Enzymes in the Environment: Activity, Ecology and Applications - Chapter 1 ppsx

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Marcel Dekker, Inc. New York
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Basel
edited by
Richard G. Burns
University of Kent
Canterbury, Kent
England
Richard P. Dick
Oregon State University
Corvallis, Oregon
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Preface
Enzymes that function within plants, animals, and microorganisms are fundamental to
life, and their contributions to metabolic pathways and processes have been studied exten-
sively. For over 100 years there has been interest in what today is called ecological or
environmental enzymology. This aspect of enzymology originates from the work of
Woods, who, in 1899, wrote about the survival and function in soil of plant peroxidases
following their release from decaying plant roots. Environmental enzymologists recognize
that the measured activity may be a composite of reactions taking place in different loca-
tions and at different rates. Thus, in addition to being intracellular, enzymes can be extra-
cellular and attached to the external surfaces of cells, associated with microbial and plant
debris, diffused or actively excreted into the solution phase, and complexed with minerals
and organic compounds.
Although most extracellular enzymes released from cells are rapidly denatured or
degraded, some will survive in solution, if only for short periods. This allows them to
complex with adjacent and appropriate substrates and hydrolyze molecules that are too
large or insoluble to pass through the cell wall or for which there are no uptake mecha-
nisms. These soluble, low molecular mass products can then be utilized as carbon and/
or energy sources by the cell. What this means is that the catalysis of a substrate does
not necessarily represent a homogeneous enzymatic reaction but may be the result of
isoenzymes derived from plants, microorganisms, or animals, and found in various loca-
tions within the soil or sediment matrix.
Much ecological enzymology research is driven by the need to understand the bio-
logical processes that are important for essential aquatic and terrestrial ecosystem func-

tions. These include: organic matter decomposition in relation to both local and global
biogeochemistry; mineralization and the release of inorganic nutrients for use by microbes,
plants, and animals; complex combinations of reactions that determine and maintain soil
Copyright © 2002 Marcel Dekker, Inc.
fertility and soil productivity; and the response to and recovery of soil and aquatic systems
from various natural and anthropogenic perturbations.
Until very recently there have been two large but rather separate camps in the study
of ecological enzymology: those involved with aquatic environments and those who have
concentrated on soil. In aquatic systems the early work included that by Fermi in 1906,
who showed proteolysis activity in stagnant pools, and Harvey in 1925, who suggested
that seawater had catalase and oxidase activity. Subsequent researchers, such as Kreps,
Elster, and Einsele in the 1930s, showed that aquatic bacteria could excrete enzymes into
solution and that these retained a portion of their catalytic activity. Pioneering soil science
work by Rotini and Waksman, among others, was focused on catalase, although the 1940s
saw a surge in influential papers on urease by Conrad and phosphatases by Rogers.
Until the 1950s ecological enzyme research made incremental progress. However,
since then there has been an ever-increasing research output on ecological enzymology,
and in the past 20 years well over 1000 papers have been published. On the aquatic side,
this rapid growth of research was initiated by Overbeck and Reichardt, who demonstrated
the role of extracellular phosphatases from bacteria in the mineralization of organic P
compounds. They showed that the released phosphate was then used by algae that lacked
the ability to directly utilize organic P, thereby showing an important microbial ecological
mechanism for extracellular enzyme activity in aquatic systems. They also carried out
pioneering research on the temporal and spatial distribution of enzyme activities in lake
water. On the soil science front, pioneering work in the 1960s by, among others, McLaren,
Kiss, Ross, Galstyan, Voets, and their coworkers gave an impetus that still drives much
of today’s research. Soil enzymology up to the late 1970s was summarized in the book
Soil Enzymes (Academic Press, 1978).
Ecological enzymology can be divided into two broad and overlapping divisions
that are both well represented in this book. The first can be categorized as microbial

ecology and biochemistry: the study of enzymatic activities in order to better comprehend
the processes or mechanisms that are operating in a given system (Chapters 1, 2, and 3).
This research may have fundamental objectives targeted toward a greater understanding
of highly complex environments such as the rhizosphere (Chapter 4), plant leaves and
shoots (Chapter 6), soil surfaces (Chapter 11), or biofilms (Chapter 12). On the other hand,
it may have explicit applied goals related to manipulating or preserving the environment
in question. These applications include: microbe–plant symbioses (Chapter 5), controlling
plant pathogens (Chapter 7), understanding organic matter decomposition and its impact
on local and global carbon and nitrogen cycles (Chapters 8, 9, and 10), and environmental
remediation of contaminated soils and sediments (Chapters 18, 19, and 20).
The second category of ecological enzymology research includes the use of enzymes
(or microbial cells) as sensors to detect microbial activity and stresses due to pollution,
management, or climatic changes in aquatic and terrestrial ecosystems (Chapters 15, 16,
and 17). In this mode, enzymes can be used to assess nutrient turnover, soil health and
the presence of plant pathogens, and the progress of remediation of polluted soils and
waters. Conventional enzyme assays are attractive as sensors because their integrative
nature, specificity of reactions, and relatively simple methodology make them feasible
for adoption by commercial environmental laboratories. Alternatively, molecular methods
using reporter systems linked to enzymatic processes are being developed for assessing
microbial diversity and function (Chapter 14).
This book, in part, was the result of the historic conference ‘‘Enzymes in the Envi-
ronment: Ecology, Activity and Applications,’’ held in Granada, Spain, in July 1999. This
Copyright © 2002 Marcel Dekker, Inc.
meeting of over 200 scientists from 34 countries was unique because it brought together
scientists from diverse backgrounds around the world who do not normally interact or
attend the same professional meetings. Those enjoying the busy sessions included bio-
chemists and microbial ecologists who study terrestrial or aquatic systems, and environ-
mental and agronomic scientists. Some of the research presented at this meeting was pub-
lished in a special issue of Soil Biology & Biochemistry (Vol. 32, Issue 13, 2000). There
will be a follow-up conference in Prague in July 2003.

