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

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-

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

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

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

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

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

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,

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

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-

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).

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.

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-

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)

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

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

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