Enzymes in the Environment: Activity, Ecology and Applications - Chapter 2 pptx

2
Ecology of Microbial Enzymes in
Lake Ecosystems
Ryszard Jan Chro
´
st and Waldemar Siuda
University of Warsaw, Warsaw, Poland
I. INTRODUCTION
During the past decade, an increasing number of ecological studies have considered the
complexity of freshwater ecosystems. One major outcome of these studies has been an
accelerated interest in the role of heterotrophic microorganisms (particularly bacteria) in
the functioning of aquatic environments and the processes by which organic matter is
made available to them (1–4). These heterotrophic microorganisms are the key trophic
level at which the metabolism of the whole ecosystem is affected, i.e., organic matter
decomposition, nutrient cycling, and structure of aquatic food webs. The demonstration
of the importance of heterotrophic bacteria as a particulate carbon source for higher trophic
levels and a major respiratory sink has created a renewed interest in the production and
utilization of organic substrates by these microorganisms.
Most organic compounds produced in natural waters have a polymeric structure
(5,6) and they are too large to be readily assimilated. The transport of organic molecules
across microbial cell membranes is an active process mediated by specific enzymes called
permeases. Only the low-molecular-weight organic molecules (monomers or oligomers)
can therefore be taken up (7). In order to be available for microbial metabolism, polymeric
compounds must be transformed into smaller molecules through enzymatic depolymeriza-
tion.
Besides the physicochemical conditions of aquatic environments, the composition
and availability of organic matter are the major factors that influence the development and
activity of heterotrophic bacteria (8,9). The heterotrophic bacteria are the only biological
populations capable of significantly altering both dissolved (DOM) and particulate (POM)
organic matter. Microbial enzymes associated with these processes are the principal cata-
lysts for a large number of biochemical transformations of organic constituents in aquatic

environments. Many of these transformations can be mediated only by heterotrophic bac-
teria because the enzyme systems required for these reactions are not found in other or-
ganisms.
Microorganisms have adopted essentially two strategies that enable them to utilize
macromolecular compounds. The macromolecule can be engulfed by the cytoplasmic
Copyright © 2002 Marcel Dekker, Inc.
membrane to form a vacuole within the cytoplasm. Enzymes are secreted into this vacuole
and the polymeric compounds are hydrolyzed and subsequently taken up. Uptake of sub-
strate solutions by this method is referred to as pinocytosis; uptake of particulate substrates
is termed phagocytosis. However, many microbial cells are unable to carry out these pro-
cesses, and therefore pinocytosis and phagocytosis are restricted to only those eukaryotic
microorganisms that lack a cell wall, e.g., many protozoa. Those eukaryotic and prokary-
otic microorganisms that possess a cell wall have developed an alternative strategy for
the assimilation of polymeric substrates. Hydrolytic enzymes are secreted outside the cyto-
plasmic membrane, where they hydrolyze macromolecules in close vicinity to the cell.
The resulting low-molecular-weight products are then transported across the cell mem-
brane and utilized inside the cytoplasm.
The hydrolysis of polymers is an acknowledged rate-limiting step in the utilization
of organic matter by microorganisms in aquatic and soil environments. Prior to incorpora-
tion into microbial cells, polymeric materials undergo stepwise degradation by a variety
of cell surface–associated enzymes and/or enzymes secreted by intact living cells or liber-
ated into the environment through the lysis of microorganisms. The importance of micro-
bial enzymatic activities to the mobilization, transformation, and turnover of organic and
inorganic compounds in freshwater and marine environments has been shown in many
studies (10–18). Results of these studies have shown that studying enzymatic processes
provides powerful information that helps in understanding basic processes of decomposi-
tion and microbial activity in both freshwater and marine ecosystems.
II. ORIGIN AND ASSOCIATION OF ENZYMES WITH AQUATIC
MICROORGANISMS
Three common terms are used for the enzymes involved in the transformation and degrada-

tion of polymeric substrates outside the cell membrane: ectoenzymes (19), extracellular
enzymes (20), and exoenzymes (21). In this chapter, the term ectoenzyme is used to refer
to any enzyme that is secreted and actively crosses the cytoplasmic membrane and remains
associated with its producer. Ectoenzymes are cell-surface-bound or periplasmic enzymes
that react outside the cytoplasmic membrane with polymeric substrates that do not pene-
trate the cytoplasm. Extracellular enzymes occur in free form dissolved in the water and/
or are adsorbed to surfaces (e.g., detrital particles, organic colloids, humic complexes,
minerals in suspension). Extracellular enzymes in water may be secreted actively by intact
viable cells, they can be released into the environment after cell damage or viral lysis,
and/or they may result from zooplankton grazing on algal cells and from protozoan grazing
on bacteria.
Ectoenzymes and extracellular enzymes (in contrast to intracellular enzymes) react
outside the cell, and most of them are hydrolases. The ectoenzymes that cleave polymers
by splitting the key linkages on the interior of the substrate molecule and form intermediate
sized fragments are called endoectoenzymes (e.g., aminoendopeptidases act on the cen-
trally located peptide bonds and liberate peptides) (22). Those ectoenzymes that hydrolyze
the substrate by consecutive splitting of monomeric products from the end of the molecule
are termed exoectoenzymes (e.g., aminoexopeptidases hydrolyze peptide bonds adjacent
to terminal α-amino or α-carboxyl groups and liberate free amino acids) (23).
There are three pools of microbial enzymes in water samples: intracellular enzymes
are located and react with substrates inside the cytoplasmic region and are mostly responsi-
Copyright © 2002 Marcel Dekker, Inc.
Figure1Percentagedistributionofcell-boundandextracellularactivityofmicrobialchitinase
(CHTase),deoxyribonuclease(DNase),5′-nucleotidase(5′-nase),alkalinephosphatase(APase),β-
glucosidase(GLCase),andaminopeptidase(AMPase)inwatersamplesfromeutrophicLakeMi-
kołajskie.(Chro
´
st,unpublished.)
bleforinternalcellmetabolism;extracellularenzymesareinthesurroundingenvironment
andcatalyzereactionswithoutcontrolfromtheirproducers;andectoenzymesarecell-

surface-boundenzymes,mostlyhydrolases,thatdegradepolymericsubstrates,yielding
readilyutilizablemonomers.Allpoolsarecomposedofbothendo-andexoenzymes.
Distributionbetweenecto-andextracellularactivityforselectedenzymes(amino-
peptidase,β-glucosidase,alkalinephosphatase[APase],5′-nucleotidase[5′-nase],deoxyri-
bonuclease[DNase],andchitinase[CHTase])hasshownthatectoenzymescontributed
onaveragefrom75%(APase)to98%(chitinase)ofthetotalactivityinlakewater(Fig.
1).Activitiesoftheintracellularandextracellularpoolenzymesarelow.Intracellular
enzymescontributedfrom0.5%(chitinase)to10.7%(aminopeptidase)tothetotalactivity
ofwatersamples.Activityoftheextracellularenzymes,dissolvedinthewater,constituted
from1%(5′-nase)to16.5%(APase)ofthetotalactivity.Aninterestingobservationis
thatextracellularenzymeactivityasapercentageoftotalactivityishigherinlakesedi-
mentsthaninthewatercolumn(Table1).Thiswasparticularlyevidentinthecaseof
chitinase and lipase activities.
Enzyme activities bound to the 0.2- to 1.0-µm-size fraction of microplankton
(mainly composed of bacteria) make up a greater fraction of activity by microorganisms
in lake water. High ectoenzyme activity found in this size fraction has correlated with
Copyright © 2002 Marcel Dekker, Inc.
Table 1 Percentage Contribution of Extracellular Enzyme Activities to the Total Activity of
Lake Water and Lake Sediment Samples
Percentage
Enzyme Lake/trophic status Water Sediment
Leucine-aminopeptidase Plußsee/eutrophic 9.5 Ϯ 3.7 12.6 Ϯ 6.4
Scho
¨
hsee/mesotrophic 8.4 Ϯ 2.1 11.2 Ϯ 5.1
α-Glucosidase Plußsee/eutrophic 10.2 Ϯ 4.9 15.1 Ϯ 7.3
Mikołajskie/eutrophic 8.7 Ϯ 2.6 13.3 Ϯ 3.8
Szymon/hypereutrophic 10.8 Ϯ 3.3 14.6 Ϯ 4.9
Lipase Jagodne/hypereutrophic 28.8 Ϯ 9.6 41.0 Ϯ 7.4
Bełdany/eutrophic 21.4 Ϯ 6.8 33.1 Ϯ 8.3

