Aerobic microbial transformation of pesticides in surface water

Abstract

Aerobic transformation by microbial organisms is a dissipation process of pesticides in surface water, but the corresponding information is much less available as compared with their microbial degradation in soil. Bacteria freely floating in water or associated with suspended particles are the key microorganisms degrading pesticides; their species and populations, however, depend on sites and seasons. The various factors related to pesticide properties, experimental conditions and characteristics of surface water are involved in the complex control of microbial processes. Bottom sediment and macrophytes with associated biofilms not only act as sink for pesticides but also provide the habitat for both bacteria and fungi that degrade pesticides. More understanding of each factor is necessary to utilize laboratory biodegradation data for the refined assessment of pesticide behavior in surface water.

Introduction

Surface water in rivers, ponds and lakes is possibly contaminated with pesticides via various routes either from a point source such as sewage plants and sewer overflows or a diffuse one along a water course, such as surface runoff and soil erosion. The contamination from point sources is reported to be dominant in several European catchments and constitutes 20–80% of the pesticide load in rivers.¹⁾ Water analysis along two German streams showed that direct runoff from farmyards due to cleaning spray equipment significantly contributed to the total pesticide load of seven insecticides and fungicides, while outlet from sewage treatment plants was an important source for 13 herbicides.²⁾ The authors also reported that the applied amount of pesticide to the field correlated with its load to the streams, but runoff from cultivated fields was a greater contributor for hydrophilic pesticides.

Pesticides are partitioned to constituents of surface water such as biota, suspended particles, and bottom sediment and transported by a water flow with concomitant chemical and biological transformation, as illustrated in Fig. 1.^1,3,4) Partition and chemical processes are individually evaluated by standard batch adsorption–desorption, hydrolysis, and photolysis studies. A small-scale laboratory water-sediment system with a typical sediment-water ratio of 1 : 3–1 : 10 (w/v) is utilized to evaluate biological processes in the assessment of pesticide behavior⁴⁾; however, aerobic transformation and mineralization by microbes in surface water have been recently proposed as an additional study.^5–7) The species and density of microbes are generally site-/season-specific and are significantly affected by the status of the water body. The seasonal stratification in lakes along with temperature changes controls the depth-dependent concentrations of oxygen and nutrients, and the change in the flow rate in rivers leads to a different concentration of particulates.⁸⁾ Sediment-water interfaces showing various physical, chemical and biological gradients are deeply involved in the recycling of elements.⁹⁾ Furthermore, sunlight with wavelengths shorter than 400 nm is reported to play a significant role in the production of dissolved organic carbons (DOC) from estuary sediments.¹⁰⁾

Fig. 1. Behavior of pesticides in surface water.

In this review, we first discuss the relative contribution of each environmental process to the dissipation of pesticide in surface water together with the importance of the microbial process especially by bacteria and fungi. After introduction of the study design assessing aerobic transformation with its kinetic analysis, the aerobic transformation profiles of pesticides are examined through a literature survey by considering the involvement of specific microbes in surface water. The factors controlling microbial transformation in surface water are discussed. Finally, an overview summary is provided, including issues that remain to be solved. The chemical structure of each pesticide appearing in this review is listed in the Appendix.

Processes Controlling Pesticide Behavior

Pesticides undergo various processes, as illustrated in Fig. 1. In rivers and streams, the pesticide concentration at an entrance point is highly diluted by the water flow by a factor of >10,¹¹⁾ while dispersion predominates in dilution for a stagnant water body such as ponds and lakes, which have a much longer hydraulic residence time.^12,13) The dilution factor is also estimated by hydrodynamic models¹⁴⁾ or monitoring the concentration of a tracer (Cl⁻) in a water system.^15,16) Depending on its physico-chemical properties, pesticide is partly adsorbed or associated with suspended particles, colloids, and dissolved organic matters (DOM),^4,17,18) reaches bottom sediment via sedimentation or diffusion,^17,19) and escapes from the water body via volatilization.^4,14) The sorption of pesticides is evaluated by an adsorption coefficient determined in a batch study, generally assuming a linear or Freundlich isotherm. Sedimentation loss is estimated for a fraction of suspended particulates adsorbing pesticides by considering an average depth of water and a settling rate.^14,20) The two-layer model is conveniently used to describe the volatilization of pesticides from water to air.^14,20)

