it is important that the resin is part of the membrane 
material, i.e., cannot be abraded by the soil, and that they are resistant to the chemicals used 
in P extractions, such as dilute HC1 or chloroform (if microbial P is to be measured). It also 
helps if they are stiff enough to be easily handled. 

The pool measured by resin extraction is very similar to that assessed with isotopic dilution 
(Amer et al. 1955). The sorption by a resin is usually complete within 20 h, and only minor 


changes are observed thereafter. Isotopic exchange also reaches a relatively constant state 
within a few hours, and the disappearance of isotope from the solution is used to estimate the 
size of the labile pool into which the isotope has been diluted. In a variation of the isotopic 
dilution method, carrier-free 32 P is added to a soil suspension, and the initial rapid removal of 
label is measured. This is followed by a determination of the continuing slow changes 
(Fardeau and Jappe 1980). These continuing changes represent the activity of less soluble 
or kinetically slower pools of soil P, which replenish available P at rates varying from days to 
years. On some soils with low or moderate P sorption, the continuing reduction in radio- 
activity in the liquid phase of the suspension has been extrapolated to times corresponding to 
seasons or longer with some success in estimating plant-available P. However, errors of 
extrapolation over long times can be large (Biihler et al. 2003; Chen et al. 2003). Phosphorus 
taking part in longer term transformations can be examined with sequential extractions, 
which first remove labile P, and then the more stable forms. 

The sequential extraction method proposed by Chang and Jackson (1957) and modified by 
Williams et al. (1967) employs, sequentially, NH4CI to extract "labile" P;, NH 4 F to dissolve 
Al-associated Pi, NaOH to extract Fe-bound P;, and dithionite-citrate to dissolve ' 'occluded' ' P; 
forms. A subsequent extraction with HC1 dissolves Ca-bound P; and the residue is analyzed by 
Na 2 C0 3 fusion for residual total P. Alternatively, the residue can be analyzed for P by ignition 
plus acid extraction before the Na2C03 fusion (Williams et al. 1967). As in all other methods 
of P determination, the amount of P is not measured directly but calculated by difference: 
acid-extractable P; is subtracted from the greater amount of Pj rendered acid extractable after 
ignition of the soil organic matter (Saunders and Williams 1955) (see Chapter 24). 

The procedure presented many interpretational problems: P; can reprecipitate during the 
fluoride extraction, the separation of Al- and Fe-associated P; is not reliable, and the 
reductant soluble or occluded P; is an ill-defined pool (Williams and Walker 1969). 
However, the sequence of alkaline followed by acid extraction gives a reliable distinction 
between Al + Fe and Ca-associated P; (Kurmies 1972). This distinction reflects the weath- 
ering stage of the soil and can be used to monitor the fate of rock phosphate fertilizer in 
weathered soils that contain little Ca-bound P. The P extracted by the procedure has usually 
been ignored although it was shown to be important in plant nutrition (Kelly et al. 1983). 

An alternative P fractionation scheme was developed by Hedley et al. (1982a) building on 
the experience with previous extractions. This sequential extraction aims at quantifying 
labile (plant-available) P;, Ca-associated P;, Fe + Al-associated P;, as well as labile and 
more stable forms of P . Labile P;, i.e., P; adsorbed on surfaces of sesquioxides or carbonates 
(Mattingly 1975), is extracted with resin and bicarbonate. Hydroxide-extractable Pi is less 
plant available (Marks 1977) and is thought to consist of amorphous and some crystalline Al 
and Fe phosphates. A more precise characterization of these P ; forms is unlikely to be 
possible since mixed compounds containing Ca, Al, Fe, P, and other ions predominate in 
soils (Sawhney 1973). Organic P extracted with bicarbonate is easily mineralizable and 
contributes to plant-available P (Bowman and Cole 1978). More stable forms of P are 
extracted with hydroxide (Batsula and Krivonosova 1973). 

Each of the extracts obtained can be assigned some role in the P transformations occurring in 
soil under incubation (Hedley et al. 1982a) or cultivation (Tiessen et al. 1983), in the 
rhizosphere (Hedley et al. 1982b), or in soil development (Tiessen et al. 1984; Roberts 
et al. 1985; Schlesinger et al. 1998; Miller et al. 2001). These empirical assignments can then 
be used to characterize P status of the soil relative to a conceptual model of P pools and their 
transformations. Little has changed in the functional assignment or characteristics attributed 


to those P extracts since the papers published in the 1980s, although some authors group 
fractions in ways that reveal interesting concepts on the function of the soil P cycle. Cross 
and Schlesinger (2001) group Pi and P fractions of the different extracts together reporting 
each extract's total P, and implying that the mode of stabilization is the most important 
characteristic, not necessarily the distinction between organic and inorganic forms. Soil 
mineralogy clearly affects the interpretation of P fractions. In semiarid, calcareous soils, 
Cross and Schlesinger (2001) identified acid-extractable P not only as true calcium phos- 
phates but also as various associations of P with carbonates. To distinguish such fractions 
more clearly, Samadi and Gilkes (1998) added (among other modifications) an ammonium 
acetate extract before the acid e 

This fractionation approach is currently the only one that can be used with moderate success 
for the evaluation of available P . Cross and Schlesinger (1995) used the ratio of bicarbonate 
P to resin plus bicarbonate total P as an index for the bioavailability of P. This is probably 
only valid in temperate soils. Due to the reactivity of mineralized P with the soil's mineral 
phase, determination of a potentially mineralizable P pool, analogous to the mineralizable N 
or S pools measured with incubation and leaching techniques (Ellert and Bettany 1988), is 
not feasible. The nature of different extractable P pools is even less well defined than that of 
the P ; pools (Stewart and Tiessen 1987). Their turnover and availability frequently depend on 
the mineralization of C during which P is released as a side product, although solubilized P 
will be rapidly mineralized by soil enzymes. Most progress on understanding soil P has 
come from organic matter studies (Tiessen et al. 1983; Stewart and Tiessen 1987). It is often 
more appropriate to determine P in physical soil organic matter fractions, than to try and 
relate a chemically extracted P to biological function. Unless one has good reasons to 
believe that an extracted organic fraction can be biologically defined, it is probably best 
to group the organic fractions and use the sequential fractionation as a multiple extractant to 
obtain as much as possible of the soil's P . 

The original fractionation (Hedley et al. 1982a) left between 20% and 60% of the soil's P 
unextracted. This residue often contained significant amounts of P that sometimes partici- 
pated in relatively short-term transformations. On relatively young Ca-dominated soils, this 
residual P can be extracted by NaOH after the acid extraction, while on more weathered 
soils, hot HC1 (Mehta et al. 1954) extracts most of the organic and inorganic residual P. The 
hot HC1 method appears to work satisfactorily on most soils, and is presented below as part 
of an extensive soil P fractionation. 

25.4.1 Equipment and Materials 

7 50 ml_ centrifuge tubes with screw caps and refrigerated high-speed centrifuge 

2 Shaker, preferably overhead type so that soils do not clump together in the round 
bottom of the centrifuge tubes 

3 0.45 |j,m membrane filter and filtration apparatus 

4 Water bath 

5 Block digester with 75 or 100 ml_ digestion tubes 

g Autoclave or household pressure cooker 
y Plastic vials for storing extracts 
q Whatman No. 40 filter paper (or equivalent) 

25.4.2 Extracting Solutions 

7 0.5 M HCI: dilute 88.5 ml_ cone. HCI to 2 L with deionized H 2 0. 

2 0.5 M NaHC0 3 (pH 8.5): dissolve 84 g NaHC0 3 + 1 g NaOH in deionized H 2 
and make to 2 L. 

j 0.1 M NaOH: dissolve 4 g NaOH in deionized H 2 and bring final volume to1 L. 

4 1 M HCI: add 1 77 ml_ cone. HCI (1 1 .3 M) to about 500 ml_ of deionized H 2 and 
bring to final volume of 2 L. 

5 H 2 2 : 30% hydrogen peroxide. 

6 Concentrated H 2 S0 4 (1 8 M). 

7 Resin strips: use anion-exchange membrane cut into strips (9 x 62 mm) and 
convert to bicarbonate form. To regenerate after the adsorbed P has been 
extracted with HCI, wash resin strips for 3 days with 6 batches of 0.5 M HCI, 
followed by washing a further 3 days with 6 batches of 0.5 M NaHC0 3 (pH 8.5). 
Then rinse well with deionized/distilled water. 

25.4.3 Extraction Procedure 

Day 1: Weigh 0.5 g soil into a 50 ml_ centrifuge tube, add 2 resin strips +30 ml_ 
deionized water, and shake overnight (16 h, and 30 rpm if using overhead 
shaker). See comments below of fineness of grinding of soil samples. 

Day 2: Remove resin strips and wash soil back into tube using deionized water. 
Place resin strip in a clean 50 mL tube, add 20 ml_ 0.5 M HCI. Set aside for 
1 h to allow gas to escape, cap and shake overnight. Determine P using 
Murphy and Riley method (see section at top of p. 301). Centrifuge soil 
suspension at 25,000 g for 1 min at 0°C. Decant water through a 0.45 |xm 
membrane filter. Discard water and wash any soil off filter back into the tube 
with a little 0.5 M NaHC0 3 (pH 8.5) solution. Add more NaHC0 3 solution to 
bring solution volume to 30 mL (by weighing) and shake suspension overnight 
(16 h). Cap the tubes and resuspend soil by handshaking before putting on 
mechanical shaker. 

Day 3: Centrifuge soil suspension at 25,000 g for 10 min at 0°C. Decant NaHC0 3 
extract through a membrane filter into a clean vial. Determine inorganic and 
total P on bicarbonate extract. Wash any soil off filter back into the tube using 
a little 0.1 M NaOH. Make volume of NaOH solution to 30 mL and shake 
suspension overnight (16 h). 


Day 4: Centrifuge suspension at 25,000 g for 10 min at 0°C. Decant NaOH extract 
through a membrane filter into a clean vial. Determine inorganic and total P 
on NaOH extract. Wash any soil off filter back into the tube using a little 1 M 
HCI. Make volume of HCI to 30 mL and shake suspension overnight. 

Day 5: 

7 Centrifuge soil suspension at 25,000 g for 10 min at 0°C. Decant HCI extract 
through a membrane filter into a clean vial. Determine P in extract. (In this step, 
any residue that right be on the filter paper is not washed back into the tube; 
decant gently so as to not lose any soil.) 

2 Soil residue heated with 1 mL cone. HCI in a waterbath at 80°C for 1 min. (Vortex 
to mix soil and HCI well and loosen caps before putting into the hot bath. The 
mixture will take about 1 min to come to temperature — check with a thermometer 
in a tube containing HCI only — i.e., the tubes will be in the hot water for a total of 
20 min.) Remove and add a further 5 mL cone. HCI, vortex and allow to stand 
at room temperature for 1 h (vortex every 15 min). Tighten caps, centrifuge at 
25,000 g for 10 min at 0°C, and decant supernatant into a 50 mL volumetric flask. 
Wash soil twice with 10 mL H 2 0, centrifuge, and add supernatant solution to 
contents in the flask. Make to volume, and if necessary filter through a Whatman 
No. 40 paper (or equivalent), and determine inorganic and total P in HCI solution. 

j Add 10 mL deionized water to soil residue and disperse soil. Transfer suspension 
into 75 mL digestion tubes using the minimum amount of water possible to transfer 
all soil residues, add 5 mL cone. H 2 S0 4 + one boiling chip (Hengar Granules, 
Hengar Co., Philadelphia, Cat. No. 1 36C), vortex and put on a cold digestion 
block. Raise the temperature very slowly to evaporate water and when 360°C is 
reached start treating with H 2 2 in the following way: remove tubes from heat and 
let cool to hand-warm; add 0.5 mLof H 2 2 ; reheat for 30 min, during which H 2 2 is 
used up. Repeat H 2 2 addition until liquid is clear (usually about 10 times). Make 
sure there is adequate heating after the final H 2 2 addition, since residual H 2 2 
interferes with the P determination. Cool, make to volume, shake, and transfer to 
vials (either filter or allow residue to settle out overnight). Determine P in solution 
(see Section at top of p. 300). (This digestion is based on Thomas et al. 1 967.) 


1 The intensity of soil grinding greatly affects the amount of Pextractable, particularly 
for the resin extraction which removes P from accessible surfaces. Interlaboratory 
testing has shown differences of an order of magnitude attributable to grinding 
between 2 mm screened and 60 mesh ground samples. The decision on how fine to 
grind should be based on a trade-off between sample homogeneity (important in a 
sample size of only 0.5 g) and preservation of the "natural" extractability of resin P. 
We have generally opted for moderate crushing of samples to 20 mesh. 

2 Sequential extraction is a lengthy procedure. A batch of samples will take a week 
(including the weekend) to process. It is therefore important to reconcile the aim 
of the study with what this fractionation can produce. If geological transform- 
ations are the target, one can probably do without the resin and bicarbonate 
extracts; if labile pools are the target, the more resistant fractions may not be 


important, and it would be more useful to include microbial P or organic matter 
separations. In many highly weathered soils, cold acid-soluble P is so little that it 
is probably a "contaminant" from the previous extract. 

25.4.5 Analysis of P in Extracts 

Reagents for P Determination 

/ Ammonium molybdate: dissolve 40.0 g ammonium molybdate in deionized H 2 
and bring to a final volume of 1 L. 

2 Ascorbic acid: dissolve 26.4 g L-ascorbic acid in deionized H2O and bring to a 
final volume of 0.5 L. 

j Antimony potassium tartrate: dissolve 1.454 g antimony potassium tartrate in 
deionized H 2 and bring to a final volume of 0.5 L. 

4 2.5 M H 2 S0 4 : slowly add 278 ml_ cone. H 2 S0 4 to 1 L of deionized H 2 and 
bring to a final volume of 2 L. 

5 Color developing reagent: to 250 ml_ 2.5 M H 2 SC>4, add 75 ml_ ammonium 
molybdate solution, then 50 ml_ ascorbate solution and finally 25 ml_ of antimony 
potassium tartrate solution. Swirl contents of flask after each addition. Dilute to a 
total volume of 500 ml_ with deionized H 2 and mix. 

6 For organic matter precipitation and pH adjustment make up: 
0.9 M H 2 S0 4 : bring 100 ml_ cone. H 2 S0 4 to 2 L with H 2 0. 
0.25 M H 2 S0 4 : bring 100 ml_ 2.5 M H 2 S0 4 to 1 L with H 2 0. 
4 M NaOH: dissolve 1 60 g NaOH and dilute to 1 L with H 2 0. 

7 p-nitrophenol, 10% (w/v), aqueous solution, 
g Ammonium persulfate, (NH 4 ) 2 S 2 O s . 

Determination of P Recovered from the Resin Strip and of Pi in HCI Extracts 

This method (Murphy and Riley 1962) is used directly for the P recovered from the resin 
strip and for P; determination in the two HCI extracts: 

7 Pipette a suitable aliquot into a 50 ml_ volumetric flask. The calibration curve 
is linear for up to a concentration of about 1 mg of P L~ 1 . Use two drops of 
p-nitrophenol as an indicator. If the extract is acid, first adjust pH with 4 M NaOH 
to yellow and then with ~0.25 M H 2 S0 4 until the indicator turns clear. For 
alkaline extracts, just acidify until solution is clear. Note that most analytical 
problems are related to the solution being adjusted too acid. 

2 Add 8 ml_ of color developing solution, make to volume, shake and read on 
spectrophotometer at 712 nm after 10 min (color is stable for several hours). 


Determination of Inorganic P in 0.5 M NaHC0 3 and 0.1 M NaOH Extracts 

7 Pipette 10 mL solution into a 50 ml_ centrifuge tube. 

2 Acidify to pH 1 .5 and set in fridge for 30 min: 

(a) to acidify 0.5 M NaHC0 3 extract use: 6 mL of 0.9 M H 2 S0 4 ; 

(b) to acidify 0.1 M NaOH extract use: 1 .6 mL of 0.9 M H 2 S0 4 . 

3 Centrifuge at 25,000 g for 1 min at 0°C. 

4 Decant supernatant into a 50 mL volumetric flask. 

5 Rinse tube carefully so as not to disturb the organic matter with a little acidified 
water and add to the solution in the flask (2 or 3 times). 

g Adjust pH and measure P by the Murphy and Riley method (see section at bottom 
of p. 300). 

Determination of Total P in 0.5 M NaHC0 3 , 0.1 M NaOH, and Cone. HCI Extracts 
(EPA 1971) 

Dissolved organic matter is oxidized with ammonium persulfate before P analysis: 
; Pipette 5 mL solution into a 50 mL volumetric flask. 

2 To 0.5 M NaHC0 3 extract: add -0.5 g ammonium persulfate + 10 mL 
0.9/VfH 2 SO 4 . 

(a) To 0.1 M NaOH extract: add ~0.6 g ammonium persulfate + 10 ml 
0.9 M H 2 S0 4 . 

(b) To cone. HCI extract: add ~0.4 g ammonium persulfate + 10 mL deionized 

The persulfate may be added by volume using a spatula with a spoon at one end 
rather than weighing every time. 

3 Cover with tinfoil (double layer for cone. HCI) and autoclave: 

NaHC0 3 and HCI extracts for 60 min, NaOH extract for 90 min. (Instead of an 
autoclave, a household pressure cooker can also be used.) 

4 Adjust pH and measure P by the Murphy and Riley method (see section at top 
of p. 300). 


7 The aliquot size of extract for the Murphy and Riley procedure may vary from 
1 mL for high P concentration acid extracts up to 40 mL in the case of very low P 
resin extracts. 


2 Most times when things go wrong, it is due to interferences in the Murphy and Riley 
colorimetry. Insufficient pH control before color development is the most common 
problem with color development. Residual oxidant from one of the digestion steps 
will of course interfere with the reduction step of the color development. In some 
soils, we have seen interference from soluble silica in the reacidified NaOH extract, 
resulting in a positive drift (i.e., increase) in absorbance. This interference is difficult 
to manage if it occurs. Very consistent absorbance reading at exactly 1 min helps 
but results will remain doubtful. 


The interpretation of data obtained from this sequential fractionation is based on an under- 
standing of the action of the individual extractants, their sequence (Figure 25.1), and their 
relationship to the chemical and biological properties of the soil. It must be remembered that, 
while the fractionation is an attempt to separate P pools according to their lability, any 
chemical fractionation can at best only approximate biological functions. Resin P is reason- 
ably well defined as freely exchangeable P;, since the resin extract does not chemically 
modify the soil solution. Bicarbonate extracts a P ; fraction, which is likely to be plant 
available, since the chemical changes introduced are minor and somewhat representative 
of root action (respiration). This fraction is not comparable to the widely used fertility test 

0.5 g soil 

Extract with resin strip 

- Bicarb-extractable P t -*- Digest, determine P, 

-+■ OH-extractable P, -+■ Digest, determine P t 
I Precipitate organic 
y matter 
^- Determine P, 

► Determine Pj 

Digest, determine P t 
Determine P, 

— ^- Digest, determine P t 

HCI-extractable P t 

FIGURE 25.1. Flow chart of the seque 


(Olsen et al. 1954) because the resin-extractable pool has already been removed at this point 
and because Olsen P is extracted over only 30 min. 

Bicarbonate-Pj and OH-P; are not really completely separate pools, particularly in acid soils, 
but represent a continuum of Fe- and Al-associated P extractable with increasing pH 
(the soils original pH to 8.5 to 13). The P extracted with these two extractants is also likely 
to represent similar pools. Since P is determined by difference between total P (P t ) and P; 
in each extract, there is a source of error. The P t determination is quite reliable, but 
Pi is determined in the supernatant after precipitation of organic matter with acid. Any 
nonprecipitated P (fulvic acid P) will not significantly react with the Murphy and Riley 
reagent, so that P; is rarely overestimated. Any P;, though, that precipitates along with the 
organic matter upon acidification would be erroneously determined as P (P t — Pi). This may 
happen with P ; associated with Fe or Al hydroxides, which are soluble at high pH but 
insoluble at low pH. It has so far been impossible to quantify the P overestimation. In soils 
with low-extractable organic matter contents (low enough not to cause precipitation in the 
acid Murphy and Riley reagent), it is possible to determine P; in the extract without prior 
acid precipitation using a blank correction for the extracts' color. 

The dilute HC1 Pj is clearly defined as Ca-associated P, since Fe- or Al-associated P that 
might remain unextracted after the NaOH extraction is insoluble in acid. There is rarely any 
P in this extract. Dilute acid is well known to be inefficient in extracting organic carbon 
from soils, and therefore, does not extract much P . 

The hot concentrated HC1 extract does not present the same problems as the other P 
extracts, since Pi is determined directly. This extract is useful for distinguishing P; and P 
in very stable residual pools. However, at the same time, P extracted at this step may simply 
come from particulate organic matter that is not alkali extractable but may be easily bioavail- 
able. Any P protected by cellulosic structure would be biologically available as a byproduct of 
cellulose breakdown, but would only become extractable in the hot concentrated acid step. 
The residue left after the hot concentrated HC1 extraction is unlikely to contain anything but 
highly recalcitrant P;. 

It is important to remember that this sequential extraction does not provide direct measures 
of biologically or geochemically important P pools. It provides circumstantial evidence that 
is more valuable if it can be corroborated by other methods such as isotope or organic matter 
studies. Particularly, for a reliable interpretation of P transformations, it is advisable to 
supplement the fractionation with a suitable characterization of soil organic matter, so that 
characteristics can be inferred from the combined results of different techniques. 

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Chapter 26 
Extractable Al, Fe, Mn, and Si 

F. Courchesne and M.-C. Turmel 

University of Montreal 
Montreal, Quebec, Canada 


The dissolution methods for extracting Al, Fe, Mn, and Si are valuable tools to help 
determine the chemical forms of these elements in soils. The results are useful in studies 
of soil classification, soil genesis, soil reactivity, and metal mobility or bioavailability in 
soils. For example, the nature and amounts of extractable organic and inorganic Al and Fe 
may reflect the pathway of soil genesis. Also, extractable soil constituents are generally fine 
grained with large specific surface area and therefore, have a marked effect on physical and 
chemical soil properties and behavior. For these reasons, extraction data, notably for Al and 
Fe, are commonly used as chemical criteria for soil classification. Moreover, extractions are 
often performed to establish the mechanisms of metal retention and fractionation in con- 
taminated soils. A variety of chemical extractants are used to approximate the amounts and 
forms of Al, Fe, Mn, and Si in soils. Five of the most commonly used extractions are 
discussed here and four methods are presented. 