An interesting observation arising from the Granada conference was that research
into such diverse microbial ecosystems as plant surfaces, soil aggregates, and biofilms of
aquatic systems or populations at 1000 meters below the surface of the ocean presented
strikingly similar methodological challenges and difficulties in the interpretation of the
information derived (Chapter 21). How do you get a representative environmental sample?
What are the appropriate assay conditions? What do the measured activities tell us about
processes in the environment? What is the microbial and macroecological significance
of extracellular enzymes? Are there commercial applications of extracellular enzymes in
remediation and nutrient provision? And are there lots of microbes and enzymes out there
waiting to be discovered and exploited (Chapter 13)? All these questions and more were
heard frequently. The multidisciplinary group also discussed the ‘‘big’’ issues and respon-
sibilities of current and future developments in environmental enzymology. Two of the
most pressing of these are adequate and sustainable food production in terrestrial and
aquatic ecosystems and counteracting global warming through carbon sequestration and
other processes in soils and aquatic systems. This book presents 21 reviews by interna-
tional experts who attempt to address all these questions and issues. Research progress
in ecological enzymology in terrestrial and aquatic ecosystems is brought into the twenty-
first century.
Richard Burns wishes to thank his wife, Wendy, for her support through this and
other writing adventures and Hugo Z., who continues to give a sense of perspective to
this confusing life. Richard Dick acknowledges Joan Sandeno for her editing assistance.
Richard G. Burns
Richard P. Dick
Copyright © 2002 Marcel Dekker, Inc.
Contents
Preface
Contributors
1. Enzyme Activities and Microbiological and Biochemical Processes
inSoil
Paolo Nannipieri, Ellen Kandeler, and Pacifico Ruggiero

2.EcologyofMicrobialEnzymesinLakeEcosystems
Ryszard Jan Chro
´
st and Waldemar Siuda
3.EcologicalSignificanceofBacterialEnzymesintheMarineEnvironment
Hans-Georg Hoppe, Carol Arnosti, and Gerhard F. Herndl
4.EnzymesandMicroorganismsintheRhizosphere
David C. Naseby and James M. Lynch
5.EnzymesintheArbuscularMycorrhizalSymbiosis
Jose
´
Manuel Garcı
´
a-Garrido, Juan Antonio Ocampo,
and Inmaculada Garcı
´
a-Romera
6.MicrobesandEnzymesAssociatedwithPlantSurfaces
Ian P. Thompson and Mark J. Bailey
Copyright © 2002 Marcel Dekker, Inc.
7. Microbial Enzymes in the Biocontrol of Plant Pathogens
andPests
Leonid Chernin and Ilan Chet
8.MicrobiologyandEnzymologyofCarbonandNitrogenCycling
Robert L. Tate III
9.EnzymeandMicrobialDynamicsofLitterDecomposition
Robert L. Sinsabaugh, Margaret M. Carreiro, and Sergio Alvarez
10.FungalCommunities,Succession,Enzymes,andDecomposition
Annelise H. Kjøller and Sten Struwe
11. Enzyme Adsorption on Soil Mineral Surfaces and Consequences for the

CatalyticActivity
Herve
´
Quiquampoix, Sylvie Servagent-Noinville,
and Marie-He
´
le
`
ne Baron
12.MicrobesandEnzymesinBiofilms
Jana Jass, Sara K. Roberts, and Hilary M. Lappin-Scott
13. Search for and Discovery of Microbial Enzymes from Thermally
ExtremeEnvironmentsintheOcean
Jody W. Deming and John A. Baross
14. Molecular Methods for Assessing and Manipulating the Diversity of
MicrobialPopulationsandProcesses
Søren J. Sørensen, Julia R. de Lipthay, Anne Kirstine Mu
¨
ller,
Tamar Barkay, Lars H. Hansen, and Lasse Dam Rasmussen
15. Bioindicators and Sensors of Soil Health and the Application of
Geostatistics
Ken Killham and William J. Staddon
16.HydrolyticEnzymeActivitiestoAssessSoilDegradationandRecovery
Tom W. Speir and Des J. Ross
17.EnzymaticResponsestoPollutioninSedimentsandAquaticSystems
Sabine Kuhbier, Hans-Joachim Lorch, and Johannes C. G. Ottow
18.MicrobialDehalogenationReactionsinMicroorganisms
Lee A. Beaudette, William J. Staddon, Michael B. Cassidy, Marc
Habash, Hung Lee, and Jack T. Trevors

19. Isolated Enzymes for the Transformation and Detoxification of Organic
Pollutants
Liliana Gianfreda and Jean-Marc Bollag
Copyright © 2002 Marcel Dekker, Inc.
20. Enzyme-Mediated Transformations of Heavy Metals/Metalloids:
ApplicationsinBioremediation
Robert S. Dungan and William T. Frankenberger, Jr.
21.EnzymesinSoil:ResearchandDevelopmentsinMeasuringActivities
M. Ali Tabatabai and Warren A. Dick
Copyright © 2002 Marcel Dekker, Inc.
Contributors
Sergio Alvarez Department of Ecology, Universidad Auto
´
noma de Madrid, Madrid,
Spain
Carol Arnosti Department of Marine Sciences, University of North Carolina, Chapel
Hill, North Carolina
Mark J. Bailey Molecular Microbial Ecology Group, Centre for Ecology and Hydrol-
ogy, Oxford, England
Tamar Barkay Department of Biochemistry and Microbiology, Cook College, Rutgers
University, New Brunswick, New Jersey
Marie-He
´
le
`
ne Baron Laboratoire de Dynamique, Interactions et Re
´
activite
´
, Centre Na-

tional de la Recherche Scientifique, Universite
´
Paris VI, Thiais, France
John A. Baross School of Oceanography, University of Washington, Seattle, Wash-
ington
Lee A. Beaudette Department of Environmental Biology, University of Guelph,
Guelph, Ontario, Canada
Jean-Marc Bollag Laboratory of Soil Biochemistry, Center for Bioremediation and De-
toxification, The Pennsylvania State University, University Park, Pennsylvania
Margaret M. Carreiro Department of Biology, University of Louisville, Louisville,
Kentucky
Copyright © 2002 Marcel Dekker, Inc.
Michael B. Cassidy Department of Environmental Biology, University of Guelph,
Guelph, Ontario, Canada
Leonid Chernin Department of Plant Pathology and Microbiology, Faculty of Agricul-
ture, The Hebrew University of Jerusalem, Rehovot, Israel
Ilan Chet Department of Plant Pathology and Microbiology, Faculty of Agriculture,
The Hebrew University of Jerusalem, Rehovot, Israel
Ryszard Jan Chro
´
st Department of Microbial Ecology, University of Warsaw, War-
saw, Poland
Julia R. de Lipthay Department of Geochemistry, Geological Survey of Denmark and
Greenland, Copenhagen, Denmark
Jody W. Deming School of Oceanography, University of Washington, Seattle, Wash-
ington
Warren A. Dick School of Natural Resources, The Ohio State University, Wooster,
Ohio
Robert S. Dungan George E. Brown, Jr., Salinity Laboratory, USDA-ARS, Riverside,
California

William T. Frankenberger, Jr. Department of Environmental Sciences, University of
California–Riverside, Riverside, California
Jose
´
Manuel Garcı
´
a-Garrido Department of Soil Microbiology, Estacı
´
on Experimen-
tal del Zaidı
´
n, CSIC, Granada, Spain
Inmaculada Garcı
´
a-Romera Department of Soil Microbiology, Estacı
´
on Experimental
del Zaidı
´
n, CSIC, Granada, Spain
Liliana Gianfreda Department of Chemical and Agricultural Sciences, University of
Naples Federico II, Portici, Naples, Italy
Marc Habash Department of Environmental Biology, University of Guelph, Guelph,
Ontario, Canada
Lars H. Hansen Department of General Microbiology, University of Copenhagen, Co-
penhagen, Denmark
Gerhard F. Herndl Department of Biological Oceanography, Netherlands Institute of
Sea Research (NIOZ), Den Burg, The Netherlands
Hans-Georg Hoppe Section of Marine Ecology, Institute of Marine Science, Kiel, Ger-
many

