A review of biodegradation kinetics in the aquatic environment

Vol.21,Nos.lO-11, Chemosphere, Printed in Great Britain

pp 1243-1284,

1990

0045-6535/90 $3.00 + -00 Pergamon Press plc

A REVIEW OF BIODEGRADATION KINETICS IN THE AQUATIC ENVIRONMENT N.S. BATTERSBY* WRc Medmenham, PO Box 16 MARLOW, Buckinghamshire SL7 2HD, U.K.

ABSTRACT This paper highlights the need for environmentally-realistic data on the kinetics of biodegradation in the aquatic environment and reviews many of the published kinetic expressions. The unsuitability of current Organisation for Economic Co-operation and European Economic Community Sixth Amendment "ready biodegradability" test procedures for generating such data is shown and an experimental approach for producing environmentally-realistic kinetic data proposed.

l.INTRODUCTION Over the past thirty years, synthetic pollution caused by persistent chemicals has highlighted the need for data on their fate in the environment, and led to national and international legislation aimed at providing this information. Much of this effort has concentrated on the fate of a chemical in the aquatic environment and during waste water treatment processes (which often lie on the route between the place where a chemical is used or spilled and the receiving water). The distribution and concentration of an organic chemical released into the aquatic environment will be determined by its physico-chemical properties (e.g. water solubility, vapour pressure, octanolpater partition coefficient) and a variety of non-biological and biological removal mechanisms. Although removal from an environmental compartment can occur by abiotic processes such as hydrolysis, photolysis, adsorption onto particulate matter and volatilisation into the atmosphere, the complete conversion of an organic chemical to principally inorganic products is invariably due to microbial activity. This process, termed "ultimate" biodegradation, results in its conversion into Cop, H20, inorganic salts, new microbial biomass and organic products associated with the normal metabolic processes of bacteria [l]. It is now nearly a decade since the Organisation for Economic Co-operation and Development (OECD) first published their guidelines [2] for testing the susceptibility of chemicals to biodegradation ("biodegradability"). Their testing strategy was later adopted by the European Economic Community (EEC) as part of a pre-marketing, notification scheme for assessing the potential hazard of new chemicals to man and the environment [3].

* Present address: Shell Research SITTINGBOURNE, Kent ME9 SAG U.K.

Limited,

Sittingbourne

1243

Research

Centre,

1244

Within the member states of the EEC the "Sixth Amendment Directive" has made the assessment of biodegradability a statutory requirement for any manufacturer or importer wishing to place over i tonne/year of a new chemical on the market. For most compounds this will involve a test for "ready biodegradability" [4] - a simple, stringent, screening test in which the attainment of a pass level of biodegradation implies that that substance should be degraded rapidly in the environment [5]. The aims of this paper are to highlight the current need for information on the rate (as opposed to extent) of biodegradation of an organic chemical in the aquatic environment and to review the various kinetic expressions used to describe that rate. The effect of test conditions on the kinetics of biodegradation and the unsuitability of current OECD and EEC screening test procedures for generating such data will be discussed, and an experimental a p p r o a c h for p r o d u c i n g e n v i r o n m e n t a l l y realistic kinetic information outlined.

2. THE NEED FOR EXPRESSIONS WHICH DESCRIBE THE RATE OF BIODEGRADATION IN THE AQUATIC ENVIRONMENT

Standard methods for determining the biodegradability of a compound in the aquatic environment produce results in terms of percentage removal after a fixed time period [2, 4]. Although this can be useful for labelling purposes (e.g. "readily biodegradable"), it is of limited use when attempting to predict the fate of a substance in the environment. There is a requirement for quantitative expressions which describe the rate of biodegradation. Such data are important components of mathematical models for predicting the distribution and concentration of synthetic chemicals in the environment, and enable biodegradation to be ranked against other competing abiotic removal processes [6-12]. The use and validation of many of the models used for predicting the fate of synthetic chemicals in aquatic ecosystems is discussed in greater detail in [13, 14]. In order to predict accurately the fate of a synthetic chemical in the aquatic environment it is important that biodegradation rates derived from the observation of microbial activity in the laboratory are sufficiently independent of the experimental conditions so as to allow extrapolation to a variety of environmental conditions. Such measurements have been termed "real world" biodegradation rates by Howard [15], who emphasises that microbial activity observed in the laboratory should relate to that occurring in the environment. Quantitative measurements of b i o d e g r a d a t i o n w h i c h w e r e test s y s t e m independent would also be useful in developing better relationships between molecular structure and the rate of biodegradation. Reliable quantitative structure-biodegradability relationships (QSBR) could enable biodegradation rates to be predicted when measured values were unavailable. This would be useful not only for environmental fate models but also during the initial screening of a range of novel chemicals for environmental acceptability [16]. Unfortunately, there is a paucity of suitable biodegradation rate data - much of the available data are from screening tests where the results are highly dependent on the test protocol and show poor reproducibility [17 1 . Unless studies on a homologous series of compounds tested under the same conditions are chosen, test conditions as opposed to chemical structure, can account for any variance in biodegradability [18]. In many QSBR's there is a

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tendency to elaborate the descriptors of molecular structure improve the quality of the biodegradability data used.

and

not

to

T h e s e a p p r o a c h e s n e c e s s i t a t e an i n v e s t i g a t i o n into the k i n e t i c s of biodegradation reactions in an attempt to produce rate equations which describe the order of the reaction and yield a specific reaction rate or rate constant (k) which can be incorporated into the mathematical model. Such s t u d i e s can also lead to an u n d e r s t a n d i n g of h o w the rate of biodegradation is a f f e c t e d by c h a n g e s in r e a c t i o n c o n d i t i o n s (i.e. conditions existing in the tests and in the aquatic environment).

3. KINETIC EXPRESSIONS FOR DESCRIBING BIODEGRADATION PROCESSES

The k i n e t i c s of b i o d e g r a d a t i o n h a v e b e e n d e s c r i b e d by a v a r i e t y of mathematical expressions, increasing in complexity as they attempt to accommodate the numerous variables which can affect the rate of biological removal of an organic chemical in the aquatic environment. Much of the published work on the determination of biodegradation rates has involved relatively simple experimental procedures which produce data on the rate of decrease in test substance or increase in degradation products (e.g. CO2). There is evidence to suggest that complex equations have been constructed in an attempt to explain a set of minimal data - a better approach would be to use improved experimental methods to follow more closely the progress of bio-degradation. However, this is easier said than done and some of the practical problems involved in the design of experiments to provide kinetic constants will be discussed later. A review of some of the more commonlyused expressions which have been used to describe biodegradation reactions is given below.

3.1 First-order reactions

If biodegradation is a first-order reaction, then the rate of biodegradation (v) will be proportional to the concentration of only one reactant the test chemical:

d[S] v

~

k I [S]

(I)

dt

where:

-d[S]/dt is the rate of test substance disappearance, t is time, k 1 is the first-order rate constant and [S] is the concentration of test substance in the system.

First-order rate constants are usually derived from experiments where a test chemical is added to an environmental sample and its disappearance with time monitored by specific analysis (or by loss of radioactivity if the compound was radiolabelled). A plot of loglo (initial [S]/[S] remaining after time t) vs___~, t yields a straight line with a slope of ki/2.303. Although k I is usually determined by linear regression, it can also be determined directly by non-linear regression analysis of the data [19]. Non-

1246

linear techniques can be modified to include asymptotes biodegradatlon curve reaches a plateau which is <100% of amount) and lag phases for biodegradation [20].

(e.g. where the the theoretical

Larson [19, 20] has also applied first-order kinetics to determine the rate of product (C02) formation using the expression:

r _J

0

for t ! c

a {I - exp [-kl(t-c)]}

for t > c

(2)

P

I where:

P is the product formed (C02) , a is the extent of biodegradation (asymptote) and c is the lag time prior to biodegradation.

First-order rate constants can be used to calculate a half-life test substance using the relationship:

th

-

in2 -kl

(t½) for the

(3)

However, care should be exercised when using the half-life concept to d e s c r i b e the b i o l o g i c a l r e m o v a l of a chemical. In o t h e r f i r s t - o r d e r processes (e.g. radioactive decay), th is independent of concentration and the time required for half the remaining increment to disappear is constant regardless of which stage of the reaction is observed. However, during biodegradation this is often not the case as at very low concentrations the rate of biodegradation can limited by the rate of entry of substrate into the cell (Section 5). First-order kinetics have been widely used to describe the biodegradation of organic chemicals in aquatic systems and some of the reported values are given in Table I. It is probably more accurate to describe many of these reactions as being pseudo first-order due to the participation of bacteria in the reaction. First-order kinetics are usually verified by demonstrating that k I does not vary over a wide (>10-fold) range of test chemical concentration [20]. However, if there is a constant concentration of biomass [B] in the test system, as is often the case with short incubation periods and a low test chemical concentration, then the reaction will appear to be first-order overall. However, the reaction may in fact be second-order with: v

-

k [S] [B]

V a r i a t i o n s in the q u a n t i t y a n d / o r q u a l i t y of b i o m a s s in environment are probably the cause of much of the reported differences in k I values shown in Table I.

(4)

the a q u a t i c between-site

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Table

i. Some published first-order rate constants for the blodegradatlon synthetic chemicals in n a t u r a l waters.

kl Chemical

(d "l)

Sample

Reference

Benzene

0.12 a

River

[21]

Cetyltrimethyl ammonium chloride

0.26

River

[22]

Chlorobenzene

0.07-0.3 b 0.04-0.2 b 0.01 b

River Estuary Marine

[23] [23] [23]

4-Chlorophenol

0.035 a

River

[21]

1.7 c 0.8-4.7 c 2.8-4.8 c

River Estuary Marine

[24] [24] [24]

Dioctadecyldimethyl ammonium chloride

0.05

River

[25]

Dodecyl nonylethoxylate

0.45

River

[20]

Hexadecyltriethoxylate

0.34

Estuary

[26]

Hexadecyltriethoxylate sulphate

0.32

Estuary

[26]

O. 30 0.02

River sediment River water

[27] [27]

0.93 0.32 0.26

River Estuary Ground water

[25] [28] [20]

C12 Linear alkylbenzene sulphonate (LAS)

0.05 0.5

River (above STW) River (below STW)

[29] [29]

1,2,4-Triehlorobenzene

0.029 b 0.026 b 0.012 b

4-Cresol

Methyl parathion

Nitrilotriacetic (NTA)

STW a b c

acid

River Estuary Marine

Sewage treatment works Calculated from t% using equation (3) Calculated using k I - Vmax/K m (Section 3.3) k I calculated after a lag period

[23] [23] [23]

of

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3.2 Monod growth,

co-metabollsm

and second-order kinetics

If the assumption is made that growth is a continuous process, and that cell mass is produced during substrate utilisation, then the rate of growth of a bacterium growing on a single, limiting substrate can be described by the following equation first put forward by Monod [30]:

I

d [B]

[B]

dt

pmax

[S]

(5)

where:

(K s + [S])

[B] is the concentration of bacteria, ~ is the specific growth rate of the bacterium degrading the sub~trate, ~max is the maximum specific growth rate, [S] is the concentration o f substrate and K s is a constant numerically equal to [S] at 0.5 ~max (i.e. the substrate concentration at which growth occurs at h a l f the maximum rate).

This relationship between bacterial growth rate and substrate concentration is shown in Figure IA. It can be seen that growth rate is a hyperbolic function of substrate concentration and approaches a maximum (~max) at high concentrations. Equation (5) has been widely used by microbiologists and describes the pattern of increase in biomass when a small number of bacteria are added to a medium containing relatively a high concentration of the growth substrate and non-limiting amounts of inorganic nutrients. As this equation describes the increase in bacteria as a function of substrate concentration it can be v i e w e d as an indirect m e a s u r e of s u b s t r a t e utilisation. If the assumption is made that a fixed amount of growth results from the metabolism of a unit quantity of substrate, then a factor "Y" (the yield coefficient which measures the efficiency of conversion of test chemical into cells) can be incorporated into equation (5) to give:

d [S]

~max

[B] [S]

(6) dt

Y (K s + [S])

This expression has been successfully used to describe the rate of removal of synthetic chemicals in laboratory studies where the chemical was the sole source of carbon and energy [31]. However, the limitations of using equation (6) to model biodegradation rates in aquatic systems should be noted. The equation assumes that microbial growth occurs on the test chemical and that this chemical is the single growth-limiting substrate in the system. In the environment growth may also be due to other substrates which may, or may not be limiting (see Section 5). This has lead Baughman et al [32] to conclude that the value of the yield coefficient Y cannot be readily determined from laboratory studies. The use of Monod kinetics also assumes that Y is a constant (at around 0.5). However, there are data which indicate that at low concentrations a large amount of the energy derived from the biodegradation of a chemical is used for maintenance energy requirements and not for growth [19].