Abiotic hydrolysis follows the pseudo-first-order kinetics at a very dilute concentration of pesticide, and its contribution is usually limited at neutral pH and low temperatures except for pesticides having an ester linkage.¹⁷⁾ In a clear water column, direct photolysis by sunlight plays a role in the dissipation of pesticides having a measurable U.V. absorption at >290 nm. A direct photolysis rate with seasonal and depth dependence can be theoretically estimated using a quantum yield of pesticide.²¹⁾ Furthermore, indirect photolysis including reactions with active oxygen species such as singlet oxygen (¹O₂) and hydroxyl radical (·OH) becomes important for some pesticides.²²⁾ Its contribution is conveniently assessed by the typical reaction rate constants and steady-state concentrations of active oxygen species in natural water.^14,20)

Microbial degradation is another important dissipation route, as is bio-sorption. Many kinds of microbes are present, freely floating in water or attached to a substratum, such as macrophytes and sediment, and they grow using different carbon and energy sources.²³⁾ Algae and protozoa can absorb and metabolize pesticides,²⁴⁾ but bacteria and fungi contribute more to pesticide dissipation through their ubiquity.^3,8,18) Bacteria are more diversely populated, with their activity depending on the habitat,²⁵⁾ while fungi tend to grow on a substratum, such as detritus, rather than in water.^26,27) Bacteria metabolize pesticides via various enzymatic reactions,²⁸⁾ and extra-cellular enzymes such as laccases,²⁹⁾ may participate in their fungal degradation. In addition to an active uptake by microbes, pesticides are adsorbed onto surfaces of microbes and biofilm, a gel-type complex mixture of microbial cells, detritus, and extra-cellular polymeric substances (EPS).^30,31) Since the constituents of EPS, such as polysaccharides and proteins, have various functional groups, they provide adsorption sites for various pesticides via hydrophobic, electrostatic, and hydrogen-bonding interactions. Microbial degradation can be studied by using the experimental designs prescribed in avalable guidelines.^5–7) The contribution of hydrolysis should be examined using a sterile control. Either a microbial community concentrated by a filtration technique or microbes isolated from surface water may be conveniently used to identify potential transformation paths.

The relative contribution of each process highly depends on the chemical structure of the pesticide. Through the dissipation of parathion (40) and malathion (49) in estuarine water, sunlight photolysis was found to be much less important than hydrolysis and microbial degradation, respectively.³²⁾ N-Methyl carbamates dissipated mainly via hydrolysis and photolysis in lake waters, while bacterial and fungal degradation played more significant roles for N-phenyl carbamates.²¹⁾ By comparison with sterile controls, a high contribution of biotic processes (>70%) was reported for chlorothalonil (4) and pendimethalin (55) in the nursery recycling pond water but less (20–50%) for organophosphorus pesticides (OPs) possibly due to the less relevant microbial activity in high salinity.³³⁾

Outdoor monitoring of pesticide dissipation is another approach to estimate the relative contribution of each process. Through a series of outdoor studies using open and closed bottles containing intact or filtered (0.45 µm) river water, the contribution of each process was assessed for thirteen OPs.³⁴⁾ Sorption to suspended particles retarded the dissipation of hydrophobic OPs. The degradation under sterile conditions was not available, but the reported hydrolysis data¹⁷⁾ showed the importance of biodegradation and photolysis with much less contribution of volatilization. A recent study on four OPs, conducted at controlled temperatures with sterilization, clarified that dominant process of dissipation is highly dependent on their chemical structures.³⁵⁾ More detailed monitoring of pentachlorophenol (2) dissipation was conducted outdoors in experimental flowing channels including aquatic plants and unglazed ceramic tiles for biofilm development.³⁶⁾ Insignificant volatilization of (2) from water and attenuation of photolysis with the water depth were confirmed. The dominant process was biodegradation (26–46%) by microbes attached to macrophytes, biofilm, and sediment. For persistent and hydrophobic pesticides, dilution or adsorption to the bottom sediment was found to be more significant in farm ditches.¹⁶⁾