Kisajno/mesotrophic 22.4 Ϯ 6.2 35.8 Ϯ 9.1
Alkaline phosphatase Plußsee/eutrophic 24.5 Ϯ 6.3 32.6 Ϯ 7.3
Chitinase Plußsee/eutrophic 2.3 Ϯ 0.7 22.8 Ϯ 6.9
Scho
¨
hsee/mesotrphic 1.7 Ϯ 0.5 23.2 Ϯ 8.4
Ϯ Standard deviation of an average value.
Source: Data from Chro
´
st, unpublished.
bacterial abundance and/or bacterial production of lake water. A variety of microorgan-
isms produce ectoenzymes in waters and sediments in freshwater ecosystems. However,
many studies have reported that bacteria are the major producers of ectoenzymes among
aquatic microorganisms (24–33).
III. CONTROL OF ECTOENZYME SYNTHESIS AND ACTIVITY
The conditions in the aquatic environment, as in the soil aqueous phase, are unfavorable
for enzymes. First, the substrate concentration is usually very low and highly variable.
Many substrates may be insoluble, exist in intimate association with other compounds,
and/or be bound to humic substances, colloidal organic matter, and detritus. Therefore,
these conditions are suboptimal for the coupling of an enzyme to its substrate. Second,
an enzyme may be lost from the parent cell and may be bound to suspended particles and
humic materials, or it may be exposed to a variety of inhibitors present in the water.
Finally, an enzyme may be denaturated by physical and chemical factors in the aquatic
environment or hydrolyzed by proteases.
Obviously, for an enzyme to be of benefit to its producer microorganism, it must
avoid degradation long enough to associate with its substrate. Moreover, even if an enzyme
overcomes these obstacles and binds with its substrate, the physical and chemical condi-
tions of the reaction medium may be unsuitable for catalysis (e.g., nonoptimal pH or tem-
perature, presence of inhibitors, absence of activators, suboptimal ionic strength). Never-
theless, there is strong evidence that various aquatic microorganisms produce ectoenzymes

in freshwaters that encounter a number of polymeric substrates (31) and that microbial
growth is dependent on the products of ectoenzymatic reactions (13,14,16,24, 34–36).
A microbial cell living in an aquatic ecosystem is influenced by a variety of environ-
mental factors. The signal for appropriate gene expression and consequent ectoenzyme
production within a cell is in response to the surrounding environment. Depending on the
Copyright © 2002 Marcel Dekker, Inc.
regulatory control of gene expression, two types of microbial enzymes are synthesized in
waters and sediments: constitutive enzymes, whose synthesis is constant regardless of the
presence or absence of the substrate in the environment, and inducible enzymes, whose
rates of synthesis are strongly dependent on the presence of their substrates (or substrate
derivatives). Many inducible enzymes are synthesized at a low basal rate (i.e., are constitu-
tive) in the absence of a substrate. When the substrate is available in the environment,
there is a dramatic increase in the production rate of the particular enzyme. Synthesis
continues at this amplified rate until the inducer is removed and/or the product of enzy-
matic catalysis accumulates (24) and it then returns to the basal rate.
Most of the ectoenzymes synthesized by aquatic microorganisms are catabolic en-
zymes involved in degradation of polymeric substrates that are not continuously available
in the water or sediments. Therefore, the constant synthesis of ectoenzymes in the absence
of substrates is unnecessary, because it requires the expenditure of energy that otherwise
may be channeled into other useful activities. Since microorganisms have been competing
with each other for millions of years, the evolutionary advantages of induction are readily
apparent. Most of the ectoenzymes found in freshwaters are inducible, and only a few
have a constitutive nature (e.g., some amylases or proteases in bacteria).
A. Induction and Repression/Derepression of
Ectoenzyme Synthesis
The efficient induction of ectoenzymes is more complicated than that of intracellular en-
zymes. First, many of the ectoenzyme substrates present in fresh water are polymeric
compounds, and they are too large to enter the cell and serve as inducers of synthesis.
Second, for an ectoenzyme to be secreted at appropriate rates, the microorganism must
be able to monitor the activity of the ectoenzyme outside the cell. We suggest that these

problems are overcome by a low constitutive rate of ectoenzyme secretion. If the substrate
is present, then low-molecular-weight products accumulate to a certain level, enter the
cell, and serve as the inducer (20). When environmental conditions inhibit an ectoenzyme
activity (e.g., unsuitable pH, absence of activating cations Mg
2ϩ
,Zn
2ϩ
), the induction of
its synthesis does not occur because the product of catalysis is not generated. However,
since the microorganisms in freshwater ecosystems are in a complex relationship with a
variety of readily utilizable compounds of autochthonous and allochthonous origin, the
induction of a particular ectoenzyme by an end product resulting only from degradation
of a single polymeric substrate seems to be questionable. Until now, it has appeared that
one ectoenzyme may have several inducing compounds (24,37).
It is well documented that synthesis of many ectoenzymes produced by aquatic
microorganisms is repressed by the end product that accumulates in the cell or in sur-
rounding environment. The repression of alkaline phosphatase synthesis by inorganic
phosphate (the end product of phosphomonoester hydrolysis) in microalgae and bacteria
is probably one of the best-known examples (11,13,38,39). In Lake Plußsee, the specific
activity of APase significantly decreased when the ambient orthophosphate concentrations
were higher than 0.5 µM (13). APase activity was inversely related to the amount of
intracellular phosphorus stored (P
st
) in algal cells. When P
st
constituted less than 10% of
the total cellular phosphorus, the algae produced alkaline phosphatase with a high specific
activity, and when P
st
was higher than 15% and the ambient orthophosphate concentrations

exceeded 0.6 µM, this activity rapidly decreased.
The synthesis of virtually all ectoenzymes in most aquatic microorganisms is re-
Copyright © 2002 Marcel Dekker, Inc.
Figure 2 Effect of water supplementation with dissolved organic matter extracted from phyto-
plankton on growth of bacteria (A) and specific activity of bacterial β-glucosidase (B) and aminopep-
tidase (C) in Lake Mikołajskiet. (Chro
´
st, unpublished.)
pressed when they are grown on sources of readily utilizable dissolved organic matter
(UDOM). This mode of regulation is called catabolic repression. When water samples
from eutrophic Lake Mikołajskie were supplemented with dissolved organic matter ex-
tracted from phytoplankton (both UDOM and polymeric compounds) the bacterial cell
numbers increased markedly during 96 hours of incubation (Fig. 2A). Contrary to that in
control samples, supplementation of lake water with phytoplankton organic matter resulted
in a significant decrease in the rates of specific activities of bacterial β-glucosidase and
aminopeptidase (calculated per bacterial cell) during the first period of bacterial growth
(6–48 hours). However, in both of these enzymes, specific ectoactivity began to increase
after 48 hours of bacterial growth. In control samples, where bacteria grew solely on
naturally present DOM in lake water, the specific activity of these ectoenzymes increased
within the incubation period (Figs. 2B, 2C).
The repression of ectoenzymes is tightly coupled to the availability of UDOM in
Copyright © 2002 Marcel Dekker, Inc.
lakewater.Figures2Band2CshowthatectoenzymesynthesisinDOM-enrichedsamples
wasnolongerrepressedwhentheconcentrationofthereadilyutilizablelowmolecular-
weightmoleculesfellbelowacriticallevel,andpolymericsubstrateshadtobeusedto
supportthegrowthandmetabolismofbacteria.Similarinsituobservationsduringphyto-
planktonbloomdevelopmentandbreakdownwerereportedforβ-glucosidaseactivityin
eutrophicLakePlußsee(24),forβ-glucosidaseandaminopeptidaseactivitiesinmeso-
trophicLakeScho
¨