1249

~m~lx

I;D

Ks

Substrate concentration [S]

Figure 1A. Effect of substrate concentration on bacterial growth rate.

V ax.

c" O

Vmax Zero-order - reaction

2

n-

First-order reaction K m

Substrate concentration [S]

Figure 1B. Effect of substrate concentration on the rate of an enzyme-catalysed reaction.

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Laboratory studies have also shown that many synthetic chemicals can be transformed by bacteria which cannot use them as a sole source of carbon and energy. This phenomenon is termed co-metabolism (sometimes called cooxidation) and has been defined by Dalton and Stirling [ 3 3 ] as: "the transformation of a non-growth substrate in the obligate presence of a growth substrate or other transformable compound". In this definition, "nongrowth substrate" refers to a compound which is unable to support cell replication (as opposed to an increase in biomass). Co-metabolism is probably due to the lack of specificity of certain microbial enzymes and has important consequences in terms of persistence of an organic chemical in the environment as can be seen from Figure 2. As the test chemical is not u t i l i s e d for b i o s y n t h e t i c p u r p o s e s there is no i n c r e a s e in the com e t a b o l i s i n g p o p u l a t i o n w i t h time as a r e s u l t of the c h e m i c a l b e i n g introduced into the system (i.e. adaptation by growth of specific degraders does not occur). The low density of these organisms means that a cometabolised chemical is usually transformed at a very slow, linear rate - in contrast to growth-linked mineralisation.

MINERALI$1NG CO-METABOUSING POPULATION

f

POPULATION

i

I

.IO0

/

Chemical

/ 9

he

8

al

Oo!

2 //

I

!

/ Populal~on

! t

Popul.~tion

I

Trne

Figure 2. Model illustratingthe populationchanges and removal of a chemical degraded by growth-linked mineralising and co-rnetabolisingmicrobial populations [34].

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The products of co-metabolism are usually structurally similar to the parent chemical and the process often does not result in detoxification. There is a paucity of direct evidence of co-metabolic processes occurring in the aquatic environment [34], a l t h o u g h data r e p o r t e d by W a n g et al [35] suggested that isopropyl N-phenylcarbamate was mineralised to CO 2 in lake water amended with 400 ng/l of the compound but was transformed by cometabolism to unidentified organic products at a much higher concentration (1 rag/l). As a c o - m e t a b o l i s e d c h e m i c a l does not s u p p o r t cell r e p l i c a t i o n , its degradation cannot be modelled by equation (6) as ~max and Y must be defined as zero [36]. Banerjee et al [37] have suggested that this problem can be circumvented by assuming that a co-metabolised chemical is utilised by the cell in an immeasurably small but finite amount. This allows the two kinetic parameters to be defined as small but non-zero, enabling equation (6) to be extended to co-metabolic processes. In most aquatic environments the concentration of a synthetic chemical will be low (< ~g/l) and much lower than Ks, which is typically of the order of 0.i to i0 mg/l. Under these conditions equation (6) can be approximated by equation (7) which takes into account the dependence of biodegradatlon rate on both substrate and active biomass concentration:

d[S]

~max

dt

Y Ks

[S] [B]

-

k 2 IS] [B]

(7)

where: k 2 is the second-order rate constant (equivalent to ~max/YKs - this equivalence has been shown in test systems where the test chemical was the sole source of carbon [38, 39]). Second-order rate constants are usually determined in die-away experiments, with a pseudo first-order rate constant (k[B]) being calculated as described in Section 3.1. The second-order rate constant k 2 is then calculated by dividing k[B] by the concentration of bacteria [B], normalising the observed pseudo first-order rate constants for biomass [20, 40, 41]. [B] has usually been quantified in terms of colony-forming units (cfu) determined by standard plate counts, although dry weight [42] and adenosine triphosphate [32] measurements have been used. When second-order kinetics are used to describe a biodegradation following assumptions are made: a) Plate counts (or other enumeration methods) the total bacterial population in different b) All, or a c o n s t a n t proportion, transforming the chemical.

of

process

detect similar percentages samples.

the

enumerated

bacteria

c) Transformation rates per cell (or average rates per microbial are approximately the same for all sites examined.

the

of

are

assemblage)

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Table 2. Mean k 2 values for the primary degradation of three pesticides in various North American waters [40].

Reaction

(I

OCH2COOC2H4OC4H9

cell ~

h'l) a

No. of sites b

OCH2COOH

Cl Butoxyethyl

Cl

ester of 2,4-D

5.4 + 0.97 x i0 "I0

31

4.5 + 0.74 x i0 "II

14

c~3o s cooc2 5 /

~H2COOC2H5

CH30

cH3o / k 2 CH30

8 ooo 2H5

CH2COOH

Malathion

HNCOOC3H 7

CI Chlorpropham

NH2

k2

CI 2.6 ± 0.72 x 10 -14

ii

a ± 95% Confidence intervals based on between-site variation. b The characteristics of the waters examined ranged from: Temperature (1-29 °C), pH (5.4-8.2), total hardness (10-420 mg CaC03/I), total N (trace-10 mg/l), total P (0.02-1.0 mg/l), total organic carbon (1.6-28.8 mg/l) and b a c t e r i a e n u m e r a t e d by i n c u b a t i o n for 48 h on tryptone-glucose-extract agar (4.8 x 105-9.8 x 108/1).

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Data published by Paris et al [40], and summarised in Table 2, indicate that the second assumption can hold as the small between-site variation in k 2 values indicates that the proportion of bacteria capable of hydrolysing malathion, chlorpropham and the butoxyethyl ester of 2,4dichlorophenoxyacetic acid (2,4-D) was relatively constant in the different microbial communities studied. These three p e s t i c i d e s are r e a d i l y transformed by hydrolytic enzymes which are constitutive in most bacteria and it would appear that the concentration of bacteria enumerated as cfu was related to the concentration of these enzymes. Similar small between-site variations in k 2 values for the primary biodegradation of phenol, parasubstituted phenols and various esters of 2,4-D in North American fresh waters were reported in later papers by the same group [43-45]. It was also found that k 2 increased as the alkyl length of the 2,4-D ester increased and could be correlated with log Kow [45]. However, Rogers et al [46], using essentially similar techniques to those reported in [40], found that k 2 for the transformation of the butoxyethyl ester of 2,4-D in other North American river waters (four rivers at seven sites) were two to three orders of magnitude higher, ranging from 1.6 x 10 .8 to 2.1 x i0 -7 i cells "I h "I. Similar differences were observed when k 2 values for 4-cresol were compared with those reported in [44]. In studies on aufwuchs samples (aquatic microbial growth attached to submerged surfaces) Lewis and Holm [47] found that k 2 for the transformation of methyl parathion and diethyl phthalate decreased as [B] (determined by standard plate counts) in the aufwuchs increased. This was thought to be due to the proportions of non-transformer bacteria increasing as [B] increased (cf. assumption b, above), and to a reduction in the rate of transformation by the specific degraders as [B] increased (cf. assumption c). In contrast to Paris et al [40], the authors found that division of the first-order rate constant k I by aufwuch surface area yielded a second-order rate coefficient ksa which showed less variation than k 2 among aufwueh samples. It is thought that substrate depletion near the surfaces of attached bacteria leads to the transformation of rapidly degraded chemicals being mass transport limited. Under such conditions, removal rates are determined by water turbulence near the biofilm surface. In a later paper [48], Lewis and co-workers derived a second-order transformation rate coefficient (ka) based on the rate of test chemical loss and a ratio of periphyton-colonised surface area to container volume at a Reynolds' number of 6 000 (Reynolds' number is dimensionless and characterises the type of liquid flow in a pipe flowing full, where the resistance to motion depends on the viscosity of the liquid and the influence of inertia). Values of k a were relatively constant under similar conditions of turbulence for several chemicals using bacteria from streams, rivers and laboratory microcosms. As an example, k a for methyl parathion was 0.92 ± 0.51 i m °2 h "I for five field sites and 1.5 ± 0.57 i m "2 h -I for 12 microcosm samples. From the preceding examples it can be seen that it certain cases secondorder transformation constants can show little between-site variation and may be of use in p r e d i c t i n g the e n v i r o n m e n t a l behaviour of e a s i l y hydrolysable chemicals transformed by free-floating or attached microorganisms (this latter case is discussed in detail by Lewis and Gattie in [49]). However, this is unlikely to be the case with recalitrant molecules or for "ultimate" biodegradation.

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In addition, research on the changes in bacterial numbers and activity in estuarine waters has shown that heterotrophic bacteria are not uniformly active, with the changes reflecting physiological differences which may be correlated with habitat changes [50]. For many synthetic chemicals then, normallsation of pseudo first-order rate constants for biomass using plate or direct counts will not take into account the activity of those specific degraders which are p e r f o r m i n g the transformation reaction under investigation.

3.3 Michaells-Menten kinetics, activity indices

heterotrophlc

uptake studies and specific

In the previous section we saw that the rate of bacterial growth under certain conditions is a hyperbolic function of substrate concentration. If the initial rate of an enzyme-catalysed reaction (e.g. the transformation of a synthetic chemical) is plotted against the concentration of substrate [S], a similar rectangular hyperbolic plot is obtained (Figure IB). The rate of this reaction (v) can be described in an analogous way to equation (5) using the general equation first proposed by Michaelis and Menten in 1913 [51]:

d[S] v

-

Vma x [S] (8)

=

(K m + I S ] )

dt

where: Vma x is the maximum initial velocity of the reaction for a given concentration of enzyme, [S] is the concentration of substrate (test chemical) and K m is the Michaelis constant numerically equivalent to the substrate concentration at which the reaction occurs at half the maximum rate. Environmental concentrations of most synthetic chemicals ([S]) will be much lower than K m which for soluble suhstrates in activated sludge is in the region of 5-10 mg/l [52]. Under these conditions equation (8) reduces to: Vma x [S] v

(9) Km

The expression Vmax/K m has the dimension time "l , and if both kinetic parameters are constant, then substrate removal will follow flrst-order kinetics with k I - Vmax/K m. However, the expression Vmax/K m represents a pseudo first-order rate c o n s t a n t as Vma x is d e p e n d e n t u p o n enzyme concentration, which in turn is affected by the concentration of active bacteria in the system. Michaells-Menten kinetics were first used over 25 years ago by Parsons and Strlckland [53] to describe the uptake of radiolabelled substrates by marine bacteria. This work was extended by Wright and Hobble [54, 55] to enable the calculation of turnover times for the naturally-occurring concentration of a given suhstrate (the turnover time being the time required for the existing microbial population to take up and/or respire an amount of substrate equal

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to the natural concentration). Pfaender and Bartholomew [56] later modified the heterotrophic uptake technique to provide a rapid and relatively simple method for determining the biodegradation rates of 14C-labelled synthetic chemicals. Their technique measured both the assimilation of l~c into microbial cells and 14C02-evolution in experiments where the water sample was incubated for a short period of time (usually < 12 h) in the presence of a low (~g/l) concentration of the test chemical. The resulting data can be used to calculate turnover times (T):

T

-

t. (- d ! i .origin_______ally a__dded ~

(i0)

~dpm in cells + dpm 1 4 C 0 2 / where: t is the time in h.

The rate of biodegradation (v) can be calculated using the equation:

v

where:

-

[S] ~ _ p m in cells + dpm 14C0_2~ ~ . . . . . . . . . t % % dpm originally a d d e d J

(II)

[S] is the concentration of test chemical added to the system.