Dithionite-citrate removes organically complexed Al, Fe, and Mn, amorphous inorganic Al, 
Fe, and Mn compounds, noncrystalline aluminosilicates as well as finely divided hematite, 
goethite, lepidocrocite, and ferrihydrite (Mehra and Jackson 1960; Guest et al. 2002). It is 
much less effective in removing crystalline oxides and hydroxides of Al. The method 
extracts virtually no Al, Fe, Mn, or Si from most crystalline silicate minerals or opal, and 
thus, provides an estimate of "free" (nonsilicate) Fe in soils. The procedure may have to be 
repeated to dissolve silt- and sand-size goethite and hematite completely (Kodama and Ross 
1991). Magnetite is not dissolved. Ross and Wang (1993) indicated that coefficients of 
variation at Fe levels of 1.4% and Al levels of 0.45% are 6.3% and 7.8%, respectively. 

Acid ammonium oxalate removes organically complexed and amorphous inorganic forms of 
Al, Fe and, to a lesser extent, Mn and noncrystalline aluminosilicates from soils (McKeague 
1967). It also dissolves poorly ordered phases like allophane and imogolite to some extent 
and their amount in soils can be estimated from oxalate-extractable Al and Si concentrations, 
taking into account that oxalate also extracts organically complexed Al (Parfitt and Henmi 
1982). Oxalate only slightly attacks crystalline Al and Fe oxides, most crystalline silicate 
minerals, opal, goethite, hematite, and lepidocrocite, but it dissolves considerable a 


magnetite (Baril and Bitton 1967) and of finely divided, easily weathered silicates, such as 
olivine. Ross and Wang (1993) indicated that coefficients of variation at Fe levels of 0.67% 
and Al levels of 0.67% are 7.2% and 4.1%, respectively. 

Hydroxylamine is closely similar to oxalate in its extraction capacity (Chao and Zhou 1983). 
It is also commonly used to extract soil Mn. Unlike ammonium oxalate, however, hydroxy- 
lamine does not dissolve magnetite and can therefore be used as an alternative to ammonium 
oxalate for soils containing magnetite (Ross et al. 1985). Ross and Wang (1993) indicated 
that the coefficients of variation at Fe levels of 0.63% and Al levels of 0.62% are 4.5% and 
3.0%, respectively. 

Tiron, 4,5-dihydroxy-l,3-benzene-disulfonic acid (disodium salt), does not dissolve 
magnetite either and its use has been suggested instead of oxalate (Kodama and Ross 
1991). Furthermore, Tiron dissolves pedogenic opaline silica (Kendrick and Graham 
2004), whereas, neither oxalate nor hydroxylamine dissolves this soil component effectively. 
Tiron is currently used mainly to remove coatings on clays (Ross and Wang 1993). However, 
it should also be suitable for soils ground to pass a 0.15 mm (100-mesh) sieve. 

Sodium pyrophosphate extracts organically complexed Al and Fe from soils. Manganese 
compounds are also dissolved. It only slightly dissolves noncrystalline inorganic forms, and 
it does not significantly attack silicate minerals and crystalline Al and Fe oxides or hydrox- 
ides (McKeague et al. 1971). The pyrophosphate solution does not dissolve opal and is a poor 
extractant for allophane or imogolite (Wada 1989). Ross and Wang (1993) indicated that the 
coefficients of variation at Fe levels of 0.64% and Al levels of 0.69% are 5.9% and 6.0%, 
respectively. The specificity of the method for organic complexes of Al and Fe has been 
challenged because amorphous and poorly ordered inorganic Al and Fe solid phases were 
found to be significantly removed by the pyrophosphate extract (Kaiser and Zech 1996). 

From the results of these methods, the following quantities can be estimated: 

A. Finely divided crystalline Fe solid phases like goethite, hematite, and lepidocro- 
cite: dithionite Fe-oxalate Fe or dithionite Fe-hydroxylamine Fe or dithionite 
Fe-Tiron Fe 

B. Noncrystalline inorganic forms of Fe including ferrihydrite: oxalate Fe-pyrophosphate 
Fe or hydroxylamine Fe-pyrophosphate Fe or Tiron Fe-pyrophosphate Fe 

C. Organic complexed Fe: pyrophosphate Fe 

Relationships B and C also hold approximately for Al; this is not the case for relationship A. 
In the case of Mn, both dithionite and oxalate attack crystalline oxide forms to some extent 
and differences between extracts are not easy to interpret. The noncrystalline forms of Si, 
such as opaline silica, are completely extracted only by Tiron (Kodama and Ross 1991). 
They are not extracted by oxalate and only partly by dithionite and hydroxylamine. Poorly 
crystalline and noncrystalline aluminosilicates, including allophane and imogolite, are 
extracted by oxalate, hydroxylamine, and Tiron. Dithionite and pyrophosphate are much 
less effective in extracting these compounds. 

A survey of the literature on the extraction of Al, Fe, Mn, and Si from soils clearly shows that 
the laboratory procedures employed vary considerably among extractions and between 


studies. Yet, results from different extractions are frequently compared in studies on soil 
genesis and metal fractionation. Moreover, the effects of the grinding of soil samples and of 
the filtration of extracts on the amounts of Al, Fe, Mn, or Si extracted are documented 
(Loveland and Digby 1984; Neary and Barnes 1993). The reduction of particle size by 
grinding is nonetheless necessary when weighing out small subsamples because it increases 
the homogeneity between subsamples and, thus, the repeatability of the extraction. In this 
context, and because of the operational character of the extraction schemes, investigators are 
strongly encouraged to report the procedures they used, notably, with respect to the prepar- 
ation of soil samples (sieving, grinding) and to the centrifugation (g force) or the filtration of 
extracts (type of membrane, pore size). In the methods proposed here, all the samples are 
ground to 0.15 mm. 


The dithionite-citrate method consists of shaking soils overnight in the presence of a reducing 
and complexing solution. Dithionite creates a reducing environment and dissolves metallic 
oxides whereas the Na-citrate chelates the dissolved metals and buffers the pH to near 7 to 
avoid the precipitation of FeS compounds. This treatment is particularly useful for dissolving 
the "free" Fe in soils. Caution must however be exercised when interpreting extracted Al. 
The overnight shaking procedure is simpler than the dithionite-citrate-bicarbonate method 
of Mehra and Jackson (1960), and it gives closely similar results (Sheldrick and McKeague 
1975). This extractant is used in the Canadian System of Soil Classification (Soil Classifica- 
tion Working Group 1998) for describing Fe accumulation in Gleysols. 

26.2.1 Reagents 

7 Sodium hydrosulfite (dithionite), Na2S20 4 

2 Sodium citrate (NaaCeHsO/ • 2H 2 0), 0.68 M (200 g L" 1 ) 

j Certified atomic absorption standards, ±1% 

26.2.2 Procedure 

j Weigh 0.500 g of <2 mm air-dry soil, ground to pass a 0.1 5 mm (1 00 mesh) sieve, 
into a 50 mL screw-cap plastic centrifuge tube (use 0.2 g for clays and 1 g for 
coarse soils). 

2 Add 25 mL of the sodium citrate solution. 

3 Add about 0.4 g of dithionite (a calibrated scoop may be used). 

4 Stopper tightly and shake overnight (1 6 h) in an end-over-end shaker. A horizontal 
shaker can also be used although interparticle abrasion can be increased. 

5 Remove stoppers and centrifuge for 20 min at 510 g (centrifuge at higher 
speed for samples rich in clay particles). Filter extracts containing suspended 


g For determining Al, Fe, Mn, and Si by atomic absorption spectroscopy (AAS), 
prepare standard solutions of these elements containing the same concentration of 
extracting solution, here Na-citrate with dithionite, as the extracting solution. 
Gently heat the solution to dissolve dithionite. Note that at high concentration, 
the precipitation of dithionite can rapidly block the AAS burner. The amount of 
dithionite added to standard solutions can be lowered to reduce this effect. 

j An air-acetylene flame is suitable for the determination of Fe and Mn, and a 
nitrous oxide-acetylene flame is used for Al and Si. 

g If it is necessary to dilute the extracts, either dilute them with the extracting 
solution or prepare standards containing the same concentration of extracting 
solution as the diluted extracts. 

26.2.3 Calculations 

in final solution x extractant (mL) x dilution ,„, „ , 

/ % Fe, Al, Mn, Si = *- 

sample weight (g) x 1 0,000 

For example, for 0.500 g of sample, 25 mL of extractant, 5 times dilution, and a 
48 jULg Fe ml_~ 1 concentration: 

% Fe in sample = 

0.500 x 10,000 


The acid NH4-oxalate method was developed in 1922 by Tamm to remove the sesquioxide 
weathering products from soils. It was revised by Schwertmann (1959), who showed that it 
could estimate noncrystalline and poorly ordered Al and Fe forms in soils. It extracts the 
amorphous Al and Fe accumulated in podzolic B horizons (McKeague and Day 1966) and is 
thus useful to identify podzolic B horizons. Oxalate also dissolves allophane and imogolite 
(Wada 1989). In the soil taxonomy, amounts of Al and Fe extracted with oxalate are criteria 
for andic soil properties (Soil Survey Staff 1990). The extraction must be conducted in the 
dark to prevent photodecomposition of the oxalate solution. 

26.3.1 Reagents 

7 Solution A: Ammonium oxalate solution (NH 4 ) 2 C 2 4 • H 2 0, 0.2 M (28.3 g L _1 ). 
2 Solution B: Oxalic acid solution H 2 C 2 4 • 2H 2 0, 0.2 M (25.2 g L _1 ). 
j Mix 700 mL of A and 535 mL of B, check pH, and adjust to 3.0 by adding A or B. 
4 Certified atomic absorption standards, ±1%. 


26.3.2 Procedure 

1 Weigh 0.250 g of <2 mm air-dry soil, ground to pass a 0.1 5 mm (1 00 mesh) sieve, 
into a 15 ml_ screw-cap plastic centrifuge tube (weigh 0.125 g for samples with 
>2% extractable Fe or Al). 

2 Add 1 ml_ of the acid ammonium oxalate solution and stopper the tube tightly. 

3 Place the tubes in an end-over-end shaker and shake for 4 h in the dark. 
A horizontal shaker can also be used although interparticle abrasion can be 

4 Centrifuge the tubes for 20 min at 51 g (centrifuge at higher speed for samples 
rich in clay particles), decant the clear supernatant into a suitable container, 
and analyze within a few days. Extracts should be stored in the dark to avoid 
the photoinduced degradation of oxalate and the subsequent precipitation of 
dissolved metals. 

5 For determining Al, Fe, Mn, and Si by atomic absorption, follow standard 
atomic absorption procedures. Consider the points mentioned in Section 26.2.2. 

26.3.3 Calculations 

sample weight (g) x 1 0,000 

For example, for 0.250 g of sample, 10 mL of extractant, 5 times dilution, and a 
12 jULg Fe mL -1 concentration: 

% Fe in sample = 

0.250 x 10,000 

(ROSS ET AL. 1985; WANG ET AL. 1987) 

The acid hydroxylamine extraction is used in geochemical studies for removing noncrystal- 
line material, notably hydrous Mn oxides, from crystalline Fe oxides with minimal dissolu- 
tion of associated Fe oxides like magnetite (Chao and Zhou 1983). Ross et al. (1985) and 
Wang et al. (1987) modified this procedure and tested it on soil samples. For Al and Fe, the 
results were similar to those obtained by oxalate extraction with the advantage that hydroxy- 
lamine did not dissolve magnetite. There was less agreement between the Si results obtained 
by the two methods. The suitability of hydroxylamine as an extractant for Mn in soils has not 
been fully tested yet. Hydroxylamine solutions are also more easily analyzed than oxalate 
solutions by AAS because the latter tend to clog the burner. 


26.4.1 Reagents 

7 Prepare a hydroxylamine hydrochloride-hydrochloric acid (0.25 M NH 2 OH • HCI, 
0.25 M HCI) solution by adding 21.5 ml_ of concentrated HCI and 
17.37 g of NH 2 OH • HCI to a 1 L volumetric flask and making to volume with 
deionized water. 

2 Certified atomic absorption standards, ±1%. 

26.4.2 Procedure 

1 Weigh 0.1 00 g of <2 mm air-dry soil, ground to pass a 0.1 5 mm (1 00 mesh) sieve, 
into a 50 ml_ screw-cap plastic centrifuge tube. 

2 Add 25 mL of the hydroxylamine solution and stopper the tube tightly. 

j Place the tubes in an end-over-end shaker and shake overnight (16 h). A 
horizontal shaker can also be used although interparticle abrasion can be increased. 

4 Centrifuge the tubes for 20 min at 51 g (centrifuge at higher speed for samples 
rich in clay particles), decant the clear supernatant into a suitable container, and 
analyze within a few days. 

5 For determining Al, Fe, Mn, and Si by atomic absorption, follow standard atomic 
absorption procedures. Consider the points mentioned in Section 26.2.2. 

26.4.3 Calculations 

„, _ ,. ,, „. iJigmL -1 in final solution x extractant(mL) x dilution 

7 %Fe, AI,Mn,Si = — ; r — (26.5) 

sample weight (g) x 1 0,000 

2 For example, for 0.1 00 g of sample, 25 mL of extractant, 5 times dilution, and a 
6 |jug Fe ml_~ 1 concentration: 

% Fe in sample = 

0.100 x 10,000 


Sodium pyrophosphate is a common extractant for Al, Fe, and Mn associated with soil 
organic matter. It does not extract opal or crystalline silicates. The method is used in the 
Canadian System of Soil Classification as chemical criteria for identifying podzolic B 
horizons, in the soil taxonomy for spodic horizons and by the FAO for classifying podzolic 
soils (Soil Survey Staff 1990; FAO 1990; Soil Classification Working Group 1998). The 
pyrophosphate extraction is strongly dependent on the centrifugation and filtration proced- 
ures because, in some cases, finely divided colloidal silicates and oxides remain dispersed 
after low-speed centrifugation. High-speed centrifugation or ultrafiltration is then necessary 
to clear the extracts (McKeague and Schuppli 1982; Schuppli et al. 1983). 


26.5.1 Reagents 

j Sodium pyrophosphate solution (Na 4 P 2 7 • 10H 2 O), 0.1 M (44.6 g L~ 1 ). 

2 Superfloc (N-100) 0.1% (1.0 g L" 1 ). Available from Cytec Canada Inc., 7900 
Taschereau Bid, A-106 Suite, Brassard, Que., J4X 1C2. 

3 Certified atomic absorption standards, ±1%. 

26.5.2 Procedure 

7 Weigh 0.300 g of <2 mm air-dry soil, ground to pass a 0.1 5 mm (1 00 mesh) sieve, 
into a 50 ml_ screw-cap plastic centrifuge tube (use 1 g for samples low in 
extractable Fe and Al). 

2 Add 30 ml_ of sodium pyrophosphate solution and stopper the tube tightly. 

j Shake overnight (1 6 h) in an end-over-end shaker. A horizontal shaker can also be 
used although interparticle abrasion can be increased. 

4 Centrifuge at 20,000 g for 10 min or, alternatively, add 0.5 mL of 0.1% superfloc 
solution and centrifuge at 510 g for 10 min. Note the following points: 

a. Concentrations of Fe and Al in sodium pyrophosphate extracts of some sam- 
ples may decrease progressively by centrifugation for longer times or at higher 

b. Ultrafiltration through a 0.025 jjim Millipore filter is recommended for 
tropical soils and for samples giving questionable results by the centrifugation 

5 Decant a portion of the clear supernatant into a suitable container and analyze 
within a few days. Extracts containing suspended materials should be filtered. 

g For determining Al, Fe, and Mn by atomic absorption, follow standard atomic 
absorption procedures. Consider the points mentioned in Section 26.2.2. 

26.5.3 Calculations 

n/ _ .. kl jjLgmL -1 in final solution xextractant(mL) 

7 % Fe, Al, Mn = — , — — 

samp e weight g)x 10,000 

For example, for 0.300 g of sample, 30 mL of extractant, and a 75 |xg Fe mL 

% Fe in sample = 

0.300 x 10,000 


Baril, R. and Bitton, G. 1967. Anomalous values 
of free iron in some Quebec soils containing 
magnetite. Can. J. Soil Sci. 47: 261. 

Chao, T.T. and Zhou, L. 1983. Extraction tech- 
niques for selective dissolution of amorphous iron 
oxides from soils and sediments. Soil Sci. Soc. 
Am. J. 47: 225-232. 

FAO (Food and Agriculture Organization of the 
United Nations). 1990. Soil Map of the World: 

Revised Legend. Rome. 

Guest, C.A., Schulze, D.G., Thompson, LA., and 
Huber, D.M. 2002. Correlating manganese x-ray 
absorption near-edge structure spectra with extract- 
able soil manganese. Soil Sci. Soc. Am. J. 66: 

Kaiser, K. and Zech, W. 1996. Defects in esti- 
mation of aluminum in humus comple> 
podzolic soils by pyrophosphati 
Sci. 161: 452-458. 

differentiating various 
/. Soil Sci. 46: 13-22. 

classes of soils. Can. 

Kendrick, K.J. and Graham, R.C. 2004. 

silica accumulation in chronosequence soils, 

Southern California. Soil Sci. Soc. Am. J. 68: 


Kodama, H. and Ross, G.J. 1 99 1 . Tiron dissolution 
method used to remove and characterize inorganic 
compounds in soils. Soil Sci. Soc. Am. ./. 55: 

Loveland, P.J. and Digby, P. 1984. The t 

of Fe and Al by 0.1 M pyrophosphate solutions: a 

comparison of some techniques. J. Soil Sci. 35: 


McKeague, J.A. 1967. An evaluation of 0.1 M 
pyrophosphate and pyrophosphate-dithionite in 
comparison with oxalate as extractants of the 
accumulation products in Podzols and some 
other soils. Can. J. Soil Sci. 47: 95-99. 

McKeague, J.A., Brydon, J.E., and Miles, N.M. 
1971. Differentiation of forms of extractable iron 
and aluminum in soils. Soil Sci. Soc. Am. Proc. 
35: 33-38. 

McKeague, J.A. and Day, J.H. 1966. Dithio: 

and oxalate-extractable Fe and Al as aids 

McKeague, J.A. and Schuppli, PA. 1982. Changes 
in concentration of Fe and Al in pyrophosphate 
extracts of soil, and composition of sediment 
resulting from ultra centrifugation in relation to 
spodic horizons. Soil Sci. 134: 265-270. 

Mehra, O.P. and Jackson, M.L. 1960. Iron oxide 
removal from soils and clays by a dithionite- 
citrate system buffered with sodium bicarbonate. 
Clays Clay Miner. 7: 317-327. 

Neary, A.J. and Barnes, S.R. 1993. The effect of 
sample grinding on extractable iron and alumi- 
num in soils. Can. J. Soil Sci. 73: 73-80. 

Parfitt, R.L. and Henmi, T. 1982. Comparison of 
an oxalate-extraction method and an infrared 
spectroscopic method for determining allophane 
in soil clays. Soil Sci. Plant Nutr. 28: 183-190. 

Ross, G.J. and Wang, C. 1993. Extractable Al, Fe, 
Mn, and Si. In M.R. Carter, Ed. Soil Sampling and 
Methods of Analysis. Lewis Publishers, Boca 
Raton, FL, pp. 239-246. 

Ross, G.J., Wang, C, and Schuppli, P.A. 1985. 
Hydroxylamine and ammonium oxalate solutions 
as extractants for Fe and Al from soil. Soil Sci. 
Soc. Am. J. 49: 783-785. 

Schuppli, PA., Ross, G.J., and McKeague, J.A. 
1983. The effective removal of suspended mater- 
ials from pyrophosphate extracts of soils from 
tropical and temperate regions. Soil Sci. Soc. 
Am. J. 47: 1026-1032. 

Schwertmann, U. 1959. Die frectionierte Extrac- 
tion der freien Eisenoxide in Boden, ihre miner- 
alogischen Formen und ihre Entstehungsweisen. Z. 

Pflanzenemaehr. Dueng. Bodeiikimd 84: 19 -I 20 i. 

Sheldrick, B.H. and McKeague, J.A. 1975. A 
comparison of extractable Fe and Al data using 
methods followed in the USA and Canada. Can. 
J. Soil Sci. 55: 77-78. 

Soil Classification Working Group. 1998. The 
Canadian System of Soil Classification. Agricul- 
ture and Agri-food Canada, Ottawa, Canada. 


Soil Conservation Service, U.S. Department Wada, K. 1989. Allophane and imogolite. In J.B. 

of Agriculture. 1972. Soil survey laboratory Dixon and S.B. Weed, Eds. Minerals in Soil 

methods and procedures for collecting soil sam- Environments. Soil Science Society of America, 

pies. Soil Survey Investigations Report No. 1 Madison, WI, pp. 1051-1087. 
(revised), U.S. Government Printing Office, 

Washington, DC. Wang, C, Schuppli, PA., and Ross, GJ. 1987. A 
comparison of hydroxylamine and ammonium 

Soil Survey Staff. 1990. Keys to Soil Taxonomy, oxalate solutions as extractants for Al, Fe, and 

4th ed. SMSS Technical Monograph No. 6. Si from Spodosols and Spodosol-like soils in 

Blacksburg, VA. Canada. Geoderma 40: 345-355. 



Chapter 27 

Determining Nutrient 

Availability in Forest Soils 

N. Belanger 

University of Sasketchewan 
Saskatoon, Saskatchewan, Canada 

D. Pare 

Natural Resources Canada 
Quebec, Quebec, Canada 

W.H. Hendershot 

McGill University 
Sainte Anne de Bellevue, Quebec, Canada 


Nitrogen (N) is the major nutrient determining tree growth, and this has been demonstrated 
abundantly in the Boreal Shield with fertilization trials or net N mineralization studies (e.g., 
Attiwil and Adams 1993; Reich et al. 1997). However, Ingestad (1979a,b) also showed that 
any other nutrient (but particularly phosphorus (P) and potassium (K)) could be limiting if 
supplied at a rate lower than tree demand, even if N was in excess. For example, 
fertilization trials with N alone or in combination with P, K, or both stimulated the growth 
of black spruce (Picea mariana (Mill.) BSP) (e.g., Wells 1994; Paquin et al. 1998) and jack 
pine (Pinus banksiana Lamb.) (e.g., Morrison and Foster 1995; Weetman et al. 1995). 
A much lower number of studies have showed the benefits of increased calcium (Ca) and 
magnesium (Mg) availability on tree nutrition and yields in Canadian forests (Hamilton 
and Krause 1985; Bernier and Brazeau 1988; Thiffault et al. 2006). A review by Binkley and 
Hogberg (1997) suggested that fertilization trials with Ca and Mg have only occasionally 
favored the growth of northern tree species. The benefits of Ca and Mg fertilization may 
actually be related to an indirect effect of liming on N availability (Nohrstedt 2001; 
Sikstrom 2002). The lack of scientific evidence about the role of soil nutrients (other 
than N) on improved tree nutrition and growth may be due to the fact that permanent site 
variables such as climate, drainage, and soil physical properties have a stronger influence 
on trees (Post and Curtis 1970). 