hsee(25),andforlipaseactivityineutrophicLakeMikołajskie(40).
Despitethewidespreadoccurrenceofcatabolicrepression,withtheexceptionof
thoseforentericbacteria,themoleculardetailsoftherepressionarepoorlyunderstood.
Somestudieshaveindicatedthatcyclicadenosinemonophosphate(cAMP),togetherwith
itsreceptorprotein,mayplayacentralroleincontrolofcatabolicrepression(41,42).
Usingtherepressionstrategyforectoenzymesynthesis,microorganismscanavoidthe
wastefulproductionofinducibleenzymes,whicharenotusefulwhentheirgrowthisnot
limitedbyUDOM(3,19,24,35).
B.InhibitionofActivity
Itisimportanttoconsiderthattherepression/derepressionofanectoenzymenotbe
equatedtothereversibleinhibitionofactivity.Evenifanectoenzymeissynthesized,its
activitymaybeinhibitedbytheaccumulationoftheendproductorbyhighconcentrations
ofthesubstrate(19).Twogeneraltypesofreversibleinhibitionareknown:competitive
andnoncompetitiveinhibition.
Competitiveinhibitionoccurswhenaninhibitingcompoundisstructurallysimilar
tothenaturalsubstrateand,bymimicry,bindstotheenzyme.Indoingso,itcompetes
withanenzyme’snaturalsubstratefortheactivesubstrate-bindingsite.Thehallmarkof
competitiveinhibitionofmanyectoenzymes(e.g.,alkalinephosphatase,β-glucosidase,
aminopeptidase)isthatitdecreasestheaffinityofanectoenzyme(anincreaseofthe
apparentMichaelisconstantisobserved)forthesubstrateand,therefore,inhibitstheinitial
velocityofthereaction(Fig.3)(13,26,37).Competitiveinhibitionisreversibleandcan
beovercomebyincreasedsubstrateconcentration,andthereforethemaximumvelocity
(V
max
)ofthereactionisunchanged(Fig.3A).
Noncompetitive inhibition generally is characterized as an inhibition of enzymatic
activity by compounds that bear no structural relationship to the substrate. Therefore, the
inhibition cannot be reversed by increasing the concentration of the substrate. It may
be reversed only by removal of the inhibitor. Unlike competitive inhibitors, reversible
noncompetitive inhibitors cannot interact at the active site but bind to some other portion of

an enzyme-substrate complex. This type of inhibition encompasses a variety of inhibitory
mechanisms and is therefore not amenable to a simple description. Noncompetitive inhibi-
tion of the activity of exoproteases by Cu
2ϩ
ions (43) and inhibition of α-glucosidase, β-
glucosidase, N-acetyl-glucosaminidase, and alkaline phosphatase by H
2
S in natural waters
have been described (12).
C. Environmental Control of the Synthesis and Activity
of Ectoenzymes
The complex environmental regulation of ectoenzyme synthesis and activity has been
demonstrated in studies of bacterial β-glucosidase and aminopeptidase in surface waters
of lakes (24,25,37). The results of these studies demonstrated that the ectoenzyme synthe-
Copyright © 2002 Marcel Dekker, Inc.
Figure 3 Competitive inhibition of β-glucosidase activity by glucose (end product of enzyme
reaction) in water samples from eutrophic Lake Mikołajskie. (A) Hyperbolic relationship between
enzyme activity and increasing substrate concentrations, (B) Lineweaver-Burk’s linear transforma-
tion of the relationship between enzyme activities (1/v) and increasing concentrations of substrate
(1/S). (Chro
´
st, unpublished.)
Copyright © 2002 Marcel Dekker, Inc.
sis and activity were under different control mechanisms, which were dependent on the
physical-chemical conditions of the habitat. There is ample evidence for general catabolic
repression of ectoenzyme synthesis in bacteria due to readily utilizable carbon sources
(24,37,44,45) as well as more specific repression by end products of enzyme catalysis
(34,46). However, control of aminopeptidases appears to be distinct and more complex
than that of other ectoenzymes. In some bacteria, amino acids, peptides, and/or proteins
seem to induce aminopeptidase synthesis (45,47). It is not known specifically how amino-

peptidase induction operates, especially since amino acids are reported to act as inducers
in some bacteria, rather than acting in their more predictable role as end-product inhibitors.
The ability of bacteria living in the euphotic zone of the lakes to produce ecto-
enzymes seems to be strongly affected by the availability of the low-molecular-weight,
readily utilizable substrates exudated by algae (eg., excreted organic carbon-EOC), which
are known to be excellent substrates for bacteria (48–50). Chro
´
st and Rai (25) found that
the rates of leucine-amino-peptidase and α-glucosidase production by aquatic bacteria
strongly depend on bacterial organic carbon demand. When the amount of EOC fulfilled
the bacterial organic carbon requirement, microorganisms did not synthesize enzymes
needed for hydrolysis of the polymeric substrates because their utilization was unneces-
sary. Moreover, the specific activity of aminopeptidase correlated negatively to the rates
of algal EOC.
During the active growth of phytoplankton, algal populations excrete into the water
a variety of photosynthetic products, including easily assimilable low-molecular-weight
substrates (51), which support bacterial growth and metabolism. These substrates inhibit
the activity and repress the synthesis of ectoenzymes in bacteria. On the other hand, when
low levels of readily available substrates limit bacterial growth and metabolism, bacteria
produce ectoenzymes with high specific activity to degrade polymers and other nonlabile
substrates. Such a situation occurs in lake water during the breakdown of phytoplankton
bloom. Senescent algae liberate, through autolysis of cells, a high amount of polymeric
organic compounds (polysaccharides, proteins, organophosphoric esters, nucleic acids,
lipids, etc.), which induces synthesis of ectoenzymes. Another mechanism that causes
repression cessation of enzyme synthesis is low level of directly utilizable organic com-
pounds in the water during bloom breakdown (52).
Bacteria living in the profundal zone of the lakes are often substrate-limited (2,53)
because the amount of substrate in deep waters depends primarily on the sedimentation
rates of the organic matter that is produced in the euphotic zone. There is no direct supply
of labile organic compounds exudated by algae. In the profundal zone, sedimentation

provides labile monomeric organic compounds that are mostly polymers that are utilized
by bacteria. Under such environmental circumstances, bacterial metabolism is strongly
dependent on the presence and amount of polymeric substrates and the activity of synthe-
sized ectoenzymes that catalyze the release of readily utilizable monomers.
Microbial ectoenzymatic activity in natural waters is also strongly dependent on
environmental factors, such as temperature, pH, inorganic and organic nutrients, ultraviolet
B (UV-B) radiation, and presence of activators and/or inhibitors (3,13,21,54–59). Several
studies have shown that ectoenzymes display the highest activities in alkaline waters of
pH 7.5 to 8.5 (24,40) or acid waters of pH 4.0 to 5.5 (55). In contrast to the pH response,
many ectoenzymes exhibit no obvious adaptation to ambient temperature, because the
optimal temperature is often considerably higher than in situ temperature of waters (13,33,
40). The optimal temperatures for alkaline phosphatase and β-glucosidase are unchanged
when they are produced by planktonic microorganisms in lake water under different in
situ temperatures (13,24).
Copyright © 2002 Marcel Dekker, Inc.
In light of these aforementioned studies, the environmental regulation of ectoenzyme
synthesis and activity is complex and usually no single factor is involved in this process. It
is important to realize that environmental regulation of ectoenzymes, induction, synthesis
repression, and inhibition are related to concentration, period of exposure, and such factors
as temperature, pH, oxygen level, and chemical characteristics of regulatory molecules.
The same molecule that is an inducer under one set of circumstances may be a repressor
under other environmental conditions, or at different concentrations.
IV. ASSAYS OF ECTOENZYME ACTIVITY
A. Methods
There are significant difficulties in measuring ectoenzyme activities in heterogeneous envi-
ronments such as natural waters and soil, which include questions about methodology
and data interpretation. For example, should assays be performed according to the well-
established principles of enzymology (e.g., excess substrate, optimal pH and temperature,
shaking of reaction mixtures) or in situ conditions encountered in an aquatic environment
(e.g., limiting and unevenly distributed substrate, suboptimal and fluctuating physical con-