If different concentrations of the test chemical are used, the kinetic parameters Vma x and K m (which incorporate both the activity of the microbial population and the effect of substrate concentration) can be calculated using a Lineweaver-Burke plot of i/v against I/[S]. The resulting straight line has a slope of Km/Vmax, an x-intercept of -I/K m and a y-intercept of i/Vma x [51]. Other approaches for calculating the two kinetic parameters can be found in [20, 41, 51]. However, care should be taken if the persistence of a synthetic chemical is predicted using Vma x values as this kinetic p a r a m e t e r is an e x t r a p o l a t e d value a s s o c i a t e d w i t h h i g h s u b s t r a t e concentrations which may not be attained in situ [28, 56]. Pfaender and Bartholomew [56] used the heterotrophic uptake technique to measure the biodegradation of 3-cresol, chlorobenzene, NTA and 1,2,4-trichlorobenzene in fresh water, brackish and marine water samples. Examples of the total uptake Vma x values obtained were II0 (freshwater), 20 (estuarine) and 0.98 (marine) ng I "I h "I for 3-cresol; and 17 (freshwater), 14 (estuarine) and 0.03 (marine) ng 1 -I h -I for chlorobenzene. These data indicated that metabolic rates for the two compounds were significantly lower under marine conditions than in the freshwater and brackish regions of the river under study. The same paper reported the results of a year-long study of biodegradation at the estuarine sampling site. T (h "l) and Vma ~ (ng 1 "I h -1) values ranged from 0.0045 to 0.022 h -I and 0 to 32 ng 1 -I h "I for 3-cresol; 0.0002 to 0.006 h -I and <0.i to 14 ng 1 "I h -I for chlorobenzene; and 0.0013 to 0.014 h -I and 850 to 2 300 ng 1 -I h -I for NTA. The rates of degradation of all the chemicals were slower in winter, however it was unclear whether these changes were due to lower water temperatures or to other seasonal factors. A later study using the same chemicals and river

256

r e p o r t e d e s s e n t i a l l y s i m i l a r results and d e m o n s t r a t e d that no simple correlation existed between 3-cresol degradation rates a n d m e a s u r e d characteristic of the microbial community such as direct and viable counts, and the rate of amino acid turnover [23]. Pseudo first-order rate constants taken from this work were given earlier in Table i. A quantitative expression of the relationship between heterotrophic activity and the bacteria responsible for that activity has been described by Wright [57]. The division of Vma x by the total number of bacteria present in the sample yields a "specific activity index" (SAI) with units of weight cell -I h "I. Wright proposed that SAI values could serve as indicators of the physiological state and m e t a b o l i c role of b a c t e r i a in the a q u a t i c environment. This approach was used by Larson and Ventullo [28] to assess the effects of salinity and dissolved organic carbon (DOC) on the kinetics of NTA biodegradation in a Canadian estuary with a history of NTA exposure. Rapid degradation of NTA was observed over a range of salinities (4-19 °/oo) and DOC levels (2-12 mg/l), with the rate of degradation being a saturable function of NTA concentration (mean Vma x over several sampling periods was 4.8 ± 2 . 8 5 ~ g I "I h-l). SAI values determined using acridine orange direct counts (AODC) varied by a maximum of ten-fold during different sampling periods (201-2 753 x 1 0 " 1 2 ~ g cell "I h -1) and by less than three-fold within a given sampling period. The variation in SAI was similar when calculated using an estimate of the numbers of specific NTA-degraders. As the numbers of specific degraders was the denominator (low compared to the total number of bacteria determined by AODC), the resulting SAI were several orders of magnitude higher and ranged from 4 to 12 x 10 -6 ~ g cell -I h -I during a sampling period. SAI (AODC) values were relatively insensitive to salinity and DOC changes, and the authors c o n c l u d e d that these e n v i r o n m e n t a l parameters had little effect on NTA degradation at the individual cell level in the estuary studied. In a study on the degradation of dodecyl [U-14C] nonylethoxylate in Ohio river and ground waters, Larson [20] analysed his biodegradation data using first-order, second order and heterotrophic bacterial activity kinetics. Saturation kinetics were observed in samples from both sites, with K m values being similar in both systems. However, the mean degradation rate in the ground water was significantly lower than in the river water (Vma x = 121 cf. 8 081 ng i -I h-r'). Degradation activity expressed as Vma x was found to be proportional to the number of viable bacteria as determined by plate counts on I/i0 to 1/100-strength nutrient agar. SAI values (Vmax/Cfu) for the ground and river water sites were similar at around 3.0-3.4 x 10 -5 ng cfu -I h "I. This variation was much less than that for the second-order rate constants k2, which varied by around 800%. Earlier work by Ladd et al [58] on the degradation of naturally-occurring substrates (glutamic acid, phenylalanine and glycollate) in Canadian ground waters and streams had shown that the large between-site differences in Vma x values were reduced when they were normalised for microbial numbers (AODC). For example, Vma x values for phenylalanine between the two sites varied by over 200-fold compared with four-fold for SAI. One or two words of caution should be made concerning the use of MichaelisMenten kinetics to predict the rate at which micro-organisms take up and degrade an organic chemical. The possibility that Vma x may not be attained in situ has already been mentioned. In addition, the assumptions must be made that the concentration of enzyme remains constant during incubation,

1257

the bacterial transport systems are responding only to the substrate, and a single set of kinetic parameters governs the processes of degradation (i.e. Vmax/K m does not vary over a wide range of [S]). Care must be taken when interpreting such data as growth of the active bacteria may occur during incubation, and the presence of other substrates in the test system (see Section 5) may affect the results. In addition, the last assumption may well be wrong, as microbial communities often possess a variety of substrate removal systems which operate optimally at different substrate concentrations and large errors (up to four orders of magnitude) can occur if kinetic parameters determined at very high substrate concentrations (g/i) are used to predict transformation rates at low ( ~ g / l ) concentrations [59].

4, CHOOSING THE MOST APPROPRIATE KINETIC MODEL TO APPLY TO A SET OF BIODEGRADATION DATA

It has already been seen that biodegradation can be modelled using a variety of kinetic expressions. Two useful approaches for determining the most appropriate kinetic expression to apply to a set of biodegradation data are discussed below and are based on either the Monod equation for microbial growth (equation 5) or the Michaelis-Menten equation for an enzyme-catalysed reaction (equation 8).

4.1 The integrated Monod equation Simkins and Alexander [60] have proposed that the variety of substrate disappearance curves encountered in biodegradability testing is the result of interactions between substrate concentration IS] and population density [B]. The authors assume that Monod kinetics (equation 5) adequately describe the r e l a t i o n s h i p between bacterial growth rate (~) and substrate concentration IS]. To accommodate changes in population density, a massbalance equation is introduced which describes the relationship between c h a n g e s in p o p u l a t i o n size and the c o n c o m i t a n t c h a n g e s in s u b s t r a t e concentration:

(12)

[So] + q[Bo] = [S] + q[B] where:

[So] is the initial concentration of substrate, [B0] is the initial concentration of bacteria and q is the cell quota (equivalent to l/Y)

The factor qB is then replaced by X which corresponds to the amount substrate required to produce a population density equal to [B], with then being equal to q[B0] (for reasons see [60]). The Monod equation

(5) and equation

1

dE

X

at

(12) can then be re-written

fmax

of X0

as:

IS] (13)

[So) + X 0

(K s + IS])

=

[S] + X

(14)

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Manipulation of equations (13) and (14) yields a differential equation which is a general expression of substrate disappearance in a system where the k i n e t i c s of b i o d e g r a d a t i o n are d e t e r m i n e d by p o p u l a t i o n d e n s i t y and substrate (i.e. test chemical) concentration:

d[S]

)Jmax

[s]

([So]

+ x o - [s]) (15)

dt

(K s + IS])

Simkins and Alexander term Extreme ratios of inoculum substrate concentration to six simplified forms shown

expression (15) the "integrated Monod equation". density to initial substrate concentration, or of Ks, allow this equation to be approximated by the in Table 3.

The conditions under which the six models apply is best explained by reference to Figure 3A. In this example, K s - I mg/l and q (the amount of substrate required for the formation of one bacterial cell) is i pg/cell. Points along the diagonal llne correspond to inoculum sizes which allow one division of the active bacteria at various initial substrate concentrations. If the initial cell density at a given test chemical concentration is above this line, then cell density can be treated as approximately constant (i.e. quantity of substrate is insufficient to support a significant increase in cells). The vertical line at i mg/l is equivalent to Ks; the broken vertical line is at a concentration one and a half orders of magnitude greater than K s and separates situations where the biodegradation rate per bacterium varies a p p r e c i a b l y w i t h substrate c o n c e n t r a t i o n (left of line), from c i r c u m s t a n c e s w h e r e the u p t a k e systems of the active d e g r a d e r s are effectively saturated until the substrate is nearly exhausted.

Table 3. Models for blodegradatlon kinetics using the variables concentration and bacterial cell density [60].

Model

A First-order

B Logistic

C Monod (no growth)

D Monod (growth)

-d[S]/dt =

Rate constant (units)

kl[S]

kl = ~max X0/Ks (h-I)

k 3 [ S ] ( [ S 0] + X 0 - IS])

k 3 - ~max/Ks (i h "I mg "I)

k 0 [S]/(K s + [S])

ko = ~max XO (mg i -I h -I)

Equation (15)

E Zero-order

F Logarithmic

of substrate

ko

~max([So]

+ Xo - [s])

~max (h-I)

ko - ~max XO (mg i -I

~max (h-I)

h -1)

1259

E

12

a Zero- J ', order

C Mo,od

.¢..

10

A .Q .c_ O

First-order

D

8

Logarithmic

Monod (growth)

o,

Uptake systems of active bacteria saturated J

i

i

1#g/I

1 mg/i

Figure 3A. KineUc models as a funclon of ini~al substrate concentration and bacterial cell density [60].

100 " " " " " "-L:,.

D

F

o) t."e

E P

50

"lime Figure 3B. Disappearance curves for a chemical degraded according to the IdnetJc models shown above [61 ].

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At substrate concentrations below K s (as would be the case for most synthetic chemicals in the aquatic environment) degradation can be described by two kinetic expressions. In Zone A of Figure 3A there is no significant increase in bacterial numbers during degradation (i.e. constant biomass concentration) and first-order kinetics can be applied. In Zone B the initial small population of active bacteria grow on the test chemical at a rate which falls constantly with its diminishing, and always limiting, concentration. This is typical of logistic kinetics, although Alexander [61] has argued that the precision of degradation data is not good enough to distinguish between this and first-order kinetics. Zone D represents the situation where a small number of bacteria are added to a system which contains the test chemical at a concentration greater than the organisms' K s value. Degradation can therefore be described using classical Monod kinetics, with growth of the active bacteria occurring during substrate disappearance. When the initial concentration of substrate is much greater than Ks, most of the substrate will disappear while the uptake systems of the cells are saturated. Under these conditions (Zone F), (K s + IS]) can be approximated by IS] until the substrate is nearly e x h a u s t e d and the k i n e t i c s of d e g r a d a t i o n can be f o l l o w e d u s i n g the l o g a r i t h m i c model. If the initial c o n c e n t r a t i o n of test c h e m i c a l is insufficient to support a significant increase in the population of the active bacteria, and if the uptake systems of these bacteria are saturated (Zone E), then substrate disappearance will be linear with time and zeroorder kinetics apply. The Monod (no growth) kinetic expression can be used to model degradation in situations where the uptake systems of the active organisms are not saturated at the start of incubation, although the initial concentration of substrate is insufficient to support a significant increase in biomass (Zone C). The Monod (no growth) expression is analogous to Michaelis-Menten kinetics for an enzyme-catalysed reaction (Section 3.3). Substrate disappearance curves for a chemical degraded according to the various kinetic models discussed above are shown in Figure 3B. Although the integrated Monod equation relies on the assumption that there is constant cell yield with time (cf. Section 3.2), and is not applicable to populations which exhibit a lag period before the onset of degradation, the approach is a useful guide to which kinetic model could be best applied to a set of biodegradation data. Although the concentrations at which the two vertical lines and the vertical intercept with the diagonal are placed would vary between different chemicals and inocula, the general pattern of Figure 3A should remain unchanged. It should be borne in mind that factors other than initial substrate concentration and cell density can influence degradation in the aquatic environment. These include the presence of other substrates, predation by protozoa and phages, toxin production by other organisms, the availability of inorganic nutrients and binding to colloidal matter [62]. The models Shown in Table 3 have been successfully used to describe the mineralisation of [U-14C] benzoate in settled sewage and by a Pseudomonas sp. [60], and the primary biodegradation (loss of methylene blue active substance) of sodium dodecyl sulphate by epilithic and planktonic bacteria from clean and polluted river water [63]. The data are fitted to each model by nonlinear regression analysis using the MARQFIT computer programme [60], which fits data by minimising the least squares differences between the data and the model curve. The basic approach of Simkins and Alexander has been e x t e n d e d in later p a p e r s d e s c r i b i n g the n o n l i n e a r e s t i m a t i o n of the

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parameters of Monod kinetics that describe the patterns of mineralisation in cultures which contain different substrate concentrations and cell densities [64], and on the modelling of the biodegradation kinetics of glucose, benzoate and phenol in sewage where the growth of active bacteria occurs at the expense of other substrates [65].