The forest floor has been the focus of many nutrition studies because it has a large fraction of 
the fine roots (Steele et al. 1997), it can be used for linking N and P turnover to tree 
productivity and nutrition (e.g., Pare and Bernier 1989; Reich et al. 1997), and it generally 
represents a large fraction of the total soil nutrient pools (Belanger et al. 2003). However, K, 
Ca, and Mg in trees are believed to be derived primarily from mineral weathering and recent 
studies suggest that parent material elemental composition and estimates of mineral weath- 
ering can be better indicators of their availability (van Breemen et al. 2000; Bailey et al. 2004; 
Thiffault et al. 2006). As for P, its availability is not only constrained by the decay process and 
biological sinks (plant and microbial uptake) but also by geochemical sinks. 

Forest soils share many characteristics with agricultural soils, but the way they are used and 
managed requires a different approach in many situations. The objective of this chapter is to 
suggest what we believe are the most acceptable analyses for determining N, K, Ca, and Mg 
availability in forest ecosystems and establishing a link with tree nutrition, growth, and 
mortality. We focus on (1) mineralizable N; (2) pH, effective cation exchange capacity 
(ECEC), and exchangeable cations; and (3) elemental Ca, Mg, and K composition and their 
release from mineral weathering. Indices of available P are mostly limited to the extraction 
of labile P and the reader should refer to Chapter 24 and Chapter 25 for more details. 
Methods for determining soil organic carbon, pH in water or CaC^ solution, electrical 
conductivity and soluble salts, carbonates (calcite and dolomite), total and fractions of sulfur, 
and pyrophosphate-extractable Fe and Al can either be found in other chapters (mostly in 
Section III) or in Kalra and Maynard (1991). These methods are not specific to agricultural 
soils and consequently can be used for forest soils. Moreover, issues related to sampling of 
forest soils and expressing data on a concentration or mass basis are discussed in Chapter 2. 


Several techniques have been used to estimate net N mineralization in the field. Each 
method has its own limitations and there is no consensus on a best method (see Binkley 
and Hart 1989). These methods could be divided into field incubation, laboratory incu- 
bation, chemical extraction, and measurements of gross N fluxes using 15 N (to better 
understand the microbial dynamics of N transformations). Binkley and Hart (1989) 
provided a comprehensive review of the components of N availability assessments in 
forest soils. In recent years, the view of the N cycle in forested ecosystem has substantially 
changed. The following findings may have a large impact on the measurement techniques 
that are considered most appropriate for forest soils as well as on the interpretation of 
the results: 

7 Ericoid and ectomycorrhizal fungi have the capacity to scavenge organic sources 
of N and P and to participate in the decomposition process (Read et al. 2004). 
Therefore, incubations that exclude active plant roots may underestimate fluxes, 
especially in boreal or coniferous forests. 

2 Organic N is the dominant form of N in soil solutions (Quails et al. 2000) and 
some plants and associated mycorrhizal fungi can absorb dissolved organic N 
(Nasholm etal. 1998). 

j Studies reporting gross N fluxes have indicated substantial rates of gross minera- 
lization and nitrification even in systems where little mineral N accumulates 
during mineralization assays. 


Given this information, net N mineralization measured with incubation techniques cannot 
be viewed as a direct measure of plant-available N but rather as an index of this process 
(see Schimel and Bennett 2004). We provide here the description of two incubation tech- 
niques that would likely be correlated with field N fluxes even though the soils are not in 
contact with living roots. The methodologies described consider periods of incubations that 
are long enough to avoid the immobilization phase typical of forest soils with high C:N ratios 
and therefore allow part of the more labile fractions of soil N to be mineralized and 
measured. The first technique is a long-term laboratory incubation that assesses the poten- 
tially mineralizable N fraction of the soil. The second method is a field incubation that is 
sensitive to field microclimatic conditions. These two techniques have been compared in 
Brais et al. (2002). 

27.2.1 Long-Term Laboratory Incubation 

The fraction of potentially mineralizable N (No) and its mineralization constant (k) can be 
assessed with long-term laboratory incubations (Stanford and Smith 1972); some related 
methods for measuring mineralizable N in agricultural soils are given in Chapter 46. The 
long-term laboratory incubation technique given here can be used to measure production of 
dissolved organic N, C, and P (Smith et al. 1998), and C0 2 (Cote et al. 2000). The effect of 
temperature on mineralization rates can be assessed with this technique to give insight on 
the reactivity of soil organic matter to changes in temperature regime (Pare et al. 2006). 
Soil disturbance during sampling and sample preparation (e.g., drying, grinding, or sieving; 
and refrigerating or freezing) can have an impact on microbial activity and this is of 
importance for obtaining indices of N turnover and availability (e.g., Van Miegroet 1995; 
Ross and Hales 2003). The field logistics and the study's objectives will determine the 
methodology used and the interpretation of the data must be done accordingly. For the sake 
of simplicity, however, these effects and the different methods used are not further 
considered in this chapter. Rather, we describe a technique using fresh moist samples that 
we believe yields reliable estimates of the potential of the soil for N and C release under 
standard conditions. 

Materials and Reagents 

7 Plastic filtration units are used (Falcon Filter units, Becton Dickinson, Model 
7102) but the original nitrocellulose filter of the microlysimeter has to be replaced 
by a glass-fiber filter (Nadelhoffer 1990). 

2 Glass wool. 

3 1 MHCI. 

4 0.005 M K 2 S0 4 . 

5 Vacuum pump (60 mm Hg). 

6 Buchner funnels. 

7 Whatman No. 42 filter paper. 

q Acid washed 1 mm (18 mesh) silica sand. 


Volumetric soil samples are collected; fresh, moist samples are sieved through 
6 and 4 mm screens for organic humus layers (FH) and mineral horizons, respect- 
ively, to remove coarse fragments and roots. The samples are then weighed. To 
maintain the soil structure of the moist samples as much as possible, the large mesh 
sizes are used while homogenizing the material. 

The soil material is weighed to obtain samples of 25 g of the fresh organic humus 
layer (about 9 g of dry FH material on average) and 100 g of fresh mineral soil 
(about 73 g of dry mineral soil on average), and inserted into the top part of the 
filtration units above a layer of prewashed (1 .0 M HCI and deionized water) glass 
wool. The soil material is then packed slightly to obtain a total volume of soil of 
70 and 100 cm 3 for the organic layer and the mineral layer, respectively. Fine- 
textured mineral soils can be mixed with acid-washed silica sand (50% soil 
volume). Silica sand is washed with 1 M HCI and rinsed until the conductivity 
falls to that of demineralized water. 

Soil samples are incubated in growth chambers at 22°C. The relative humidity 
level is maintained around 85% to keep the soil moist. The microcosm remains 
open to air exchange inside the growth chamber unless a respiration measure- 
ment is taken over for short periods (24-48 h). Water content is verified by 
weighing and adjusted to 85% of field capacity weekly. 

Soils are flushed monthly with 100 ml_ 0.005 M K 2 S0 4 . Solution is gently 
added to the soil with a burette (2 x 50 ml_) to limit disturbance to the soil 
structure. Soils are allowed to drain freely; the excess solution is removed 
by applying vacuum. If the solution contains soil particles, it can be refiltered 
using a Buchner funnel and filter paper. Samples are transferred to the refri- 
gerator at4°C and should be analyzed for ammonium, nitrate, and total N within 
2 weeks. 

Cumulative mineralized N through time (N t ) is fitted to the following simple negative 
exponential model (Stanford and Smith 1972): 

where No is potential mineralizable N, and k is mineralization rate and is unitless. Nq can be 
expressed on a total N basis to estimate organic matter quality or on an area basis to give the 
reserve of potentially mineralizable N for a given soil depth. See Chapter 46 for more details. 


Higher temperatures of incubation are often used (e.g., 35°C). These high temperatures 
often provide a better fit and convergence of first-order models. However, Equation 27.1 
parameters {Nq and k) are sensitive to temperature (MacDonald et al. 1995; Pare et al. 2006) 
and we recommend using a soil temperature that is high (e.g., 22°C), but not outside the 
range of temperatures observed in surface soils under a closed forest canopy. 


27.2.2 Field Incubation 

Field incubation techniques include incubating a soil sample in the field with the least 
possible disturbance and estimating the net amounts of ammonium and nitrate that accumu- 
lated in the sample. The incubation period varies from a week to a year. These techniques 
originate from in situ buried bags (Eno 1960) where samples are incubated in a polyethylene 
bag. The main drawback of this method is the disturbance to the soil sample, which can 
increase mineralization rates from 2- to 10-fold according to Raison et al. (1987). The latter 
authors have described the use of in situ incubations in closed-top tubes perforated on the 
sides and open at the bottom. This technique limits disturbance of the soil samples while 
allowing the use of the samples to a greater depth (40 cm). In addition, Di Stefano and Gholz 
(1986) have proposed the use of a resin core above and below the incubated core. The resin 
core above is discarded at the end of the measurement period. Its only use is to prevent 
contamination with atmospheric N inputs. Nitrogen mineralization is estimated as the net 
amount of N mineralized in the soil core in addition to the N captured by the bottom resin 
core. This technique may provide conditions that more closely mimic those in intact soils 
because it allows water movement in the soil core as well as the removal of the products of 
mineralization. A simpler alternative is the use of closed-top cores; since there is no water 
flux into the top of the cores, it is assumed that there is no leaching loss from the bottom. We 
describe here closed-top field incubations. 

Materials and Reagents 

1 ABS cores, 30 cm long, 4.5 cm in diameter, capped 

2 2 M KCI solution 

j Whatman No. 42 filter paper 

4 Reciprocating shaker 

5 Erlenmeyer flasks (250 ml_) 
g Funnels 


1 Two tubes are brought to the field. The first one is used to collect a soil sample for 
initial determination of mineral N content. The second one is inserted in the soil to 
the required depth near the first tube and is left in the field for the incubation 
period (i.e., 1 week to 1 year; 6 weeks incubations gave reproducible results on a 
rich soil in the boreal mixedwood although it was too short to measure net 
mineralization in black spruce sites). 

2 Tubes collected from the field are kept in a cooler and should be extracted 

The soil samples are separated into forest floor and mineral soil samples. They are 
sieved through 6 and 4 mm screens for organic humus layers and mineral 
horizons, respectively. The total wet weight of the soil that is kept is weighed. 


4 A subsample is dried at 65°C for organic horizons and at 105°C for mineral soil 
horizons to estimate water content, and these subsamples are also used for 
determination of total N and total C preferably on a CN analyzer (see Chapter 
21 and Chapter 22). 

5 An amount of fresh soil that corresponds to 10 g dry weight (about half for strongly 
organic samples) is placed into a 250 mL Erlenmeyer flask. Then add 100 mL of 
KCI solution. Flasks are capped and shaken for 1 h and then filtered through a 
Whatman No. 42 filter. Solutions are analyzed for NH 4 -N and NO3-N (see 
Chapter 6). 


The difference between final and initial concentrations is used to express net N mineraliza- 
tion, net nitrification, net ammonium production, or all the above. Production rates are 
expressed in N weight, on time, and on either soil dry weight, total N or C basis to express 
the quality of the soil organic matter, or on an area basis to get an estimate of nutrient fluxes. 


7 We would advise the use of long incubation periods or the use of laboratory 
incubations in soil with high C:N, high organic matter content, low N turnover, 
little net nitrification, little nitrate in soil solution, or presence of ericoid and 
ectomycorrhizal fungi. On the other hand, short-term incubations would be 
suitable for forests with thin or nonexistent organic layers that undergo net 
nitrification (such as sugar maple [Acer saccharum Marsh.] forests). 

2 We often found very low and negative rates of net mineralization (net immobili- 
zation) in boreal black spruce forests with thick organic layers (D. Pare, unpub- 
lished data). Such results are frequent and not often published. In all cases, it is 
advisable to compare results with the nutrient budget. 

j Estimates of N in litterfall and immobilization in biomass provide estimates of N 
mineralization that are totally independent of incubation estimations and that 
should match them. Although it is not always possible to obtain such an estimate, 
the comparison of incubation results with budget estimates should be done on a 
few plots within the forest and soil types investigated. 


27.3.1 SonpH 

Because of the variations in ionic strength of agricultural soils, the most common method of 
measuring their pH in Canada is the 0.01 MCaCi2 method. By measuring the pH in an 
electrolyte of known concentration, the effects of variable ionic strength of the soil solutions 
are largely eliminated. Forest soils, on the other hand, tend to have close to the same ionic 
strength throughout the year, except as influenced by variations in water content. For this 
reason, many researchers choose to use pH measured in water. Since the ionic strength of the 
measurement solution is lower, the pH obtained will be closer to that observed by plants 


growing in the field. Researchers should be aware that the disturbance caused by sampling 
and drying soils does have an effect on the measured pH (Courchesne et al. 1995). A 
discussion on the choice of pH methods can be found in Chapter 16 along with a detailed 
description of the methods themselves. 

27.3.2 Effective Cation Exchange Capacity and Exchangeable Cations 

The use of an unbuffered BaCb solution is now generally preferred for determination of 
ECEC. The BaC^-compulsive exchange procedure (Gillman and Sumpter 1986) is recom- 
mended for determining CEC on all soils (except soils containing salts, carbonates, or 
zeolites) (Sumner and Miller 1996). Similarly, Chapter 18 suggests the use of a simplified 
BaCb extraction. The BaCl2 extraction has the ability to displace trivalent cations at lower 
ionic strength without being preferentially adsorbed compared to the NH4CI or KC1 extrac- 
tions. Although the extraction can modify pH because the ionic strength of 0.3 mol L -1 (for 
0.1 M BaCl2) of the solution is still about three orders of magnitude higher than the soil 
solution of a sandy Podzol, it causes smaller changes in pH than the more concentrated 
solutions (e.g., 1 mol L ' for 1 M KC1 or NH4CI). In this method, ECEC is calculated by 
summing exchangeable cations (Ca, Mg, K, Na, Al, Fe, and Mn). 

The method of Chapter 18 is strongly recommended for acid forest soils such as Podzols as 
well as Sombric and Dystric Brunisols (or Dystric Cambisols according to the Food and 
Agriculture Organization of the United Nations (1974)). For boreal plain forests with higher 
soil pH values (above pH 5.5-6.0) and low levels of exchangeable aluminum and manga- 
nese (e.g., Melanic and Eutric Brunisols [or Eutric Cambisols], Gray Luvisols [or Albic 
Luvisols], and Chernozems), the unbuffered NH4CI extraction is also acceptable and is 
commonly used (Kalra and Maynard 1991). However, although most agronomists are 
interested in determining the amount of exchange sites for management or control of soil 
pH, usually by liming, forest soil scientists are also interested in knowing the cation species 
(base vs. acid) held on these exchange surfaces. The method proposed here is therefore a 
one-step extraction that uses an unbuffered NH 4 C1 solution and that allows the measurement 
of ECEC as well as the individual contribution of Ca, Mg, K, and Na to ECEC (and Al, Fe, 
and Mn if needed). 

Materials and Reagents 

1 Centrifuge tubes (50 mL) with screw caps. 

2 Ultracentrifuge accepting 50 mL tubes. 

3 End-over-end shaker. 

4 Ammonium chloride, 1 M: dissolve 53.5 g of NH 4 CI with double deionized (d.d.) 
water and make to volume in a 1000 mL volumetric flask. 

5 Standards of Ca, Mg, K, Na, Al, Fe, and Mn are prepared using atomic absorption 
reagent-grade standards of 1 000 mg L~ 1 . The matrix in the standards must correspond 
to the NH4CI concentration of the analyzed sample (diluted or nondiluted matrix). 

6 Lanthanum solution, 100 mg L~ 1 : dissolve 53.5 g of LaCI 3 • 7H 2 in a 200 mL 
volumetric flask and make to volume (for analysis by atomic absorption spectroscopy 


7 Cesium solution, 100 g L~ 1 : dissolve 25.2 g CsCI in a 200 ml_ volumetric flask 
and make to volume (for analysis by AAS). 

q Whatman No. 41 filter paper. 

1 Weigh out about 2.5 g of dry organic soil (FH samples) or fine-textured mineral 
soil and about 12.5 g of dry (<2 mm) coarse-textured mineral soil into a 50 ml_ 
centrifuge tube. Record the exact weight of soil used to the nearest 0.001 g. 
Include blanks, duplicates, and quality control samples. 

2 Add 25.0 ml_ of 1 M NH 4 CI to each tube and shake slowly on an end-over-end 
shaker (15 rpm) for 2 h. 

3 Ultracentrifuge (1 5 min, 7000 g) and filter the supernatant with Whatman No. 41 
filter paper. 

4 Analyze Ca, Mg, K, Na, Al, Fe, and Mn in the supernatant solution with AAS 
or other suitable instrument. Dilution (10- or 100-fold) will likely be required 
for Ca, K, Mg, and Na. The addition of 0.1 ml_ of La solution and 0.1 ml_ of 
Cs solution to a 10 ml_ aliquot of diluted extract is required for the determina- 
tion of Ca, Mg, and K by AAS (for detailed instructions on this and other aspects 
of analysis refer to the manual for your AAS). If needed, preservation of samples 
by acidifying to 0.2% HNO3 will prevent the loss of Fe and Al. 


1 Exchangeable cations: 

M + cmol c kg~ 1 = C cmol c L~ 1 x (0.025 L/wt. soil g) x 1000 g kg~ 1 x DF 

where M + is the concentration of an adsorbed cation (cmol c kg~ ); C is the 
concentration of the same cation measured in the NH 4 CI extract (cmol c L~ ); 
and DF is the dilution factor (if applicable). 

Effective CEC cmol c kg" 1 = 2 cmol c Ca, Mg, K, Na, Fe, Al, Mn kg (27.3) 

See Section 18.2.4 in Chapter 18 for details on quality controls, standards, and the effects of 
different soil: solution ratios on results. 

27.3.3 Contribution of Exchangeable H + to Effective Cation Exchange 

It is difficult to account for the amount of H + coming from the exchange reaction and its 
contribution to ECEC from NH4CI or BaCLi extractions because some H + in the extract may 
come from sources other than exchangeable H + (e.g., dissociation of organic acids) or be 


produced or consumed in reactions involving Al-OH complexes or hydrolysis of free Al 3+ 
(Thomas and Hargrove 1984). The salt solution, ionic strength, and the soil: solution ratios 
have an influence on the amount of exchangeable H + displaced from exchange sites. 
Therefore, the contribution of exchangeable H + to ECEC or base saturation is operationally 
defined from titration of a 1 M KC1 extract as suggested by Thomas (1982) (see Chapter 18 
for details on methodology). Exchangeable H + is relatively abundant in acidic organic 
horizons (e.g., forest floor material), but acidic mineral soils such as Bhf and Bf horizons 
also have high enough amounts to draw our attention (Ross et al. 1996; Belanger et al. 2006). 
Belanger et al. (2006) noted that "fundamentally, any valid measure of ECEC must therefore 
include some estimate of exchangeable H + concentration or a demonstration that it is 
negligible". Unfortunately, the direct measurement of exchangeable H + is time-consuming 
and not practical for routine analysis. Therefore, Belanger et al. (2006) have used soil pH in 
water (as proposed in Chapter 16) and ECEC (using BaCl2 as described in Chapter 18) to 
estimate exchangeable H + concentrations in FH and podzolic (spodic) B samples of acidic 
forest soils developed from granitic bedrock or parent material. Although Equation 27.4 
provides good estimates of the proportion of exchangeable H + on the exchange complex of 
organic and podzolic B horizons from all types of forests, we recommend the readers build 
specific relationships using their own samples if greater predicting power is required: 

log(exch. H+)/ECEC = 0.682 - (0.308 x soil pH in water); R 2 = 0.691 (27.4) 


It has long been recognized that Ca and Mg in trees are derived primarily from Ca and Mg 
released into the soil solution from mineral weathering (van Breemen et al. 2000; Blum et al. 
2002), and additional studies suggest that parent material elemental composition can be a 
reliable indicator of tree Ca and Mg nutrition. For example, Thiffault et al. (2006) examined 
soil and foliar nutrient status of black spruce and balsam fir {Abies balsamea (L.) Mill.) 
stands in Quebec subject to whole-tree and stem-only harvesting and found that total mineral 
parent C elemental content was more indicative of nutrient limitations than surface soil- 
available nutrient concentrations: the signal of this low Ca and Mg availability was very 
weak in the upper soil layers, including the forest floor, probably because the chemistry of 
these layers is largely controlled by litter material with relatively well-balanced nutrient 
ratios (Knecht and Goransson 2004). Additional studies suggest that parent material ele- 
mental composition may be an important predictor of tree mortality as well as Ca and Mg 
nutrition. For example, van Breemen et al. (1997) showed that sugar maple mortality in the 
northeastern United States increased with decreasing elemental Ca in the parent C material. 
Sugar maple foliar Ca and Mg status and mortality were also more strongly linked to B 
horizons compared to forest floor Ca and Mg chemistry (Bailey et al. 2004). 

Bailey et al. (2004) further suggested that a model that calculates release of Ca and Mg from 
soil mineral weathering of the parent C material would likely be successful in predicting 
stand nutrition and productivity. Many indices of soil mineral weathering have been deve- 
loped in the past (Birkeland 1999), but one of the preferred approaches compares the 
concentration of elements in the various soil horizons to the concentration of elements on 
the presumably unaltered parent material in the C horizon (Kirkwood and Nesbitt 1991; Bain 
et al. 1994; Hodson 2002). Adjustments are made to consider additions of organic matter and 
the leaching of elements that are not of interest in the study because both will affect the 
concentration of the elements studied. Therefore, an equation using an element resistant to 


weathering (most often zirconium and titanium) is used to normalize the data for mobile 
elements. Assuming the age of the soil is known, the release rate of a mobile element in a 
particular horizon can be calculated using the following equations (Hodson 2002): 

£,.* = Ei x C PM /C ; (27.6) 

where R is the element release rate (|xg m~ 2 year~ ), E PM is the concentration of element 
E in the parent material ((jug g _1 ), E* is the adjusted concentration of element E 
in horizon i (jxg g _1 ), £, is the concentration of element E in the horizon i (jxg g _1 ), p is 
the horizon density (g m~ 3 ), Z is the horizon thickness (m), t is the soil age (years), Cpm 
is the concentration of immobile element C in the parent material (|xg g _1 ), and C, is the 
concentration of immobile element C in horizon i (jxg g _1 ). 