Copyright © 2002 Marcel Dekker, Inc.
Jana Jass Department of Molecular Biology, Umea
˚
University, Umea
˚
, Sweden
Shung-Chang Jong Department of Microbiology, American Type Culture Collection,
Manassas, Virginia
Ellen Kandeler Institute of Soil Science, University of Hohenheim, Stuttgart, Germany
Ken Killham Department of Plant and Soil Science, University of Aberdeen, Aberdeen,
Scotland
Annelise H. Kjøller Department of General Microbiology, University of Copenhagen,
Copenhagen, Denmark
Sabine Kuhbier Institute of Applied Microbiology, Justus Liebig University, Giessen,
Germany
Hilary M. Lappin-Scott Department of Biological Sciences, Exeter University, Exeter,
England
Hung Lee Department of Environmental Biology, University of Guelph, Guelph, On-
tario, Canada
Hans-Joachim Lorch Institute of Applied Microbiology, Justus Liebig University,
Giessen, Germany
James M. Lynch School of Biological Sciences, University of Surrey, Guildford, Sur-
rey, England
Anne Kirstine Mu
¨
ller Department of General Microbiology, University of Copenha-
gen, Copenhagen, Denmark
Paolo Nannipieri Scienza del Suolo e Nutrizione Della Planta, Universita
´
degli Studi

de Firenze, Firenze, Italy
David C. Naseby Department of Biosciences, University of Hertfordshire, Hatfield,
Hertfordshire, England
Juan Antonio Ocampo Department of Soil Microbiology, Estacı
´
on Experimental del
Zaidı
´
n, CSIC, Granada, Spain
Johannes C. G. Ottow Institute of Applied Microbiology, Justus Liebig University,
Giessen, Germany
Herve
´
Quiquampoix Laboratoire Sol et Environnement, Institut National de la
Recherche Agronomique, Montpellier, France
Lasse Dam Rasmussen Department of General Microbiology, University of Copenha-
gen, Copenhagen, Denmark
Copyright © 2002 Marcel Dekker, Inc.
Sara K. Roberts Department of Periodontics, College of Dentistry, University of Illi-
nois–Chicago, Chicago, Illinois
Des J. Ross Landcare Research, Palmerston North, New Zealand
Pacifico Ruggiero Dipartimento di Biologie e Chemical Agro-Forestale et Anibieritale,
Universita
´
degli Studi di Bari, Bari, Italy
Sylvie Servagent-Noinville Laboratoire de Dynamique Interactions et Re
´
activite
´
, Cen-

tre National de la Recherche Scientifique, Universite
´
Paris VI, Thiais, France
Robert L. Sinsabaugh Department of Environmental Science, University of Toledo,
Toledo, Ohio
Waldemar Siuda Department of Microbial Ecology, University of Warsaw, Warsaw,
Poland
Søren J. Sørensen Department of General Microbiology, University of Copenhagen,
Copenhagen, Denmark
Tom W. Speir Institute of Environmental Science and Research, Porirua, New Zealand
William J. Staddon Department of Biological Sciences, Eastern Kentucky University,
Richmond, Kentucky
Sten Struwe Department of General Microbiology, University of Copenhagen, Copen-
hagen, Denmark
M. Ali Tabatabai Department of Agronomy, Iowa State University, Ames, Iowa
Robert L. Tate III Department of Environmental Science, Rutgers University, New
Brunswick, New Jersey
Ian P. Thompson Molecular Microbial Ecology Group, Centre for Ecology and Hydrol-
ogy, Oxford, England
Jack T. Trevors Department of Environmental Biology, University of Guelph, Guelph,
Ontario, Canada
Copyright © 2002 Marcel Dekker, Inc.
1
Enzyme Activities and Microbiological
and Biochemical Processes in Soil
Paolo Nannipieri
Universita
´
degli Studi di Firenze, Firenze, Italy
Ellen Kandeler

University of Hohenheim, Stuttgart, Germany
Pacifico Ruggiero
Universita
´
degli Studi di Bari, Bari, Italy
I. INTRODUCTION
It is well known that soil organisms, particularly microbiota, play an essential role in the
cycling of elements and stabilization of soil structure (1,2). The mineralization of organic
matter is carried out by a large community of microorganisms and involves a wide range
of metabolic processes. For this reason, it is important to relate ecosystem structure and
function to species and functional diversity (3). However, the relationships between ge-
netic diversity and taxonomic diversity are not well understood and even less is known
about the manner in which these two properties affect microbial functional diversity (3–
5). Microbial species diversity is related to richness (i.e., the number of different species),
evenness (i.e., the relative contribution that individuals of all species make to the total
number of organisms present), and composition (i.e., the type and relative contribution
of particular species present) (3). According to the well-known and much used BIOLOG
approach, microbial functional diversity is related both to the rates of substrate utilization
and to the presence or absence of utilization of specific substrates. A decrease in microbial
diversity may reduce microbial functionality of soil if ‘‘keystone species,’’ such as nitrifi-
ers and nitrogen-fixing microorganisms, are negatively affected (6). However, this is not
generally the rule because rarely are there only a few species that perform a singular
function. Several processes, such as organic carbon mineralization, are carried out by a
large number of microbial species and a reduction in any group of species has little effect
on overall soil processes since other organisms can fulfill these functions (7–9). This spare
capacity or resilience is a feature of most soil ecosystems.
The molecular techniques used today have identified myriad microbial genes. How-
ever, the ecological importance of these genes is largely unknown because it is difficult
Copyright © 2002 Marcel Dekker, Inc.
to quantify their biochemical and microbial expression in situ (10). One significant ad-