4.2 ~ichaells-Henten kinetics The Michaelis-Menten equation describing the rate of an enzyme-catalysed reaction was discussed in Section 3.3. At very low concentrations of substrate, the transformation rate will appear to be first-order with respect to substrate. Higher substrate concentrations may allow bacterial growth to occur, with a corresponding increase in enzymic activity and m i x e d - o r d e r k i n e t i c s (such " S - s h a p e d " b i o d e g r a d a t i o n c u r v e s and their significance are discussed in greater detail in [19]). If the concentration of substrate is very high, v approaches Vma x and the rate of transformation will be constant and zero-order with respect to substrate. Experimental examples of such first-order, mixed-order and zero-order kinetics for the degradation of radiolabelled linear alcohol ethoxylates in estuarine water have been reported [26]. Michaelis-Menten kinetics have been used in the US Environmental Protection Agency's Exposure Analysis Modelling System (EXAMS) in an attempt to predict the rate of biotransformation of an organic pollutant in the aquatic environment. EXAMS has been used to predict the steady-state concentrations of linear alkylbenzene sulphonates (LAS) in a US fresh water stream using both an aqueous biodegradation rate constant for LAS (water column and sediment interstitial water) and a sediment rate constant [66]. Although there was an excellent agreement between predicted and measured LAS levels, this was only achieved by assigning a relatively arbitrary value to the dispersion coefficient at the sediment-water interface. Unfortunately, the model calculations were most sensitive to the least understood variables: this dispersion coefficient and the sediment biodegradation rate constant. It was c o n c l u d e d that f u r t h e r r e s e a r c h is n e e d e d to b e t t e r d e f i n e biodegradation processes in sediments (see Section 7). The use of Michaelis-Menten kinetics in EXAMS, and the assumptions inherent in this approach, are discussed in detail by Lewis et al [48] and summarised in Figure 4. The authors concluded that the kinetic models are best suited to p r e d i c t i n g transformation rates at e i t h e r trace, environmental concentrations (first-order kinetics) or under spill or discharge conditions (zero-order, although consideration may have to be given to toxicity or adaptation effects on the microbial population). Unfortunately, most degradation studies (and all EEC and OECD tests for "ready biodegradability") are conducted using test chemical concentrations of i0 -° to 10 .4 M. The predictive use of rate data derived from such tests is therefore much reduced by the possibilities of mixed-order kinetics, toxic effects and adaptation to the test chemical. For example, Spain and van Veld have demonstrated that aquatic bacteria can become adapted to xenobiotic chemicals at concentrations as low as 10 .7 M [67].

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Concentration of chemical Spill or discharge concentrations

(M)

10-2

BACTERIAL TRANSFORMATION RATES ARE:

- zero-order with respect to [S] - often affected by substrate toxicity - often subject to population adaptation

10-3

10-4

BACTERIAL TRANSFORMATION RATES ARE:

10-5 Optimum analytical concentrations

- multiphasic with respect to [S] (series of i st, mixed- & 2nd-order) - sometimes affected by toxicity - sometimes subject to adaptation

i0-6

i0-7

10-8

BACTERIAL TRANSFORMATION RATES ARE:

Environmental concentrations

first-order with respect to [S] rarely affected by toxicity rarely subject to adaptation

Figure 4. Application of Mlchaelis-Menten kinetics to predict microbial transformation rates (primary biodegradatlon) in the aquatic environment [48].

5. OTHER EFFECTS OF CONCENTRATION ON THE KINETICS OF BIODEGRADATION

If a flrst-order rate constant has been obtained for the degradation of a chemical in a laboratory test, and if degradation in situ follows firstorder kinetics, then the rate of biodegradatlon in si~u will be proportional to the environmental concentration of substrate (i.e. test chemical). This linear relationship between rate and concentration has been demonstrated in lake waters for the mineralisation of phenol [68]. However, there have been several reports of threshold concentrations below which known biodegradable compounds are persistent, an occurrence which is not predicted by classical

1263

M o n o d and M i c h a e l i s - M e n t e n kinetics. Threshold concentrations for degradation appear to be substrate dependent and range from I0 ~g/l for a Pseudomonas sp. with 4-nitrephenol [69] and 2 ~g/l for 2,4-dichlorophenoxyacetate in stream water [70], to < l~g/l for linear alcohol ethoxylates (LAE) and LAE sulphates in estuarine water [26]. It should be noted that these threshold concentrations are of the same order of m a g n i t u d e as r e p o r t e d e n v i r o n m e n t a l concentrations for m a n y o r g a n i c "micropollutants". Even known natural, biodegradable compounds such as glucose, glycerol and lactate are known to have threshold concentrations below which they are not degraded [71]. Natural waters and waste waters contain low levels of many naturally-occurring chemicals which are known to be biodegradable and the presence of a low concentration of a synthetic chemical in a sample does not necessarily mean that it is not biodegradable. There have also been reports of threshold concentrations below which there is no adaptation of aquatic microbial communities to 4-nitrophenol [72]. Research on biofilm kinetics has shown that for a single organic compound under steady-state conditions the m i n i m u m c o n c e n t r a t i o n of s u b s t r a t e ([Smin]) r e q u i r e d to s u p p o r t the b a c t e r i a l p o p u l a t i o n d e g r a d i n g that compound is given by the following equation [73]:

[Smin]

-

Ks

(16) (Y ut - b)

where: b is the first-order decay coefficient and ut is the maximum specific rate of substrate utilisation by the bacteria.

If the concentration of substrate (e.g. synthetic chemical) is below [Smin], then the biofilm cannot be sustained and will eventually decay. Equation (16) is a useful expression as attached bacteria are key components of waste water treatment systems such as trickling filters and rotating biological contactors, and the natural state of most aquatic bacteria is thought to be as sessile organisms attached to surfaces [74]. As typical values for [Smin] are in the mg/l range, many trace organics would apparently fail to be utilised by a biofilm. However, the presence of a primary substrate at a concentration greater than its [Smin] could provide the microbial population with energy and carbon for growth, enabling degradation of the trace organic to occur. Experimental evidence for the "secondary utilisation" of trace concentrations (8-30 ~ g / l ) of o r g a n i c compounds under aerobic and methanogenic conditions "inthe presence of a higher concentration of primary substrate (1-100 mg/l acetate) have been reported [73]. As the concentration of the secondary substrate was very low (<
d [Sf]

ut Xf [Sf]

(17) dt

where:

Ks

[Sf] is the concentration of the secondary substrate within the biofilm and Xf is the active cell density.

at

a

point

1264

Another theoretical approach to predicting threshold concentrations, this time for the growth of an individual bacterlum~ has been described by Schmidt et al [75]. It is assumed that the rate of substrate uptake and metabolism (i.e. biodegradation) by a bacterial cell is dependent on the rate of substrate diffusion to the cell surface. At very low substrate concentrations this sets a limit on growth and can be described by the following expression:

I/Yma x (Rd2 - Rb2)/2 r

=

(18) DAB Cb/p - (m/in2)

where:

(Rd 2 - Rb2)/2

r is the maximum diffusion-limited doubling time, Ymax is the true yield coefficient [cf. Y, equation 6: I/Y = i/Yma x + (mr/in2)], R d is the radius of the bacterial cell at division, R b is the radius of the cell at its first appearance, DAB is the diffusivity of the substrate in the s u s p e n d i n g fluid, C b is the b u l k c o n c e n t r a t i o n of the substrate, p is the dry weight of a single cell/volume and m is the maintenance coefficient of the organism.

Using published values for the above parameters, and assuming that a growth threshold was reached when r approached infinity, the authors calculated that for a typical organism the threshold concentration for growth would be around 0.2 ~g/l. This apparently conflicts with published reports of biodegradation in natural waters at sub-~g/l concentrations [76, 77]. Schmidt et al suggest that bacteria from ollgotrophic environments may have maintenance energy requirements which are up to 100-fold lower than those usually reported in the literature this would then lower the predicted threshold concentration by approximately the same amount. As we have already seen, bacteria in biofilms can metabolise a chemical at a concentration b e l o w its t h r e s h o l d if they s i m u l t a n e o u s l y use other substrates. Research summarised by Harder and Dijkhuizen [78] has shown that under the nutrient-limited conditions found in many natural waters versatile bacteria which can simultaneously utilise a mixture of substrates are at a competitive advantage. The significance of mixed-substrate utilisation in the degradation of xenobiotic chemicals has been demonstrated in laboratory studies by Bouwer [see above] and LaPat-Polasko et al [79]. The latter workers found that in batch and continuously-fed reactors, a pseudomonad utilised trace (~g/l) amounts of dichloromethane whilst simultaneously using acetate (mg/l) as the primary substrate for maintenance and growth. The results of pure culture studies lead Schmidt and Alexander [80] to conclude that the rate and extent of biodegradation of many synthetic chemicals in the environment may be controlled by the presence of higher concentrations of other substrates. A threshold concentration for b i o d e g r a d a t i o n can also occur if the concentration of a s y n t h e t i c c h e m i c a l is too low induce the e n z y m e s necessary for uptake or metabolism. Schmidt et al [69] found that a Pseudomonas sp. isolated from soil could not mineralise 4-nitrophenol (4-NP) at concentration of i0 ~g/l (mineralisation occurred at concentrations > 50 ~g/l). In the presenc~ of a relatively high concentration (20 mg/l) of a second substrate (glucose, glutamate, acetate or phenol) the organism was

1265

still unable to utilise 4-NP at concentrations below the I0 ~g/l threshold. This was despite the fact that these second substrates promoted the growth of the organism, and glucose increased the rate of degradation of higher concentrations (i mg/l) of 4-NP by stimulating growth. It was concluded that these findings supported the hypothesis that the threshold concentration of 4-NP was too low to induce degradative enzymes.

6. THE EFFECT OF TEMPERATURE ON THE RATE AND EXTENT OF BIODEGRADATION

Within physiological limits the rate of a biological reaction increases as the temperature is increased. This means that the size of a biodegradation rate constant k is temperature dependent. The relationship between k and the absolute temperature is defined by the Arrhenius equation:

k where:

=

Ae'E/RT

(19)

A is constant, E is the activation energy of the reaction, gas constant and T is the temperature in K.

Equation

(19) can also be written in a logarithmic

R is the

form as:

E

In k

=

In A - -RT

(20)

This shows that the logarithm of k is a linear function of the reciprocal of the absolute temperature and a plot of in k against I/T gives a straight line with a slope of -E/R. Using equation (20) Larson [19] demonstrated that the rates of degradation of nitrilotriacetie acid and linear alkylbenzene sulphonate were proportional to temperature within the range 2-35 °C. This approach can be used to predict the rate of biodegradation at in situ temperatures from rates measured at the more commonly used test temperatures of 20-25 °C. However, care must be taken as psychrophilie bacteria (optimum growth temperature <15 °C) present in the environment may exhibit greater efficiencies for degradation than those predicted and rate predictions based on the Arrhenius equation may be conservative. Another problem is the observation that while certain compounds such as branched and linear alkylphenol ethoxylates are d e g r a d e d e x t e n s i v e l y in a c t i v a t e d sludge simulation tests at 15-20 °C, removal can be erratic at 5-8 °C [81, 82].