In forest soils, the concentration of immobile elements tends to decrease with depth; there is 
a concentration effect from bottom to top because of the loss of mobile elements and 
accumulation of organic matter in upper soil horizons (Melkerud et al. 2000; Courchesne 
et al. 2002; Hodson 2002). Research has shown that some of the elements that are resistant to 
weathering can nonetheless be eluviated and we recommend that anyone applying this 
technique study the results of authors such as those mentioned above. 

Weathering rates in the >50 p,m fraction are sometimes assumed to be negligible because of the 
relatively low surface area and lack of easily weatherable minerals in that fraction (Kolka et al. 
1996). Therefore, the method is sometimes employed on the silt fraction (2-50 |xm) alone after 
wet sieving to remove sand and by multiple centrifugations to remove clay. In this case, the 
expression (£p M — E*) is multiplied by the silt mass in that horizon, which can be measured 
after determination of soil bulk density, particle size distribution, and horizon thickness. 

Wavelength dispersive x-ray fluorescence spectroscopy on fused beads is generally 
the preferred approach to determine the elemental composition of soil samples (e.g., van 
Breemen et al. 1997; Melkerud et al. 2000), but this can also be determined by inductively 
coupled plasma (ICP) or AAS on samples digested using hydrofluoric acid. 

Attiwil, P. and Adams, M.A. 1993. Nutrient eye- upland granitic till catchment in Scotland. Water 

ling in forests. New Phytol. 124: 561-582. Air Soil Poll. 73: 1 1-27. 

Bailey, S.W., Horsley, S.B., Long, R.P., and Belanger, N., MacDonald, J.D., Pare, D., 

Hallett, R.A. 2004. Influence of edaphic factors Thiffault, E., Claveau, Y., and Hendershot, W.H. 

on sugar maple nutrition and health on the Alle- 2006. The assessment of exchangeable hydrogen 

gheny Plateau. Soil Sci. Sot: Am. J. 68: 243-252. ions in Boreal Shield soils of Quebec. Can. J. Soil 

Sci. 86: 513-521. 
Bain, D.C., Mellor, A., Wilson, M.J., and 

Duthie, D.M.L. 1994. Chemical and mineral- Belanger, N., Pare, D., and Yamasaki, S.H., 2003. 

ogical weathering rates and processes in an The soil acid-base status of boreal black spruce 


Bernier, B. and Brazeau, M. 1988. Magnesium 
deficiency symptoms associated with sugar 
maple dieback in a Lower Laurentians site in 
southeastern Quebec. Can. J. Forest Res. 18: 

Binkley, D. and Hart, S.C. 1989. The components 
of nitrogen availability assessments in forest 
soils. Adv. Soil Sci. 10: 57-112. 

Binkley, D. and Hogberg, P. 1997. Does atmos- 
pheric deposition of nitrogen threaten Swedish 
forests? Forest Ecol. Manag. 92: 119-152. 

Birkeland, P.W. 1999. Soils and Geomorphology, 
3rd ed. Oxford University Press, New York, 430 pp. 

Blum, J.D., Klaue, A., Nezat, C.A., Driscoll, C.T., 
Johnson, C.E., Siccama, T.G., Eagar, C, Fahey, 
T.J., and Likens, G.E. 2002. Mycorrhizal weath- 
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base-poor forest ecosj stems. Nature 417: 729-731. 

Brais, S., Pare, D., Camire, C, Rochon, P., 

and Vasseur, C. 2002. Nitrogen net mineralization 
and dynamics following whole-tree harvesting and 
winter windrowing on clayey sites of northwestern 
Quebec. Forest Ecol. Manag. 157: 119-130. 

Cote, L., Brown, S., Pare, D., Fyles, J., and 
Bauhus, J. 2000. Dynamics of carbon and nitro- 
gen mineralization in relation to stand type, stand 
age and soil texture in the boreal mixedwood. Soil 
Biol. Biochem. 32: 1079-1090. 

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59 pp. 

Gillman, G.P. and Sumpter, E.A. 1986. Modifi- 
cation to the compulsive exchange method to for 
measuring exchange characteristics of soils. Aust. 
J. Soil Res. 24: 61-66. 

Hamilton, W.N. and Krause, H.H. 1985. Relation- 
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in New Brunswick plantations. Can. J. Forest 
Res. 15: 922-926. 

Hodson, M.E. 2002. Experimental evidence for 
mobility of Zr and other trace elements in soils. 
Geochim. Cosmochim. Acta 66: 819-828. 

Ingestad, T. 1979a. Nitrogen stress in birch seed- 
lings. II. N, P, Ca, and Mg nutrition. Physiol. 
Plant. 45: 149-159. 

Ingestad, T. 1979b. Mineral nutrient requirement 
of Pinus silvestris and Picea ahies seedlings. 
Physiol. Plant. 45: 373-380. 

Kalra, Y.P. and Maynard, D.G. 1991. Methods 
Manual for Forest Soil and Plant Analysis. Infor- 
mation Report NOR-X-319. Forestry Canada, 
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Kirkwood, D.E. and Nesbitt, H.W. 1991. Forma- 
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Courchesne, F., Halle, J.-P., and Turmel, M.-C. 
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meridional. Ge'ogr. Pins. Quatern. 56: 5-17. 

Courchesne, F., Savoie, S., and Dufresne, A. 
1995. Effects of air-drying on the measurement 
of soil pH in acidic forest soils of Quebec, 
Canada. Soil Sci. 160: 56-68. 

Di Stefano, J. and Gholz, H.L. 1986. A proposed use 
of ion exchange resin to measure nitrogen mineral- 
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Eno, C. 1960. Nitrate production in the field by 
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Soc. Am. Proc. 24: 277-279. 

Knecht, M.F. and Goransson, A. 2004. Terrestrial 
plants require nutrients in similar proportions. 
Tree Physiol. 24: 447-460. 

Kolka, R.K., Grigal, D.F., and Nater, E.A., 1996. 
Forest soil mineral weathering rates: use of mul- 
tiple approaches. Geoderma 73: 1-21. 

MacDonald, N.W., Zak, D.R., and Pregitzer, K.S. 
1995. Temperature effects on kinetics of micro- 
bial respiration and net nitrogen and sulphur 
mineralization. Soil Sci. Soc. Am. J. 59: 233-240. 

Melkerud, P.A., Bain, D.C., Jongmans, A.G., and 
Tarvainen, T. 2000. Chemical, mineralogical and 
morphological characterization of three podzols 
developed on glacial deposits in Northern Europe. 
Geoderma 94: 125-148. 


Morrison, I.K. and Foster, N.W. 1995. Effect of Reich, P.B., Grigal, D.F., Aber, J.A., and Gower, 

nitrogen, phosphorus and magnesium fertilizers S.T. 1997. Nitrogen mineralization and pro- 

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Nadelhoffer, K.J. 1990. Microlysimeter for meas- 
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Am. J. 54: 411-415. 

Niisholm, T., Ekblad, A., Nordin. A., Giesler, R., 
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Nohrstedt, H.-6. 2001. Effects of liming and 
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Air Soil Pollut. 139: 343-354. 

Paquin, R., Margolis, H.A., and Doucet, R. 1998. 
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layers and planted seedlings in response to nutrient 
addition in the boreal forest of Quebec. Can. 
J. Forest Res. 28: 729-736. 

Pare, D. and Bernier, B. 1989. Origin of the 
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Can. J. Forest Res. 19: 24-34. 

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nutrient fluxes in clearcut and mature deciduous 
forests. Soil Sci. Soc. Am. J. 64: 1068-1077. 

Raison, R.J., Connell, M.J., and Khanna, P.K. 

1987. Methodology for studying fluxes of soil 
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Read, D., Leake, J.R., and Perez-Moreno, J. 2004. 
Mycorrhizal fungi as drivers of ecosystem pro- 
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Can. J. Bot. 82: 1243-1263. 

Ross, D.S., David, M.B., Lawrence, G.B., and 
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Chapter 28 

Chemical Properties 

of Organic Soils 

A. Karam 

Laval University 
Quebec, Quebec, Canada 


Organic soils are rich in fresh plant material or organic materials at various stages of 
decomposition, namely fibric, hemic, and sapric materials (Soil Survey Staff 2003). These 
soils usually form under conditions of water saturation. Organic soils include muck and peat 
soils or histosols (Canada and United States of America), the tundras, the Irish peat bogs, the 
moor peats (Australia), les sols hydromorphes organiques (France), and earthy peat soils in 
Great Britain (Okruszko and Ilnicki 2003). Common organic soil parent materials may 
include mosses (such as sphagnum), gyttja, dy, marl, volcanic ash, cattails, reeds, sedges, 
pondweed, grasses, and various "water-loving" deciduous and coniferous shrubs and trees. 
Organic soils can contain silicate minerals from trace to appreciable amounts. The charac- 
teristics of organic soils depend mainly on the nature of the vegetation that was deposited in 
the water and the degree of decomposition (Mokma 2005). 

Although the chemical properties of organic soils are different from those of their mineral 
counterparts, the chemical methods given for mineral soils are also applicable to organic 
soils; therefore, these analytical procedures are not repeated here. The analyst may choose an 
appropriate method from other chapters. 

28.2.1 Introduction 

Soil testing is complicated by the larger range of volume percentage of the solid phase, 
computed as the ratio of bulk density to particle density, and water contents in samples of 
organic soils compared with mineral soils, and with a greater influence of organic soil drying 
on soil chemical properties (Parent and Khiari 2003). According to Watson and Isaac (1990), 
quantitative analysis of soil samples can be broken down into six steps: (i) handling and 


preparation of the samples, (ii) weighing samples, (iii) dissolution of samples/extraction of 
elements, (iv) pretreatment or removal of interferences, if needed, (v) measuring a property 
of the sample, and (vi) calculating and reporting of concentration of analyte. Handling and 
preparation of organic soils before soil analysis are critical. Chemical analyses may be 
conducted on fresh samples (field moisture content) or on air-dried samples. It should 
be noted that drying organic soils increases dry bulk density, volume of the solid phase 
(Ilnicki and Zeitz 2003), and mineralization of organic P (Daughtrey et al. 1973), decreases 
pH measured in water, 0.01 M CaCl2, and 1 M KC1 compared with the field-moist condition 
(van Lierop and Mackenzie 1977), and may lead to higher soil test levels of certain nutrients, 
such as available P and K (Daughtrey et al. 1973; Anderson and Beverly 1985). These latter 
authors postulate that screened organic soils are more easily compacted upon drying, and 
conversely expand upon rehydration. Parent and Khiari (2003) noted that air-dried pristine 
peat contained more P in available form than fresh peat. Anderson and Beverly (1985) 
recommend that organic soils be sampled on a volume basis in order to ensure uniformity of 
results. Harrison (1979) suggested that where soils vary in bulk density, soil data should be 
expressed in terms of soil volume. 

28.2.2 Materials 

/ Analytical balance 

2 Blender, high speed 

3 Large flat pan or equivalent 

4 Spoon or spatula 

28.2.3 Procedure 

/ Mix organic soil sample thoroughly and weigh a 100 to 300 g representative 
sample. Determine the mass of the sample and spread evenly on a large flat pan, 
square rubber sheet, or paper. Crush soft lumps with a spoon or spatula and let the 
sample come to moisture equilibrium with room air, not less than 24 h. 

2 Stir occasionally to maintain maximum air exposure of the entire sample. 

j When the mass of the sample reaches a constant value, calculate the moisture 
removed during air drying as a percentage of the as-received mass. 

4 Grind a representative portion of the air-dried sample 1 to 2 min in a high-speed 
blender. Determine the amount, in grams, of air-dried sample equivalent to 50 g 
of as-received sample as follows: 

Equivalent sample mass, g = 50.0 - [(50 x M)/100] 

where M is the percent of moisture removed in air drying. 
Place the sample in a moisture-proof container. 



28.3.1 Introduction 

The simplest method for the direct determination of moisture content is the gravimetric 
method, which involves the measurement of water lost by weighing a soil sample (as-received) 
before and after it is dried at 105°C-1 10°C in an oven. The moisture content is expressed either 
as a percent of the oven dry mass or of the as-received mass. This method may not be suitable 
when a dried soil sample is used to assess nitrogen, pH, cation exchange, and other soil 
chemical properties. An alternative method that removes the total moisture and provides a 
more stable sample, the air-dried sample, has been suggested by the ASTM Committee 
(ASTM 1997). This method includes two steps: (1) evaporation of moisture in air at room 
temperature (air-drying) and (2) the subsequent oven drying of the air-dried sample at 105°C. 

There are basically two procedures involved in the determination of the ash (inorganic 
fraction) of a peat or organic sample: dry-ashing methods and wet-ashing methods. 

The dry-ashing method involves the removal of organic matter by combustion of the sample 
at medium temperature (375°C to 800°C) in a temperature-regulated muffle furnace. The 
principal errors in dry-ashing arise through incomplete combustion when the temperature or 
time allowed for combustion is insufficient, and through losses resulting from the use of too 
high a temperature (Allen 1989). 

If necessary, samples are dried (105°C-110°C) before ashing. The substance remaining after 
ignition is the ash and includes mineral impurities such as sand. The weight lost on ignition is 
calculated and considered as an approximate measure of the organic content of acid organic 
soils and noncalcareous peatlands. Vessels suggested for ashing are porcelain, quartz, or 
platinum dishes. Sample weights used vary from 0.25 to 2.00 g. The ash may be further 
dissolved in an acid solution for elemental analysis. 

28.3.2 Materials 

7 Muffle furnace — controlled to ±5°C for ashing at 600°C 

2 High-form porcelain, 30 mL crucible 

j Porcelain crucible cover or aluminum foil, heavy duty 

4 Desiccator cabinet or nonvacuum, desiccator with desiccant 

5 Analytical balance and spoons 

g Electric drying oven: regulated to a constant temperature of 105°C 

28.3.3 Procedure 

7 Weigh a 2 g sample of 2 mm oven-dried soil (105°C) into a tared high-form 
porcelain, 30 mL crucible with 0.1 mg accuracy. Determine the mass of the 
covered high-form porcelain crucible. Remove the cover and place the crucible 
in a muffle furnace. 


2 Gradually bring the temperature in the furnace to 370°C and maintain it for 1 h 
and then ash the sample either at 550°C for 16-20 h (Andrejko et al. 1983) or at 
600°Cfor 6 h (Goldin 1987). 

3 Remove the crucible from the furnace, cover, place it in a desiccator, allow to 
cool, and weigh with 0.1 mg accuracy. Save the crucible and its contents for 
metal ion determination. 

28.3.4 Calculation 

Calculate the ash content as follows: 

Ash, g/100 g = [(a - c)/(b - c)] x 100 (28.2) 

where a is the final weight (g) of crucible and ash; b is the weight (g) of crucible and sample; 
and c is the weight (g) of empty crucible. 

The procedure described above can be used to determine the amount of organic matter as 

% Organic matter = 100 — % mineral content (ash) (28.3) 


/ Using the above procedure for determining ash content it has been shown 
by Andrejko et al. (1983) that a temperature setting of 550°C is satisfactory for 
most purposes. The standard method approved by the ASTM Committee (ASTM 
1997) proposes a temperature setting of 440°C and heating until the sample is 
completely ashed (no change of mass occurs after a further period of heating). 

2 Dry ashing may overestimate the amount of organic matter in the soil. Positive 
errors are dependent on soil properties, such as the amount of carbonates 
and the amount and type of clay present in the mineral fraction of the soil 
(Goldin 1987). 

j The procedure outlined above measures the mass percentage of ash and 
organic matter in organic soil, including moss, humus, and reed-sedge types 
(Day etal. 1979). 

4 Samples should be placed in the muffle furnace cold and the temperature allowed 
to rise slowly to avoid volatilization losses, which are aggravated by violent 

5 Use high-form porcelain crucibles with covers or equivalent if ashes are retained for 
elemental analysis. These crucibles eliminate possible contamination of the ash by 
boron, which may volatilize from the furnace walls (Williams and Vlamis 1 961 ). 

6 Values derived from loss-on-ignition results should only be considered as 
approximate (Allen 1989). 



28.4.1 Introduction 

Most methods that have been developed for the determination of total elements in organic 
soils involve a two-step procedure, namely: (i) the complete destruction of both organic and 
inorganic fractions of the soil matrix by various digestion/oxidation procedures in order to 
liberate all elements in solution and (ii) the determination of soluble elements by various 
techniques. The chemical procedure involved in the destruction of organic materials 
(peat, plants, sediments, soils) falls basically into two groups: (a) dry-ashing methods and 
(b) wet-ashing (or digestion) methods. 

In the dry-ashing procedure, the organic material is ignited in an electrically controlled tem- 
perature muffle furnace with fume disposal at low (400°C) or medium temperature (550°C- 
660°C) to oxidize organic matter and the ions are extracted from the ash with an acid solution: 
1 .5 M HC1 (Ali et al. 1988), 6 M HC1 (Kreshtapova et al. 2003), or 2 M HN0 3 (Day et al. 1979). 
Dry-ashed sample may be heated on a hot plate with dilute HC1 to dissolve the residues and then 
with concentrated hydrofluoric (HF) acid to destroy any silicates present (Papp and Harms 1985). 

Wet digestion involves complete dissolution of the organic material to convert elements to 
soluble forms by heating with concentrated acids in either open or closed vessels. This phase is 
then followed by determination of the liberated ions. Dissolution technique can be performed 
by hot plate, hot block digestion, or pressured digestion systems. Open or closed vessels can be 
used in microwave systems while open vessels are usually used in block digestion. Disadvan- 
tages of open vessel digestion systems include the greater risk of loss of volatile elements. 
Microwave digestion is a commonly used practice in many laboratories. Important variables in 
a microwave digestion procedure are the microwave energy power profiles (power, time, and 
pressure), the volume and combination of acids used, and the acid-to-sample ratio. 

Recently, a new wet digestion procedure for the determination of As in biomasses, coal, and 
organic-rich sediment samples using hydride generation-atomic fluorescence spectrometry 
(HG-AFS) has been developed (Chen et al. 2005). This method involves digestion of 200 mg 
sample aliquots with 3 mL HN0 3 (65%) + 0.1 mL HBF 4 (-50%), heating in a microwave 
autoclave up to a temperature of 240°C. After digestion, no evaporation of HNO3 to remove 
acid from the digests is needed before arsine generation can be carried out. 

The following microwave acid digestion procedure is a modification and synthesis from 
methods proposed by Papp and Harms (1985), Weiss et al. (1999), and Morrell et al. (2003). 
It employs microwave heating of the sample, first in HNO3 + H2O2 to oxidize organic 
matter and then in HNO3 + HF to decompose any remaining organic material and complete 
dissolution of the inorganic fractions of the peat or organic soil. The first gentle phase of the 
digestion program, the organic, carbon-rich matrix components are slowly converted to CO2 
to avoid foaming (Krachler et al. 2002). 

28.4.2 Materials 

7 Microwave oven-assisted sample digestion system, with closed vessels 
2 Nitric acid (HNO3), concentrated (trace metal grade), 65% 

j Hydrogen peroxide (H2O2), 30% 

4 Hydrofluoric acid (HF), 40% or 48%, as recommended by the instrument 

5 Polyfluoroethylene (PTFE) Teflon vessels 
g Polypropylene volumetric flask, 50 mL 

j Screw-capped polypropylene bottles, 60 mL 

28.4.3 Procedure 

7 Weigh a 250 mg of air-dried and finely ground soil (1 00 mesh) of known moisture 
content in a 60 mL Teflon digestion tube with a cap. 

2 Add 4 mL of trace metal-grade concentrated HNO3 and 1 mL of H2O2. 

3 Place the vessel in the microwave and digest the soil for 30 min at 296 W. 

4 Seal the vessel with the cap and digest for 1 5 min at 296 W. 

5 Cool for approximately 35 min well below the boiling point of the acid at 
atmospheric pressure, and then open the reaction chamber. 

6 Add 4 mL concentrated HNO3 + 1 mL concentrated HF. Seal the vessel with the 
cap and digest as follows: 4 min at 250 W, 8 min at 565 W, 4 min at 450 W, 4 min 
at 350 W, 5 min at 250 W, and vent for 35 min. Other operating power and 
temperature parameters can be set as specified by the instrument manufacturer. 

j After cooling, loosen the vessel cap in order to expel the interior gas into a fume 
hood. Remove the cap and allow the vessel to stand for ca. 2 min to remove any 
further gas. 

g Transfer the tube contents into a 50 mL polypropylene volumetric flask. Wash 
the inside of the tube and cap, and adjust the volume to 50 mL with 
distilled/deionized water. A colorless digestion solution is an indication of 
efficient destruction of the organic matter. 

9 Store the sample solutions in 60 mL screw-capped polypropylene bottles before 
analysis for metals and other elements of interest. 

7Q Perform a blank containing all reagents used in the sample digestion. 


7 This procedure does not purport to address all of the safety problems associated 
with its use. It is the responsibility of the user of this procedure to esta- 
blish appropriate safety and health practices and determine the applicability of 
regulatory limitations before use. The analyst should read carefully all warnings 


and follow all hints and instructions provided with the instruction manual issued 
by the instrument manufacturer to ensure correct and safe operation of the 

Peat samples containing mixtures of fine-grained matter and plant or fibrous 
material should be dried in an oven at 50°C and then mixed thoroughly in a 
low-speed blender to preserve all parts of the sample. To achieve the proper 
homogeneity with the use of small amounts of samples, dried soil or peat samples 
have to be ground to pass through 1 00 mesh sieve. The portion left on the sieve 
should be ground again for a short period, sieved, and so on until the complete 
sample could pass through the sieve. 

All vessels have to be checked for metals and other elements contamination 
before use. Avoid using commercial detergents containing phosphate or other 
elements. The reagents and filter paper selected should be as free of metals and 
P as possible. It is essential to use reagents and distilled water of suitably low 
metal content, taking into consideration that the concentrated mineral acids are 
generally used in amounts several times that of the sample. 

Low sample amounts may result in a good decomposition result, but may impair 
the analytical accuracy. Excessive sample amounts may lead to a poor decom- 
position result. 

Add the nitric acid slowly, with swirling, to the sample. More HNO3 may be 
needed to achieve the complete oxidation of organic matter. Nitric acid may 
react violently with some samples containing high organic material. Hydrogen 
peroxide has a high oxidization potential and can produce very strong reactions. 

Addition of acids and sample digestion must be conducted in a fume hood with 
adequate ventilation. 