Study Design of Aerobic Microbial Transformation

Excellent reviews are available on various biodegradation methods, each of which was developed to simulate a specific environmental situation.^37–39) The important experimental factors are the origin of the inoculum, the concentration of a chemical, the inorganic or organic nutrients used, and the system conditions such as aerobicity, temperature, mixing, and the duration of incubation. The changes in DOC, total organic carbon (TOC) and biochemical oxygen demand (BOD), and formation of carbon dioxide are generally used as a substrate-independent index to examine the ultimate biodegradation of a chemical, while substrate-specific methods such as gas (GC) and liquid (LC) chromatography are applied to assess the primary biodegradation. In contrast to screening and inherent biodegradability tests at high concentrations of a chemical and biomass (activated sludge or sewage plant secondary effluent), the microbial process of a chemical in surface water should be examined at an environmentally relevant ppb–ppm level using microbial community therein. From this viewpoint, modified die-away methods have been proposed using a concentrated microbial community by filtration of river water and validated for various simple aromatics with TOC as a degradation index.^40,41) By using these methods, the biodegradation of various pesticides in Japanese river and pond waters at 20–30°C was examined using GC in the presence of 0.1–0.2% organic nutrients.^42–44) Their biodegradation half-lives significantly varied among surface waters sampled at different sites and seasons. The nutrient effects in this system were examined through the biodegradation of chlornitrofen (57) using river waters.⁴⁵⁾ The moderate biodegradation of (57) (80% after 18 days) was not affected by the addition of a mixture of inorganic salts, while the presence of 0.2% polypeptone resulted in complete degradation after 5 days.

To simulate the biodegradation of pesticides at an environmentally relevant concentration, Cripe et al.⁴⁶⁾ developed a modified river die-away test under shaking using fresh river water without additional nutrients and examined the effect of sediment (0.5 g L⁻¹). A comparison of dissipation under non-sterile and sterile conditions showed that the degradation of parathion-methyl (41) was markedly enhanced by sediment-associated microbes while partition to sediment predominated in the dissipation of more hydrophobic methoxychlor (9). Sediment-mediated biodegradation of fenthion (43) was dominant in salt-marsh waters by using the same method, as compared with hydrolysis and biodegradation in water.⁴⁷⁾ The usage of radio-labeled (¹⁴C) compound gives information on a product distribution in a test system as well as the extent of mineralization. Dissociative pentachlorophenol (2) was partly adsorbed to sediment particles (10–20%) in river waters, but the presence of sediment enhanced the degradation of (2) with a shorter acclimation period, showing the involvement of sediment-associated microbes.⁴⁸⁾ The slower degradation of 2,4-d (10) but with shorter acclimation periods than those of (2) and no mineralization of atrazine (52) in the same river water indicated a different population of microbes specifically degrading each pesticide.⁴⁹⁾

There are three test guidelines available for pesticides, and the details are summarized in Table 1. In all cases, freshly collected surface water and sediment are used, and the decline of a pesticide is monitored to determine the degradation rate constant, which can be used in modeling for assessment. OECD 309⁵⁾ requires two low concentrations of a pesticide, most likely to examine the possible concentration effect on degradation, as discussed in more detail later. Agricultural Canada T-1-255⁶⁾ characteristically requires that the study be carried out under illumination with a fluorescent light unless a pesticide is photo-labile. U.S. EPA OPPTS 835.3170⁷⁾ recommends the collection of water 6 cm below the surface, which may exclude unexpectedly high adsorption or degradation by neustons and surface microlayer constituents.⁴⁾ The top sediment should be collected to represent the effect of aerobic microbes as well as biofilm. Identification of major metabolites with their formation rates is important to assess the impact on aquatic species.

Table 1. Study design of aerobic degradation

Guideline	OECD 309	OPPTS 835.3170	Agricultural Canada
	Simulation Biodegradation	Shake Flask Die-Away Test	Guideline T-1-255
Chemical	Radiolabel or non-radiolabel	Radiolabel or non-radiolabel	Radiolabel
Concentration	Two (differ by a factor of 5–10)≤0.1 ppm and preferably <1–10 ppb)	Typically, 0.2 ppm	One or two (10-fold difference)
Co-solvent	<1% (volatile organic)	Volatile (acetone)	Minimum volume (acetone or ethanol)
Medium	Surface water (fresh, brackish, marine) or+sediment (top 5 mm; high and low OC; 0.01–1 g L−1)	Surface water (6 cm below surface; river, lake, estuary)±sediment (top 5–10 mm; 0.5 g L−1)	Pelagic (water only)+sediment (if it is a major sink for pesticide)
Agitation	ca. 100 rpm or gentle stirring	140–150 rpm	Static, shaking or bubbled with air
Illimination	Dark (preferred) or diffuse light	n.d.	Fluorescent light for cultivation
Temperature	20–25°C (±2°C)	25 (or site temperature)±2°C	3–8°C and 20–30°C
Sterilized control	Autoclaving (121°C, 20 min) or+NaN3, HgCl2 or formalin or γ-ray irradiation	Effect, examined by addition of ca. 8% formalin	Necessary, with standard plate counts
Study period	≤60 days (max., 90 days)	≤28 days or more	≤ 12 months
Sampling	≥5	≥6	≤16
Mass balance	90–110% (14C), 70–110% (non-14C)	n.d.	Should be determined
Degradate identification	≥10% or continuously increasing and CO2	n.d.	≥10% and CO2
Kinetics	Rate constant, lag phase	1st- and 2nd (microbial biomass-based)-order rate constants, sorption coefficient	Rate constant (parent, major metabolite)