7. BIODEGRADATION IN SEDIMENTS

Most natural waters are low in nutrients. It has long been known that in such environments ions and various macromolecules accumulate at solid-liquid interfaces and that attached bacteria often exhibit an elevated rate of mineralisation and growth compared with their planktonic counterparts. This stimulation of microbial activity is thought to be due to the elevated concentration of adsorbed nutrients and to other surface-assoclated effects such as cross-feeding, co-metabolism and interspecies hydrogen transfer [83]. In fact it has been suggested that the natural stat4 of most aquatic

1266

bacteria is as sessile organisms attached to surfaces [74]. The activities of such microbial communities are utilised and optimised during waste water treatment by trickling filters and rotating biological contactors. Surfaces for m i c r o b i a l a t t a c h m e n t can be small s u s p e n d e d p a r t i c l e s , submerged solid supports (e.g. rocks and plants) or bottom sediments. The latter are particularly important as their microbial populations are usually several orders of magnitude greater than the free-floating planktonic bacteria, and their contribution to the mineralisation of organic compounds and the geochemical cycling of nitrogen, sulphur and phosphorous is much greater than that occurring in the water column [84]. In addition, aquatic sediments often act as sinks for synthetic organic chemicals [85]. It is therefore surprising that so little research has been reported on the biodegradatlon of synthetic chemicals in freshwater, estuarine and marine sediments. The need for reliable rates for sediment-associated biodegradation processes was emphasised by the results of the validation e x e r c i s e on the EPA's EXAMS model m e n t i o n e d p r e v i o u s l y . M u c h of the published data on the biodegradation of sediment-associated chemicals is reviewed in [86, 87]. The latter paper cited studies where biodegradation was enhanced by the presence of sediments, studies where the presence of sediments had little effect on biodegradation rates and other work which concluded that sorbed chemicals were unavailable for biodegradation. Shimp and Young [87] examined the bioavailability of sediment-adsorbed organic chemicals using two 14C-labelled model compounds: dodecyltrimethylammonium chloride (TMAC) a highly sorptive, cationic q u a t e r n a r y a m m o n i u m surfactant, and phenol a neutral, r e l a t i v e l y hydrophobic chemical which undergoes moderate adsorption. Sediment columns were prepared by allowing a slurry of river water and surface (2-4 cm) sediment to settle, these were then spiked with 10-20 000 ~g/l test compound and incubated for 12-24 h. The amount of test chemical in the overlying water, interstitial water and sediment, and the amount of 14C02 produced on acidification were determined at the end of incubation. Biodegradation rate data for TMAC were fitted to a flrst-order model (equation I) and those for phenol to a Michaelis-Menten model (equation 8). It was found that the biodegradation rate for TMAC was a function of the concentration of chemical dissolved in the interstitial water (i.e. unadsorbed TMAC). Biodegradation of adsorbed TMAC only occurred when this material became desorbed as the interstitial water became depleted of the chemical due to biodegradation. In contrast, the rate of phenol biodegradation was a function of the total concentration of the material present which suggested that some of the adsorbed material was available for biodegradation. The authors concluded that chemical structure and related adsorption mechanisms may be important factors controlling the rate of biodegradation of adsorbed chemicals in sediments. It is known for example, that the sorption of charged organics is probably controlled by interactions with ion exchange sites on the surface of the sediment particle or the fibrous mat of polysaccharide fibres (often called the glyeocalyx) formed by many attached organisms. Adsorption of neutral chemicals is likely to be controlled by hydrophobic and/or van der Waal's interactions [88]. The mineral composition of the sediment particles can also have a marked effect on the availability of organic compounds for degradation [89]. Baughman et al [32] used a modified version of the second-order rate expression (equation 7) to study the biodegradation of synthetic chemicals

1267

in suspended, sediment systems. Total microbial biomass was estimated by adenosine triphosphate measurement (this was a surrogate measure of the population of active degraders the assumption being made that these organisms were a l w a y s a constant fraction of the total m i c r o b i a l population). Sediment sorption effects were incorporated into equation (7) by incorporating the distribution of chemical between sorbed and dissolved forms:

d[S]T

k [B] [S]T

dt

where:

-

kap p [B] [S] T

(21)

I + pKp

[S]T is the total concentration of substrate (test chemical) per unit aqueous volume of the system, p is the sediment-water mass ratio, Kp is the partition coefficient of test chemical between sediment and water, and kap p is the apparent second-order rate constant.

Using this approach Baughman and co-workers found that the pesticides methoxychlor and chlorpropham, and several phthalic acid esters, were less available for biodegradation when sorbed to suspended sediments. Equation (21) is similar to that proposed by Furmidge and Osgerby [1967, cited in 41] for describing biodegradation in soil, in which the dissolved chemical is degraded according to first-order kinetics and the adsorbed material is protected from microbial attack. An attempt to incorporate the effect of sediments on biodegradation rates into a routine test procedure ("Chemical/toxicity Abatement Test") is described in [27]. The CTA Test is a modified river die-away test which determines the primary biodegradation of relatively low levels (around 0.2 mg/l) of test substance, and assesses the toxicity of any degradation products towards mysids or Daphnia magna. The use of slurries of 0.5 g surface (top 2 cm) sediment/l river water and sterile controls yield information on the effects of sediment-associated and abiotic factors on biodegradation rates (kl). Limitations of the test protocol are the sitespecific nature of the data obtained and the fact that the use of mixed slurries containing a relatively low amount of sediment may not be a realistic model of biodegradation processes in settled, bottom sediments

[87]. The modelling and prediction of biodegradation rates in sediments is complicated by the fact that the nature of the mineralisation process changes as the sediment redox potential (Eh) decreases. In most unpolluted sediments there is an oxic layer w h e r e o x y g e n c o n s u m e d d u r i n g the mineralisation of organic matter is replenished by diffusion from the overlying water. However, a depth is eventually reached where the removal of oxygen by aerobic respiration exceeds replenishment and anoxic conditions occur. Within this region degradation can continue either by anaerobic f e r m e n t a t i o n or by the use of a l t e r n a t e inorganic terminal electron acceptors to 02 . Microbial fermentations can occur at any depth or Eh within the anoxic sediment [84]. Anaerobic bacterial respiration on the other hand generally follows a vertical sequence down the sediment as the available

1268

electron as:

acceptors

become

less oxidising.

This sequence

SEDIMENT SURFACE

AVERAGE DEPTH IN SEDIMENT

can be generalised

Eh

I ram

02

> 2H20

+ 600 mV

i

NO 3-

> N2/NH4 +

+ 350 mV

S042"

~ H2S

- I00 mV

CO 2

> CH 4

- 200 mV

cm

I0 cm im

In estuarine and coastal sediments, dissimilatory sulphate reduction is often the predominant means of anaerobic mineralisation and can account for half the organic carbon mineralisation [90]. This is due to the high levels of sulphate present in the sediment interstitial water (up to 29 mM) and to the low concentration and rapid reduction of energetically-favoured electron acceptors such as 02 and NO3-. However, in most freshwater sediments the concentration of sulphate is low and denitrification and methanogenesis assume more importance. For example, in surface sediment from a UK lake the contribution of the various respiration types to CO2-accumulation was: aerobic respiration 42%, denitrification 17%, sulphate reduction 2% and methanogenesls 25% [91]. It is clear that within sediments Eh (determined by organic input, sediment particle size and hydrographic conditions) and the availability of electron acceptors will play crucial roles in determining microbial activity and diversity, which in turn determine the potential, rate and extent of sediment-associated biodegradation. In studies using anaerobic slurries from methanogenic and sulphate-reducing aquifer sites Suflita and Miller [92] observed that material from the methanogenic site readily dehalogenated 4-chlorophenol and 2,5-dichlorophenol to phenol and 3chlorophenol, respectively whilst samples from the sulphate-reducing site could mineralise phenol but failed to transform the chlorophenols tested. A later paper reported that pre-adapted slurries from the sulphate-reducing site could degrade 4-cresol up to 18-times faster than methanogenic samples. The activities of anaerobic bacteria can also have other effects on the environmental fate of a synthetic chemical. It is known that anaerobic organisms can remove CI atoms directly from the ring of aromatic compounds [93,94]. This reductive dechlorination process does not appear to occur under aerobic conditions and is important as it renders the chemical less toxic and more amenable to biodegradation. Saldick [95] reported that cyanuric acid was persistent in screening tests under highly aerobic conditions but was rapidly degraded at dissolved oxygen concentrations below 3 mg/l. It therefore seems prudent, when designing test strategies to predict the fate of a synthetic chemical in the aquatic environment, to include systems which take into account the interactions between the test chemical, the water column, the sediment and its associated microflora under low-sulphate (freshwater) and high-sulphate (marine) conditions. Promising approaches to this include the "eco-core" t e c h n i q u e w h e r e a s e d i m e n t core and its overlying water are incubated with the test chemical and disappearance monitored [72, 96] and the intact core incubation technique of Christensen [97].

1269

8. DO "READY B I O D E G R A D A B I L I T Y " CONDITIONS ?

TESTS H A V E

"ENVIRONMENTALLY

REALISTIC"

In S e c t i o n 2 the n e e d for k i n e t i c data which, t h o u g h d e r i v e d under laboratory-controlled conditions, were sufficiently test system independent, so as to allow extrapolation to the environment was discussed. Research and method development on biodegradability testing in Europe and elsewhere during the past decade has tended to concentrate on the assessment of "ready biodegradability". The reasons for this are probably the relative simplicity of the test methods, their incorporation into internationally-accepted guidelines [2, 4] or legislation [3], the paucity of laboratory tests which simulate the conditions found in the aquatic environment, the small number of new chemicals requiring a "higher" level of testing (see [5]), an accumulating data base based on the results of ready tests, and interlaboratory testing programmes co-ordinated by the OECD and EEC designed to evaluate new ready test procedures and revise existing ones [98-101]. Unfortunately, over recent years there has been a tendency to look upon the various ready biodegradability tests as models of the aquatic environment and to extrapolate rate "constants" derived from such tests in order to predict the rate of b i o d e g r a d a t i o n in a p a r t i c u l a r environmental compartment. A comparison of "ready biodegradability" test conditions with values often reported for UK and Northern European waters is given in Table 4. It is apparent that there are large differences, particularly in the elevated concentrations of the synthetic chemical under investigation and inorganic mineral salts. "Ready biodegradability" test methods were developed with a view to assessing the mineralisation ("ultimate biodegradability") of a wide variety of c o m p o u n d s of d i f f e r e n t structures and p h y s i c o - c h e m i c a l properties. Test substance removal had therefore to be determined by the use of non-specific analytical techniques such as dissolved organic carbon (DOC) removal, O 2 - c o n s u m p t i o n or C 0 2 - e v o l u t i o n , and the c a l c u l a t i o n of the difference in these parameters between test vessels containing a known concentration of the test chemical and controls "identical" except for the absence of test substance ("blanks"). This necessitated the use of a high concentration of test material dissolved in a reproducible (i.e. synthetic) medium. This medium had to have a low concentration of other organic components so that "blank" values would be low and have sufficient buffering capacity for pH control if the high concentration of test substance was degraded. The test conditions in "ready biodegradability" tests tend to have a lower biodegradation potential than most aquatic environments and as a consequence it is assumed that a "readily biodegradable" chemical will degrade "rapidly in the environment" [5]. It is worth noting that several of the test conditions are arbitrary and many of variations between the various test protocols reflect historical differences. In nearly all the tests, for example, the medium has been derived from traditional BOD dilution water multiplied by a particular factor to account for the higher test substance concentration [i14]. The high concentration of test substance used in "ready biodegradability" tests can have an inhibitory effect on the active microflora which would not occur at in situ concentrations. Larson [25] compared biodegradation results obtained with two screening tests (MITI I and a proposed American Society for Testing and Materials CO2-evolution test) and an acclimated activated sludge, with those measured by incubating a low concentration of 14C-

1270

Table 4. A

comparison

of conditions in ready biodegradability tests with

those found in the aquatic environment. Table compiled using data from [102-114] and sources cited in the text.