vance using comparatively recent developments in molecular techniques (based on deoxy-
ribonucleic acid [DNA] extraction, purification, amplification, and analysis) has allowed
recording and monitoring the so-called nonculturable microorganisms (11). However, the
determination of microbiological activities requires detecting metabolic gene transcripts
(messenger ribonucleic acids [mRNAs]) in conjunction with modern sensitive assays of
metabolites and mRNA translation products, such as enzymes. These approaches are rap-
idly improving our understanding of the distribution and extent of microbe-mediated pro-
cesses in the field. It has been proposed that the monitoring of enzyme activities can be
used to determine the effect of genetically modified microorganisms on soil metabolism
(12,13).
Enzymes are proteins many of whose activities can be measured in soil. The assays
of soil enzymes are generally simple, accurate, sensitive, and relatively rapid. A range of
enzyme activities, and a large number of samples, can be analyzed over a period of a few
days using small quantities of soil. It is well known that changes in enzyme activities
depend not only on variations of gene expression but also on changes of environmental
factors affecting the considered activity (14,15). The expression of genes may occur in
natural samples, but numerous factors might effectively prevent the actual enzyme process
from taking place (Fig. 1). It has been hypothesized that the microbial composition of a
soil determines its potential for substrate catalysis since most of the processes occurring
in soil are microbe-mediated and are carried out by enzymes (14).
The objective of this chapter is to discuss the potential of soil enzyme assays to
determine soil microbial functional or process diversity and, when possible, identify future
research needs. The carbon substrate utilization approach is compared with enzyme mea-
surements for monitoring soil microbial functional diversity. Since soil microbial func-
tional diversity encompasses several metabolic activities, this approach requires assays of
many hydrolytic and oxidative enzymes. Therefore, the discussion includes the use of
composite indices or multivariate statistical analysis for integrating enzyme data sets for
comparing soil samples. Methods that distinguish between the contributions of extracellu-
lar and intracellular enzyme reactions are discussed.
Figure 1 Scheme showing the possible relations among taxonomic diversity, genetic diversity,

functional diversity, and enzyme activity. (From Ref. 14.)
Copyright © 2002 Marcel Dekker, Inc.
II. CARBON SUBSTRATE UTILIZATION PATTERNS AND
MICROBIAL FUNCTIONAL DIVERSITY
The BIOLOG system, originally developed to assist with the taxonomic description of
bacteria in axenic culture, is based on the ability of bacterial cells to oxidize up to 95
different carbon substrates in microtiter plates (16). The plates are incubated for a suitable
period of time (generally 72 h), and the oxidation of the substrate is monitored by measur-
ing the reduction of a tetrazolium dye. The rate and density of the color change depend
upon the number and activity of microbial cells in the well of the microtiter plate. The
BIOLOG system also has been applied to assess the functional metabolic diversity of
microbial communities from different habitats (17) and soil types (18–20), including the
rhizosphere (21,22) and grassland soils (23). Microbial communities produced habitat-
specific and reproducible patterns of carbon source oxidation, and thus this technique was
suggested to be sensitive for detecting temporal and spatial differences among soil micro-
bial communities (24). The carbon substrate utilization profiles have been shown to be
sensitive to heavy metal pollution (25), organic pollutants (26), soil types (26,27), crop
type, and crop management (28). The last mentioned study showed that, of the expectants
tested, 0.01 M NaCl yielded the highest well color development (28).
The BIOLOG approach presents several advantages. First, there is no doubt that the
utilization of carbon is a key factor in governing microbial growth in soil, and, for this
reason, this technique has been considered ecologically important (Table 1). In addition,
the technique is very rapid and simple. Thus it has become a very popular method of
assessing soil metabolic diversity and therefore soil functionality. In 1997 an international
meeting was devoted almost completely to the subject (29).
However, there are several drawbacks that limit the accuracy of the method in as-
sessing the metabolic diversity of soil microbiota (Table 1). First, the inoculum is a mixture
of organisms extracted from soil rather than a single species from axenic culture. Thus,
only the culturable minority of microorganisms is capable of oxidizing the organic sub-
strates (30). It is well established that only 1–10% of soil microflora are culturable by a

range of conventional techniques and media (31). Furthermore, reproducible results can
be obtained only if replicates present a similar inoculation density (24). Because BIOLOG
plates were developed for a different type of microbiological analysis, not all of the 95
organic substrates offered are ecologically relevant, and it could be important to choose
organic compounds that are appropriate for the microorganisms in their specific habitats.
For example, organic carbon compounds commonly present in root exudates were tested
to discriminate between the functional activities of microbial communities in rhizosphere
Table 1 Advantages and Disadvantages of the BIOLOG Technique
Advantages Disadvantages
Simple and rapid Fungi are not involved in the carbon substrate
Use of organic carbon, a key factor in govern- utilization profiles
ing microbial growth in soil Only culturable microorganisms provide infor-
Patterns of carbon source oxidation are repro- mation
ducible and habitat-specific Changes in the microbial community structure
may occur during incubation
Copyright © 2002 Marcel Dekker, Inc.
soil (32). Insam (33) reduced the number of substrates to 31 of those reported in soil to
allow three replicates of each substrate (plus a control) in a 96-well plate. Another problem
is that the multivariate statistics used (principal component analyses, discriminant analy-
ses, and detrended correspondence) to analyze BIOLOG data may mask the analytical
problems described. Changes in the microbial composition of the inoculum may occur
duringtheincubation(Table1).Indeed,notallconstituentsoftheinoculumcontribute
to the color development, and significant changes in the community structure occur over
a 72-h period (34). By comparing DNA melting profiles (denaturing gradient gel electro-
phoresis and temperature gradient gel electrophoresis) at the beginning and at the end of
the incubation, it was shown that the structure of the microbial community changed in
potato rhizosphere soil. In particular, it was observed that fast-growing bacteria become
dominant during the incubation period. However, the changes were not observed in an
activated sludge reactor amended with glucose or peptone (34). It was postulated that in
this case the dominant bacteria had been selected for rapid growth on readily utilizable

carbon sources because the activated sludge had been continuously fed with glucose and
peptone. In contrast, the dominant bacterial population of the potato rhizosphere was se-
lected in an environment characterized by a much lower content of organic substrates
than in activated sludge. Therefore, the dominant bacterial population (equivalent to K
strategists) of the rhizosphere inoculum was displaced by a more competitive microbial
population (equivalent to r strategists). Another significant limitation is that the community
profiles obtained with the use of the BIOLOG procedure do not include fungi because of
their comparatively slow growth rate (19).
Degens and Harris (35) have attempted to overcome the many problems of the
BIOLOG approach by measuring the patterns of in situ catabolic potential of microbial
communities. Differences in the individual short-term respiration responses (or substrate
induced respiration [SIR]) of soils to the addition to 36 simple substrates were used to
assess microbial communities. Despite the fact that this approach seems more accurate
than the BIOLOG technique, it has not been used widely since it was first published in
1997.
III. ENZYME ACTIVITIES AND SOIL MICROBIAL FUNCTIONAL
DIVERSITY
A. Limitations of a Single Enzyme Activity as an Index of
Microbial Activity
Enzyme activities can be measured and used as an index of microbiological functional
diversity if they reflect changes in microbial activities. Since microbial functional diversity
includes many different metabolic processes, theoretically a large number of different
enzymes should be measured. Data should be integrated in an attempt to calculate an
index of microbial functional diversity and to compare the microbial functional diversity
of different soil samples. Because this task is not possible, a representative set of enzyme
activities is needed. One approach might be to measure only the enzyme activities that
control the key metabolic pathways. Generally, these are rate-limiting exergonic steps and
are the targets of metabolic regulation. In the case of glycolysis (one of the main metabolic
pathways present in almost every microbial cell and transforming 1 mole of glucose to
2 moles of pyruvate), phosphofructokinase 1 is one of the 10 enzymes involved and cata-