n.d., no description. OC, organic carbon.

Kinetic Analysis

The apparent biodegradation rate constant is estimated from the decline curve of a relevant parameter such as a pesticide concentration, assuming various kinetic models.^38,50,51) When an acclimation or lag period (T_a≥0) is observed before the initiation of apparent biodegradation, kinetic analysis using “t−T_a” as a time scale should be conducted. The simplest expression on the biodegradation of a substrate (S) is the pseudo-first-order reaction kinetics below, under the constant concentration of biomass (B).

  

(1)

k₁ and k_b are the apparent first-order and biomass-normalized second-order rate constants, respectively. The bacterial biomass is estimated by using a microscope to count cell numbers after staining, counting colonies on a plate inoculated with serially diluted samples based on the most-probable-number (MPN) technique (colony-forming unit, c.f.u.), and using the turbidometric method or a dry-weight measurement.⁵²⁾ The fungal biomass can be estimated by converting the amount of extracted ergosterol to the carbon weight.²⁷⁾ Paris et al.⁵³⁾ have examined the microbial degradation of three pesticides at ppm levels in surface waters collected from 40 U.S. sites. The coefficient of variation for the estimated k_b values was less than 65% at all sites with insignificant site-variability. The effect of inorganic salts as nutrients on the k_b value was examined through the biodegradation of five pesticides in the waters of ponds and rivers.⁵⁴⁾ Most microbes detected as colonies on enumeration plates could degrade each pesticide, and both k₁ and biomass increased in parallel with the nutrient amendment, resulting in an almost constant k_b. However, the meaning of k_b should be carefully considered, since the obtained biomass does not always represent specific degraders of a pesticide. When the biomass of non-degraders only increases with keeping k₁ almost constant, the calculated k_b value decreases with the microbial growth.^51,55)

Most of the biodegradation data were analyzed using Eq. 1, but many other kinetic models have been applied to explain deviations from the first-order kinetics. The microbial growth can be described by the Monod equation.⁵⁰⁾

  

(2)

μ_max and K_s are the maximum specific growth rate and the substrate concentration at the growth rate of 0.5 μ_max, respectively. By introducing a yield coefficient (Y) that means the conversion efficiency of a substrate into cells, the reaction rate can be described.

  

(3)

Under the usual environmental situation for contaminants, that is [S]≪K_s, Eq. 3 can be approximated to be the second-order reaction kinetics.

  

(4)

Other models, such as the zero-order, logarithmic, and logistic models, have also been applied.^50,51) When a small number of active degraders grow on a substrate at [S]≪K_s, the logistic kinetics is applied. Under the condition of [S]≫K_s, zero-order or logarithmic kinetics explains the biodegradation when the substrate uptake by microbes is saturated and will be saturated, respectively. The applicability of each kinetic model depends on the concentrations of a substrate and biomass. The mineralization study of phenols in lake water indicates that there is a shift from the first-order and logistic kinetics to the Monod and logarithmic ones with an increasing substrate concentration; however, this is dependent on the season for sampling the lake water.⁵⁶⁾ When a specific enzyme reaction represents the primary biodegradation of a substrate, the Michaelis–Menten kinetics, which resembles Eq. 3, describes the decline of a substrate, for example when specific degrading bacteria and fungi are used.^21,57,58)

These kinetic analyses are applied to microbial degradation studies in closed bottles. When a flow-through reactor is utilized to investigate the biodegradation and sorption of a substrate by microbes, the change of the substrate concentration in effluents with time should be kinetically analyzed. Schrap et al.⁵⁹⁾ developed a chemostat system by constantly introducing a growth medium with dicloran (5) to surface water and estimated its first-order degradation rate. By using river biofilms grown on polycarbonate slides in the flow-through bioreactor, the rate and coefficient of linear adsorption for seven pesticides were successfully estimated.⁶⁰⁾