Typical environmental values a Ready test

Parameter

Concentration of synthetic chemical

2-100 mg/l

Concentration of other substrates (as C)

ib mg/l

River water

Sea water

Waste water

ng-~g/l

ng-~g/l

<~g-mg/l

1-20 mg/l

0,3-3 mg/l

200 mg/l

1-20 ~g/l

20 mg/l

Concentration of phosphate (as P)

12-242 mg/l

<0.1-2 mg/l

Concentration of ammonia (as N)

0.5-86 mg/l

0.1-5 mg/l

<1-50 ~g/l

40 mg/l

105-1010

108-1010

108-109

i010-i011

Concentration of bacteria (cells/l) c Possibility of the development of a population of acclimated organisms ? Predominant microbial population

Low

High

High

High

Planktonic

Attached

Attached

Planktonic Flocs d Attached e

Nature of system

Static

Dynamic

Dynamic

Dynamic

Temperature (°C)

20-25

2-20

3-15

8-20

7.2-7.5

7.1-8.0

H2P04- -HP042"

HC03" -CO 2

pH Buffer system

a b c d e

7.8-8.2 HC03--CO 2

7.2-7.8 HC03 - _CO 2

Considerable local variations occur (especially in C, P and N levels) Nominally absent Direct microscopic'counts Activated sludge reactors Attached to particulate matter and solid supports (e.g. trickling filters, rotating biological contactors)

1271

labelled compound with a range of river waters. The extent of degradation of 50 mg/l nitrilotriacetic acid (NTA) in the MITI test and 20 mg/l LAS (C12 average alkyl chainlength) in the CO2-evolution test was comparable to that in pre-exposed and unexposed river water samples incubated with 5 0 ~ g / l of radiolabelled test chemical; although degradation rates (expressed as kl) tended to be lower in the screening tests. However, a model cationic surfactant (octadecyltrimethylammonium chloride) was toxic to the test organisms at i0 mg/l in the CO2-evolution screening test but was degraded at i0 ~g/l in river water. Shimp and Young [116] compared the biodegradation of 50 ~ g / l [U-14C] benzoic acid (as DOC) in nutrient-supplemented sea water with that occurring in a sea water version of the modified OECD screening test [I01]. In unpolluted sea water the measured degradation of 20 mg DOC/I benzoic acid in the screening test was poor and highly variable. At a concentration of 50 ~g DOC/I degradation was rapid, less variable and extensive with between 70-80% of the chemical being converted into 14C02 or microbial biomass after 7 days incubation. However, in samples taken from an estuary receiving large quantities of municipal wastewater the biodegradation of benzoate was rapid at both 20 mg DOC/I and 5 0 ~ g DOC/I. It is clear then that for certain chemicals the h i g h c o n c e n t r a t i o n of test s u b s t a n c e u s e d in "ready biodegradability" test methods can lead to an underestimation of its likely biodegradability in all but the most polluted waters. Other problems in using these tests to generate been discussed and can be summarised as follows:

kinetic

data have

already

a) If growth-linked biodegradation of the test substance occurs over the 28 day incubation period then the test system is in effect a batch enrichment which will select microbial specialists with high growth rates at these unnaturally high substrate concentrations. The growth of more versatile organisms, which may be the active degraders in environments which contain low levels of a variety of substrates, will be discouraged. b) The e l e v a t e d incubation temperatures e s t i m a t i o n of the rate and e x t e n t of temperatures found in situ.

can result in biodegradation

an at

incorrect the lower

c) The test systems fail to address the effects of sorbed chemicals, sediments or other particulate matter on biodegradation. Many of the test protocols contain instructions to coarse filter the inoculum before use. It is known, for example, that the removal of suspended particles (>3 ~ m diameter) from lake water can alter the kinetics of biodegradation [117]. d) The use of high test substance often results in unpredictable Kinetic data obtained dependent on the test rate of blodegradation be useful however, in chemical in a spill or

concentrations or mixed-order

and long incubation periods kinetics.

using "ready biodegradability" tests will be highly conditions and as such cannot accurately predict the in a variety of aquatic environments (such data may predicting the behavlour of a high concentration of point source discharge [36, 118].

1272

9. " E N V I R O N M E N T A L L Y R E A L I S T I C " TEST C O N D I T I O N S

From the preceding discussion it is clear that the generation of "real world" b i o d e g r a d a t i o n k i n e t i c data [15] requires tests w h i c h e m p l o y sensitive analytical methods to determine biodegradation at environmentallyrealistic concentrations of test chemical (i.e. [S] << Ks/Km), and "realworld" media such as natural or waste waters or other samples (e.g. surface sediments). Incubation should take plac e at environmental temperatures and will probably last for hours and not days. However, while short incubation periods facilitate the application of kinetic models, problems may arise with chemicals that require a period of adaptation (lag-phase) before biodegradation can begin. At the time of w r i t i n g the most p r o m i s i n g a p p r o a c h is to a m e n d an environmental sample with a small amount of uniformly 14C-labelled chemical and monitor the change with time of one or more of the 14C-fractions shown in Table 5. The most appropriate kinetic model is then fitted to the data by regression techniques to yield biodegradation rate constants. Incubation enclosures used in 14C die-away tests have included: shake flasks and aqueous C02-traps [70, 116]; vials and impregnated paper CO2-traps [56, 119, 120]; batch activated sludge units and aqueous C02-traps [121]; double-vial and C02-tra p respirometers [122]; and "eco-cores" [72]. Other papers using these techniques are cited elsewhere in this review. The choice of enclosure for incubation will be dependent on the system being studied (e.g. water column, attached organisms, sediment) and the most appropriate testing system should be evaluated experimentally.

Table 5. Data provided by 14C die-away tests.

Fraction measured

Method of collection

Particulate 14C

Filtration (0.2 ~m) or centrifugatfon

Evolved 14C021

Aspect of test chemical metabolism measured

Assimilation into biomass

Trapped in KOH, NaOH or 2-methoxyethanolamine either continuously &/or after acidification of filtered sample

Mineralization

Evolved 14CO21 (sterile)

As above

Loss by volatilisation

Particulate 14C (sterile)

Filtration (0.2j~m) or centrifugatlon

Loss by adsorption

14C remaining in solution

Determined after filtration, acidification & gassing-off of 14CO 2

i May include volatile organic material

Mass balance

1273

Two difficulties in the use of 14C die-away tests are the high cost of custom synthesis and the difficulties in synthesising a uniformly 14Clabelled, representative sample of the test substance particularly for commercial products which may be mixtures or contain a number of isomers. While specific analysis has been used widely in biodegradation studies its usefulness in environmental fate prediction is limited as the kinetic data derived are for "primary" hiodegradation and not mineralisation. Unfortunately, given our current poor understanding of the factors affecting the rate and extent of biodegradation in the aquatic environment, the generation of data for use in predicting the rate of biological removal in a variety of different compartments in the aquatic environment may well have to use tests which quantify the effects of the following variables: a) The nature of the sampling site (e.g. fresh surface water, estuarine, marine, groundwater, sewage, activated sludge, etc.). Many first-order rate " c o n s t a n t s " are in fact s i t e - s p e c i f i c - a r e f l e c t i o n of the variation in biomass concentration and population diversity between sites. There is an urgent need for techniques which can quantify the numbers of active degraders. Promising approaches to this problem are the 14C-most-probable-number method [123, 124] and the technique of Anderson et al [125] which uses gel zymography to determine the proportion of "nonselectively" isolated bacteria which can produce alkylsulphatase (assumed to equivalent to the number of bacteria capable of degrading alkyl sulphate surfactants). This latter method has the disadvantage that active organisms may not be recovered during the initial plating process. With the 14C-MPN technique the use of uniformly [U-I~c] and side-chain labelled compounds enables different populations of active bacteria to be quantified. The use of gene probes to identify specific groups of micro-organisms in biodegradability studies is limited as the technique detects potential and not in situ activity (e.g. the genes may not be expressed as enzymic activity), and by the need to cultivate the organisms (although techniques for DNA extraction from heterogeneous populations are improving rapidly). b) The concentration of inorganic and organic nutrients. Chemical speciation (e.g. m e t a l - o r g a n i c chemical complexes) is also k n o w n to a f f e c t mineralisation [126]. c) The p r e s e n c e compounds).

of

particulate

matter

(especially

for

highly

sorbed

d) Temperature. e) Oxygen concentration, to 0 2 .

redox potential

and alternative

electron

acceptors

f) Can the microbial population become adapted to the test chemical ? Such effects can often be detected from the shape of the biodegradation curve and an increase with time in the numbers of active degraders. g) Concentration

of test chemical

(is there a threshold

?).

Many of the above variables are of course site-specific and the challenge to experimenters is to understand how physico-chemical conditions (which are relatively easy to quantify) affect biodegradation.

1274

i0. CONCLUSIONS

The biodegradability of a chemical is not a fundamental constant that can be quantified in the same way as a physico-chemical property such as melting point or vapour pressure. It is a variable affected by the compound's molecular properties and the environmental compartment where it is found. In devising and performing testing strategies we are attempting to predict environmental acceptability from laboratory studies. Perhaps the best that we can hope to achieve in the foreseeable future is a greater understanding of the factors that affect, and a measurement of, the rate and extent of biodegradation in a number of "typical" environmental compartments.

ACKNOWLEDGEMENT

This review is based on a Progress Report to the Department of the Environment (DoE 2161-M, April 1989) conducted at WRc Environment, Medmenham Laboratory as part of contract PECD 7/8/114 from the Central Directorate of Environmental Protection (CDEP), U.K. Department of the Environment. The author wishes to acknowledge this support and the Directorate's kind permission to publish.

REFERENCES

[i] Swisher R.D. (1987). Surfactant Biodegradation, Dekker Inc: New York.

Second Edition. Marcel

[2] Guidelines for Testin~ of i Chemicals. Organisation for Co-operation and Development. Paris, 1981.

Economic

[3] Council Directive 79/831/EEC Amending for the Sixth Time Directive 6 7 / 5 4 8 / E E C on the C l a s s i f i c a t i o n , "Packaging and L a b e l l i n g of Dangerous Substances. Official Journal of the European Communities L259 i0, 1979. [4] Part C: Methods for the Determination of Ecotoxicity. Official Journal of the European Communities L251 27, 160-211, 1984. [5] G u i d i n g P r i n c i p l e s for a S t r a t e g y for B i o d e g r a d a b i l i t y Testing. Guidance Document of the Competent Authorities for the Implementation of Directive 79/831/EEC. XI/841/86 final, 1986. [6] Mackay D. (1979). Finding fugacity feasible. Environmental Science and Technology 13, 1218-1223. [7] Baughman G.L. and Burns L.A. (1980). Transport and transformation of chemicals: a perspective. In The Handbook of Environmental Chemistry Vol. 2 Part A, edited by O. Hutzlnger, pp 1-17. Springer-Verlag: Berlin.

1275

[8] Mackay D. and Paterson S. (1981). Calculating fugacity. Environmental Science and TechnoloEv 15, 1006-1014. [9] Jacobsen B.N. (1987). A mathematical model for behaviour of xenobiotic compounds in an activated sludge reactor. Water Pollution Renort 6. Behaviour of Organic MicroDollutants in Biological Waste Water Treatment, pp 185-195. Proceedings of COST 641 Workshop, Copenhagen May 1987. EUR 11356. [i0] Reed M. (1990). The physical fates component of the natural resource damage assessment model system. Oil and Chemical Pollution 5, 99-123. [ii] Feng S-S, Reed M. & French D.P. (1989). The chemical database for the natural resource damage assessment model system. Oil and Chemical Pollution 5, 165-193. [12] Burns L.A. (1985). Models for predicting the fate of synthetic chemicals in aquatic ecosystems. In Validation and Predictability of Laboratory Methods for Assessin~ the Fate and Effects of Contaminants in Aquatic Ecosystems, ASTM STP 865, edited by T.P. Boyle, pp 176-190. American Society for Testing and Materials: Philadelphia. [13] Dickson K.L., Maki A.W. & Cairns J. (1982). Modeling the Fate of Chemicals in the Aouatic Environment. Ann Arbor Science Publishers: Michigan. [14] Swan R.L. & Eschenroeder Environment. ACS Symposium Washington.