Hydrofluoric acid is normally used in the acid mixture to dissolve silicates, which 
are present in the samples and more HF will be required for the decomposition of 
peat or organic soils high in silicate minerals. However, HF can give rise to 
problems in glassware and torch damage of some spectrometers, in particular 
inductively coupled plasma-mass spectrometry (ICP-MS) (Melaku et al. 2005). 
This problem can be avoided by using an HF-resistant nebulizing system and 
plasma torch (Swami et al. 2001). Special safety instructions must be observed 
when handling HF. Avoid the use of Pyrex glass materials or quartz vessels. 

In the HNO3/HF treatment, some elements such as Ca, Mg, Al, and rare earth 
elements may form insoluble fluorides that easily precipitate (Krachler et al. 2002; 
Wang et al. 2004); H3BO3 solution is often added to digestion mixtures to dissolve 
slightly soluble fluorides. In such cases, proceed as follows: add 10 ml_ of H3BO3 
solution (5%, m/v) per 1 ml_ of HF (Swami et al. 2001 ) to the decomposed sample 
(step 7), seal the vessel again and subject it to a second decomposition run at high 
temperature or power rating for 1 0-1 5 min. After cooling, loosen the vessel cap in 
order to expel the interior gas into a fume hood. Remove the cap and allow the 
vessel to stand for ca. 2 min to remove any further gas. In cases of incomplete 
dissolution, continue to microwave until the sample is dissolved. Transfer the 
sample to a polypropylene volumetric flask and dilute with distilled/deionized 


water to a fixed volume of 50 ml_. This dissolution method is not suited for the 
determination of B in the soil digest. 

g Selection of the most suitable digestion method must be based on local require- 
ments and facilities. Digestion mixture options include: HNO3 + HCIO4 + HF 
(Papp and Harms 1985), HN0 3 + H 2 2 + HF and HN0 3 + H 2 2 + HCIO4 + HF 
(Weiss et al. 1999), HN0 3 + HBF 4 (Krachler et al. 2002), HNO B + HCI and 
HN0 3 + HCI + HF (Burt et al. 2003), HNO B + HCI0 4 ; hot plate, microwave or 
block digestion, open or closed vessel. An HN0 3 + HCI0 4 treatment of peat or soil 
samples is not always complete and a residue (siliceous materials) might remain. 
Filtration using Whatman No. 42 filter paper is desirable to keep the solution free of 
solid particles that cause clogging of the capillary tip of spectrometers. Acid 
digestion procedure using HCI0 4 requires a HCI0 4 fume hood. Perchloric acid is 
a very strong oxidizing agent that bears many risks and should not be used alone, 
but only in combination with other acids. As a safety precaution, it is recommended 
that organic samples be digested in HN0 3 before proceeding with HN0 3 /HCI0 4 
digestion. Only use HCI0 4 in microwave oven for processes that have been 
approved by the manufacturer. Perchloric acid can react with explosive force if 
the digestion mix approaches dryness. In general, perchlorates are easily soluble 
and the use of HCI0 4 can considerably reduce the amount of HN0 3 required and 
complete the oxidation in a shorter time. 

1q The analyst may use a suitable dilution factor depending on the detection limit of 
the instrument and the concentration of the element. 

7 7 Metals in the digestion solution may be determined by atomic absorption spec- 
troscopy (AAS) and La is added to the extracting solution (Ca and Mg determin- 
ations) as a suppressant. Sodium and potassium are commonly determined on a 
flame emission spectrophotometer. Low content at ppb level of some elements 
may be determined by using graphite furnace atomic absorption spectroscopy 
(GFAAS). The majority of elements may be determined by inductively coupled 
plasma-atomic emission spectrometry and -mass spectrometry (ICP-AES and 
ICP-MS). Selenium can be determined by using a hydride-generating system 
attached to an ICP emission spectrometer. If B is one of the elements of interest, 
it should be determined in H 3 B0 3 -free digestion solution. Phosphorus, sulfur, and 
boron may be determined by spectrophotometric methods. 

Ali, M.W., Zoltai, S.C., and Radford, F.G. 1988. potassium and bulk density of organic and min- 
A comparison of dry and wet ashing methods for eral soils of the Everglades. Soil Sci. Soc. Am. 
the elemental analysis of peat. Can. J. Soil Sci. J. 49: 362-366. 
68: 443-447. 

Andrejko, M.J., Fiene, F., and Cohen, A.D. 
Allen, S.E. 1989. Chemical Analysis of Eco- 1983. Comparison of ashing techniques for 
logical Materials, 2nd edn. Blackwell Scientific determination of inorganic content of peats. In 
Publications, Oxford, UK. P.M. Jarret, Ed. Testing of Peats and Organic 

Soils. ASTM STP 820. American Society 
Anderson, D.L. and Beverly, R.B. 1985. The for Testing and Materials, Philadelphia, PA, 
effects of drying upon extractable phosphorus, pp. 5-20. 


ASTM (American Society for Testing and Mater- 
ials). 1988 (02974). Annual Book of ATM Stand- 
ards, Volume 04.08. ASTM, Philadelphia, PA. 

ASTM (American Society for Testing and Mater- 
ials). 1997. Annual Book of ATM Standards, 
Volume 11.05. ASTM, Philadelphia, PA. 

Burt, R., Wilson, M.A., Mays, M.D., and Lee, C.W. 
2003. Major and trace elements of selected pedons 
in the USA. /. Environ. Qual. 32: 2109-2121. 

Chen, B., Krachler, M., Gonzalez, Z.I., and 
Shotyk, W. 2005. Improved determination of 
arsenic in environmental and geological specimens 
using HG-AFS. /. Anal. At. Spectrom. 20: 95-102. 

Daughtrey, Z.W., Gilliam, J.W., and Kamprath, 
E.J. 1973. Soil test parameters for assessing plant- 
available P of acid organic soils. Soil Sci. 115: 

Day, J.H., Rennie, P.J., Stanek, W., and Raymond, 
G.P. 1979. Peat testing manual. Associate commit- 
tee on geotechnical research. National Research 
Council of Canada. Technical Memorandum No. 
125, Ottawa, Canada. 

Goldin, A. 1987. Reassessing the use of loss-on- 
ignition for estimating organic matter content in 
noncalcareous soils. Commun. Soil Sci. Plant 
Anal. 18: 1111-1116. 

Harrison, A.F. 1979. Variation of four phosphorus 
properties in woodland soils. Soil Biol. Biochem. 
11: 393-403. 

Ilnicki, P. and Zeitz, J. 2003. Irreversible loss of 
organic soil functions after reclamation. In L.-E. 

Parent and P. Ilnicki. Eds. Organic Soils and Peat 
Materials for Sustainable Agriculture. CRC 
Press, Boca Raton, FL, pp. 15-32. 

Krachler, M., Mohl, C, Emons, H., and Shotyk, W. 
2002. Analytical procedures for the determi- 
nation of selected trace elements in peat and 
plant samples by inductively coupled plasma 
mass spectrometn . Spectrochim. Acta, Part B, 57: 

Melaku, S., Dams, R., and Moens, L. 2005. 
Determination of trace elements in agricultural soil 
samples by inductively coupled plasma-mass spec- 
trometry: microwave acid digestion versus aqua 
regia extraction. Anal. Chim. Acta 543: 1 17-123. 

Mokma, D.L. 2005. Organic soils. In D. Hillel, 
J.L. Hatfield, D.S. Powlson, C. Rosenzweig, 
K.M. Scow, M.J. Singer, and D.L. Sparks, Eds. 
Encyclopedia of S. he Environ) i >1. 3 

Elsevier Academic Press, Amsterdam, pp. 1 18-129. 

Morrell, J.J., Keefe, D., and Baileys, R.T. 2003. 
Copper, zinc, and arsenic in soil surrounding 
Douglas-Fir poles treated with ammoniacal 
copper zinc arsenate (ACZA). /. Environ. Qual. 
32: 2095-2099. 

Okruszko, H. and Ilnicki, P. 2003. The moorsh 
horizons as quality indicators of reclaimed 
organic soils. In L.-E. Parent and P. Ilnicki, Eds. 

Organic Soils and Pent Materials for Sustainable 
Agriculture. CRC Press, Boca Raton, FL, pp. 1-14. 

Papp, C.S.E. and Harms, T.F. 1985. Comparison 
of digestion methods for total elemental analysis 
of peat and separation of its organic and inorganic 
components. Analyst 110: 237-242. 

Parent, L.E. and Khiari, L. 2003. Nitrogen and 
phosphorus balance indicators in organic soils. In 
L.-E. Parent and P. Ilnicki, Eds. Organic Soils 
and Peat Materials for Sustainable Agriculture. 
CRC Press, Boca Raton, FL, pp. 105-136. 

Soil Survey Staff. 2003. Keys to Soil Taxonomy, 
9th edn. USDA. Natural Resources Conservation 
Service, Washington, DC. 

Swami, K., Judd, CD., Orsini, J., Yang, K.X., and 
Husain, L. 2001. Microwave assisted digestion of 
atmospheric aerosol samples followed by induct- 
ively coupled plasma mass spectrometry deter- 
mination of trace elements. Fresen. J. Anal. 
Chem. 369: 63-70. 

van Lierop, W. and Mackenzie, A.F. 1977. Soil 
pH and its application to organic soils. Can. J. Soil 
Sci. 57: 55-64. 

Kreshtapova, V.N., Krupnov, R.A., and Uspenskaya, 
O.N. 2003. Quality of organic soils for agricultural 
use of cutover peatlands in Russia. In L.-E. Parent 

ind P. Ilnicki. Ed ' o aid Peat Materials 

for Sustainable Agriculture. CRC Press, Boca Raton, 
FL, pp. 175-186. 

Wang, J., Nakazato, T., Sakanishi, K., Yamada, O, 
Tao, H, and Saito, I. 2004. Microwave diges- 
tion with HNO3/H2O2 mixture at high temperat- 
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R.L. Westerman, Ed. Soil Testing and Plant 300-305. 
Analysis, 3rd edn. Soil Science Society of 

America, Madison, WI, pp. 691-740. Williams, D.E. and Vlamis, J. 1961. Boron con- 
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Section Editors: E. Topp and C.A. Fox 


Chapter 29 

Cultural Methods for Soil 

and Root-Associated 


J.J. Germida and J.R. de Freitas 

University of Saskatchewan 
Saskatoon, Saskatchewan, Canada 


Soil is an ecosystem that contains a variety of microbial populations whose members 
represent many physiological types. For example, some microorganisms, such as fungi, are 
aerobic chemoorganotrophs (heterotrophs) and use organic compounds as a source of carbon 
and energy. Others, such as nitrifying bacteria, are aerobic chemolithotrophs (autotrophs) 
using CO2 as a carbon source and oxidizing reduced inorganic N compounds to obtain 
energy. Some microorganisms require special growth factors, a specific environmental pH, 
low O2 levels, or the absence of O2 (i.e., anaerobes) for optimum growth. The chemical, 
physical, and biological characteristics of a particular soil, as well as the presence of growing 
plants, will influence the numbers and activities of its various microbial components. 
Furthermore, because of the heterogeneous nature of soil, many different physiological 
types of organisms will be found in close proximity to one another. The microbial commu- 
nity in soil is important because of its relationship to soil fertility and the biogeochemical 
cycling of elements, and the potential use of specific members for industrial applications. 
Thus, there is a need to enumerate and isolate major and minor members of the microbial 
community in soils. 

The nonselective enumeration and isolation of soil microorganisms is relatively straightfor- 
ward, but the final result is not necessarily meaningful (Parkinson et al. 1971; Wollum 1994). 
On the other hand, selective enumeration and isolation of specific physiological types of 
microorganisms can provide useful and meaningful data (e.g., Lawrence and Germida 1988; 
Germida 1993; Koedam et al. 1994). Methods to enumerate and isolate soil microorganisms 
are constantly changing as our knowledge of the types of microorganisms present in soil 
expands (Vincent 1970; Woomer 1994; Pepper and Gerba 2005). This chapter provides basic 
principles and references on enumeration procedures and culture media for representative 
types of soil microorganisms. 



Enumeration of viable soil microorganisms may be accomplished by the plate count tech- 
nique or most probable number (MPN) technique. The underlying principles are (i) disper- 
sing of a sample in a suitable diluent, (ii) distributing an aliquot to an appropriate growth 
medium, (iii) incubating inoculated plates under suitable conditions, and (iv) counting the 
developed colonies or MPN tubes. These procedures are fairly standard and may be used to 
enumerate populations in bulk (Germida 1993) and rhizosphere (Goodfellow et al. 1968; Kucey 
1983) soils along with root-associated and endophytic populations (Germida et al. 1998). 

The composition of the growth medium used to enumerate microbial populations is import- 
ant as it will affect the final result. Growth media may be selective or nonselective, although 
no medium is truly nonselective (James 1958). Selective media contain components, which 
allow or favor the growth of a desired group of organisms. Nonselective media should 
encourage growth of as many diverse groups of organisms as possible. To enumerate a 
specific physiological type of microorganism it is usually possible to design a growth 
medium which, when incubated under appropriate conditions of atmosphere and tempera- 
ture, reduces interference from nondesired populations. For example, chitin-degrading 
actinomycetes may be enumerated on medium containing chitin as the sole source of carbon 
and nitrogen (Lingappa and Lockwood 1962; Hsu and Lockwood 1975), and inclusion of 
specific antibiotics prevents growth of undesired organisms (Williams and Davies 1965). 
Similar selective media are available for many different soil microorganisms, e.g., phos- 
phate-solubilizing bacteria (Kucey 1983), siderophore-producing microorganisms (Koedam 
et al. 1994), free-living nitrogen-fixing bacteria (Rennie 1981; Knowles and Barraquio 
1994), nitrifying bacteria (Schmidt and Belser 1994), or sulfur-oxidizing organisms (Post- 
gate 1966; Germida 1985; Lawrence and Germida 1988). 

Although the plate count and MPN techniques are simple to perform, their usefulness will be 
limited by a number of key factors (Germida 1993; Wollum 1994; Woomer 1994). Many 
times choice of media, problems with dispersion, and even adsorption of microbes to pipette 
walls can interfere with standardization of these procedures. It should be pointed out that 
consistency and adequate use of replicate samples will help to minimize some of these 
problems. Investigators must realize that microorganisms are not uniformly dispersed within 
the soil environment, and that numbers of any particular microorganism are not synonymous 
with its importance. 


Enumeration of microbial populations by the spread plate method is a simple and rapid 
method to count viable microbial cells in soil. However, counts obtained are generally 10- to 
100-fold less than those determined by microscopic counts of soil smears (Skinner et al. 
1952). Reasons for this discrepancy include measurement of viable and nonviable counts in 
soil smears, and the inability to provide adequate or appropriate nutrients in the growth 
media for spread plate counts (Germida 1993). Basically, this method consists of preparing a 
serial dilution (e.g., 1:10 dilutions) of a soil sample in an appropriate diluent, spreading an 
aliquot of a dilution on the surface of an agar medium, and incubating the agar plate under 
appropriate environmental conditions. These spread plates may be used not only for counting 
microbial populations but also as a starting source for isolation of organisms. In this latter 
case, an isolated colony is picked and repeatedly streaked on a suitable growth medium to 
check for purity. After several such transfers it may be cultured and preserved for future 


study and identification. Selective or nonselective media may be used, depending on the 
nature of the desired microorganisms. Soil extract agar (James 1958) is commonly used as a 
nonselective medium for enumerating soil bacteria. Recently proposed alternatives to soil 
extract agar include a defined "soil solution equivalent medium" (Angle et al. 1991), 
"trypticase soy agar and/or R2A medium" — two commercially available complex media 
(Martin 1975; Reasoner and Geldreich 1985). 

29.3.1 Materials 

7 Petri plates containing ca. 20 ml_ of an appropriate agar medium, e.g., soil extract 

2 Dilution bottles (e.g., 50 x 160 mm; 200 mL capacity) and (or) test tubes (e.g., 
1 8 x 1 50 mm) containing appropriate diluent such as sterile tap water 

j Sterile 1 and 10 mL pipettes 

4 Glass spreader (i.e., glass rod shaped like a hockey stick) 

5 Glass beaker containing 95% ethanol (ETOH) 

29.3.2 Preparation of Agar Petri Plates 

7 Agar media may be prepared from commercially available dehydrated compon- 
ents or from recipes found in the literature. The American Type Culture Collection 
(2005) and Atlas (1995) are an excellent source of media recipes and relevant 
references. Prepare media according to directions, sterilize in the autoclave at 
1.05 kg cirr 2 and 121°C for 20 min, cool to a pouring temperature of ca. 48°C. 
Some components of a medium may be heat labile and must be filter-sterilized 
and then added to the autoclaved agar medium (cooled to ca. 48°C-49°C) just 
before pouring plates. 

2 Distribute ca. 20 mL of media into sterile glass or disposable, presterilized plastic 
Petri plates and allow the agar to solidify. The plates should be allowed to sit at 
room temperature for 24-48 h, allowing excess surface moisture to be absorbed 
into the agar; this helps prevent microbial colonies from spreading over the agar 
surface. Poured plates not used immediately may be stored under refrigeration 
(2°C-5°C) for up to 2 weeks in plastic bags. These stored plates should be removed 
and allowed to warm to room temperature before use. 

29.3.3 Preparation of Soil Dilutions 

7 Samples should be collected, handled, and stored with due consideration to their 
ultimate use, and the effects these steps will have on microbial populations. 

2 Pass representative soil samples through a 2 mm mesh sieve and mix thoroughly. 

3 Weigh out a 1 0.0 g portion of the soil into a dilution bottle containing 95 mL of a 
diluent. Glass beads (ca. 25x2 mm beads) may be added to this dilution blank to 
facilitate mixing. Cap the bottle, place on a mechanical shaker, and shake for 


10 min. Alternatively, shake by hand moving the bottle through specified arc a 
number of times (e.g., a 45° arc at least 50-1 00 times). This first dilution represents 
a 1:10 or a 10~ 1 dilution. 

4 After removing the bottle from the shaker, shake vigorously before removing 
aliquots. To prepare a serial 1:10 dilution series of the soil sample, transfer a 
10 ml_ sample to a 90 ml_ dilution blank cap and shake the dilution bottle 
(alternatively it is possible to transfer 1 ml_ samples to 9 ml_ dilution blanks 
[prepared in 1 8 x 1 50 mm test tubes]). Continue this sequence until a dilution of 
10~ 7 is reached. Subsequent spread plating of a 0.1 ml_ aliquot of this dilution will 
allow enumeration of up to 3 x 1 10 colony-forming units (cfu) per g soil. Experi- 
ence will indicate the appropriate range of dilutions for samples being analyzed. 

29.3.4 Preparation of Dilutions from Root-Associated Bacteria 

For bacteria isolation from the root interior (endophytic), root material is collected from soil 
and maintained at ca. 5°C until processed in the laboratory (Foster and Rovira 1976). 
Approximately 10.0 g of fresh root material is shaken to eliminate adhering soil and then 
washed in sterile tap water. Subsequently, roots are placed in a 250 mL Erlenmeyer contain- 
ing 100 mL of 1.05% sodium hypochlorite (NaCIO) and shaken (200 rpm) for 10 min. Roots 
are washed (4x) in sterile tap water and aseptically blended in a sterile Waring blender 
containing 90 mL sterile phosphate buffered saline (PBS) — see recipe below. This will result 
in 1/10 dilution, i.e., 10.0 g roots and 90 mL sterile water. The root suspension is serially 
diluted in PBS and aliquots of appropriate dilutions spread plated onto selective and/or 
nonselective nutrient media depending on the nature of the study. 

29.3.5 Preparation of Agar Spread Plates 

/ Select a range of four dilutions that will adequately characterize the microorgan- 
isms in the sample. Transfer 0.1 mL aliquots to a separate plate from the 
highest dilution. Note that a 0.1 mL aliquot from a 10~ 7 dilution corresponds to 
an actual dilution of 10~ 8 on the plate. Repeat the process, transferring 0.1 mL 
aliquots from each of the next three successive and lower dilutions onto each of 
triplicate plates for each dilution. 

2 Spread the suspension on the agar surface using a sterile glass spreader for each 
plate. The glass spreader is kept submerged in a beaker of ETOH and excess 
ETOH burned off before use. In the spreading step start with the highest dilution 
and progress to the next lower dilution, continuing the sequence until all the 
plates have been spread. Alcohol flame the glass spreader between each plate. 
Invert the plates and place in an incubator at an appropriate temperature. 

3 Incubation conditions will depend on the facilities and the purpose of the study. 
When possible, one should try to mimic environmental conditions. Generally, 
spread plates samples are incubated in the dark, in an aerobic environment at a 
temperature between 24°C and 28°C. Incubation periods and conditions will vary 
depending on the nature of the organisms being enumerated. 

4 After a suitable incubation interval, plates are removed from the incubator and 
those containing 30 to 300 colonies are counted. Plates with spreading or 


swarming organisms should be excluded from the final count. The colonies can 
be counted manually or by an automated laser colony counter. 

29.3.6 Calculations 

Average the number of colonies per plate for the dilution giving between 30 and 300 
colonies. Determine the number of cfu g _1 of dry soil (DW) as follows: 

dry weight soil, initial dilution 


/ % moisture, soil sample\ 
Dry weight soil = (weight moist soil, initial dilution) x I 1 I 

V ioo / 



Because bacteria may exist in soil as groups or clumps of cells, it is often desirable to 
disperse these cells so that colonies on spread plates arise from one cell. This may be 
accomplished by shaking on a mechanical shaker or by hand, through application of a 
high shearing force as with Waring blender, by sonic vibration (Stevenson 1959), mechanical 
vibration (Thornton 1922), and through the use of deflocculating agents. 

29.3.8 Type of Dilutions 

A number of diluents may be used. In most cases, tap or distilled water is adequate. Other 
diluents routinely used include: 

7 Physiological saline: NaCI, 8.5 g; distilled water, 1 L. 

2 PBS: NaCI, 8.0 g; KH 2 P0 4 , 0.34 g; K 2 HP0 4 , 1 .21 g; distilled water, 1 L. Adjust pH 
to 7.3 with 0.1 M NaOH or HCI. 

3 Peptone water: Peptone, 1.0 g; distilled water, 1 L. 

29.3.9 Types of Media 

The choice of medium depends on the type of organism desired. Media may be made 
selective by omitting or altering a component, or by incubation conditions. For enumeration 
of total heterotrophic populations in soil, a general nonselective medium is usually 
employed. The following are examples of growth media commonly used to enumerate 
total soil bacteria, root-associated bacteria, actinomycetes, and fungi. Additional examples 
of specific media are described in Section 29.5. 