ditions, stationary incubation)? How are the optimal assays related to those assays done
under more ‘‘realistic’’ conditions?
A variety of methods are available for monitoring the enzyme activities when work-
ing with microbial cultures or isolated enzymes in biochemical laboratories. However,
most classical enzymatic methods cannot be applied directly in aquatic environments. The
enzyme amount and activity in natural waters are usually much lower than those measured
in cultures or in enzyme extracts, and therefore the classical biochemical methods often
are inadequate for measuring low ectoenzyme reaction velocity. Furthermore, the environ-
mental conditions of ectoenzyme assays in water samples often are suboptimal (e.g., un-
suitable temperature, pH, presence of interfering compounds) and the choice of substrate
used to study ectoenzymes of natural microbial assemblages in aquatic environments often
is problematic.
Depending on the chemical nature of the ectoenzyme substrate, there are three cate-
gories of methods for measurement of ectoenzyme activity in aquatic environments: spec-
trophotometric, fluorometric, and radioactive. The most commonly used in the past were
spectrophotometric methods (60–63). The major disadvantage of spectrophotometric
methods is long incubation time necessary for enzyme reactions, which is due to their
relatively low sensitivity (micromolar [µM ] to millimolar [mM ] concentrations of the
final product of enzyme reaction are required). However, spectrophotometric assays can
be used when measuring high enzyme activity in samples, or when working with purified
and/or concentrated enzymes.
During the last two decades, fluorometric methods have been widely used for en-
zyme activity determinations in aquatic environments (3,21,24,33,52,64,65). Fluorometric
assays are very sensitive, and they measure the final products of enzymatic reactions in
nanomolar (nM) to micromolar (µM ) concentrations. When using a modern spectroflu-
orometer to measure enzyme activity in water samples, the incubation time for monitor-
ing substrate-enzyme reaction can be shortened to a few minutes. Several authors have
applied radiometric methods for enzyme activity determination in aquatic environments
Copyright © 2002 Marcel Dekker, Inc.
(66–69). Although these methods are extremely sensitive, they are seldom used because

of greater costs and precautions needed when handling radioactive materials.
B. Substrates
When studying the significance of ectoenzyme activities in relation to in situ substrate
turnover in aquatic (and in soil) environments, one should be able to determine the real
rates of the process. In this case, the substrate should have an affinity for the ectoenzyme
similar to that of the natural substrates in situ. Moreover, the enzyme should be assayed
by using low substrate concentrations comparable to the concentrations of the natural
substrate. Application of low substrate concentrations results in low levels and difficult
detection of end products. Generally, in enzyme reactions, the length of incubation re-
quired for end product formation is related to product detection sensitivity.
A large variety of commercially produced ectoenzyme substrates are now available.
Depending on their chemical structure and the enzyme assay, two types of organic com-
pounds can serve as substrates: natural and artificial substrates. Natural substrates are
native compounds (nonlabeled) or their chemical structure is only slightly modified by
labeling with chromophores, fluorophores, or radiolabeling with
14
C,
3
H,
32
P,
35
S, or
125
I.
Most natural substrates have an affinity for the enzyme that is complementary to that of
the natural substrates in aquatic samples. Monitoring of enzyme activity by means of
natural nonlabeled substrates requires a sensitive analytical method to measure the end
product or substrate remaining after incubation time (70). Modern analytical methods offer
precise and rapid determination of several natural compounds that can be used as enzyme

substrates or products (e.g., amino acids, proteins, deoxyribonucleic acid [DNA], carbohy-
drates). The application of the labeled natural substrates requires very sensitive and accu-
rate methods for quantitative determination of the label bound to the substrate molecule
(e.g., spectrophotometry, fluorometry, radiometry). Most suitable natural substrates are
radiolabeled compounds because their end products can be measured after a short incuba-
tion period (minutes) (66–69). Until now, this approach has been limited by the reduced
availability of radiolabeled substrates and the high handling costs of radioactive materials.
Except for some cases of analytical difficulties, natural substrates are promising for study-
ing ectoenzymes in aquatic environments.
Artificial substrates are synthesized in laboratories and their chemical structure (e.g.,
chemical bonds) only mimics that of natural compounds. Ectoenzymes react with artificial
substrates by splitting specific chemical bonds between an organic moiety and its chromo-
phore or fluorophore, yielding colored or fluorescent products, respectively. Because these
are not natural substrates, enzyme activities obtained are not necessarily identical to those
measured by using natural substrates. However, their application allows for low costs and
simpler and more rapid measurements of ectoenzymatic activity.
In the past, chromogenic artificial substrates were used intensively in the studies of
ectoenzyme activity in fresh waters (10,11,60–63,71). It is advantageous to use chromo-
genic substrates because they can be measured easily by spectrophotometry. However,
low sensitivity is a major disadvantage of this technique, and long incubation times of 72
to 96 hours often are required (62,71). This may result in microbial proliferation and
ectoenzyme synthesis during the assay, changes, which must be prevented. They usually
are avoided by adding plasmolytic or antiseptic agents to assays, such as toluene or chloro-
form (10,38,71). However, these agents change the membranes, thereby leading to release
Copyright © 2002 Marcel Dekker, Inc.
of ecto- and intracellular enzymes. In cases in which some enzymes are located intra- and
extracellularly (e.g., phosphatase, arylsulfatase), ectoenzyme activity may be significantly
overestimated (72).
Recently, fluorophore-labeled artificial substrates have been commonly used for
sensitive assays of ectoenzyme activity in aquatic environments (13–16,21,25,33,36,37,

54,55,64,65) and are advantageous when it is necessary to perform a large number of
assays. Fluorogenic substrates yield highly fluorescent, water-soluble products with optical
properties significantly different from those of the substrate. Many substrates are degraded
to products that have longer wavelength excitation or emission spectra. Therefore, these
fluorescent products typically can be quantified in the presence of an unreacted substrate
by using a fluorometer. Three types of substrates derived from water-soluble fluorophores
are commercially available: blue, green, and red (73).
Hydroxy- and amino-substituted coumarins are the most widely used fluorogenic
substrates. Coumarin-based substrates produce highly soluble, intensely blue fluorescent
products. Phenolic dyes such as 7-hydroxycoumarin (umbelliferone) and the more com-
mon 7-hydroxy-4-methylcoumarin (methylumbelliferone) are not fully deprotonated and
therefore not fully fluorescent unless the reaction mixture has pH Ͼ 10 (64). Substrates
derived from these fluorophores are not often used for continuous measurement of enzy-
matic activity. Products of substrates containing aromatic amines, including the commonly
used 7-amino-4-methylcoumarin, 7-amino-4-trifluoromethylcoumarin, and 6-aminoquino-
line, are partially protonated at pH Ͻ 5 but fully deprotonated at neutral pH. Thus, their
fluorescence is not subject to variability due to pH-dependent protonation/deprotonation
when assayed near or above physiological pH.
Substrates derived from water-soluble green fluorophores, fluorescein, and rhoda-
mine, provide significantly greater sensitivity in fluorescence-based enzyme assays. In
addition, most of these longer-wavelength fluorophores have excitation coefficients that
are 5 to 25 times that of coumarins, nitrophenols, or nitroanilines, making them potentially
useful as sensitive chromogenic substrates (73). Substrates based on the derivatives of
fluorescein and rhodamine usually incorporate two moieties, each of which serves as a
substrate for the enzyme. Consequently, they are cleaved first to the monosubstituted ana-
log and then to the free fluorophore. Because the monosubstituted analog often absorbs
and emits light at the same wavelength as that of the ultimate hydrolysis product, this initial
hydrolysis complicates the interpretation of hydrolysis kinetics (often biphasic kinetics are
observed). However, when highly purified, the disubstituted fluorescein- and rhodamine-
based substrates have virtually no visible-wavelength absorbance or background fluores-