A. (1983). Fate of Chemicals in the Series 225. American Chemical Society:

[15] Howard P.H. (1985). Determining "real world" biodegradation Environmental Toxicology and Chemistry 4, 129-130.

rates.

[16] Vaishnav D.D., Boethling R.S. & Babeu L. (1987). Quantitative structure-blodegradability relationships for alcohols, ketones and alicyclic compounds. Chemosphere 16, 695-703. [17] Howard P.H., Hueber A.E. & Boethling R.S. (1987). Biodegradation data evaluation for structure/biodegradability relations. Environmental ToxicoloEv and Chemistry 6, I-I0. [18] Boethling R.S. (1986). Application quantitative structure-biodegradability Toxicology and Chemistry 5, 797-806.

of molecular relationships.

topology to Environmental

[19] Larson R.J. (1980). Role of biodegradation kinetics in predicting environmental fate. In Biotransformation and Fate of Chemicals in the Aouatic Environment, edited by A.W Maki, K.L. Dickson & J. Cairns, pp 67-86. American Society for Microbiology: Washington. [20] Larson R.J. (1984). Kinetic and ecological approaches for predicting biodegradation rates of xenobiotic organic chemicals in natural ecosystems. In Current Perspectives in Microbial Ecology, edited by M.J. Klug & C.A. Reddy, pp 677-686. American Society for Microbiology: Washington.

1276

[21] Lee R.F. & Ryan C. (1979). Microbial degradation of organochlorine compounds in estuarine waters and sediments. In Proceedings of the Workshop: Microbial Degradation of Pollutants in Marine Environments, edited by A.W. Bourquin and P.H. Prltchard, pp 443-450. EPA-600/9-79012.

[22] Larson R.J. & Perry R.L. (1981). Use of the electrolytic respirometer to measure biodegradation 15, 697-702.

in natural waters.

Water Research Bulletin

[23] Bartholomew G.W. & Pfaender F.K.

(1983). Influence of spatial and temporal variations on organic pollutant biodegradation rates in an estuarine environment. Applied and Environmental Microbiology 45, 103109.

[24] Spain J.C. (1983). Unpublished data, cited in reference

[41].

[25] Larson R.J. (1983). Comparison of biodegradation rates in laboratory screening studies with rates 159,171.

in natural waters.

Residue

Reviews

85,

[26] Vashon R.D. & Schwab B.S. (1982). Mineralization of linear alcohol ethoxylates and linear alcohol ethoxy sulfates at trace concentrations in estuarlne water. Environmental Science and Technology 16, 433-436.

[27] Cripe C.R., Walker W.W.,

Pritchard P.H. & Bourquin A.W. (1987). A shake-flask test for estimation of biodegradability of toxic organic substances in the aquatic environment. Ecotoxicology and Environmental Safety 14, 239-251.

[28] Larson R.J. & Ventullo R.M.

(1986). Kinetics of biodegradation of nitrilotriacetic acid (NTA) in an estuarine environment. Ecotoxicology and Environmental Safety 12, 166-179.

[29] Larson R.J. & Payne A.G. (1981). Fate of the benzene ring of linear alkylbenzene Microbiology

sulfonate in natural waters. 41, 621-627.

Applied

and

Environmental

[30] Monod J. (1949). The growth of bacterial cultures. Annual Reviews in Microbiolo~v 3, 371-394.

[31] Paris D.F., Steen W.C. & Burns L.A. (1982). Microbial transformation kinetics of organic compounds. In The Handbook of Environmental Chemistry Vol. 2 Part B, edited by O. Hutzinger, pp 73-81. SpringerVerlag: Berlin. [32] Baughman G.L., Paris D.F. & Steen W.C. (1980). Quantitative expression of biotransformation rate. In Biotransformation and Fate of Chemicals in the Aquatic Environment, edited by A.W Maki, K.L. D i c k s o n & J. Cairns, pp 105-111. American Society for Microbiology: Washington.

[33] D a l t o n H. & S t i r l i n g

D.I. (1982). C o - m e t a b o l i s m . Philosophical Transactions of the Royal Society of London B 297, 481-496.

1277

[34] Alexander M. (1981). Biodegradation concern. Science 211, 132-138.

of

chemicals

of

environmental

[35] Wang Y., Subba-Rao R.V. & Alexander M. (1984). Effect of substrate concentration and organic and inorganic compounds on the occurrence and rate of mineralization and co-metabolism. ~Dolled and Environmental Microbiolo~v 47, 1195-1200. [36] Howard P.H. & Banerjee S. (1984). Interpreting results from biodegradability tests of chemicals in soil and water. Environmental Toxicology and Chemistry 3, 551-562. [37] Banerjee S., Howard P.H., Rosenberg A.M., Dombrowki A.E., Sikka H. & Tullis D.L. (1984). Development of a general kinetic model for b i o d e g r a d a t i o n and its application to chlorophenols and related compounds. Environmental Science and Technology 18, 416-422. [38] Paris D.F., Lewis D.L., Barnett J.T° & Baughman G.L. (1975). Cited in reference [32]. [39] Smith J.H., Mabey W.A., Bohonos N., Holt B.R., Lee S.S., Bomberger D.C. & Mill T. (1978). Cited in reference [32].

Chiu T-W,

[40] Paris D.F., Steen W.C., Baughman G.L. & Barnett J.T. (1981). Secondorder model to predict microbial degradation of organic compounds in natural waters. Applied and Environmental Microbiology 41, 603-609. [41] Kle~ka G.M. (1985). Biodegradation. In Environmental Exposure from Chemicals Vol. I, edited by W. Brock Neely & G.E. Brau, pp 109-155. CRC Press Inc: Florida. [42] Kollig H.P., Parrish R.S. & Holm H.W. (1987). An estimate of the variability in biotransformation kinetics of xenobiotics in natural waters by aufwuchs communities. Chemosphere 16, 49-60. [43] Paris D.F., Wolfe N.L. & Steen W.C. (1982). Structure-activity relationships in microbial transformation of phenols. Applied and Environmental Microbiology 44, 153-158. [44] Paris D.F., Wolfe N.L., Steen W.C. & Baughman G.L. (1983). Effect of phenol molecular structure on bacterial transformation rate constants in pond and river samples. Applied and Environmental Microbiology 45, 1153-1155. [45] Paris D.F., Wolfe N.L. & Steen W.C. (1984). Microbial transformation of esters of chlorinated carboxylic acids. Applied and Environmental Microbiolo~v 47, 7-11. [46] Rogers J~E., Li S-M.W. & Fellce L.J. (1984). Microbial Transformation Kinetics of Xenobiotics in Aquatic Environment. Report to U.S. Environmental Protection Agency EPA-600/3-84-043. [47] Lewis D.L. & Holm H.W. (1981). Rates of transformation of methyl parathion and diethyl phthalate by aufwuchs microorganisms. Applied and Environmental Microbiology 42, 698-703.

1278

[48] Lewis D.L., Holm H.W. & Hodson R.E. (1984). Application of single- and multiphasic Michaelis-Menten kinetics to predictive modeling for aquatic ecosystems. Environmental Toxlcolo~y and Chemistry 3, 563-574. [49] Lewis D.L. & Gattle D.K. (1988). Prediction of substrate removal rates of attached microorganisms and of relative contributions of attached and suspended communities at field sites. Applied and Environmental Microblolo~v 54, 434-440. [50] Wright R.T. (1978). Measurement and significance of specific activity in the heterotrophic bacteria of natural waters. APPlied and Environmental Microbiology 36, 297-305. [51] Lehnlnger A.L.

(1972). Biochemistry. Worth Publishers Inc: New York.

[52] Painter H.A. (1983). Metabolism and physiology of aerobic bacteria and fungi. In Used Water Treatment Vol. 2, pp 11-75. Academic Press: London. [53] Parsons T.R. & Strickland J.D. (1962). particulate organic carbon by heterotrophic Deep Sea Research 8, 211-222.

On the production of processes in sea water.

[54] Wright R.T. & Hobble J.E. (1965). The uptake of organic lake water. Limnolo~v and Oceano~raDhv i0, 22-28.

solutes

in

[55] Wright R.T. & Hobble J.E. (1966). Use of glucose and acetate bacteria and algae in aquatic ecosystems. Ecology 47, 447-464.

by

[56] Pfaender F.K. & Bartholomew G.W. (1982). Measurement of aquatic biodegradation rates by determining heterotrophic uptake of radiolabeled pollutants. APPlied and Environmental Microbiology 44, 159-164. [57] Wright R.T. (1978). Measurement and significance of specific activity in the heterotrophic bacteria in natural waters. A~Dlied and Environmental Microbiology 36, 297-305. [58] Ladd T.I., Ventullo R.M., Wallis P.M. & Costerton J.W. (1982). Heterotrophic activity and biodegradation of labile and refractory compounds by groundwater and stream microbial populations. ADDlied and Environmental Microbiology 44, 321-329. [59] Lewis D.L., Hodson R.E. & Hwang H. (1988). Kinetics of mixed microbial assemblages enhance removal of highly dilute organic substrates. APPlied and Environmental Microbiology 54, 2054-2057. [60] Simkins S. & Alexander M. (1984). Models for mineralisation with variables of substrate concentration and population Applied and Environmental Microbiology 47, 1299-1306. [61] Alexander M. (1985). Biodegradation of organic Environmental Science and Technology 18, 106-111.

kinetics density.

chemicals.

1279

[62] Schonborn W. (1986). Historical developments and ecological fundamentals. In Biotechnology. Vol. 8 Microbial Degradations, edited by W. Sch~nborn, pp 3-41. VCH: Weinheim. [63] Anderson D.J., Day M.J., Russell N.J. & White G.F. (1990). Die-away kinetic analysis of the capacity of epilithic and planktonic bacteria from clean and polluted river water to biodegrade sodium dodecyl sulfate. Applied and Environmental Microbiology 56, 758-763. [64] Simkins S. & Alexander M. (1985). Nonlinear estimation of the parameters of Monod kinetics that best describe mineralization of several substrate concentrations by dissimilar bacterial densities. Applied and Environmental Microbiology 50, 816-824. [65] Simkins S., Mukherjee R. & Alexander M. (1986). Two approaches to modeling kinetics of biodegradation of growing cells and application of a two-compartment model for mineralization kinetics in sewage. Applied and Environmental Microbiology 51, 1153-1160. [66] Games L.M. (1982). Field validation of Exposure Modeling System (EXAMS) in a flowing stream. In Modelin~ the Fate of Chemicals in the Aouatic Environment, edited by K.L. Dickson, A.W. Maki and J. Cairns, pp 325-346. Ann Arbor Science Publishers: Michigan. [67] Spain J.C. & van Veld P.A. (1983). Adaptation of natural microbial communities to degradation of xenobiotic compounds: effects of concentration, exposure time, inoculum, and chemical structure. APPlied and Environmental Microbiology 45, 428-435. [68] Rubln H.E., Subba-Rao R.V. & Alexander M. (1982). Rates of mineralisation of trace concentrations of aromatic compounds in lake water and sewage samples. Applied and Environmental Microbiology 43, 1133-1138. [69] Schmidt S.K., Scow K.M. & Alexander M. (1987). Kinetics of p-nitrophenol mineralization by a Pseudomonas sp.: effects of second substrates. Applied and Environmental Microbiology 53, 2617-2623. [70] Boethling R.S. & Alexander M. (1979). Effect of concentration of organic chemicals on their biodegradation by natural microbial communities. Applied and Environmental Microbiology 37, 1211-1216. [71] Jannasch H.W. (1967). Growth of marine bacteria at limiting concentrations of organic carbon in seawater. Limnology and Oceanography 12, 264-271. [72] Spain J.C., Pritchard P.H. & Bourquin A.W. (1980). Effects of adaptation on biodegradation rates in sediment/water cores from estuarine and fresh water environments. Applied and Environmental Microbiology 40, 726-734. [73] Bouwer E.J. (1985). Secondary utilization of trace organic compounds in biofilms. Environmental Progress i, 43-45.