Microbes in Surface Water

Bacteria and fungi play significant roles in the biodegradation of pesticides.²³⁾ Bacteria having various shapes grow with either inorganic or organic carbon, depending on the trophic level of water. Freshwater bacterial assemblage is distinct from that in terrestrial communities, as demonstrated by the survey of 16S ribosomal RNA sequences, and dominant taxa are different from marine ones.⁶¹⁾ Actinobacteria commonly found in lakes may comprise a large fraction of bacterioplankton in a water column, while some bacteria in an aerobic epilimnion, such as Proteobacteria (α and β), are associated with suspended particles. Different bacterial species were distributed between a water column and bottom sediment of the eutrophic lake.⁶²⁾ In the water columns of rivers,^63,64) ponds⁴³⁾ and lakes,^62,64–66) Gram-negative and rod-shaped bacteria, such as Pseudomonas, Aeromonas, and Alcaligenes sp., are reported to be the dominant species, with both temporal and spatial variations in bacterial communities. Although the seasonal changes in the flow regimes of water are important, the temperature should be the most critical factor controlling the variation via bacterial growth, and the resultant mixing and stratification of a water column are primary causes especially in lakes.⁶¹⁾ Edwards et al.⁶⁷⁾ reported marked seasonal changes in the bacterial biomass by a factor of >10 in a eutrophic lake in the U.K. with depth dependence, most likely due to stratification in summer. The biological interactions are also important, as indicated by the fact that the blooming pulse of cyanobacteria and diatoms are followed by the increase of bacterial counts, likely due to the supply of organic matters from planktons.⁶⁸⁾ Both the temperature dependence and biological interactions in the production and abundance of bacterioplanktons have been confirmed through 8-year monitoring in a hypertrophic lake.⁶⁹⁾ By analyzing several bacterial profiles for 80 bacteria isolated from three ponds, Hashizume et al.⁴³⁾ reported marked horizontal variation in the distribution of these bacteria with their seasonal changes.

Incidentally, bacterial population depend on the nutrient level of a water body, and more counts of bacterial plankton in eutrophic lakes are observed by about an order of magnitude than oligotrophic ones.⁷⁰⁾ By the molecular microbial analysis of lake water amended with four levels of inorganic nutrients, Knapp et al.⁷¹⁾ showed good correlation between the universal small sub-unit ribosomal RNA level of all microbes and the total phosphorus concentration of the water. A similar increase of bacterial cells with the addition of PO₄³⁻ was observed for channel water.⁷²⁾ Microbe growth in clear river water increased in the presence of inorganic nutrients.⁷³⁾ It is noteworthy that oligotrophic bacteria can grow at low concentrations of nutrients under specific cultivation conditions.

Fungi are mostly filamentous and grow well in association with solid surfaces in water body. The static littoral zone, especially the surfaces of leaf litters and sediments, is known as hot spots for fungi that participate in degrading particulate organic matters.^26,27) Niemi et al.⁷⁴⁾ reported mesophilic fungi and Actinomycetes as common species in Finnish rivers and lakes, with their occurrence being related to runoff from soil. Several degradative enzymes, including phenol oxidases, contribute to the fungal decomposition of litters, and the products are utilized by co-existing bacteria as organic nutrients for their growth.⁷⁵⁾

Uptake and Sorption of Pesticides

Uptake of pesticide molecules by microbes is basically prerequisite for aerobic transformation and, therefore, molecular diffusion from a boundary layer around the microbe to its interior through the microbial surface becomes a key process.⁷⁶⁾ When pesticides are taken up, either energy-mediated active transport or passive sorption to microbial surfaces can be postulated. Not only comparison of the uptake between living and dead cells but reversibility in sorption is convenient to distinguish these processes. The uptake rate by living microbes vs. the substrate concentration plot showed saturation kinetics in the biodegradation of four simple organic compounds including m-cresol.⁷⁷⁾ Under similar kinetics, phenol was incorporated into cellular macromolecules of living microbes.⁷⁸⁾ Lal and Saxena⁷⁹⁾ reported the rapid species-dependent uptake of organochlorine pesticides by microbes, likely under the balance between sorption and metabolism. Sorption of these pesticides to microbes was mostly independent of cell viability, indicating a passive process.⁸⁰⁾ The sorption coefficient of pesticide to microbes is generally estimated by using linear and Freundlich isotherms and is known to correlate well with the n-octanol/water partition coefficient (K_ow), as reported for algae.^24,80) Pesticide sorption to fungi has been investigated in relation to the removal of contaminants from water by biosorbents. Studies using Rhizopus anhizus have shown that the sorption isotherm of several pesticides could not be explained by assuming a monolayer surface coverage.⁸¹⁾ Furthermore, the partition of pesticides to the fungal cell walls did not fully account for their sorption and the participation of cytoplasmic components leaking from the cells was highly suspected.^82,83)