cence, making them extremely sensitive substrates.
Substrates derived from water-soluble red fluorophores (long-wavelength fluoro-
phores) often are preferred because background absorbance and autofluorescence generally
are lower when longer excitation wavelengths are used. Substrates derived from the red
fluorescent resorufin and dimethylacridinone contain only a single hydrolysis-sensitive
moiety, thereby avoiding the biphasic kinetics.
The majority of fluorophore-labeled substrates produce very low background fluo-
rescence and can be used without any loss in sensitivity at the high concentrations (milli-
molar) that are sometimes needed for enzyme saturation (65). It also is possible to work
with substrate concentrations in the nanomolar range, close to the presumed range of
natural substrate concentrations in aquatic environments. It has been shown that substrates
linked to fluorophores provide a very sensitive system for detecting and quantifying many
specific and nonspecific hydrolases in aquatic environments (21). The potential ectoenzy-
Copyright © 2002 Marcel Dekker, Inc.
maticactivityofwaterorsedimentsamplescanbemeasuredoverashortincubation
timewithoutproblemsofmicrobialproliferation,lowactivity,andnonsaturationofthe
ectoenzyme.Inspiteofthisadvantageinusingthefluorescentsubstratesinectoenzyme
assays,theiruseiscontroversial(asarechromogenicsubstrates),becauseoftheirunknown
affinityfortheectoenzymesincomparisontothatofnaturalsubstrates.
C.PotentialEnzymeActivity—KineticApproach
Ifinformationaboutthepotentialactivityoftheectoenzymeintheaquatichabitatis
required,therearereasonsforusinghighconcentrationsofthesubstrateinassays.The
enzymeshouldbesubstrate-saturated.Possiblecompetitionwithco-occurringnaturalsub-
stratesshouldbeprevented,asshouldcompetitiveinhibitionofthesubstratewithinhibi-
torsinsamples(Fig.3).
ManyhydrolyticectoenzymesfollowMichaelis-Mentenkinetics:
vϭ(V
max
ϫ[S])/(K
m

ϩ[S])
whereaplotoftheinitialvelocityofreaction(v)againstincreasingconcentrationsof
substrate([S])givesarectangularhyperbola.Forsuchassays,thekineticapproachis
recommended;itallowscalculationfromtheexperimentaldataofthekineticparameters
characterizinganenzyme-substratereaction.TheyareV
max
,themaximalvelocityofen-
zymecatalysisthattheoreticallyisattainedwhentheenzymehasbeensaturatedbyan
infiniteconcentrationofsubstrate,andK
m
,theMichaelisconstant,whichisnumerically
equaltotheconcentrationofsubstrateforthehalf-maximalvelocity(V
max
),whichindicates
theenzymeaffinitytothesubstrate(74).Thekineticapproachrequiressubstrateconcen-
trationsrangingfromlowtohighforfirst-order(thereactionvelocityincreaseslinearly
withtheincreaseinsubstrateconcentrations)andzero-order(reactionvelocityremains
constant,notaffectedbytheconcentrationofsubstrate)enzymereactions.
Atypicalectoenzymekineticexperimentmaybedescribedasfollows:Dataare
collectedasafunctionofatleastfivetriplicatereactantconcentrationsofsubstrate,and
theexperimentaldependenceonthisfunctionisdeterminedandplottedgraphically.The
resultsdependessentiallyontheshapeofthehyperboliccurvedescribedbythedata,thus
makingdeterminationofV
max
andK
m
difficult(Fig.3A).Toobtainthesekineticparame-
ters,theMichaelis-MentenequationoftenisrearrangedtothelinearformandV
max
and

K
m
areobtainedfromtheslopeandintercept(Fig.3B)(74).
Such graphical methods produce correct values for the parameters only in the ab-
sence of error. Unfortunately, all the measurements are subject to some degree of impreci-
sion, and therefore use of linearized equations such as that of Lineweaver-Burk, Eadie-
Hofstee, and Woolf may give inaccurate or biased experimental data (75,76). The best
solution to this problem is to perform a nonlinear regression analysis on the original experi-
mental data. The kinetic parameters then can be calculated from the direct plot of reaction
velocity (v) versus substrate (S) concentration by using a computer program to determine
the best fit of the rectangular hyperbola (77).
D. In Situ Enzyme Activity—Direct Approach
True ecological information requires the detection of environmental processes under in
situ conditions, which cannot be fully controlled and, therefore, cannot be simulated in
the laboratory. The composition of naturally occurring substrates in water samples usually
Copyright © 2002 Marcel Dekker, Inc.
is unknown, and concentrations may vary widely over short sampling times. This condition
complicates the choice of the substrate concentration being monitored in ectoenzyme
assays because of the potential interference or competition with natural substrates and/or
inhibitors.
Ideally, to prevent these problems and to measure the real in situ rates of ectoenzyme
activity, one should follow the decrease in naturally occurring substrate concentration or
the increase in ectoenzyme product formation under in situ conditions. Because of the
analytical difficulties, this approach is very seldom used in aquatic studies. Moreover, the
increase in concentration of ectoenzyme product in samples simply cannot be measured
because liberated product is simultaneously utilized by microorganisms. To overcome this
problem and to be able to measure the amount of product released from its substrate, it
is necessary to inhibit product assimilation by intact living microorganisms. Several inhibi-
tory agents that do not inhibit enzyme activity can be used to prevent the microbial assimi-
lation of low-molecular-weight products of ectoenzymatic hydrolysis of polymeric sub-

strates (e.g., antibiotics or chemotherapeutics blocking active transport systems, some
fixing agents such as sodium azide).
Figure 4 Direct estimation of enzymatic hydrolysis of natural DNA by means of decrease in
substrate concentration in water samples from eutrophic Lake Mikołajskie. (A) Concentration of
remaining DNA in samples after incubation times, (B) amount of DNA hydrolyzed (see text for
description of kinetic parameters). (Chro
´
st, unpublished.)
Copyright © 2002 Marcel Dekker, Inc.
Figure4presentsanexampleofdirectestimationofnaturalDNAhydrolysisin
eutrophicLakeMikołajskie.Topreventmicrobialgrowthandutilizationofproductsof
DNAdegradationinthecourseofitshydrolysis,watersampleswerefixedwith0.3%
sodiumazide.TheconcentrationofDNAdecreasedfrom4.75Ϯ0.08µgL
Ϫ1
(attime
zero)toaplateauof3.79Ϯ0.17µgL
Ϫ1
(after55hours)(Fig.4A).Aplotoftheamount
ofDNA-hydrolyzed[s]versustimeofhydrolysis[t]gavearectangularhyperbola(Fig.
4B):
s ϭ [S
n
] ϫ [t])/(k
hydrolysis
ϩ [t])
By applying a nonlinear regression analysis to the experimental data it was possible to
estimate hydrolysis parameters: [S
n
], concentration of DNA naturally present in water
samples, which theoretically is attained after infinite time of hydrolysis, and k

hydrolysis
,the
hydrolysis constant, i.e., the hydrolysis time of the half-concentration of natural DNA
(S
n
). The preceding data provide an example for determining hydrolysis parameters of a
naturally occurring enzyme substrate by analysis of substrate concentration evolution in
water samples during the course of its enzymatic degradation.
Using a direct approach, it also is possible to estimate the hydrolysis parameters
characterizing in situ enzymatic degradation of natural substrates when the concentrations
of the final product are determined during the course of hydrolysis. In situ hydrolysis of
proteins by proteolytic enzymes yielded increasing concentrations of free, dissolved amino
acids in lake water samples when microbial uptake was inhibited by 0.3% sodium azide
(Fig. 5).
Figure 5 Direct estimation of enzymatic hydrolysis of natural protein by means of release of
reaction products (amino acids) in water samples from eutrophic Lake Mikołajskie. (Siuda and Kier-
sztyn, unpublished.)
V. TEMPORAL AND SPATIAL DISTRIBUTION OF ECTOENZYME
ACTIVITY
Both spatial and seasonal ectoenzymatic activities fluctuate markedly in lake waters
(13,24,28,29,36,38). The production of ectoenzymes by microorganisms is strongly corre-
Copyright © 2002 Marcel Dekker, Inc.
Figure6Seasonalaminopeptidaseactivityandchlorophyll
a
concentrationinthesurfacewater
samples(0-to1-mdepth)fromLakePlußsee.(Chro
´
st,unpublished.)
latedtotheinfluxofpolymericorganicmatterand/orthedepletionofreadilyutilizable
UDOMintheenvironment(3,24,52).

