TM
Marcel Dekker, Inc. New York
•
Basel
edited by
Richard G. Burns
University of Kent
Canterbury, Kent
England
Richard P. Dick
Oregon State University
Corvallis, Oregon
Copyright © 2002 by Marcel Dekker, Inc. All Rights Reserved.
Copyright © 2002 Marcel Dekker, Inc.
ISBN: 0-8247-0614-5
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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

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

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

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

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

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

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,

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

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

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

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

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.

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

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

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

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

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