1280

[74] Costerton J.W., Geesey G.G. & Cheng R-J Scientific American 238, 86-95.

(1978).

How bacteria

stick.

[75] Schmidt S.K., Alexander M. & Shuler M.L. (1985). Predicting threshold concentrations of organic substrates for bacterial growth. Journal of Theoretical BioloKy 114, 1-8. [76] Subba-Rao R.V., Rubin H.E. & Alexander M. (1982). Kinetics and extent of mineralization of organic chemicals at trace levels in freshwater and sewage. ADvlied and Environmental Microbiology 43, 1139-1150. [77] R u b i n H.E., S u b b a - R a o R.V. & A l e x a n d e r M. (1982). Rates of mineralization of trace concentrations of aromatic compounds in lake water and sewage samples. Applied and Environmental Microbiology 43, 1133-1138. [78] Harder W. & Dijkhuizen L. (1982). Strategies of mixed substrate utilization in microorganisms. Philosovhlcal Transactions of the Royal Society of London B 297, 459-480. [79] LaPat-Polasko L.T., McCarty P.L. & Zehnder A.J.B. (1984). Secondary substrate utilization of methylene chloride by an isolated strain of Pseudomonas sp. Aovlied and Environmental Microbiology 47, 825-830. [80] Schmidt S.K. & Alexander M. (1985). Effects of dissolved organic c a r b o n and s e c o n d s u b s t r a t e s on the b i o d e g r a d a t i o n of organic compounds at low c o n c e n t r a t i o n s . Applied and Environmental Microbiology 49, 822-827. [81]

S t i f f M.J., R o o t h a m R.C. & Culley G.E. (1973). The effect of temperature on the removal of non-ionic surfactants during small-scale activated-sludge treatment - I. Comparison of alcohol ethoxylates with a branched-chain alkyl phenol ethoxylate. Water Research 7, 1003I010.

[82] Stiff M.J. & Rootham R.C. (1973). The effect of temperature on the removal of non-ionic surfactants during small-scale activated-sludge treatment - II. Comparison of linear alkyl phenol ethoxylate with a branched-chain alkyl phenol ethoxylate. Water Research 7, 1407-1415. [83] Wardell J.N., Brown C.M. & Flannigan B. (1983). Microbes and surfaces. In Microbes in their Natural Environments, edited by J.H. Slater, R. W h i t t e n b u r y & J.W.T. Wimpenny, pp 351-378. S o c i e t y for G e n e r a l Microbiology and Cambridge University Press: Cambridge. [84]

Jones J.G. (1982). Activities of aerobic bacteria and anaerobic bacteria in lake sediments and their effect on the water column. In Sediment Microbiology, edited by D.B. Nedwell & C.M. Brown, pp 107145. Society for General Microbiology and Academic Press: London.

[85] H a m e l i n k J. (1980). B i o a v a i l a b i l i t y of c h e m i c a l s in aquatic environments. In Biotransformation and Fate of Chemicals in the Aquatic Environment, edited by A.W. Maki, K.L. Dickson & J. Cairns, pp 56-62. American Society for Microbiology: Washington.

1281

[86] Pritchard P.H. (1987). Assessing the biodegradation of sediment associated chemicals. In Fate and Effects of Sediment-Bound Chemicals in Aquatic Systems, edited by K.L. Dickson, A.W. Maki & W.A. Brungs, pp I09-135, Society of Environmental Toxicology and Chemistry and Pergamon Press: New York. [87] Shimp R.J. & Young R.L. (1988). Availability of organic chemicals for b i o d e g r a d a t i o n in settled bottom sediments. Ecotoxicolo~y and Environmental Safety 15, 31-45. [88] Pavlou S.P. (1980). Thermodynamic aspects of equilibrium sorption of persistent organic molecules at the sediment-water interface: a framework for predicting distributions in the aquatic environment. In Contaminants and Sediments, Vol. 2, edited by R.A. Baker, pp 323-332. Ann Arbor Science Publishers: Michigan. [89] Weber J.B. & Coble H.D. (1968). Microbial decomposition of diquat adsorbed on montmorillonite and kaolinite clays. Journal of Agricultural and Food Chemistry 16, 475-478. [90] J~rgensen B.B. (1977). The sulfur cycle of a coastal marine sediment (Limfjorden, Denmark). Limnolo~v and Oceano£raDhv 22, 814-832. [91] Jones J.G. & Simon B.M. (1980). Decomposition processes in the profundal region of Blelham Tarn and the Lund Tubes. Journal of Ecolo~v 68, 493-512. [92] Suflita J.M. & Miller G.D. (1985). The microbial ehlorophenolic compounds in ground water aquifers. Toxicology and Chemistry 4, 751-758.

metabolism of Environmental

[93] Mikesell M.D. & Boyd S.A. (1985). Reductive dechlorination of the pesticides 2,4-D, 2,4,5-T, and pentachlorophenol in anaerobic sludges. Journal of Environmental Quality 14, 337-340. [94] Mikesell M.D. & Boyd S,A. (1986). Complete reductive dechlorination and mineralization of pentachlorophenol by anaerobic microorganisms. Applied and Environmental Microbiology 52, 861-865. [95] Saldick J. (1974). Biodegradation Microbiology 28, i004-I008.

of

cyanuric

acid.

Applied

[96] Spain J.C., van Veld P.A., Monti C.A., Pritehard P.H. & Cripe C.R. (1984). Comparison of p-nitrophenol biodegradation in field and laboratory test systems. Applied and Environmental Microbiology 48, 944-950. [97] Christensen D. (1984). Determination of substrates oxidized by sulfate reduction in intact cores of marine sediments. Limnology and Oceanography 29, 189-192. [98] Painter H.A. & King E.F. (1985). Final Report of the Commission of the European Communities Degradation/Accumulation Sub-Grouo Ring Test Programme 1983-4 on the Assessment of Biodegradability of Chemicals in Water by Manometric Respirometry. DG XI 283/82, REV 5.

1282

[99] Hashimoto M., Kitano M. & Takatsuki M. (1988). 1988 OECD Ring-Test of M ~ h o d s for D e ~ m l n i n g Ready Biodegradability, Chairman's Report of the Meeting of OECD Experts held in Tokyo on 12-14 October 1988 and Final Report on the Evaluation of the Results of the Ring-Test. [i00] Blok J. (1988). Renort of International Round Robin Test for the RDA Biodegradation Method (Determination of Dissolved Oxvgen in Two-Phase C~osed Bottles). [i01] Nyholm N. & Kristensen P. (1987). Screening Test Methods for Assessment of Biodegradabilitv of Chemical Substances in Sea Water: Final Report of CEC Ring-Test Programme 1984-1985. VKI: Denmark. [102] Department of the Environment Digest of Environmental Statistics No, 2 (1979). HMSO: London.

Pollution

[103] Department of the Environment Digest of Environmental Protection and Water Statistics No, 8 (1985). HMSO: London. [104] Fielding M. & Hunt D.T.E. (1984). The nature and concentration of pollutants in water. In Environmental Protection: Standards. Compliance a~d Costs, edited by T.J. Lack, pp 33-54. Water Research Centre and Ellis Horwood Ltd.: Chichester. [105] Herbert R.A. (1982). Nitrate dissimilation in marine and estuarine sediments. In Sediment MicrobioloK¥ , edited by D.B. Nedwell & C.M. Brown, pp 53-71. Society for General Microbiology and Academic Press: London. [106] Northumbrian Water Collated Data on Water Quality 1986-1987 (1987). [107] Riley J.P. & Chester R. Academic Press: London.

(1971).

Introduction

to Marine

Chemistry.

[108] Stanley S.O., Pearson T.H. & Brown C.M. (1978). Marine microbial ecosystems and the degradation of organic pollutants. In The Oil Industry and Microbial Ecosystems, edited by K.W.A. Chater & H.J. Somerville. Heydon and Son: London. [109] Sorokin Y.I. & Kadota H. (1972), Techniques for the Assessment of Microbial Production and Decomposition in Fresh Waters. International Biological Programme Handbook No. 23. Blackwell Scientific Publications: Oxford. [ii0] Austin B. (1988). Cambridge.

Marine Microbiology.

Cambridge

University

Press:

[iii] Analvsis of Organic Micropollutants in Water (COST 64b bis). A Comprehensive List of Polluting Substances Which Have Been Identified in Various Fresh Waters. Effluent Discharges, Aouatic Animals and Plants. and Bottom Sediments. Third Edition 1979, Vol. I and Vol. II. Commission of the European Communities XII/992, i & 2/79-EN.

1283

[112] Buisson R.S.K., Kirk P.W.W. & Lester J.N. (1988). The behaviour of selected chlorinated organic micropollutants in the activated sludge process: a pilot plant study. Water, Air and Soil Pollution 37, 419432.

[113] Bishop P., Arvin E. & Mortensen E. (1987). Biodegradation of phenoxy acids in biofilms. Water pollution Research Report 6, Behaviour of Organic MicroDollutants in Biological Waste Water Treatment, pp 39-64. Proceedings of COST 641 Workshop, Copenhagen May 1987. EUR 11356. [114] SAC Scientific UK Waters.

(1987).

Survey of Potentially Dangerous

Substances

in

[115] ECETOC (1985). Technical Report No. 18: Harmonisation of Ready Biodegradability Tests. European Chemical Industry Ecology and Toxicology Centre: Brussels. [116] Shimp R.J. & Young R.L. (1987). Comparison of radiolabeled substrate methods for measuring biodegradation in marine environments. Ecotoxicolo~v and Environmental Safety 14, 223-230. [117] Jones S.H. & Alexander M. (1986). Kinetics of mineralization of phenols in lake water. Applied and Environmental Microbiology 51, 891-897. [118] Lindgaard-J~rgensen P. (1987). Use of biodegradation tests to predict the fate of chemicals in waste water treatment plants, the experiences of the Water Quality Institute. Water Pollution Research Report 6. Behaviour of Organic Microvollutants in Biological Waste Water Treatment, pp 88-96. Proceedings of COST 641 Workshop, Copenhagen May 1987. EUR 11356. [119] Aellon C.M., Swindoll C.M. & Pfaender F.K. (1987). Adaptation to and biodegradation of xenobiotic compounds by microbial communities from a pristine aquifer. Applied and Environmental Microbiology 53, 22122217. [120] Swindoll C.M., Aelion C.M., Dobbins D.C., Jiang O., Long S.C. & Pfaender F.K. (1988). Aerobic biodegradation of natural and xenobiotic organic compounds by subsurface microbial communities. Environmental Toxicology and Chemistry 7, 291-299. [121] Blanchard F.A., Takahashi I.T. & Alexander H.C. (1976). Biodegradability of [14C] methylcellulose by activated sludge. Applied and Environmental Microbiology 32, 557-560. [122] McKinley V.L., Federle T.W. & Vestal J.R. (1983). Improvements in and environmental applications of double-vial radiorespirometry for the study of microbial mineralization. Applied and Environmental Microbiology 45, 255-259. [123] Lehmicke L.G., Williams R.T. & Crawford R.L. (1979). 14C-mostprobable-number method for the enumeration of active heterotrophic microorganisms in natural waters. ADDlied and Environmental Microbiology 38, 644-649.

1284

[124] Somerville C.C., Monti C.A. & Spain J.C. (1985). Modification of the 14C-most-probable-number method for use with nonpolar and volatile substrates. APplied and Environmental Microbiology 49, 711-713. [125] Anderson D.J., Day M.J., Russell N.J. & W h i t e G.F. (1988).Temporal and geographical distributions of epilithic sodium dodecyl sulfatedegrading bacteria in a polluted South Wales river. Applied and Environmental Microbiolo~v 54, 555-560. [126] Madsen E.L. & Alexander M. (1985). Effects of chemical speciation on the mineralization of organic compounds by microorganisms. Applied and Environmental MierobioloKy 50, 342-349.

(Received

in G e r m a n y

1 November

1990)