Media for Total Heterotrophic Bacteria 

1 Soil extract agar (James 1958): One kg of soil is autoclaved with 1 L of water for 
20 min at 1 .05 kg crrr 2 . The liquid is strained and restored to 1 Lin volume. If it is 


cloudy, a little CaS0 4 is added and after being allowed to stand, it is filtered 
through Whatman paper No. 5. The extract may be sterilized and solidified with 
agar (1 .5%) as it is, or after the addition of other nutrients, e.g., 0.025% K 2 HP0 4 
or 0.1% glucose, 0.5% yeast extract, and 0.02% K 2 HP0 4 . 

2 Tryptic soy agar (Martin 1 975): Add 3.0 g of tryptic soy broth and 1 5.0 g of agar to 

I L distilled water. Sterilize the medium by autoclaving. 

j R2A (Reasoner and Geldreich 1 985): Yeast extract, 0.5 g; Proteose Peptone No. 3, 
0.5 g; casamino acids, 0.5 g; glucose, 0.5 g; soluble starch, 0.5 g; K2HPO4, 0.3 g; 
MgS04 • 7H 2 0, 0.05 g; sodium pyruvate, 0.3 g; agar, 15.0 g; distilled water, 1 L. 
Adjust the pH to 7.2 with crystalline K 2 HP0 4 or KH 2 P0 4 and sterilize the 
medium by autoclaving. 

Media for Actinomycetes 

Starch-casein agar (Kiister and Williams 1966): Starch, 10.0 g; casein (vitamin free), 0.3 g; 
KNO 3 ,2.0g;NaCl,2.0g;K 2 HPO 4 ,2.0g;MgSO 4 ■ 7H 2 0, 0.05 g;CaC0 3 , 0.02 g;FeS0 4 • 7H 2 0, 
0.01 g; agar, 15.0 g; distilled water, 1 L; pH, 7.2. Sterilize in autoclave as described above. Media 
can be improved by addition of fungistatic agents (Williams and Davies 1965). 

Media for Fungi 

, Czapek-Dox agar: Solution I: Sucrose, 30.0 g; NaN0 3 , 2.0 g; K 2 HP0 4 , 0.1 g; KCI, 
0.5 g; MgS0 4 • 7H 2 0, 0.5 g; FeS0 4 , trace; distilled water, 500 ml_; pH, 3.5 using 
1 0% lactic acid. Solution II: Agar, 1 5.0 g; distilled water, 500 ml_. If desired, 0.5 g 
of yeast extract may be added (Gray and Parkinson 1968). Sterilize solutions I and 

II by autoclaving separately. Cool both solutions to pouring temperature, pour 
solution I aseptically into solution II and use immediately. 

2 Streptomycin-rose bengal agar (Martin 1950): Glucose, 10.0 g; peptone, 5.0 g; 
KH 2 PC> 4 , 1 .0 g; MgS0 4 • 7H 2 0, 0.5 g; rose bengal, 0.03 g; agar, 20.0 g; tap water, 
1 L. Autoclave, cool medium to about 48°C and add 1 ml_ of a solution of 
streptomycin (0.3 g 10 ml_~ 1 sterile water). The final concentration of strepto- 
mycin in medium should be about 30 (xg ml_~ 1 . 


This method employs the concept of dilution to extinction in order to estimate the number of 
microorganisms in a given sample (Taylor 1962; Woomer 1994). It is based on the presence 
or absence of microorganisms in replicate samples in each of several consecutive dilutions of 
soil. For example, if a series of test tubes containing broth medium are inoculated with 
aliquots representing a dilution series from 10~ 4 through 10~ 7 , and the highest dilution 
exhibiting growth is 10~ 5 , then the number of cells present may be estimated to be between 
10 4 and 10 5 . The key is that the desired organism must possess a unique characteristic or 
metabolic trait, which can be detected. Thus, this technique can be used to count micro- 
organisms based on growth (i.e., turbidity), metabolic activity such as substrate disappear- 
ance and product formation. Other uses for the MPN technique include enumeration 
of infective vesicular-arbuscular mycorrhizae (VAM) propagules in soil (see Chapter 
30) or nodule-forming rhizobia in soil (see Chapter 31). 


t total heterotrophic counts of a soil sample the MPN procedure is similar to the 
spread plate count method, except that aliquots of dilutions are inoculated into test tubes of 
liquid medium. Alternatively, multiwell microtiter plates (e.g., 24 wells per plate) may be 
used in place of test tubes; this allows a savings of materials, reagents, incubator space, and 
allows for increased replication. 

29.4.1 Materials 

1 Twenty five test tubes containing appropriate culture medium or 1 disposable 
sterile microtiter plates — 24 or 96 wells per plate 

2 Water dilution blanks (see Section 29.3.1) 

3 Sterile 1 and 10 ml_ pipettes (see Section 29.3.1) 

29.4.2 Procedures 

7 Prepare medium appropriate for the desired organism. 

2 Dispense aliquots of medium in test tubes and sterilize or dispense aliquots of 
sterile medium into presterile microtiter plates. 

3 Prepare a serial 1:10 dilution sequence of the soil sample (see Section 29.3.2). 

4 Select a range of dilutions that will adequately characterize the organisms in the 
sample. Transfer 0.1 ml_ aliquots to each separate well in five replicate microtiter 
plate wells, starting with the highest dilution. Repeat the procedure, transferring 
0.1 mL aliquots from each of the next successive and lower dilutions into each of 
the five replicate wells for each dilution. 

5 Incubate MPN assay tubes and/or plates under appropriate conditions. 

g After suitable incubation, score wells positive for growth or physiological reaction. 

29.4.3 Calculations 

The MPN of organisms in the original sample is calculated by reference to an MPN table 
(e.g., Cochran 1950). Designate as p\ the number of positive tubes in the least concentrated 
dilution in which all tubes are positive or in which the greatest number of tubes is positive. 
Let p2 and p^ represent the numbers of positive tubes in the next two higher dilutions. Refer 
to Table 29.1 and find the row of numbers in which p\ and p2 correspond to the values 
observed experimentally. Follow that row of numbers across the table to the column headed 
by the observed value of p^. The figure at the point of intersection is the MPN of organisms 
in the quantity of the original sample represented in the inoculum added in the second 
dilution. This figure is multiplied by the appropriate dilution factor to obtain the MPN value 
for the original sample. 

As an example, consider the instance in which a 10-fold dilution with 5 tubes at each dilution 
yielded the following numbers of positive tubes after incubation: 5 at 10~ 4 , 5 at 10~ 5 , 4 at 
10~ 6 , 2 at 10~ 7 , and at 10~ 8 . In this series, p x =5, p 2 = 4, and p 3 = 2. For this 


TABLE 29.1 Table of Most Probable Numbers for Use with 10-Fold Dilutions and 5 Tubes per 

Most probable number for indicated values of p 3 


0.0 i 8 

































































































































































































Source: From Cochran, W.G., Biometrics, 5, 105, 1950. With permission. 

combination of p\,pi, and/73, Table 29.1 gives 2.2 as the MPN of organisms in the quantity 
of inoculum applied in the 10~ 6 dilution. Multiplying this MPN by the dilution factor 10 6 
gives 2.2 million as the MPN value for the original sample. 

As a second example, consider the same situation as above except that the most concentrated 
dilution is 10~ 6 . Under these circumstances, p\ = 4, pi = 2, and pj, = 0. For this combin- 
ation of pi, P2, and P3, Table 29.1 gives 0.22 as the MPN of organisms in the quantity of 
inoculum applied in the 10~ 7 dilution. Multiplying 0.22 by 10 7 yields 2.2 million organisms 
as the MPN value for the original sample, as before. 


TABLE 29.2 Factors for Calculating the Confidence Limits for the Most Probable Number Count 
Number of tubes Factor for 95% confidence limits with indicated dilution ratios 

per dilution (n) 











































1 .95 


Source: From Cochran, W.G., Biometrics, 5, 105, 1950. With permission. 

The 95% confidence limits for MPN values can be calculated from prepared tables. A 
compilation of factors keyed to rate of dilution and to number of tubes per dilution is 
shown in Table 29.2. To find the upper confidence limit at the 95% level, multiply the MPN 
value by the appropriate factor from the table. To find the lower limit, divide the MPN value 
by the factor. In the first example above, the factor is 3.30, and the confidence limits are 

(2.2)(3.30) = 7.26 

(2.2)/(3.30) = 0.66. 


The MPN method is usually employed to enumerate and isolate organisms that will not 
readily grow on solid agar medium, or those that cannot be readily identified from the 
background community. For example, autotrophic-nitrifying bacteria (Schmidt and Belser 
1994) and sulfur-oxidizing bacteria (Postgate 1966; Germida 1985; Lawrence and Germida 
1988) are routinely enumerated using the MPN assay. The medium employed may be used to 
measure total growth, and hence optical density is satisfactory measurement. Alternatively, 
a physiological reaction may be monitored. For example, oxidation of sulfur to an acidic 
end product will alter pH and the difference may be recorded by using an appropriate pH 
indicator. The procedure may be used to provide a relative estimate of the numbers of many 
diverse physiological groups of organisms in soils. Choice of media and incubation condi- 
tions is limited only by our knowledge of specific physiological groups. 


29.5.1 Media for Isolation Heterotrophic Bacteria 

7 Peptone yeast extract agar (Goodfellow et al. 1 968): Peptone, 5.0 g; yeast extract, 
1.0 g; FeP0 4 , 0.01 g; agar, 15.0 g; distilled water, 1 L; pH, 7.2. 


2 Nutrient agar: Yeast extract, 1.0 g; beef extract, 3.0 g; peptone, 5.0 g; sodium 
chloride, 5.0 g; agar, 15.0 g; distilled water, 1 L; pH, 7.3. 

j Fluorescent pseudomonads (Sands and Rovira 1970; Simon et al. 1973): Proteose 
Peptone, 20.0 g; agar, 1 2.0 g; glycerol, 1 0.0 g; K 2 S0 4 , 1 .5 g; MgS0 4 • 7H 2 0, 1 .5 g; 
distilled water, 940 mL. Adjust pH to 7.2 with 0.1 M NaOH before autoclaving. 
Sterilize by autoclaving. Add 1 50,000 units of penicillin G, 45 mg of novobiocin, 
75 mg of cycloheximide, and 5 mg of chloramphenicol to 3 mL of 95% ethanol. 
Dilute to 60 mL with sterile distilled water, and add (filter-sterilized using a sterile 
0.45 |Jim Millipore filter) to the cooled (48°C) medium before pouring. Prepared 
plates should be dried overnight before using and may be stored in the refrigerator 
for several weeks before use. 

29.5.2 Media for Isolation of Specific Physiological 
Groups of Organisms 

Microorganisms Involved in Carbon Transformations 

/ Cellulose agar (Eggins and Pugh 1961): NaN0 3 , 0.5 g; K 2 HP0 4 , 1.0 g; 
MgS0 4 • 7H 2 0, 0.5 g; FeS0 4 • 7H 2 0, 0.01 g; cellulose (ball-milled), 12.0 g; 
agar, 15.0 g; distilled water, 1 L. 

2 Chitin agar: Ball-milled, purified chitin, 10.0 g; MgS0 4 • 7H 2 0, 1.0 g; K 2 HP0 4 , 
1 .0 g; agar, 1 5.0 g; distilled water, 1 L. 

j Starch agar: 0.2% soluble starch may be added to any suitable growth medium as 
an alternative or additional carbohydrate. Starch hydrolysis is shown by flooding 
incubated plates with an iodine solution and then noting clear zones. 

Microorganisms Involved in Nitrogen Transformations 

j Combined carbon medium — Free-living putative nitrogen-fixing bacteria (Rennie 
1981): Solution I: K 2 HP0 4 , 0.8 g; KH 2 P0 4 , 0.2 g; Na 2 FeEDTA, 28.0 mg; 
Na 2 Mo0 4 • 2H 2 0, 25.0 mg; NaCI, 0.1 g; yeast extract, 0.1 g; mannitol, 
5.0 g; sucrose, 5.0 g; Na-lactate (60% v/v), 0.5 mL; distilled water, 900 mL; 
agar, 1 5.0 g. Solution II: MgS0 4 • 7H 2 0, 0.2 g; CaCI 2 , 0.06 g; distilled water, 1 00 
mL. Solution III: Biotin, 5.0 (jug mL~ 1 ; p-aminobenzoic acid (PABA), 
10.0 (jug mL~ 1 . Autoclave solutions I and II, cool to 48°C and mix thoroughly, 
then add (filter-sterilized using a sterile 0.45 \im filter) 1 mL L~ 1 of solution III. 

2 Azotobacter enrichment broth: MgS0 4 • 7H 2 0, 0.2 g; K 2 HP0 4 , 1.0 g; 
FeS0 4 • 7H 2 0, 0.02 g; CaCI 2 , 0.02 g; MnCI 2 • 7H 2 0, 0.002 g; NaMo0 4 • 2H 2 0, 
0.001 g; distilled water, 1 L; pH, 7.0; ethyl alcohol (95%), 4.0 mL (add to autoclaved 
and cooled media). 

j Azotobacter chroococcum and A. agilis (Ashby's medium): These two Azotobacter 
species utilize mannitol as their only carbon source. MgS0 4 • 7H 2 0, 0.2 g; K 2 HP0 4 , 
0.2 g; NaCI, 0.2 g; CaS0 4 • 7H 2 0, 0.1 g; CaCO s , 3.0 g; Na 2 Mo0 4 • 2H 2 0, 25.0 mg; 
mannitol, 10.0 g; agar, 15.0 g; distilled water, 1 L. For isolation of A. indicus, 
substitute mannitol by glucose (5.0 g L~ 1 ). 


4 Yeast extract mannitol medium (Allen 1957): Mannitol, 10.0 g; K 2 HP0 4 , 0.5 g; 
NaCI, 0.1 g; MgS0 4 • 7H 2 0, 0.2 g; CaC0 3 , 3.0 g; yeast extract, 0.4 g; agar, 1 5.0 g; 
distilled water, 1 L. 

5 Nitrifying bacteria (Lewis and Pramer 1958): Na 2 HP0 4 , 13.5 g; KH 2 P0 4 , 0.7 g; 
MgS0 4 • 7H 2 0, 0.1 g; NaHC0 3 , 0.5 g; (NH 4 ) 2 S0 4 , 2.5 g; FeCI 3 ■ 6H 2 0, 1 4.4 mg; 
CaCI 2 • 7H 2 0, 18.4 mg; distilled water, 1 L; pH, 8.0. 

Microorganisms Involved in Sulfur Transformations 

1 Thiobacillus thiooxidans or T. thioparus (Postgate 1966): (NH 4 ) 2 S0 4 , 0.4 g; 
KH 2 P0 4 , 4.0 g; MgS0 4 • 7H 2 0, 0.5 g; CaCI 2 , 0.25 g; FeS0 4 , 0.01 g; powdered 
sulfur, 1 0.0 g or Na 2 S 2 7 , 5.0 g; distilled water, 1 L; pH, 7.0. This medium can be 
made selective for T. thiooxidans-\\ke bacteria by using S° as the sulfur source and 
adjusting the initial pH to <3.5. 

2 T. denitrificans (Postgate 1966): KN0 3 , 1.0 g; Na 2 HP0 4 , 0.1 g; Na 2 S 2 7 , 2.0 g; 
NaHC0 3 , 0.1 g; MgCI 2 , 0.1 g; distilled water, 1 L; pH, 7.0. This medium may be 
used for agar plates, or dispensed into test tubes containing small Durham 
fermentation tubes to capture gas. Incubate under anaerobic conditions. 

Allen, O.N. 1957. Experiments in Soil Bacteri- 
ology, 3rd ed. Burgess Publishing Company, Min- 
neapolis, MN. 

American Type Culture Collection 2005. Microbial 
media formulations, web page address: http://www. 
(last verified April 2006), Rockville, MD. 

Angle, J.S., McGrath, S.P., and Chaney, R.L. 
1991. New culture medium containing ionic con- 
centrations of nutrients similar to concentrations 
found in soil solutions. Appl. Environ. Microbiol. 
57: 3674-3676. 

Atlas, R.M. 1995. Handbook of Media for Envir- 
onmental Microbiology. CRC Press, Boca Raton, 
FL, 544 pp. 

Cochran, W.G. 1950. Estimation of bacterial 
densities by means of the "most probable num- 
ber". Biometrics 5: 105-116. 

Eggins, H.O.W. and Pugh, G.J.F. 1961. Isolation 

of cellulose-decomposing fungi from soil. Nature 
(London) 193: 94-95. 

Foster, R.C. and Rovira, A.D. 1976. Ultrastructure 
of wheat rhizosphere. New Phytol. 76: 343-352. 

Germida, J.J. 1985. Modified sulfur containing 
media for studying sulfur oxidizing microorgan- 
isms. In D.E. Caldwell, J.A. Brierley, and C.L. 
Brierly, Eds. Planetary Ecology. Van Nostrand 
Reinhold, New York, NY, pp. 333-344. 

Germida, J.J. 1993. Cultural methods for soil 
microorganisms. In M.R. Carter, Ed. Soil Sam- 
pling and Methods of Analysis. A Special Publi- 
cation of the Canadian Society of Soil Science. 
Lewis Publishers, Boca Raton, FL, pp. 263-275. 

Germida, J.J., Siciliano, S., de Freitas, J.R., and 
Seib, A.M. 1998. Diversity of root-associated bac- 
teria associated with field-grown canola (Brassica 
napus L.) and wheat ( Triticum aestivum L.). FEMS 
Microbiol. Ecol. 26: 43-50. 

Goodfellow, M., Hill, I.R., and Gay, T.R.G. 1968. 
Bacteria in a pine forest soil. In T.R.G. Gray and D. 
Parkinson, Eds. The Ecology of Soil Bacteria. Uni- 
versity of Toronto Press, Toronto, ON, Canada, 
pp. 500-515. 


Gray, T.R.G. and Parkinson, D., eds. 1968. The 
Ecology of Soil Bacteria. An International Sym- 
posium. University of Toronto Press, Toronto, 
ON, Canada. 

Hsu, S.C. and Lockwood, J.L. 1975. Powdered 
chitin as a selective medium for enumeration of 
actinomycetes in water and soil. Appl. Microbiol. 
29: 422-426. 

James, N. 1958. Soil extract in soil microbiology. 
Can. J. Microbiol. 4: 363-370. 

Knowles, R. and Barraquio, W.L. 1994. Free- 
living dinitrogen fixing bacteria. In R.W. Weaver 
et al., Eds. Methods of Soil Analysis, Part 2 — 
Microbiological and Biochemical Properties. 
Soil Science Society of America, Madison, WI, 
pp. 180-197. 

Koedam, N, Wittouck, E., Gaballa, A., Gillis, A., 
Hofte, M., and Cornells, P. 1994. Detection and 
differentiation of microbial siderophores by iso- 
electric focusing and chrome azurol S overlay. 
Biometalsl: 287-291. 

Kucey, R.M.N. 1983. Phosphate-solubilizing bac- 
teria and fungi in various cultivated and virgin 
Alberta soils. Can. J. Soil Sci. 63: 671-678. 

Kiister, E. and Williams, S.T. 1966. Selection of 
media for isolation of streptomycetes. Nature 
(London) 202: 928-929. 

Lawrence, J.R. and Germida, J.J. 1988. 
Most-probable number procedure to enumerate 
S°-oxidizing, thiosuli'aie producing heterotrophics 
in soil. Soil Biol. Biochem. 20: 577-578. 

Lewis, R.F. and Pramer, D. 1958. Isolation of 
Nitrosomonas in pure culture. J. Bacterial. 76: 

Lingappa, Y. and Lockwood, J.L. 1962. Chitin 
media for selective isolation and culture of 
actinomycetes. Phytopathology 52: 317-323. 

Martin, J.K. 1975. Comparison of agar media for 
counts of viable soil bacteria. Soil Biol. Biochem. 
7: 401^102. 

Martin, J.P. 1950. Use of acid, rose bengal and 
streptomycin in the plate method for estimating 
soil fungi. Soil Sci. 69: 215-232. 

Parkinson, D., Gray, T.R.G., and Williams, S.T. 
1971. Methods for Studying the Ecology of Soil 

Microorganisms. IBP Handbook No. 19. Black- 
well Scientific Publications, Oxford, UK. 

Pepper, I.L. and Gerba, C.P. 2005. Environmental 
Microbiology: A Laboratory Manual, 2nd ed. 
Elsevier, Amsterdam, the Netherlands, 209 pp. 

Postgate, J.R. 1966. Media for sulphur bacteria. 
Lab. Pract. 15: 1239-1244. 

Reasoner, D.G. and Geldreich, E.E. 1985. A new 
medium for the enumeration and subculture of 
bacteria from potable water. Appl. Environ. 
Microbiol. 49: 1-7. 

Rennie, R.J. 1981. A single medium for the isol- 
ation of acetylene-reducing (dinitrogen-fixing) 
bacteria from soils. Can. J. Microbiol. 27: 8-14. 

Sands, D.C. and Rovira, A.D. 1970. Isolation of 

fluorescent pseudomonads with a selective med- 
ium. Appl. Microbiol. 20: 513-514. 

Schmidt, E.L. and Belser, L.W. 1994. Autotrophic 
nitrifying bacteria. In R.W. Weaver et al., Eds. 
Methods of Soil Analysis, Part 2 — Microbiological 
and Biochemical Properties. Soil Science Society 
of America, Madison, WI, pp. 159-179. 

Simon, A., Rovira, A.D., and Sands, D.C. 1973. 
An improved selective medium for the isolation 
of fluorescent pseudomonads. J. Appl. Bacterid. 
36: 141-145. 

Skinner, F.A., Jones, P.C.T., and Mollison, J.E. 
1952. A comparison of a direct- and a plating- 
counting technique for the quantitative estimation 
of soil microorganisms. /. Gen. Microbiol. 6: 

Stevenson, I.L. 1959. The effect of sonic vibra- 
tion on the bacterial plate count of soil. Plant Soil 
10: 1-8. 

Taylor, J. 1962. The estimation of numbers of 
bacteria by ten-fold dilution series. J. Appl. Bac- 
terid. 25: 54-61. 

Thornton, H.G. 1922. On the development of a 
standardized agar medium for counting soil bac- 
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spreading colonies. Ann. Appl. Biol. 9: 241-274. 

Vincent, J.M. 1970. A Manual for the Practical 
Study of the Root-Nodule Bacteria. IBP Hand- 
book No. 15. Blackwell Scientific Publications, 
Oxford, UK. 


Williams, S.T. and Davies, F.L. 1965. Use of and Biochemical Properties. Soil Science Society 

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251-261. Woomer, P.L. 1994. Most probable number 
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Methods of Soil \ 'art 2— ft iological America, Madison, WI, pp. 59-79. 