lyzes the conversion of fructose–6-phosphate to fructose–1,6-bisphosphate. This regulates
Copyright © 2002 Marcel Dekker, Inc.
the rate of glycolysis (36). Therefore, determination of the phosphofructokinase-1 activity
should be an indication of the potential rate of glycolysis in soil.
The amount of soil N available to plants depends on many processes, but N mineral-
ization-immobilization turnover is considered to play the main role (37). The immobiliza-
tion process (the conversion of N-NH
4
ϩ
to organic N) includes several reactions catalyzed
by different enzymes. The first reaction converts N-NH
4
ϩ
to an amino acid and involves
the formation of glutamine from glutamate catalyzed by glutamine synthetase (GS). The
deamination of 2-oxoglutarate results in the formation of glutamate, which is catalyzed
by glutamate dehydrogenase (GDH) followed by the amination of aspartate with the for-
mation of asparagine by asparagine synthetase (38). The K
m
of GS is lower than that of
the other two enzymes, and at an ammonia ion concentration Ͻ 0.1 mM, ammonia is
incorporated into glutamine only, according to the reaction catalyzed by GS (39). There-
fore, the determination of GS activity can give useful information on the potential rate
of the N immobilization in soil.
It has been suggested (42) that β-glucosidase activity is a sensitive indicator for
assessing the effect of long-term burning and fertilization on the biological activity of
tallgrass prairie soil. According to Staddon and coworkers (43), acid phosphatase can be
used as an index for assessing the impact of fire on soils. Usually such hypotheses derive
from the fact that the measured enzyme activity was significantly correlated with some
general microbial parameter such as respiration or microbial biomass (15,44–46). In the

1950s and 1960s, invertase, protease, asparaginase, urease, phosphatase, and catalase ac-
tivities were assumed to be valid indicators of soil fertility (40). According to Skujins (40),
such suggestions produced conflicting and confusing data, not only because the adopted
methodologies were sometimes questionable but also because it is conceptually wrong to
use a single enzyme activity to determine plant productivity, microbiological activity, or
soil fertility. However, the use of a single enzyme activity, whether or not in a crucial
metabolic process, as an index for the microbiological activity or fertility of soil has been
criticized (40,41). Nannipieri (15) explains why reliance on a single assay is wrong.
1. Enzyme activities catalyze specific reactions and generally are substrate-
specific. Thus, they cannot be related to the overall microbiological activity of soil, which
includes a broad range of enzyme reactions. The synthesis of a particular enzyme can be
repressed by a specific compound while the overall microbiological activity of soil or crop
productivity is not affected. Chunderova and Zubers (47) showed that after 4 years of
cropping, high P concentrations depressed phosphatase activity. Microbial phosphatase is
a repressible enzyme, as shown by its decrease when microorganisms are transferred from
a deficient to a normal phosphate medium (48).
2. Measurements of microbial biomass have been used as indexes of microbial
activity in soil. Of course, biomass size and activity are different properties, but they often
are confused in soil studies (41). Nonetheless, microbial biomass can be correlated with
microbial processes, such as respiration rate, or with specific enzyme activities under
particular environmental conditions, but this does not mean that this relationship is valid
for every soil under every environmental condition.
3. As discussed later, the overall activity of any single enzyme in soil may depend
on enzymes in different locations including extracellular enzymes immobilized by soil
colloids. The activity of these immobilized enzymes may not be as sensitive to environ-
mental factors as are these directly associated with microbiological activity. For example,
enzymes in humic complexes are more resistant to thermal denaturation and proteolysis
than are the respective free enzymes (41).
Copyright © 2002 Marcel Dekker, Inc.
Thereisthepotentialtouseanintegratedmeasurementofanumberofenzyme

activities(asdiscussedlater),inconjunctionwithotherphysical,chemical,andmicrobio-
logicalmeasurements,inassessingsoilquality.Somehydrolaseactivitiesdonotshow
wideseasonalvariation,probablybecauseofthelargeamountofactivityassociatedwith
enzymesstabilizedbysoilcolloids.Thisprovidesagreatadvantageovermicrobiological
measurementssuchasrespiration,whichvarysowidelywithinaseasonoronayear-to-
yearbasisandmakeitdifficulttofindtrendsoridentifytheimpactsofdifferentmanage-
mentsystems.
Dehydrogenaseactivityisanintracellularprocessthatoccursineveryviablemicro-
bialcellandismeasuredtodetermineoverallmicrobiologicalactivityofsoil(40,41).The
problemwiththisisthattheelectronacceptors(2,3,5-triphenyltetrazoliumchloride[TTC]
(seealsop.19)or2-p-Iodophenyl-3-p-nitrophenyl-5-phenyltetrazoliumchloride[INT]
(seealsop.13))usedintheassaysarenotveryeffective,andthusthemeasurementsmay
underestimatethetruedehydrogenaseactivity(41).
Anotherpotentiallyconfusingaspectofthesestudiesarisesasaconsequenceofsoil
collectionandpretreatment.AccordingtoNannipieriandcoworkers(41),enzymeactivity
measurementsarecarriedoutafterremovalofvisibleanimalsandplantdebrisandon
sievedsoilsamplesunderlaboratoryconditions.Thusthemeasuredactivitiesofthese
samplesmaydependonthemetabolicprocessesorenzymeactivitiesassociatedonlywith
themicrobialcells.Conversely,whenratesofmetabolicprocesses,suchasrespiration,
aremeasuredinthefield,thecontributionsoflivingrootsandanimalsaswellasmacro-
scopicorganicdebrisarerecorded.Inotherwords,totalbiologicalratherthanmicrobiolog-
icalactivitiesaremeasured(41).
B.EnzymeActivities:MethodologyandInterpretation
AccordingtothereviewbySkujins(40),Woodsin1899suggestedthatextracellular
enzymescouldbepresentandactiveinsoil.Thefirstmeasurementsofenzymeactivities
insoilweredoneoncatalaseandperoxidaseactivitiesfrom1905to1910(40).Since
then,theactivityofdozensofenzymeshasbeendetectedinsoil.Obviouslythenumber
ofenzymesisconsiderablygreaterthanthosemeasuredbecauseofthemultitudeofmicro-
bial,faunal,andplantspeciesinhabitingsoil(46).Inaddition,theactivitymeasuredby
manyassayscannotbeascribedtotheactionofasingleenzyme.Thusdehydrogenase