Ectoenzymeproductionandactivityshowmarkedseasonalvariationinbothsurface
anddeepwatersoflakes.Insurfacewaters,themaximalectoenzymaticactivitiesoccur
duringthelatestageofphytoplanktonbloomandafteritsbreakdown,andminimalactivi-
tiesoccurduringtheclearwaterphaseinlakes(Fig.6)(13,24,38,78).
Ectoenzymaticactivityisespeciallylowerduringsummerthermalstratificationin
thehypolimnionthanintheepilimnionofalake,andtheactivityisstronglydependent
uponthesedimentationratesofdetritusproducedintheeuphoticzone(Fig.7).Usually,
alagperiodisobservedbetweenthemaximalectoenzymaticactivityinsurfaceandthat
indeepwatersofalake(Fig.7B,D)(78).
Therealsoareknownhighdiurnalfluctuationsinenzymeactivityinlakewater
(78,79)becausedynamicenvironmentalfactorsaffectenzymeproductionand/oractivity,
aswellasmicrobialgrowth,metabolism,andbiomassofmicrobialenzymeproducers
(3,25,36).InthesummerepilimnionofeutrophicLakeMikołajskie,APaseactivityvaried
byafactor1.8during24hours(Fig.8).MinimalratesofAPaseactivityweremeasured
during the night period, and the enzyme activity continuously increased from morning
to evening. The rates of enzyme activity were positively correlated with fluctuations in
chlorophyll
a
concentrations, indicating that phytoplankton was a major producer of alka-
line phosphatase in an epilimnion of the lake. In contrast to alkaline phosphatase activity,
the highest rates of aminopeptidase activity in the surface water layer of eutrophic Lake
Głe
˛
bokie were recorded during the night when the maximal concentrations of the total
and dissolved proteins were determined (78).
VI. ROLE OF PHOSPHOHYDROLASES IN PHOSPHORUS CYCLING
Current research clearly shows that orthophosphate ions (Pi) are major factors for control-
ling microbial primary and secondary production in many freshwater environments (80–
Copyright © 2002 Marcel Dekker, Inc.
Figure 7 Aminopeptidase (A, B) and β-glucosidase (C, D) activities in the thermal stratified water

column of Lake Mikołajskie during summer phytoplankton bloom (A, C) and after bloom breakdown
(B, D) in the epilimnion (epi-), metalimnion (meta-), and hypolimnion (hypo-). (Chro
´
st, unpub-
lished.)
85). As confirmed by several independent approaches, the ambient Pi concentration is far
too low to meet plankton phosphorus (P) requirements in the euphotic zone of lakes, and
therefore most (80–90%) of the P used for production of microbial biomass originates
from dephosphorylation of P organic compounds during their degradation.
A variety of aquatic organisms (bacteria, algae, cyanobacteria, protozoa, macrozoo-
plankton, benthic animals, and aquatic angiosperms) release Pi from organic compounds.
Although contributions of the last two groups of aquatic organisms to P cycling can be
important in some fresh waters (86), these are not discussed here.
In this review, enzymatic microbial cycling of P is defined as the process of dephos-
phorylation of organic P compounds by hydrolytic enzymes produced by microorganisms
that leads to the release of Pi into the environment surrounding microbial cells. This defi-
Copyright © 2002 Marcel Dekker, Inc.
Figure 8 Summer diurnal fluctuations of alkaline phosphatase activity (A) and concentration of
chlorophyll
a
in the surface water samples (0–0.5 m) from eutrophic Lake Głe
˛
bokie. (Chro
´
st, unpub-
lished.)
nition excludes the release of Pi and dissolved organic phosphorus (DOP) compounds by
zooplankton (87,88). However, since these planktonic animals can affect significantly the
whole microbial community as well as dynamics of P compounds in aquatic environments,
it is necessary to discuss selected aspects of their influence on enzymatic Pi release.

Only 30%, or less, of the total organic P pool in freshwaters is composed of easily
hydrolyzable dissolved or colloidal P constituents (86). The remainder constitutes particu-
late organic P that is not directly available for microbial metabolism but can be utilized
after ingestion of food particles by herbivorous zooplankton. Transformation of particulate
P into DOP compounds (incomplete digestion of food particles, zooplankton grazing)
effectively accelerates enzymatic Pi release by increasing the substrate pool for phospho-
hydrolases in an environment (89). One interesting and extensively studied aspect of P
recycling by zooplankton entails the production of specific phosphohydrolases by these
planktonic animals and the liberation of the intracellular phosphohydrolytic enzymes from
grazed phytoplankton (67).
Almost all (except phosphoamides) natural DOP compounds in aquatic environ-
ments are chemically stable phosphate esters. Phosphorus in these compounds is not
readily available for microorganisms because the majority of phosphate ester molecules
cannot be transported directly through microbial cell membranes. Limited quantities of
Copyright © 2002 Marcel Dekker, Inc.
β-glycerophosphate and several phosphorylated monosaccharides can be taken up by some
aquatic bacteria (90,91); orthophosphate ions, however, are the dominant forms of phos-
phorus for microbial assimilation. Therefore microbial utilization of Pi from almost all
its organic compounds must be preceded by their enzymatic dephosphorylation (3,13,68).
Release of Pi into aquatic ecosystems is affected by a great variety of abiotic and
biotic environmental factors and processes. The most important are activity of phosphohy-
drolytic enzymes (3,11), zooplankton grazing (92,93), viral and spontaneous lysis of mi-
croplankton cells (94,95), and UV light (96). However, it is now well founded that enzyme-
mediated hydrolysis of naturally occurring phosphate esters is the most significant mecha-
nism for P release in aquatic environments.
There are three main groups of hydrolytic enzymes responsible for Pi release: non-
specific and/or only partially specific phosphoesterases (mono- and diesterases), nucleo-
tidases (mainly 5′-nucleotidase), and nucleases (exo- and endonucleases). Most of them are
typical ectoenzymes (3,67). However, some of phosphohydrolytic enzymes are actively
secreted by planktonic microorganisms into surrounding water (e.g., extracellular phos-

phoesterases and some nucleases) (Fig. 9).
Figure 9 Conceptual model of enzymatic decomposition of various organic phosphorus com-
pounds in lake water. Pathways that are crucial for Pi regeneration in scale of the whole ecosystem
are illustrated by bold arrows. ‡@, 5′-nucleotidase; ‡A, alkaline and acid phosphatases; ‡B, exo-
nucleases; ‡C, endonucleases; ‡D, phytase; ‡E, cyclic 3′,5-nucleotide phosphodiesterases and 2′,3-
nucleotide phosphodiesterases; ‡F, liberation and release of DOP compounds from disrupted and
living cells; ‡G, direct uptake of organic P source. (Siuda, unpublished.)
Copyright © 2002 Marcel Dekker, Inc.
A.Phosphomonoesterases(Phosphatases)
Phosphatases(nonspecificphosphomonoesterases)arethemostintensivelystudiedgroup
ofphosphohydrolasesparticipatinginPirelease.Thepresenceofactivephosphohydrolytic
enzymesthatwereexcretedbyzooplanktoninwatersamplesandtheircapacityforPi
releasefromorganicPcompoundswerefirstmentionedbySteinerin1938(97).But
theresearchofOverbeckandReichardt(60,61,98,99)arethefoundationsforthecurrent
knowledgeoftheecologicalroleofphosphatasesinaquaticenvironments.Theresultsof
hundredsofstudiesinthelast40yearshavecontributedgreatlytothepresentknowledge
ofphosphatases,whichareprobablynowthebest-knownphosphohydrolyticenzymesin
aquaticecosystems(3,10–13,26,37,39,55,60–63,68,71,98,99).Avarietyofalkalineand
acidphosphatasesareproducedbyalmostallmembersoftheplanktoncommunity,includ-
ingbacteria,algae,cyanobacteria,fungi,protozoa,andzooplankton(11).
Alkalinephosphatases(APases)arethegroupofadaptativeisoenzymesthatreact
optimallyinpHrange7.6–9.6.TheyliberatePifrommonophosphateestersofprimary
andsecondaryalcohols,sugaralcohols,cyclicalcohols,phenols,andaminesbutnotfrom
phosphodiesters(100).TheratesofAPasesynthesisareregulatedbyrepression/derepres-
sionmechanisms,andPiactsasarepressor(3,11,13,101).APaseactivityalsoisregulated
bycompetitiveinhibitionPi(3,11,13).
SincethepHoflakesoftenisalkaline(pH7.2–9.5),phosphatasesthatexhibittheir
maximalactivityinacidwater(acidphosphatases[AcPases])probablyhaveonlyaminor
importanceinalkalinelakes.SeveralstudiesreportedhighAcPaseactivitiesduringhyd-
rolysisoforganicPcompoundsinacidifiedlakes(55,102).ContrarytoAPase,acidphos-