Chapter 30 
Arbuscular Mycorrhizae 

Y. Dalpe 

Agrk allure and Agri-rood Canada 
Ottawa, Ontario, Canada 

C. Hamel 

Agriculture and Agri-Food Canada 
Swift Current, Saskatchewan, Canada 


Most plant species live in a symbiotic association with mycorrhizal fungi whose establish- 
ment in roots increases the water supply and mineral nutrition. These soil-borne fungi 
colonize the root cortex and develop external filaments, making connecting bridges between 
roots and soil. They are recognized as improving plant fitness and soil quality. Widely 
distributed under all ecosystems, they are the most common type of symbionts involved in 
agricultural systems, influencing both plant production and plant protection. 

The mycorrhizal fungi can be subdivided into three major categories: (1) the arbuscular 
mycorrhizal (AM) fungi, obligate symbionts, which belong to the Glomeromycota associated 
with the majority of herbaceous and cultivated plants and with some deciduous trees, (2) the 
ectomycorrhizal (EM) fungi taxonomically associated with Basidiomycetes and Ascomy- 
cetes found in symbiosis mostly with trees, and (3) the ericoid mycorrhizal (ERM) fungi, a 
symbiosis between mainly Ascomycetes and plants from the Ericaceae family (e.g., blue- 
berry, rhododendron, and heather-type plants). 


Several procedures used in mycorrhizal research are time consuming. Thus, sampling plans 
will often have to be a compromise between a desire for precision and limited resources. More 
intensive or targeted sampling protocols are required in heterogeneous sites to allow for the 
accurate measurement of variables. Therefore, knowledge of the conditions causing variation 
in the distribution of AM structures in soil may help develop efficient sampling plans. 

Sampling strategies must be planned carefully according to the experimental goal, the 
conditions of the study area, and knowledge on how these conditions may influence AM 
fungi distribution in soil. The AM fungi occurrence in soil is largely driven by the plant 


distribution. Some plant species do not host AM fungi. These non-host species are found 
particularly in the families Polygonaceae, Juncaceae, Brassicaceae, Caryophyllaceae, Che- 
nopodiaceae, Amaranthaceae, Cistaceae, Pinaceae, Fagaceae, Ericaceae, and Ranuncula- 
ceae. Furthermore, different plant species may selectively augment different AM species 
in their surroundings, and in some plant communities, AM fungi species distribution may 
follow plant species distribution (Gollotte et al. 2004). 

AM spore abundance generally decreases with soil depth and spores are generally absent 
below the root zone. Approximately 75% of the AM spores and hyphae found in tilled and 
no-tilled fields in eastern Canada are found in the top 15 cm of the soil, but these structures 
are still present at 20-25 cm depth (Kabir et al. 1998b). Very few propagules are found 
below 40-50 cm (Jakobsen and Nielsen 1983). AM populations at depth are not only thinner, 
but their composition may also differ from that of the top soil populations. These populations 
may be more stable than those in the top soil layer, which is influenced by crop management 
(Oehl et al. 2005). AM fungi tend to follow plant root distribution and, in row crops, they are 
more abundant under the row than in between rows (Kabir et al. 1998a). AM root coloniza- 
tion development is a three-phase process including a lag phase, an exponential phase, and a 
plateau (McGonigle 2001). The length of the lag phase depends on the mycorrhizal potential 
of the soil and the mycorrhizal dependency of the host plant and may be increased by lower 
soil temperature. For example, root colonization peaks at flowering in maize and decreases 
thereafter up to host senescence (Kabir et al. 1998a). 

Sampling strategy must be adapted to the requirements of the problems studied. The 
evaluation of AM biodiversity under mixed plant populations would require that samples 
be taken in the vicinity of all plant species. The evaluation of the abundance of AM fungi in 
a given area would require a more random sampling plan. For example, a stratified random 
sampling plan would be the most efficient strategy to generate a soil sample representa- 
tive of an experimental monocultured plot at a given point in time. This procedure is 
given below. 

30.2.1 Materials and Reagents 

1 Labeled plastic bags 

2 Graduated bulb planter or soil probe 

3 Bucket 

30.2.2 Procedure 

/ Take an equal number of soil cores at random in each of the following predeter- 
mined zones: (a) directly on the row and (b) in between rows. In crops with wide 
interrows, one may decide to sample (a) on the row, (b) between rows, and 
(c) midpoint between the row and midrow. The number of separate sampling 
sets will depend on plot size and homogeneity. 

2 Place cores in a bucket as you go, up to completion of the plot sampling exercise. 

j Pour the content of the bucket into an appropriately labeled plastic bag. The soil 
cores can be conveniently pooled to produce a single representative sample for a 
plot, which can be sieved, mixed, and subsampled for different analyses. 


4 Avoid leaving plastic bags with their contents under the sun as they may heat up 
quickly. The use of coolers is recommended. 


Soils are commonly sampled to 15 cm depth, but the selection of a sampling depth may be 
based on specific considerations. For example, one may like to sample the plow layer, i.e., 
the top 20, 15, or 7.5 cm, depending on the tillage system used, or up to a known rooting 
depth. The number of cores required depends on the core size — the larger the core the 
smaller the number of samples — and site uniformity. Heterogeneous sites require more 
intensive sampling. In an apparently homogeneous field plot, it is advisable to take at least 
three core sets if a large core is used (e.g., a bulb planter). Using a soil probe, 10-12 cores 
can rapidly be taken from a plot and mixed in a bucket. 


Root clearing and staining reveal the intraradical phase of AM fungi and allow the evaluation 
of the extent of AM root colonization. Clearing is usually done by boiling root samples in a 
solution of KOH on a hot plate or in the autoclave, to remove alkali-soluble tannins. Tender 
roots like those of cucumber, corn, or onion may require only 5-10 min, whereas clearing 
some tree roots may require more than 1 h in a boiling 10% KOH solution. Roots can be 
cleared at room temperature as well. This method produces high quality root material, but 
requires several hours or days of soaking in the KOH solution. Further soaking in 30% H2O2 
(Phillips and Hayman 1970) or in 3% NaOCl acidified with a few drops of 5 M HC1 (Bevege 
1968) solutions has also been used to remove some residual tannins in roots that are difficult 
to clear. 

Staining solutions have evolved along a decreasing toxicity gradient from being lactophenol- 
based (Phillips and Hayman 1970) to lactoglycerol-based (Brundrett et al. 1994) and 
household vinegar-based (Vierheilig et al. 1998) or mild acids such as dilute hydrochloric 
acid (HC1) or even tonic water (Walker 2005). At last, ordinary permanent ink has been 
proposed as a replacement for possibly carcinogenic stains, such as chlorazol black E, trypan 
blue, and acid fuchsin (Vierheilig et al. 1998). The ink and vinegar staining technique is 
given here, as it is a safe and inexpensive method. Staining with trypan blue is given as an 
alternative. The performance of different stains varies with the quality of the AM root 
material under evaluation and with the plant species. Information on other commonly used 
stains can be found in an article by Brundrett et al. (1984) who compared the performance of 
chlorazol black E, trypan blue, acid fuchsin, and aniline blue. 

30.3.1 Materials and Reagents 

1 Aqueous solution of 10% (w/v) KOH. 

2 Black Shaeffer ink solution: 5% black Shaeffer ink (Ft. Madison, Iowa) in household 
vinegar (5% acetic acid); ortrypan blue solution: 0.5 g trypan blue in 500 ml_ glycerol, 
450 ml_ H 2 0, and 50 ml_ 1% HCI with a destaining solution: 500 ml_ 
glycerol, 450 ml_ H 2 0, and 50 ml_ 1% HCI. 

3 Water acidified with vinegar or 1% HCI. 


4 Sample staining cassettes (Omnisette Embedding Cassettes, Fisher Scientific, 
Nepean, Ontario). In absence of cassettes, vials, beakers, or Erlenmeyer flasks 
can be used as recipients for roots. 

5 Lead pencil. 

6 1 L beaker. 
j Hot plate. 
30.3.2 Procedure 

7 Cut roots into 1 cm fragments and take a representative sample that is small 
enough to lay somewhat loosely in the sample staining cassettes in such a way 
as to permit the circulation of liquids around the roots being processed. A 
representative sample can be achieved by mixing the root pieces in a volume of 
water. Ensure that no soil adheres to the root pieces to be stained. For a repre- 
sentative evaluation of root colonization level, a minimum of 50 and an optimum 
of 100 mounted root segments are required. 

2 Place the root samples in the sample staining cassettes labeled with a lead 
pencil (ink will vanish with KOH). In the absence of sample staining cassettes, 
the whole procedure can be performed using vials, beakers, or Erlenmeyer 
flasks. Root segments are introduced into the containers, soaked in the pre- 
scribed solutions, and recovered manually or by filtration to a 50 |i,m mesh 
nylon sheet. 

^ Place sample staining cassettes in a 1 L beaker. The beaker should not be more 
than half full. 

4 Cover with the KOH solution. 

5 Place the beaker with its contents on a hot plate and boil for the time it takes to 
clear the roots. It usually takes 10-30 min, but it can also take hours, depending 
on the plant species and the quality of the root material. 

6 Discard the KOH solution and rinse the cassettes several times with tap water. 

7 It is then advisable to rinse in acidified water if Shaeffer ink is used for staining or 
in 1% HCI, in the case of trypan blue. 

g Boil gently in the ink-vinegar stain for 3 min or in the trypan blue solution for 
15-60 min. 

9 Discard the ink-vinegar solution and rinse the cassettes in tap water acidified with 
a few drops of vinegar. Trypan blue stained roots are placed in a destaining 
solution: 500 ml_ glycerol, 450 ml_ H 2 0, and 50 ml_ 1% HCI over night. The 
trypan blue staining solution can be carefully filtered to remove root debris and 
reused for subsequent root staining. Store the stained root cassettes or the vials 
containing roots in acidified tap water or in destaining solutions at 4°C. 


Very small root fragments can be secured in between two pieces of nylon screen or filter 
paper before being placed in the sample staining cassette to prevent loss of material. In this 
case, care must be taken to ensure proper rinsing between the clearing and staining solutions. 
Toth et al. (1991) proposed a way to calculate AM fungal biomass from colonized root 
length and root radius. 


Stained roots are most often scored for colonization using the grid-line intersect method 
(Giovannetti and Mosse 1980). McGonigle et al. (1990) modified this method into the 
"magnified intersection method" in which roots are examined at 200 x magnification. 
This method involves the inspection of intersections between the microscope eyepiece 
crosshair and roots. Closer examination allows for accurate recognition and recording of 
arbuscular, hyphal, and vesicular colonization. Hyphal and vesicular colonization should be 
interpreted with caution as they can be produced by nonmycorrhizal fungi. This warning also 
holds for the grid-line intersect method. With the slide method (Giovannetti and Mosse 
1980), 50 to 100 1 cm root sections are mounted on slides in polyvinyl lactoglycerol (PVLG) 
mounting media (Omar et al. 1979) (166 g polyvinylalcohol high viscosity — 24-32 cP — 
dissolved in 10 mL H2O is added to 10 mL lactic acid and 1 mL glycerol). The length of 
colonized root tissue is measured and compared to the total length of root observed. Results 
are expressed as a percent. 

Biochemical methods have also been used to determine AM fungal colonization of roots. 
These sometimes present the advantage of discriminating between AM fungi and other root 
endophytes in addition to allowing the evaluation of lignified or large roots, which cannot be 
cleared. Chitin (Hepper 1977), 24-methyl/methylene sterols (Fontaine et al. 2004), and 
phospholipid fatty acid (PLFA) C16:lco5 (van Aarle and Olsson 2003) were proposed as 
indicators of AM colonization. Biochemical methods of measurements of intraradical AM 
colonization have some limitations. Chitin is not specific to AM fungi; it is present in the cell 
wall of zygomycetous fungi. The relative abundance of sterols and fatty acid indicators is not 
consistent among AM fungal species. Thus, use of biochemical indicators of AM coloniza- 
tion is not recommended for the evaluation of field-grown plant roots as colonization, in this 
case, is likely the result of a mixed fungal population. 

The grid-line intersect method is simple, relatively rapid, and appropriate for most routine 
determinations of the mycorrhizal colonization of roots. This method is described below. 

30.4.1 Materials and Reagents 

1 A plastic Petri dish on the underside of which lines were lightly etched with a 
sharp scalpel blade. Lines can be random or arranged in a grid pattern. Gridded 
dishes can also be directly purchased. 

2 Wash bottle containing water acidified with a few drops of vinegar or HCI. 

3 Tweezers and needle to disperse the roots. 


4 Dissecting microscope. 

5 Two-key desktop counter for keeping track of point counts. 

6 Cleared and stained roots. 

30.4.2 Procedure 

/ A stained root sample is placed in the etched Petri dish and dispersed using a jet of 
acidified water from the wash bottle, tweezers, and needle. 

2 Scanning the grid-lines under the dissecting microscope, the total number of 
points where a root intersects a line is recorded using the key-counter. In parallel, 
the number of these intersects bearing AM colonization is also recorded. For 
example, if AM colonization is found in 42 out of a total of 100 intersects, AM 
root colonization would equal 42%. 


Root length can easily be assessed concurrently with root scoring, using the relationship of 
Newman (1966): 

R=An/2H (30.1) 

where R is the root length, A the area of the Petri dish, n the total number of root x grid-line 
intersects, and H the sum of all etched lines' lengths. 

When root measurement is sought, care must be taken not to lose roots in the processes of 
staining, clearing, washing, and extracting roots from a known amount of soil. Colonized and 
total root length densities are best expressed as lengths of total or colonized roots per volume 
of soil. 


There are four methods for assessing the mycorrhizal potential of the soil: 

/ The most probable number (MPN) method was applied to estimate AM fungal 
propagules in soil by Porter (1979). The method involves repeated serial dilutions 
with pasteurized volumes of the soil to be tested, and growth of a trap plant. Trap 
plant roots are examined for presence or absence of AM colonization, and values 
of MPN are derived from published statistical tables (Fisher and Yates 1963; 
Woomer 1994). The MPN method is laborious and yields only imprecise esti- 
mates of propagule numbers. Furthermore, the intense mixing of the soil under 
study dictated by the method disrupts AM hyphal networks that may also be 
involved in soil infectivity. Thus this method considers the MPN of infective 
spores and vesicles, underestimating soil infectivity. 

2 An improved method for mycorrhizal soil infectivity (MSI) determination 
was proposed by Plenchette et al. (1989) as an alternative to the MPN method. 


With the MSI, soil dilution series are made similarly to the MPN method, but 
population of 10 plants, rather than single plants, are grown. The relationship 
between the percentage of mycorrhizal plants and the minimum amount of 
nonsterile soil in a dilution allows for a calculation of the amount of soil required 
to induce mycorrhizal infection in 50% of the plants, which is one unit of MSI. 
This method is more precise than the MPN but involves growing a plant popula- 
tion rather than single plants as with the MPN, and thus requires the examination 
of hundreds and thousands of root systems. 

3 Franson and Bethlenfalvay (1989) have proposed to count infection units formed 
on a trap plant root system directly extracted from the substrate as the expression 
of the mycorrhizal infectivity of a soil. This requires great skill and precision 
because infection points rapidly become sources of additional infections in 
addition to being hard to distinguish. It is unlikely that all AM fungal propagules 
in a soil sample will be synchronized in initiating root infection and, thus, this 
method will likely underestimate the number of propagules in a soil. 

4 A simple infectivity assay can be more conveniently conducted. Trap plants are 
grown in the soil under examination for 2-4 weeks, a period of time long enough 
for colonization to occur but short enough to avoid mycorrhizal development to 
reach its full potential, a point at which plants may become uniformly colonized 
(see McGonigle 2001). Brundrett et al. (1994) proposed to grow trap plants in 
undisturbed cores to maintain an intact AM mycelium in the soil under evalu- 
ation. In contrast to the MPN and MSI methods, the intact core method accurately 
evaluates soil infectivity. The limitation of this method is that its end result is not a 
convenient number of propagules, but the percentage of colonization of a trap 
plant after a number of days. This method is appropriate to compare the mycor- 
rhizal potential of soils or plots within the framework of an experiment. Further- 
more, AM fungi are filamentous, a type of growth that makes it impossible to 
determine where a propagule starts and ends. Thus, rate of colonization expresses 
soil AM infectivity more realistically than a number of propagules. This method is 
reported below. 

30.5.1 Materials and Reagents 

1 Appropriately labeled steel cylinders with a tapered and sharpened lower end, 
which sends the pressure outward and preserves the soil core when it is pushed 
into the soil. These cylinders serve as growth containers. As many cylinders as 
there are plots multiplied by the number of desired subsamples per plot are 

2 Small wooden board to cover and push cylinders in the soil. 

3 Trowel to lift the cylinders. 

4 Knife to level off the lower end of the soil core. 

5 Nylon mesh and high-gauge rubber bands to close the lower end of the cylinder 
while allowing drainage. 


5 Growth chamber to control temperature, humidity, and lighting conditions to 
allow repetition of the assay at other times. 

j Perforated plastic trays with corrugated bottom that will allow cores to drain 
excess water while maintaining the soil in place upon watering. 

g Germinated seeds or seedlings of the desired species of trap plant. Note that more 
than one species can be grown simultaneously in each core. The number of plants 
of each species should be kept constant among growth cylinders. Clover seedlings 
inoculated with Rhizobium (Brundrett et al. 1994) have been used, but any 
mycotrophic plant species may be used. 

g Polyester wool may be used to cover the surface of cores, particularly in the case 
of soil rich in clay, in order to protect the soil surface structure from water damage 
during watering events. 

30.5.2 Procedure 

/ Cover the cylinders with the wooden board and push it in the soil until filled. 

2 Carefully dig the cylinders out of the soil. 

j Invert the cylinders holding the surface with your palm or on the wooden board, 
level off the lower surface of the core, cover it with the nylon mesh, and secure 
this mesh with a high-gauge rubber band. Avoid exposing the cores to excessive 
heat during the collection process. 

4 Carefully carry the cores to a clean greenhouse work bench. 

5 Insert germinated seeds or plantlets in the center of the cores. Seeds can be placed 
in pairs and thinned to one per pair after a few days. 

6 Cover the soil surface with polyester wool to protect the soil structure. 

7 Place in corrugated trays in growth chamber at required preset settings. 

q Water very gently to field capacity when required, to avoid soil structure degrad- 

9 Grow the trap plants for about 2-4 weeks (see Section 30.5.3). 

10 Clear, stain, and assess the roots, as per the methods given in Section 30.3, for 
their percentage of mycorrhizal colonization using the grid-line intersect method 
(see Section 30.4), and determine simultaneously total root length. 

30.5.3 Comment 

It is worthwhile having extra cores available to assess periodically the status of root 
colonization before harvest and ensure that trap plants have adequate amount of mycorrhizal 



Several methods have been used to quantify the extraradical mycelium of AM fungi. Soil 
chitin measurement has been used to estimate AM fungal biomass in soil (Bethlenfalvay and 
Ames 1987). Chitin measurement does not give an estimate of the active AM extraradical hyphal 
biomass. Chitin is also abundant in invertebrates, exoskeletons, and zygomycetous fungi. 

The spread of AM fungi extraradical hyphae in soil was studied using root exclusion 
chambers. Sequential sampling in compartmentalized growth containers allowed the com- 
parison of the spread of AM fungal species from a root barrier into a hyphal compartment 
(Schuepp et al. 1987; Jakobsen et al. 1992). A rotating wire system was proposed to facilitate 
the task of extracting extraradical hyphae from mineral soil samples with low clay content 
(Vilarino et al. 1993; Boddington et al. 1999). The method of root exclusion chambers has 
been the most popular way to evaluate the size of the extraradical AM hyphae in soil, even 
though there are some limitations to direct measurement of hyphae extracted from soil. AM 
fungal hyphae are generally nonseptate and some researchers found morphological features 
typical of AM mycelia such as hyphal size (Ames et al. 1983), angle of branching, and wall 
characteristics. But considering the different morphologies found among AM genera, the 
range of possible AM extraradical hyphae size (from <1 |jim in the fine endophytes to 
18 |JLm in Glomus manihotis (Dodd et al. 2000)) and, on the other hand, the diversity of other 
soil fungi, one must conclude that the identity of coenocytic hyphae extracted from a soil 
sample is at best uncertain, especially in hyphal pieces bearing no branching. Also, much of 
the AM hyphae extracted from soil are neither viable nor functional and vital staining 
techniques must be applied if the amount of active hyphae is sought. 

Inserted membrane techniques in which a membrane of a material resilient to decomposition 
is inserted in the soil to trap the hyphae that will cross it were proposed by Wright and 
Upadhyaya (1999) and Balaz and Vosatka (2001). These methods are simple and rapid. They 
give a measure of total hyphae cross-section plates. This method can be used with immu- 
nodetection of the AM fungi-specific protein glomalin (Wright and Upadhyaya 1999; Wright 
2000) to assess the proportion of AM and saprophytic fungi. Inserted membrane techniques 
should be considered where data on hyphal density are not necessary. 

Fatty acids are often specific to taxonomic groups. The phosphate group of phospholipids, 
the lipids making up membranes, is rapidly cleaved in soil, and PLFA measurement reflects 
the occurrence of living or recently dead organisms. The measurement of AM fungi PLFA 
indicator thus provides information on the functionality of the organisms. The PLFA 16:lw5 
is the preferred indicator of AM fungal biomass (Balser et al. 2005). The fatty acids 16:lco5, 
18:lco7, 20:4, and 20:5 were proposed as indicative of AM fungi (Olsson et al. 1995; Olsson 
1999). AM fungi do not have completely specific fatty acids; 20:4 and 20:5 are present in 
algae and protozoa but are rare in non-AM fungi and bacteria, and 16:lw5 and 18:lw7 
occur in some bacterial genera but are not normally found in other fungi. Background 
level of the PLFA 16:lco5 ranging from 30% to 60% was attributed to the presence of 
bacteria in soil. The fatty acid 16:lw5 is dominant in many AM fungal species although it was 
absent from several Glomus species and from most Gigaspora species (Graham et al. 1995). 
Whole-cell fatty acid (WCFA) 16:lco5, which is a more specific indicator of AM fungi 
than PLFA 16:lw5, was correlated with hyphal length but the relationship varied seasonally 
(Gryndler et al. 2006). A nonmycorrhizal control can be used to correct for background levels of 
the fatty acid 16:lco5. PLFA data on bacterial biomass, which can be generated simultaneously, 


can be used to interpret changes in the abundance of the fatty acid 16:lw5. Neutral lipid fatty 
acid (NLFA) 16: 1 w5 should also be monitored because it highly dominates reserve fatty acids of 
all AM fungi tested, and because bacteria produce very little neutral lipids. The measurement of 
NLFA 16:lco5 can be used to support observation on the variation in fatty acid 16:lw5 from the 
PLFA fraction. 