activityisdeterminedbythemultipleenzymereductionofasyntheticsubstrate(TTCor
INT)duetoanoxidationofgenerallyunknownendogenoussubstrateswhoseconcentra-
tionisalsounknown(46).Casein-hydrolyzingactivitiesaremeasuredwithoutspecific
identificationofthebondhydrolyzedorofallproductsformed.Itisimportanttoempha-
sizethatevenwhenallthecomponentsofthereactionareknown,forexample,inthe
caseofureaseassays,differentenzymesfromdifferentsources(microbial,plant,oranimal
cells)catalyzethesamereaction.
Tables2and3(99–105)reportarangeofactivitiesofenzymescommonlyinvesti-
gated in soil. The ranges are generally very broad, possibly as a result of differences in
methods and soil types. In the late 1990s, several authors suggested standardized proce-
dures (106–108) based on conventional enzymology protocols. Tscherko and Kandeler
(109) proposed a classification for microbiological variables based on the activity classes
of the 30th and 70th percentiles. Using 30th and 70th percentiles gave similar widths of
the three categories, which were attributed to low, normal, and high activities. Different
sites could be classified according to the land use.
Copyright © 2002 Marcel Dekker, Inc.
Table 2 Some Enzyme Activities Involved in Organic C and N Transformations with Their
Range of Activities in Soil
Enzyme Range Reference
Enzymes involved in C trans-
formations
Xylanase 1.33–3125.00 µmol glucose (14, 49, 50, 51, 52, 53, 54,
g
Ϫ1
24 h
Ϫ1
55)
Cellulase 0.4–80.0 µmol glucose g
Ϫ1
(14, 56, 57)

24 h
Ϫ1
Invertase 0.61–130 µmol glucose g
Ϫ1
h
Ϫ1
(52, 53, 58, 59)
β-Glucosidase 0.09–405.00 µmol p-nitrophe- (42, 56, 57, 60, 61, 62, 63,
nol g
Ϫ1
h
Ϫ1
64, 65, 66)
β-Galactosidase 0.06–50.36 µmol p-nitrophenol (56, 57, 61)
g
Ϫ1
h
Ϫ1
Enzymes involved in N trans-
formations
Protease (casein-hydrolyzing 0.5–2.7 µmol. p-tyrosine g
Ϫ1
(14, 50, 51, 67, 68, 69, 70)
proteases) h
Ϫ1
Dipeptidase 0.08–1.73 µmol leucine g
Ϫ1
(60)
h
Ϫ1

Arginine deaminase 0.07–0.86 µmol N-NH
3
g
Ϫ1
h
Ϫ1
(14, 42, 69, 71, 72, 73, 74)
l-Asparaginase 0.31–4.07 µmol N-NH
3
g
Ϫ1
h
Ϫ1
(60, 65, 75, 76)
Amidase 0.24–12.28 µmol N-NH
3
g
Ϫ1
(60, 65, 75)
h
Ϫ1
l-Glutaminase 1.36–2.64 µmol N-NH
3
g
Ϫ1
h
Ϫ1
(75, 77)
Urease 0.14–14.29 µmol N-NH
3

g
Ϫ1
(42, 69, 78, 79, 80, 81, 82,
h
Ϫ1
83, 84)
Nitrate reductase 1.86–3.36 µgNg
Ϫ1
h
Ϫ1
(14)
Many enzyme activities have been detected in soil, but a reliable assay either has
not been developed or has been developed, but long after the initial report. For example,
hydrolysis of laminarin and inulin occurs in soil (110–112), but there is no specific assay
protocol. l-glutaminase, which catalyzes the hydrolysis of l-glutamine, yielding l-glu-
tamic acid and NH
3
, was first detected in soil by Galstyan and Saakyan (113), but a sim-
ple and rapid method was developed much later by Frankenberger and Tabatabai (77).
l-Asparaginase activity, which catalyzes the hydrolysis of l-asparagine producing l-
aspartic acid and NH
3
, was detected in soil by Drobni’k (114), but the simple and sensitive
method was developed much later (76). In fact, Tabatabai and coworkers have been re-
sponsible for the development of many assays for enzyme activities in soil (91,115,116).
The enzyme assay has to be simple and rapid, but above all sensitive and accurate
(117,118). This requires an efficient extraction and then an accurate determination of the
substrate or the reaction products from soil. Since most of the procedures for determining
either product formation or substrate disappearance are based on colorimetric reactions,
it is preferable to use buffers, which, in general, do not extract organic matter from soil.

Appropriate substrate concentration, pH, and temperature and optimal pH have to be found
for the assay of any soil enzyme (119). At a substrate concentration exceeding the value
limiting the reaction rate, the period of time selected should assure a linear substrate
disappearance or product formation and should be the shortest one (only a few hours at
Copyright © 2002 Marcel Dekker, Inc.
Table 3 Range of Some Hydrolase and Oxidase Activities in Soil
Enzyme Range Sources
Enzymes involved in
organic S transfor-
mations
Arylsulfatase 0.01–42.50 µmol p-nitrophenol g
Ϫ1
h
Ϫ1
(14, 42, 43, 63, 69, 85,
86, 87, 88, 89, 90, 91)
Enzymes involved in
organic P transfor-
mations
Alkaline phosphatase 6.76–27.34 µmol p-nitrophenol g
Ϫ1
h
Ϫ1
(14, 42, 43, 51, 86, 92,
93, 94, 95)
Phosphatase at pH 6.5 6.76–27.34 µmol p-nitrophenol g
Ϫ1
h
Ϫ1
(96, 97)

Acid phosphatase 0.05–86.33 µmol p-nitrophenol g
Ϫ1
h
Ϫ1
(42, 43, 69, 86, 94, 95,
98, 99, 100, 101)
Phospholipase C 5.02–8.15 µg p-nitrophenol g
Ϫ1
h
Ϫ1
(14)
Other enzyme activities
Dehydrogenase 0.002–1.073 µmol TPF g
Ϫ1
24 h
Ϫ1a
(14, 42, 50, 61, 69, 92,
93, 102, 103, 104,
105)
0.003–0.051 µmol INF g
Ϫ1
24 h
Ϫ1a
(102, 105)
Fluorescein diacetate 0.12–0.52 µmol fluorescein g
Ϫ1
h
Ϫ1
(60)
hydrolysis