phatasescompriseisoenzymesinwhichsynthesisoftenisnotrepressedbyPipresentin
theaquaticenvironment(103).
DespiteconsiderableknowledgeofspatialandtemporalchangesinpotentialAPase
activitiesinlakes,thequantitativeaspectsofPireleasemediatedinsitubyphosphatases
arepoorlyunderstood.AlthoughAPaseactivitiesinsurfacewatersofmesotrophicand
eutrophiclakes,measuredbysensitivefluorometricmethods,arecommonlyhigh(5–100
nmolPO
4
3Ϫ
L
Ϫ1
and20–500nmolPO
4
3Ϫ
L
Ϫ1
,respectively),theydonotdemonstratethe
importanceofPireleaseintheseenvironments.ToestimatetherealinsituratesofPi
releasebyAPase,theambientconcentrationsofPiandnaturalAPasesubstrates(phospho-
monoesters[PMEs])inwatersamplesshouldbeconsidered.
Duringthesummerstratificationperiod,Piconcentrationsinwatersamplesfrom
eutrophiclakesextendoverawiderange.Theyvaryfrom0µMinsurfacesamplesto15
µM,orevenmore,inthehypolimnion(Fig.10).Concentrationandchemicalcomposition
ofPMEinlakewaterusuallyarenotknown.However,somestudiesreportedthecon-
centrationsofenzymaticallyhydrolyzableP(EHP)inaquaticecosystems(104–107).As-
sumingthatthemajorityofPMEfractioniscomposedofEHPcompounds,onecansup-
posethatthePMEpooldoesnotexceed0.2–0.4µmolL
Ϫ1
.HighPMEconcentrations
usuallywerefoundineutrophiclakesduringperiodsofbreakdownofphytoplankton

blooms.
Orthophosphateisawell-knowncompetitiveinhibitorofAPaseinaquaticenviron-
ments(Fig.11).Therefore,insituactivityoftheseenzymesisdependentoninhibitor/
substrate ([I]/[S]) ratio. More than 80% of APase activity is inhibited when the value of
the [I]/[S] ratio in lake water increases above 2.5 (107). Determination of ([I]/[S]) ratios
in surface waters of various mesotrophic and eutrophic lakes showed that in the majority
Copyright © 2002 Marcel Dekker, Inc.
Figure10(A,C)Concentrationsoforthophosphateion(Pi)andenzymaticallyhydrolyzablephos-
phate(EHP),and(B,D)relationshipbetween[Pi]/[EHP]ratioandpercentageofcompetitiveinhibi-
tionofalkalinephosphataseactivity(APase)inthephoticandprofundalzoneofeutrophicLake
Głe
˛
bokie.(DatamodifiedfromRef.38.)
ofthestudiedlakes,the[I]/[S]ratiovariedfrom3.3to29.1andonlyoccasionallydropped
below1duringtheperiodsofmaximalPidepletion.Similarcalculationsweremadefor
depthprofilesofthelakeduringasummerstratificationperiod.Theyshowedthatsurface
watershad[I]/[S]ratiosthatfluctuatedaround1andincreasedrapidlyinthehypolimnion
toϳ252(Table2).TheseobservationsstronglysuggestedthatefficientPiregeneration
by APase in deep stratified lakes probably is restricted exclusively to the thin layer of the
surface waters and periods of maximal Pi depletion (67). In the water column of the
moderately deep lakes (10- to 30-m depth), APase may have only minor importance for
the decomposition of organic P compounds.
Phosphatases of lake microplankton are represented by a group of enzymes charac-
terized by different biochemical properties (half-saturation constants, temperature and pH
optima, substrate specificity, susceptibility to the presence of activators and/or inhibitors).
It should be emphasized that the role of APase in Pi release processes in freshwater ecosys-
tems probably is more complicated than may be expected from simple models of the
synthesis and activity regulation based on mechanisms of the repression/derepression and
Copyright © 2002 Marcel Dekker, Inc.
Figure 11 Relationship between alkaline phosphatase activity (APase) and orthophosphate (Pi)

concentration in the surface water of eutrophic Lake Głe
˛
bokie. (Data modified from Ref. 38.)
Table 2 Competitive Inhibition of Alkaline Phosphatase (APase) Activity by Orthophosphate
(Pi) in the Water Column of Mesotrophic Lake Constance and Eutrophic Lake Schleinsee
(µgP-PO
4
3Ϫ
L
Ϫ1
)
APase
Depth Pi EHP inhibition
a
Lake (m) Pi/EHP (%)
Constance
2 1.39 1.64 0.85 77
10 1.80 1.67 1.08 80
12 1.79 1.05 1.70 82
20 6.70 2.08 3.22 91
50 28.49 0.80 35.61 97
190 59.30 4.53 13.09 94
Schleinsee
1 1.30 2.51 0.52 20
3 1.80 2.20 0.82 40
5 2.98 0.99 3.01 85
7 161.51 0.93 173.67 98
9 297.60 2.08 143.07 96
12 327.63 1.30 252.03 99
APase, alkaline phosphatase; Pi, orthophosphate ion; EHP, enzymatically hydrolyzable P.

a
The decrease of APase activity was calculated from enzyme kinetics and inhibitor/substrate ratio. APase was
measured by means of methylumbelliferyl-phosphate (MUFP) as a substrate under saturation condition; the
affinity of MUFP and EHP to APase was assumed to be the same.
Source: Adapted from Ref. 26.
Copyright © 2002 Marcel Dekker, Inc.
competitive inhibition by Pi. Information on alternative mechanisms of the regulation of
APase activity and synthesis in freshwaters is limited. A few studies described high APase
activity in deep oceanic waters in the presence of high (ϳ3.5-µM) Pi concentrations
(108,109). Similar observations were found in the profundal zone of deep eutrophic lakes
(Siuda, unpublished).
It is commonly believed that APase activity in deep regions of the lakes originates
from the surface water and is exported down to the hypolimnion by rapidly sinking parti-
cles. There is also some evidence that APase activity in Pi-rich layers of an aquatic ecosys-
tem originates from bacteria producing Pi-resistant APase (109,110). Quantitative partici-
pation of bacterial, Pi-resistant APase in Pi release in freshwaters is unknown and needs
further intensive investigation. Several studies suggested that bacterial APase, contrary to
algal APase that is produced under Pi limitation, has multifunctional properties that can
alter both organic P decomposition and C and N mineralization from dissolved organic
compounds (13,50,107,109).
B. 5′-Nucleotidase and Nucleases
Free nucleic acids dissolved in water (DNA and ribonucleic acid [RNA]) represent proba-
bly the most significant reservoir of P potentially available for planktonic microorganisms
in aquatic ecosystems. The distribution of extracellular, dissolved DNA (dDNA) in both
freshwater and marine environments is relatively well known. Minear (111) found from
4to30µg dDNA L
Ϫ1
in oligotrophic and eutrophic lowland ponds. Similar dDNA concen-
trations in various oligotrophic and mesotrophic environments (0.2–44.0 µgL
Ϫ1