A method for lipid extraction from soil and measurement of PLFA and NLFA 16:lw5 
(J.M. Clapperton, personal communication, Agriculture & Agri-Food Canada, Lethbridge, 
Alberta) is given below. 

30.6.1 Materials and Reagents 


7 Weighing boats 

2 35 ml_ glass centrifuge tubes 

3 Dichloromethane (DMC) 

4 Methanol (MeOH) 

5 Citrate buffer 

6 Saturated NaCI solution 

7 7mL glass vials 

8 Pipette 

g Nutating shaker 

10 Centrifuge 

7 7 N 2 -gas flow drying manifold (we use a Reacti-Vap III) 

72 Hot plate at 37°C (we use a Reacti-Therm III) 

Lipid-Class Separation 

7 Clamp-holder construction with 10 clamps (to hold columns) 

2 Pasteur pipettes filled with silica gel up to 2 cm from the top (columns) 

j Pasteur pipettes fitted with pipetting bulb 

4 4 ml_ glass vials 

5 DCM 

f. Acetone 


7 MeOH 

8 N 2 -gas flow drying manifold (we use a Reacti-Vap III) 

9 Hot plate at 37°C (we use a Reacti-Therm III) 

Transmethyl Esterization 

7 N 2 -gas flow drying manifold (we use a Reacti-Vap III) 

2 Hot plate at 37°C (we use a Reacti-Therm III) 

j Pasteur pipettes fitted with pipetting bulb 

4 Micropipette with tips 

5 MeOH 

g H2SO4 (concentrated) 

7 Water bath 

g Hexane 

9 Vortex mixer 

10 Ultrapu re water 

77 Methyl nonadecanoate (19:0; Sigma, Aldrich) 

72 200 |xL glass syringe with needle 

73 1 00 |xL tapered glass inserts and gas chromatograph (GC) vials 

Gas Chromatography Measurement of Fatty Acids 

7 1 6:1 w5 standard fatty acid (from MJS Biolynx #MT1 208). 

2 Gas chromatograph with flame ionization detector (FID). We use a Varian 3900 
GC equipped with a CP-8400 autosampler, helium as carrier gas (30 mL min -1 ), 
and a 50 m Varian Capillary Select FAME #cp7420 column. 

30.6.2 Procedure 

To extract total soil lipids, shake 4 g (dry weight equivalent) of frozen orfresh soil in 
9.5 mL DMC:MeOH:citrate buffer (1 :2:0.8 v/v) for 2 h in glass centrifugation tubes. 

Add 2.5 mL of DMC and 10 mL of a saturated NaCI solution to each tube and 
shake for five more minutes. 


j Centrifuge tubes at 1 500 g for 10 min. 

4 Pipet the organic fraction into clean vials. 

5 Add 5 ml_ of DCM:MeOH (1 :1 v/v) to the tubes, 
g Shake for 15 min. 

7 Centrifuge for 1 min at 1 000 g. 

g Combine the organic fractions in the corresponding vials and dry under a flow of 
N 2 at 37°C in the fume hood. 

9 Dissolve samples in 2 ml_ of DCM. 

1Q Samples can be stored at -20°C for a short time, if necessary. 

Lipid-Class Separation 

Lipid-class separation is conducted in silica gel columns made with Pasteur pipettes. 

7 Using a pipette, load samples onto columns washing the vials twice with a small 
amount of DCM and adding the wash to the columns. Care must be taken to keep 
solvent level above the silica gel at all times. 

2 Elute the neutral lipid fraction first by leaching columns with approximately 2 ml_ 
of DCM, collecting the eluent in 4 ml_ vials. 

j Elute the glycolipid fraction by leaching columns with approximately 2 ml_ of 
acetone, collecting the eluent in other 4 ml_ vials. 

4 Elute the phospholipid fractions by leaching columns with approximately 2 mL of 
MeOH, collecting also the eluent in 4 mL vials. 

5 Discard the glycolipid fraction. 

g Dry the neutral and phospholipid fractions under a flow of N 2 at 37°C in the fume 

y Dissolve the dried fractions in a few mL of MeOH for PLFA or DCM for NLFA and 
store at -20°C. 

Transmethyl Esterization 

Fatty acid methyl esters are created through mild acid methanolysis as follows: 

/ Dry neutral and phospholipids fractions under a flow of N 2 at 37°C in the fume 

2 Add half a Pasteur pipette full of MeOH/H 2 S0 4 (25:1 v/v) to the vials. 


3 Place vials in an 80°C water bath for 10 min. 

4 Cool to room temperature. 

5 Add 1 Pasteur pipette of hexane, vortex vials for 30 s, and leave to settle for 5 min. 
g Discard the lower fraction. 

j Add 1 ml_ of ultrapure water, vortex vials for 30 s, let stand for 5 min. 

g Discard the aqueous fraction entirely. 

g Add 10 |xL of methyl nonadecanoate, the internal standard. 

10 Dry samples under a flow of N 2 at 37°C in the fume hood. 

1 -j Wash vials with 50 julL of hexane using a glass syringe. 

12 Transfer the samples into 1 00 |xL tapered glass inserts, and place inside a GC vial. 

Gas Chromatography Measurement of Fatty Acids 

1 Sample (2 |xL) injection is in 5:1 split mode. 

2 In our program, for example, the injector is held at 250°C and the FID at 300°C. 
The initial oven temperature, 140°C, is held for 5 min, raised to 210°C at a rate of 
2° C min~ 1 , then raised from 21 0°C to 250°C at a rate of 5° C min~ 1 , and finally 
held for 12 min. 

Peak Identification 

Identification of peaks is based on comparison of retention times to a known 16:lw5 
standard. Amounts are derived from the relative area under specific peaks, as compared to 
the 19:0 peak value, which is calibrated according to a standard curve made from a range of 
concentrations of the 19:0 FAME standard dissolved in hexane. The abundance of individual 
PLFAs is expressed as micrograms PLFA per gram dry soil. 

The amount of fatty acid is calculated with the following formula: 

16: lco5 = (A 1Ma)5 /Ai Stid )C istd D (30.2) 

where 16:lto5 is the calculated concentration of the AM fungal indicator (moles or weight 
per unit volume), Ak,^^ is the GC area of the AM fungal indicator, Ai str( j is the GC area of 
the internal injection standard as determined by the GC data system (unitless), Ci st( j is the 
concentration of the internal injection standard given, and D is the appropriate dilution 



Solvents used throughout the procedure are HPLC grade, and tubes and vials are made of 
glass and their screw-top caps lined with Teflon. Organic solvents are toxic and must be 


handled in the fume hood. The use of parafilm is prohibited. Care must be taken to avoid 
contamination with extraneous lipids. For example, the use of gloves is recommended. A 
bacterial and fungal saprobe fatty acid indicators mix (Supelco Bacterial Acid Methyl Esters 
#47080-U) can be used in conjunction with the AM fungal indicator 16:lw5 to simultane- 
ously obtain information on the whole soil microbial community. 


A number of methods have been used to study the extraradical mycelium of AM fungi. The 
cultivation of AM fungi on transformed root cultures has generated considerable knowledge 
on the physiology of AM fungi. This body of work was reviewed by Fortin et al. (2002). This 
system, in which the plant component of the AM symbiosis is reduced to a root often 
transformed by the Agrobacterium rhizogenes plasmid, is artificial. Giovannetti and her 
group successfully used a membrane sandwich method from which much knowledge on the 
extraradical phase of AM fungi was also gained (Giovannetti et al. 1993, 2001). This method 
is closer to reality as whole plants are used, although the environment of the symbioses 
formed is artificial and bidimensional. In this method clean AM spores are germinated in 
between two Millipore7 membranes (0.45 |jim diameter pores) placed on moist sterile quartz 
grit in 14 cm diameter Petri dishes. Clean plantlets are added to the sandwich. Sandwiches 
are harvested at intervals to monitor mycorrhizal development. Rillig and Steinberg (2002) 
have used glass beads of different sizes to simulate different hyphal growing space condi- 
tions to show the large influence of the environment on hyphae length and glomalin 
production. Friese and Allen (1991) have used root observation chambers to describe runner 
hyphae, hyphal bridges, absorptive hyphal networks, germ tubes, and infection networks 
produced in soil by spores and root fragments. Although a root observation chamber allows 
the study of the morphology of arbuscular mycorrhizae formed in soil, this system may not 
be representative of the field situation. The film method in which a soil-molten agar 
suspension was poured into films was proposed by Jones et al. (1948). These agar films 
can be dried and stained to facilitate the enumeration of entrapped organisms under 
the microscope. A modification of this method was used to document AM hyphal links 
formed between the roots of different plant species. This method can be useful for examin- 
ing interactions between roots, AM hyphae, and soil microorganisms in the field; it is 
described below. 

30.7.1 Materials and Reagents 

j Warm 1 .5% water agar 

2 Microscope slides 

3 Tra Y 

4 Thin plastic film (Saran wrap) 

5 Colored plastic flags to facilitate slides recovery in the field 

g A staining solution made of 15 ml_ phenol (5% aqueous), 1 ml_ of aniline blue 
W.S. (1% aqueous), and 4 mL of glacial acetic acid (use fume hood, gloves, and 
eye protection) 


7 Euparal mounting medium (Bioquip, Gardena, California) 

8 95% Ethanol 

30.7.2 Procedure 

1 Place the microscope slides side-by-side in a tray. 

2 Pour the water agar on the slides to produce a thin agar coat on the slides. 

3 When cold and solidified, free and remove the agar-coated slides from the tray 
using a scalpel. Store the fresh agar-coated slides wrapped in plastic film at 4°C. In 
the field, remove the plastic wrap from the slides. Bury the slides vertically in the 
rhizosphere at a chosen distance from the plant roots. 

4 Mark the location of each buried slide with a plastic flag. 

5 After a period of time, carefully dig out the buried slides cutting off the soil around 
the slide with a scalpel and any fine roots that might have grown in the agar coat. 
Agar-coated slides can remain buried for a few months. 

g In the laboratory, delicately wash the bulk of adhering soil off in a water bath. 

7 Dry the agar films at 60°C for 1 5-20 min. 

g Under the fume hood, immerse the dried films for 1 h in the staining solution. 

9 Wash and dehydrate in 95% ethanol and permanent mount in Euparal. 


Phenol and phenol-containing solutions should be handled under the fume hood using gloves 
and eye protection. Good results are also obtained with fuchsin acid staining. 


Soil sampling strategies may generate the need to analyze in detail the composition of 
harvested soil samples for their AM spore population, spore abundance, and spore species 
diversity. Moreover, the extracted spores may provide starting inoculum in the form of 
isolated spores that may be used to obtain mixed or purified species inoculum. 

According to the type of soil worked on, different approaches may be taken to facilitate the 
bulk isolation of spores from a soil substrate. The density gradient centrifugation method for 
spore extraction together with spore sieving and decanting is probably the most common 
method for AM spore extraction, especially for biodiversity and taxonomical studies (Khan 
1999). However, it is very time consuming and less appropriate for most basic AM soil 
species and soil population investigations; the spore extraction method proposed below is the 
result of an adaptation of combined methodologies of soil sieving, decanting, sucrose 


centrifugation, and filtrating. Literature provides descriptions and evaluated methodologies 
that may help in refining the techniques when special requirements are needed. These 
requirements mainly concern diverse gradient methods (Ohms 1957; Allen et al. 1979; 
Furlan et al. 1980; Kucey and McCready 1982) and wet-sieving approaches (Gerdemann 
and Nicolson 1963; Daniels and Skipper 1982; Singh and Tiwari 2001). Book chapters and 
review articles have also been dedicated to the evaluation and description of procedures 
(Tommerup 1992; Brundrett et al. 1996; Clapp et al. 1996; Jarstfer and Sylvia 1997; Johnson 
et al. 1999; Khan 1999). 

Once spores are extracted from soil material, they can be used directly as starting 
inoculum or classified by morphotypes as done for population studies (Smilauer 2001; 
Brundrett 2004). Spores may then be separated according to their size, their color, and 
their subtending hyphae morphology before being mounted on microscopic slides for AM 
fungi diversity assessment. When extracted spores are expected to serve as starting 
inoculum for the propagation of AM fungal strains, a preliminary spore vitality test is 
highly recommended. This can be done by a time-consuming evaluation of the germin- 
ation potential of isolated spores. Moreover, dehydrogenase-activated stains such as 
tetrazolium bromide stain (3-(4,5-dimethylthiazol-yl) 2-5-diphenyl-2H-tetrazolium 
bromide (MTT)) and tetrazolium chloride stain (2-(/?-iodophenyl)-3-(/?-nitrophenyl)-5- 
phenyl-2H-tetrazolium chloride (INT)) have been frequently tested and used for the 
evaluation of AM spore vitality (An and Hendrix 1988; Meier and Charvat 1993; Walley 
and Germida 1995). Even though the interpretation of such staining approaches may be 
confusing, sometimes they remain the easiest and least time-consuming methods to estimate 
the spore vitality of a population. Procedures for spore extraction and viable staining of 
spores are given below. 

Because of the laborious process associated with AM fungi spore extraction from soil 
samples, the limited species- specific spore morphological characters, and the incapacity to 
identify AM fungi from colonized roots, molecular-based techniques have been developed 
to study AM fungal communities. Several polymerase chain reaction (PCR)-based methods 
have been developed and applied to the detection of AM fungi and to the study of genetic 
diversity either directly from soil samples or from colonized root segments (Claassen et al. 
1996; Vandenkoornhuyse et al. 2002). The nested PCRs (two steps amplification) allow a 
rapid and efficient method for the study of soil and root AM fungal communities (van 
Tuinen et al. 1998; Jacquot et al. 2000; Jansa et al. 2003; de Souza et al. 2004). Polymerase 
chain reaction-denaturing gradient gel electrophoresis (PCR-DGGE) can also be used for 
direct analysis of amplified DNA from soil and root samples. It allows the separation of 
DNA products of the same length and the distinction of single-base substitution 
between different nucleotide sequences, does not require a cloning process (Ma et al. 
2005; Sato et al. 2005), and can discriminate between species (Kowalchuk et al. 2002; 
de Souza et al. 2004). 

Finally, quantitative real-time PCR based on the detection of a fluorescent signal produced 
proportionally to amplification of a PCR product allows not only detection but also quan- 
tification of genomic DNA (Filion et al. 2003; Isayenkov et al. 2004). 

General discussion on the comparative value and applications of PCR-based method for the 
study of soil fungi communities and investigations with AM fungal diversity can be 
consulted respectively in Anderson and Cairney (2004) and Clapp et al. (2002). 


30.8.1 Extraction of AM Spores from Soil by Sieving and Sucrose 

Materials and Reagents 

i Scale 

3 Sieves set (suggested mesh size: 1000 |xm to intercept gravel, soil debris; 500, 
1 50, and 50 jjim to recover sporocarps and spores of different sizes) 

4 Centrifuge tubes: round bottom 100 mL 

5 Centrifuge 

5 Vacuum filter apparatus (vacuum source, Buchner funnel and side-arm flask (2 L)) 

7 Filter paper (Whatman No. 1 (40 |xm)) 

q Plastic Petri dishes 

9 Sucrose 50% (w/v) 

1q Tween 80 (optional) 


7 Weigh 50 g of soil and pour in a 1 L flask with 200-300 mL of water. 

2 Shake vigorously and then allow the soil to soak for 30 min to 1 h. 

3 Pour the water and soil mix through the sieves piled in a decreasing order of mesh 
size (largest mesh on the top to retain debris), ensure recovery of the entire soil 
mix by carefully rinsing the flask. This allows the recovery of all soil material for 
spore extraction. 

4 Wash the soil with running water, manually breaking soil aggregates if required, 
being careful not to clog the small mesh sieve. At this step, root pieces can also be 
recovered for either root colonization evaluation or inoculum material. 

5 Recover the entire soil from each sieve, distribute the soil material in centrifuge 
tubes (max 1 mL of soil volume), fill the tube with a 50% (w/v) sucrose solution, 
thoroughly mix the tube content with a glass or metal rod, and centrifuge at 800 g 
for 4 min. (Optional step, see next section.) 

6 Recover supernatants on a 40 |xm paper filter and wash carefully to dilute sucrose 
concentration as it affects the spore wall morphology for the subsequent identifi- 
cation process. 


7 Pour spores in water into a plastic Petri dish, or vacuum filter the spores on filter 
paper for examination under a dissecting microscope. 


When an evaluation of the spore abundance is required, it is necessary to measure soil moisture 
content in order to express spore abundance as the number of spores recovered by weight in 
grams of dry soil weight. For steps 2-A, a drop of dispersant such as Tween 80 can be added to 
water to facilitate the separation of spores from soil debris. The dispersant does not seem to affect 
either spore morphology for further species identification, or spore germination potential. 
Depending on the soil texture, sucrose extraction and centrifugation (step 5) can be skipped 
especially when working with sandy soils because spores are easily separated from silica 
particles by a vigorous shaking of the soil-water-Tween mix. In this case, vacuum filtration is 
recommended and should be performed 2-3 times in order to recover a maximum number of 
spores. On the other hand, AM spore extraction from organic soil containing an abundance 
of partially decomposed plant debris requires the combination of sieving and sucrose 
extraction processes with, in some cases, the repetition of the extraction step 5 at least twice. 
The newly proposed use of low concentration of HC1 or hydrofluoric acid for cleaning and 
separating spores from their surrounding organic material (Garampalli and Reddy 2002) yielded 
cleaner spores suitable for microscopic observations and in vitro culture propagation. However, 
extreme care should be taken in the use of such chemicals and the extraction should be performed 
under a fume hood to avoid inhalation. For hydrofluoric acid special fume hoods are needed. 

30.8.2 Extraction of Vesicles through Enzymatic Digestion of Roots 

Vesicles are excellent AM fungal propagules. Strullu and Plenchette (1991) proposed their 
use in alginate beads as high quality root inoculum. Monoxenic culture of spores can also 
provide high quality inoculum, but only a few species can be grown in vitro. Vesicle 
extraction from root is another way to produce clean inoculum of the AM species that 
cannot be cultivated in vitro. Another application for extracted vesicles is the initiation of 
root organ cultures because intraradical vesicles are devoid of attached organic debris and 
cleaner than soil extracted spores. 

Materials and Reagents 

7 Scalpel 

2 Enzyme solution made of 0.2 g L~ 1 of macerozyme, 0.5 g L~ 1 of driselase, and 
1.0 g L _1 of cellulase 

3 250 ml_ beaker 

4 Blender 

5 50 |xm sieve 


7 Cut 1 g of roots in 3-1 mm segments. 

2 Place root pieces in 100 ml_ of the enzyme solution. 


Incubate overnight at room temperature. 

Wash digested tissues with demineralized water. 

Homogenize in a blender. 

Filter the homogenized sample on a 50 fjurn sieve and recover clusters of vesicles 
attached to hyphae under the dissecting microscope. 

Jabaji-Hare et al. (1984) used mortar and pestle, a homogenizer, filtration, and density 
centrifugation to quantify and recover vesicles for biochemical analysis. 

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Chapter 31 

Root Nodule Bacteria and 

Symbiotic Nitrogen Fixation 

D. Prevost 

Ayic. allure and Agri-rood Canada 
Quebec, Quebec, Canada 

H. Antoun 

Laval University 
Quebec, Quebec, Canada 


Symbiotic nitrogen fixation in plants occurs in root nodules of legumes and nonlegumes. The 
bacterium rhizobium is one of the most studied symbiotic nitrogen-fixing bacteria because 
it nodulates legumes, which are environmentally significant in soil N fertility management of 
cultivated lands. The majority of nonleguminous nodules belong to the Alnus-type symbiosis, in 
which the actinomycete Frankia is the microsymbiont. The cyanobacteria Nostoc or Anabaena 
nodulate the Cycadales, while the bacterium rhizobium forms Parasponia-type symbioses. 

This chapter focuses on the methodology developed to study the rhizobium-legume symbi- 
osis. The global success of legume production is due to the development of inoculation 
technologies and cropping systems by multidisciplinary teams. Microbiologists, soil scien- 
tists, plant physiologists, plant breeders, and agronomists contributed to this breakthrough. 
Increased knowledge in rhizobial ecology is mainly due to the development of molecular 
techniques. Moreover, the taxonomy of the microsymbiont rhizobium has considerably 
changed since the last edition of this chapter (Rice and Olsen 1993). The use of genotypic 
and phenotypic approaches, applied to isolates obtained from a large number of legume 
species and from different regions, resulted in reclassification of known rhizobial species and 
in an increased number of new species. 

Symbiotic rhizobia belong to the a-subclass of Proteobacteria (a-rhizobia). However, some 
tropical legumes are nodulated by strains of Burkholderia and Ralstonia species belonging to the 
p-subclass of Proteobacteria. These strains evolved from diazotrophs through multiple lateral 
nod gene transfers, and this phenomenon seems to be widespread in nature (Chen et al. 2003). 
The current taxonomy of rhizobia (RhizobiaJTaxonomy 2006) includes the genera Rhizobium 


(14 species), Mesorhizobium (10 species), Azorhizobium (1 specie). Sinorhizobium, which 
could be renamed as Ensifer (11 species), Bradyrhizobium (5 species), and six other genera 

(Methylhacteriitin. BurklioUleria. Ralstonia. Dcvosia, Blastobacter, and Ochrobacterium). 

In this chapter, the general term "rhizobia" will be used for the designation of bacteria that 
form nodules on legumes root and stem. Table 31.1 shows the rhizobial species associated to 
some indigenous and cultivated legumes. Recent classification of rhizobia isolated from 
legumes in tropic regions is not included. 

TABLE 31.1 Some Indigenous and Cultivated Legum 
Rhizobial Species 

n Canada and Their Nodulating 

Rhizobial specie! 

Arachis hypogae 


Astragalus cicer 

Cicer milkvetch 

Astragalus sinicus 

Astragalus adsurgens 

Cicer arietinum 



Goat's rue 

Glycine max 


Lathyrus spp. 

Flat pea, tangier pea 

beach pea 

Lathyrus sativus 

Chickling vetch, grass 

Lathyrus pratensis 

Yellow vetchling 

Lens culinaris 


Lotus corniculatus 

Birsfoot trefoil 

Lupinus spp. 

Lupine (white, blue, 


Medicago spp. 


Melilotus spp. 

Sweetclover (white, 


Onobrychis vivifolia 


Oxytropis sp.