Catalase 61.2–73.9 µmoles O
2
g
Ϫ1
24 h
Ϫ1
(63)
a
TPF, triphenyl formazan; INF, iodonitrotetrazolium formazan.
most) to produce a measurable value of activity. Complications due to microbial growth,
intracellular catalysis, and new enzyme synthesis can occur in assays with long incubation
times. The buffer solution must maintain the pH to that required for optimal activity
throughout the incubation period (120).
It is important interpreting of enzyme activities to understand that potential rather
than in situ activity is often being determined. This is because the incubation conditions
are chosen to ensure a rapid rate of substrate catalysis (41,120). In addition, enzyme assays
employ soil slurries to reduce diffusional limitations. These assay conditions are very
different from those occurring in soil, where moisture and temperature fluctuate widely,
the substrate concentration is generally not in excess, and pH is rarely optimal. When
possible, it is worthwhile to compare potential enzyme activities measured by the enzyme
assays involving soil slurries with hydrolysis of natural substrates added to soil samples
and incubated under the realistic in situ conditions. In the case of urea hydrolysis, it is
easy to compare urease activity determined by the conventional enzyme assay with the
rate of hydrolysis of solid urea added to soil and incubated in a range of relevant tempera-
tures and in the absence of buffer. This might be useful for predicting urea fertilizer hydro-
lysis under field conditions for specific soil types.
It has been discussed in several reviews that the foremost problem in interpreting
measurements of enzyme activities is to distinguish among many components contributing
to the overall activity (15,41,121,122). The activity of any particular enzyme in soil de-
pends on enzymes that can have different locations: (1) active and present intracellularly

in living cells, (2) in resting or dead cells, (3) in cell debris, (4) extracellularly free in the
Copyright © 2002 Marcel Dekker, Inc.
Figure 2 Various activities contributing to the overall enzyme activity measured in soil with those
affecting the microbial functional diversity in soil.
soil solution, (5) adsorbed by inorganic colloids, or (6) associated in various ways with
humic molecules. In addition, abiotic transformations, the so-called enzyme-like reactions,
can contribute to the overall activity (Fig. 2). Intracellular enzymes are present in plant,
animal, and microbial cells; however, since visible animals and vegetable remains are
largely removed prior to an assay and those that have been released from lysed cells are
rapidly degraded by microorganisms, it is reasonable to suppose that the most important
intracellular enzymes of soil are those in living microbial cells. Thus, the determination
of intracellular enzyme activity can give important information about the processes medi-
ated by the current microbial inhabitants. By determining the intracellular enzyme activi-
ties of soil samples, it is possible to have information on the microbial functional diversity
of soil. On the other hand, of the three extracellular locations (free enzymes, enzymes
adsorbed by inorganic complexes, or those associated with organic colloids) the first are
supposed to be short-lived (15,46,121), whereas the other two are characterized by a
marked resistance to thermal and proteolytic degradation (123). With the present methods
it is difficult if not impossible to determine the different locations of the enzyme activities.
C. Soil Minerals as Catalysts (Pseudoenzymes) in
Biochemical Reactions
Soil minerals can affect the fate of biochemical compounds in soil in at least three main
ways: (1) incorporation of N-, P-, and S-bearing organics into the structural network of
mineral colloids and adsorption of these organics to their surface: consequently the dynam-
Copyright © 2002 Marcel Dekker, Inc.
ics and bioavailability of these nutrients may be modified (124); (2) adsorption of enzymes
on clay and/or clay-organic complexes: these immobilized enzymes are active and stable,
but they exhibit activities quite different from those of free enzymes (123,125); and (3)
abiotic transformation of natural organic components: this means that the mineral compo-
nent should be considered not only as a support for adsorption and binding of organics

(enzymes and substrates), but also as a reactive surface for many transformation processes.
Clays demonstrate the ability to catalyze electron transfer reactions. In the same
way, metal oxides/hydroxides, such as Fe
3ϩ
and Mn
3ϩ,4ϩ
oxides, quite common in soils,
are able to catalyze oxidation reaction of organic compounds. Birnessite (δ-MnO
2
), in
particular, is considered to be an ‘‘electron pump’’ for a wide range of redox reactions
(126). In summary, clay minerals and metal oxides and hydroxides are reactive in promot-
ing abiotic degradation of natural organics.
A large part of the research into the catalytic role of clay minerals has been devoted
to studying the oxidative transformation of phenols and polyphenols and other natural
organic components and the subsequent formation of humic substances (127,128). It is
noteworthy that phenolic acids are degradation products of lignin and constitute an impor-
tant fraction of the exudates released by plants under stress conditions (129).
Shindo and Huang (130) found that montmorillonite, vermiculite, illite, and kaolin-
ite accelerated the synthesis of humic substances from hydroquinone, as the precursor,
whereas non-tronite- (Fe
3ϩ
–bearing smectite), in the presence of O
2
, showed a synergistic
effect that greatly enhanced the polymerization of hydroquinone (131,132). Similar results
were obtained when pyrogallol was used as the precursor (133). In systems containing
phenolic compounds and amino acids, Ca-illite catalyzed the formation of N-containing
humic acids (134). The yields and nitrogen contents depended on the kind of amino acids
used. In 1997 Bosetto and colleagues (135) studied the formation of humiclike polymers

from l-tyrosine and homoionic clays and showed that the amount produced depended on
the type of interlayer cation.
Besides the clay minerals, Mn
2ϩ/4ϩ
and Fe
3ϩ
oxides oxidize phenolic compounds
rapidly. Their relative effectiveness in the synthesis of humic substances has been studied
extensively (127,128,136). Hydroquinone, resorcinol, and catechol were used as sub-
strates. The catalytic effect of birnessite was higher than that of Fe oxide; however, the
relatively high content of the Fe oxides in soils suggests that their role in the abiotic
formation of humic substances should not be overlooked. The synthesis of humic sub-
stances was obtained also by using aluminas as catalysts (137). McBride and associates
(138) proposed an oxidation mechanism by which soluble Al tended to stabilize o-semiqui-
none radicals at low pH, directing subsequent radical polymerization. Pyrogallol-derived
polymer formation was strongly promoted by birnessite (139) and the cross-polarization,
magic angle spinning-nuclear magnetic resonance
13
C-nuclear magnetic resonance
(CPMAS-
13
C NMR) spectrum of humic acids formed resembled those of humic acids
extracted from natural soils. Birnessite also was able to cleave the ring structure of pyrogal-
lol, releasing CO
2
. The abiotic ring cleavage of polyphenols might, in part, form aliphatic
fragments contributing to the aliphatic nature of humic substances in the environment.
The amount of CO
2
released from the ring cleavage of pyrogallol and the quantity of

humic polymers formed were directly related to the catalytic activity of clay minerals
such as nontronite, bentonite, and kaolinite saturated which Ca
2ϩ
(139).
Further research has compared the activity of biotic and abiotic catalysts in the
transformation of phenolic substrates. Mn oxide influenced the darkening of hydroquinone
and resorcinol to a larger extent than did the enzyme tyrosinase, whereas the reverse was
Copyright © 2002 Marcel Dekker, Inc.
true for catechol (140). The yields of humic acids were influenced significantly by the
kind of catalyst and the type of diphenols. The comparison between biotic and abiotic
catalysts has also been conducted by examining the reaction products that resulted from
the transformation reactions. Birnessite, bentonite, tyrosinase, horseradish peroxidase, and
the laccases of Trametes versicolor and Rhizoctonia praticola were used separately in the
oxidation of guaiacol (141). All reaction products were analyzed for the presence of guaia-
col and guaiacol-derived oligomers. The same seven products (five dimers and two tri-
mers) were produced by using the various oxidizing agents, whereas the amounts of the
respective oligomers varied. The differences in the amounts obtained were likely due to
different reaction conditions.
The rate of 2,6-dimetoxyphenol transformation was considerably higher in reactions
catalyzed by tyrosinase or laccase than in those by birnessite (141). At the same time,
significant differences in oxygen consumption were observed. When continuous additions
of catechol as substrate were employed, it was shown that tyrosinase and laccase trans-
formed catechol after each addition, whereas birnessite was active only in the first incuba-
tion with catechol.
The reaction products of catechol oxidation by tyrosinase had a higher degree of
aromatic ring condensation relative to those of the birnessite-catechol system (142). In
addition, the products of birnessite catalysis contained various fragments, including ali-
phatic components, with lower molecular weights than those produced by tyrosinase. In
contrast, Birkel and Niemeyer (143) obtained similar reaction products when comparing
enzymatic reaction products of humic precursors with products obtained at clay mineral