and 0.5–
25.6 µgL
Ϫ1
, respectively) were documented by later studies (32,112–115). An extremely
high content of dDNA (88 µgL
Ϫ1
) was found by Karl and Bailiff (116) in an eutrophic
Hawaiian pond. RNA, one of the basic cell components, constitutes about 10–20% of cell
biomass; free dissolved RNA, however, has not been found in lake water. It suggests that
turnover time of dissolved RNA is extremely short because of its rapid enzymatic hydroly-
sis by ribonuclease (RNase).
The absolute quantities of Pi release resulting from extracellular nucleic acids are
subject to considerable uncertainty as a result of methodological limitations. Nevertheless,
they may be important Pi sources in many aquatic environments. Cautious calculations
show that enzymatically hydrolyzable extracellular DNA contributes from 10% to 60%
of P to the total dissolved organic P pool (32,117). Therefore, enzymatic liberation of Pi
from dDNA by DNase and subsequently by 5′-nucleotidase action can be one of the most
effective pathways of Pi release.
Three main groups of microplankton enzymes mediate the processes of Pi liberation
from nucleic acids: nucleases (exo- and endonucleases), 5′-nucleotidase (5′-nase), and
phosphatases. Literature information on nucleases and their activity and distribution
among various microplankton components in freshwaters is scarce. Siuda and Gu
¨
de (117)
found that in the epilimnion of Lake Constance, DNase activity was mainly extracellular
and/or similar to 5′-nase activity in being coupled to the plankton size fraction (0.1–1.0
µm). In a eutrophic part of Tokyo Bay, Maeda and Taga (118) concluded that DNA-
hydrolyzing bacteria were distributed widely in seawater and that DNase activity was
bound mainly to suspended particles or microbial cells.
DNA-degrading capacity of microbial communities is usually estimated by measur-

ing the loss of DNA integrity added to water samples in quantities similar to those naturally
present in the environment. Kinetic data give only rough approximation of the velocity
Copyright © 2002 Marcel Dekker, Inc.
of nucleic acid degradation in situ. Various independent approaches suggest that half-life
of extracellular DNA is relatively short and usually varies between 4.2 and 15 hours in
oligotrophic and eutrophic ecosystems (113,117). Considering the relatively high extracel-
lular DNA concentration in aquatic environments and the fact that DNase activity remains
relatively unchanged during the whole summer period, constant supplementation of lake
waters with nucleotides can be expected (117).
In earlier studies, only APase was regarded as the main enzyme responsible for
enzymatic Pi release in natural waters. However, Azam and Hodson (119) showed the
potential role of 5′-nase activity in Pi release in marine environments. This bacterial,
membrane-bound enzyme is largely specific for various 5′-nucleotides and does not de-
phosphorylate other phosphate esters. In contrast to activity of APase, the activity of 5′-
nase is not dependent on Pi concentrations (67,120,121). Although the function of 5′-nase
in Pi release processes in marine environments is relatively well known (67,119,122,123),
few papers have discussed the role and importance of this enzyme in fresh waters (3,
117,124,125). Orthophosphate ions released by the action of 5′-nase may be either immedi-
ately taken up by bacteria producing this enzyme or mixed with the bulk water. The fate
of the released Pi strongly depends on ambient Pi concentration in the environment and
on Pi demand of microplankton. In oligotrophic and other Pi-poor waters, release of Pi
from 5′-nucleotides by 5′-nase is tightly coupled with its uptake and more than 50% of
released Pi can be taken up by the bacteria (3,19). Under high Pi concentrations, in highly
polluted waters or in deep regions of the lakes, however, only 10–15% of the Pi resulting
from 5′-nase activity is assimilated by microorganisms; an excess of enzymatically liber-
ated Pi mixes with the existing Pi in the bulk phase (67,107,124,125).
Estimation of the quantitative contribution of APase and 5′-nase activities to Pi
release into aquatic environments is difficult. Studies of various DOP compounds suggest
that P nucleotide generally is assimilated by aquatic bacteria more efficiently than P bound
to other phosphate esters. Moreover, comparative studies on kinetic parameters of both

enzymes showed that liberation of Pi by 5′-nase appears more efficient than Pi release by
Apase, especially in Pi-rich environments (Table 3) (107,117). However, it should be
stated that in situ 5′-nase activity may be substantially affected by the rate of 5′-nucleotide
Table 3 Comparison of the Turnover Time of 5′-
Nucleotidase Substrates (Adenosine Monophosphate,
Adenosine Triphosphate, and Alkaline Phosphatase Substrate
glucose-6-phosphate) in Surface Water Samples from
Mesotrophic Lake Constance
Alkaline
5′-Nucleotidase phosphatase
Method AMP ATP G6P
32
P 1.11 0.45 12.22
Colorimetric 0.30 2.40 19.20
AMP, adenosine monophosphate; ATP, adenosine triphosphate; G6P,
glucose-6-phosphate.
Source: Recalculated data from Ref. 107.
Copyright © 2002 Marcel Dekker, Inc.
supply(117)andthathydrolysisofextracellularnucleicacidsmustberegardedasthe
limitingstepfortheenzymaticregenerationofPifromthenucleotidepool.
C.OtherPhosphohydrolases
Asubstantialpartofthehigh-molecular-weightDOPfraction(Ͼ10,000D)isthoughtto
becomposedofinositolphosphatesboundtoproteins,lipids,orfulvicacid.Moreover,
someoflow-molecular-weightDOPcompounds(Ͻ1,000D)inaquaticecosystemsfre-
quentlyhavebeenidentifiedasinositolphosphates(126–130).Thesefindingssuggest
thatphytasemaybeanother(probablyadaptative)phosphohydrolyticenzymethatpoten-
tiallymaybeimportantinPireleaseprocesses.Herbesandassociates(127)foundthat
upto50%ofDOPinlakewaterwashydrolyzedbyphytaseunderitsoptimalpHcondi-
tions.Ontheotherhand,studiesofDOPdecompositioninthenaturalpHoflakewaters
(pH7.2–8.6)showedthatphytaseactivitydecomposedonly15%ofthetotalDOPwithin

30days(129).SincephytasehasanoptimalpHaround5.0(127),itsactivitymaypresum-
ablysupportPireleasemainlyinnaturallyorartificiallyacidifiedenvironments.
Aswithphytase,theecologicalsignificanceofcyclicnucleotidephosphodiesterase
ispoorlyelucidated.AccordingtoBarfieldandFrancko(69),cyclicnucleotidephospho-
diesteraseactivityinlakewaterisaresultofactivitiesofagroupofseasonallydifferent
isoenzymeswithanoptimalpHbetween7.0and8.0.Althoughparticipationofthisen-
zymeinPireleaseinlakesseemstobealmostinsignificantfromaqualitativeperspective,
thecyclicnucleotidephosphodiesteraseprobablyisoneofthemostimportantenzymes
controllingcAMPconcentrationinaquaticenvironments,thuspotentiallyaffectingavari-
etyofphysiologicalprocessesofmicroplanktonmediatedbycAMP(42).
D.EnzymaticReleaseofPiinLakeWater—Conclusions
SincetheamountofreadilyassimilablePiinthemajorityofnonpollutedlakesdoesnot
fulfillPrequirementsformicroplankton,itmustbereleasedbymicroorganismsfrom
organicPcompounds.Forthispurpose,aquaticmicroorganismsdevelopedtwomainen-
zymaticPireleasesystems(Fig.9).
The first, adaptative mechanism, activated relatively rapidly (by induction/derepres-
sion) during Pi limitation periods, is based on activity of nonspecific phosphomonoester-
ases and (APases) produced by almost all members of the plankton community. In eutro-
phic environments, this mechanism is mediated mainly by algal phosphatases. In
oligotrophic and mesotrophic ecosystems, however, APase of bacterial origin seems to
play a more important role. As APase activity is strongly dependent on dynamically chang-
ing Pi concentrations in the environment, its participation in Pi release processes in lake
water is restricted to the trophogenic zone of the lake and short intervals of Pi depletion
during the summer stratification period.
The second mechanism, exclusively bacterial, involves interaction of various types
of nucleases and 5′-nucleotidase that liberate Pi from nucleic acids and from nucleotides.
Release of Pi from nucleic acids must be preceded by endo- and exonuclease reactions
that liberate 5′-nase substrates (5′-nucleotides). Although 5′-nucleotides can be hydrolyzed
by both APase and 5′-nase, it seems that the role of APase in their decomposition is of
minor importance. Orthophosphate release is mediated by nucleases and 5′-nase probably

is constitutive in the majority of aquatic environments. And those processes are more
Copyright © 2002 Marcel Dekker, Inc.