surfaces (montmorillonite) through abiotic reactions.
Phenolic acids with higher methoxy substituents were oxidized more rapidly in the
presence of Mn and Fe oxides (144,145). By enzymatic polymerization of syringic and
vanillic acids, soluble oligomers (from dimers to hexamers) were found as oxidation prod-
ucts (146,147), whereas tests with ferulic acid, incubated with MnO
2
, showed that the
soluble products obtained did not contain any oligomers because they were rapidly sorbed
to MnO
2
surfaces (144). In 1998 Hames and coworkers (148) reported the first efficient
modification/degradation of in situ lignin by MnO
2
and oxalic acid, either produced by
fungi or resulting from oxidative degradation of cell wall components. The MnO
2
/oxalate
system appeared to oxidize the lignin macromolecule selectively and to play an important
role in the transformation of the lignin polymer into humus and/or its precursors.
The results of these studies indicate that the catalytic effects of soil minerals and
enzymes on the oxidation of phenols need more investigation. The similarities and differ-
ences when comparing both activities might be due to the reaction conditions (pH,
substrate/catalyst ratio, rate of oxidation) and/or to the reaction mechanisms. It is not
clear, for instance, whether the metal oxides meet the definition of a catalyst. A true
catalyst cannot be altered by the reaction it catalyzes. For both abiotic and biotic agents,
ions such as Cu
2ϩ
,Mn
4ϩ
, and Fe

3ϩ
act as electron acceptors. However, whereas in the
protein complex (enzyme) an electron is transferred from the reduced metal to oxygen in
a cyclic manner, in metal oxides, in particular birnessite (MnO
2
), the metal remains in
the reduced form (Mn
2ϩ
) even for long periods of incubation and Mn
2ϩ
recovers to its
initial state fairly slowly in the presence of O
2
. The result is that the enzymes behave as
a catalyst and birnessite as an oxidizing agent.
Phenols, together with amino acids, have been used to synthesize nitrogenous poly-
mers resembling humic acids in the presence of both polyphenol oxidases and birnessite
and nontronite (149–151). It was assumed that ring cleavage of phenol and release of
Copyright © 2002 Marcel Dekker, Inc.
CO
2
and NH
3
occurred and were followed by polycondensation of semiquinone radicals,
aliphatic fragments, and amino acids to form humic polymers. Similar mechanisms have
been proposed in systems involving phenoloxidase enzymes.
The deamination of amino acids, such as serine, phenylalanine, proline, methionine,
and cysteine by birnessite, and the role of pyrogallol in influencing their mineralization
have been investigated (152,153). Nitrogen mineralization was inhibited by pyrogallol,
whereas S mineralization of S-containing amino acids was not, except in the iron oxide–

methionine system. Deamination and decarboxylation of amino acids as catalyzed by soil
minerals may constitute a pathway of C turnover and N transformation in nature.
Some natural organic compounds, other than polyphenolics, also have been shown
to be oxidized by metal oxides. The reactions of malic acid, an important constituent of
root exudates, with the hydrous oxides of Mn and Fe were studied (154). The reactions
followed two pathways, depending on the pH-controlled adsorption of oxalacetic acid (the
first product of the malate oxidation) on the oxide surfaces. The production of NH
4
ϩ
in the
glutamic acid–treated birnessite suspensions was attributed to direct chemical oxidative
deamination of glutamic acid by the manganese oxide (155).
In the presence of Na-montmorillonite, the isocitric acid was oxidized and trans-
formed into α-ketoglutaric acid (156). Isocitrate oxidative decarboxylation comprised sev-
eral steps but always started with a protonation reaction. A certain analogy exists between
enzymatic and clay mineral catalysis, provided that in both cases transformation began
with a protonation of a chemical function. For enzymatic catalysis, it was principally
the coenzyme (NADH ϩ H
ϩ
) of the isocitrate dehydrogenase that supplied the proton.
Nevertheless, the reaction rate in the presence of clay was very much lower than in the
presence of enzyme.
Glutamic acid was selectively deaminated by a combination of pyridoxal phosphate
(PLP), a cofactor in enzyme systems important in amino acid metabolism, and Cu
2ϩ
-
smectite with production of ammonia and α-ketoglutaric acid (157). The system exhibited
specificity for glutamic acid in comparison to some other amino acids. One possible expla-
nation was that glutamic acid reacted with PLP to form a Schiff base, which then com-
plexed with Cu

2ϩ
on the mineral surface. This was followed by hydrolysis to ammonia,
α-ketoglutaric acid, and regenerated PLP. The catalyst system was not stereoselective
because it was equally effective for both L- and D- glutamic acid. The PLP-Cu
2ϩ
-smectite
has acted as a ‘‘pseudoenzyme’’ wherein the PLP was active and independent of the
protein matrix of the enzyme and the silicate structure substituted for the apoenzyme
(157).
The deamination of glutamic acid also occurred in the presence of montmorillonite
saturated with various cations and in the absence of any cofactor. The reaction prod-
ucts were α-hydroxyglutaric acid and traces of butyric acid, whereas in the presence of
the enzyme glutamate dehydrogenase and oxidized nicotinamide-adenine dinucleotide
(NAD
ϩ
), the glutamic acid yielded α-ketoglutaric acid (158). The catalytic activity of the
montmorillonite surface depended on pH and remained lower than that of the enzyme.
Nevertheless, the montmorillonite had an advantage over the enzyme as it displayed a
larger activity pH range and a nonspecific activity. The abiotic catalytic role of montmoril-
lonite also was demonstrated in deamination of aspartic acid by aspartase-Ca-montmoril-
lonite systems (159). Deamination of l- and d-glutamic and aspartic amino acids and of
their DL racemic mixtures in the presence of Na-montmorillonite showed a stereoselectiv-
ity of the clay mineral for the L-isomer and implicitly a structural chirality character of
the clay mineral (160).
Copyright © 2002 Marcel Dekker, Inc.