Aerobic Transformation of Pesticides

1. Co-metabolism

Aerobic biotransformation of pesticides by microbes can be classified into two types, metabolism for microbial growth and co-metabolism.^3,28,38) Pesticides are only transformed by enzyme(s) attacking the growth substrate in the latter process and not used as an energy source. Therefore, the microbial populations responsible for the transformation do not increase in numbers and biomass. Under this condition, the transformation rate follows the first-order kinetics, as expressed in Eq. 1. Co-metabolism can be conveniently confirmed by the changes in either transformation rate or biomass. MCPA (12) and disulfoton (48) were degraded by bacteria in channel and river waters, respectively, but without any increase of bacterial cell number.^14,72) More than 2-fold dissipation of 2,3,6-trichlorobenzoic acid by microbes in lake water was observed by the addition of sodium benzoate as a growth substrate but without any increase of bacteria in its absence.⁸⁴⁾ Novick and Alexander⁸⁵⁾ reported a doubled yield of products through the incubation of propachlor (33) in eutrophic lake water by addition of glucose, but no microbes using (33) as carbon and energy sources could be isolated. Recently, Pileggi et al.⁸⁶⁾ isolated Pantoea ananatis from Brazilian freshwater and showed its co-metabolic activity on mesotrione (58) by varying the nutrient composition of the medium. The mineralization of propham (20) in lake water was enhanced by ca. 20% in the presence of non-chlorinated hydrocarbons, showing the contribution of co-metabolism.⁸⁷⁾

2. Transformation in surface water

The aerobic transformation and mineralization of pesticides have been examined in various surface waters, roughly in accordance with the experimental conditions prescribed in the test guidelines.^5–7) The contribution of microbes is conveniently assessed by using membrane-filtered or sterile water, and its relative importance as compared with other processes, such as photolysis, has been sometimes examined in outdoor studies.^88–90) Kanazawa⁹¹⁾ proposed a convenient system to grasp the dissipation profiles of pesticides, using mixtures of activated sludge and water extracts of soil and sediment as the inoculum in the nutrient culture medium. The aerobic transformation of many pesticides in darkness has been reported using various surface waters free from large suspended particles, as briefly summarized in Table 2. When pesticide degrades only via hydrolysis and biodegradation, the contribution of biodegradation (B%) can be expressed by Eq. 5.

  

(5)

k_ns is the apparent first-order rate constant estimated under non-sterile conditions. k_s and k₁ are the first-order rate constants of hydrolysis under sterile conditions and biodegradation, respectively. t_1/2 is the half-life calculated from each rate constant.

Table 2. Aerobic degradation of pesticide in non-sterile natural water

No.	Pesticide	NW (pH)1)	Conditions2)	Deg/Miner, L3)	r4)	Ref.
(1)	lindane	M (7.7)	2 ppm, S, 21	5.5w, —	ca. 0.1	89
(2)	pentachlorophenol	Po (7–8)	2 ppm, S, f	80d, —	—	152
		R (N)	1–100ppb, *M, 15	13d, 56d	—	49
(10)	2,4-D	R (8–9)	2 ppb, S, 29	6% (2w)#, -	—	93
		R (ca. 8)	50 ppm, S, 25	10–>50d, 6–12d	—	94
(12)	MCPA	R, E (8.7)	10 ppb, M, 25	8–40d, —	0.02	20
(13)	dichlorprop	R, E (8.7)	10 ppb, M, 25	7–46d, —	0.02	20
(16)	propanil	R (8–9)	1 ppm, M, -	3–4d, 3–4d	—	51
		6NW (7–8)	0.1 and 1 ppm, S,17–22	ca.1d, <7d	n	97
(17)	deet	R (N)	15 ppm, 38	15d, —	—	101
(18)	fenhexamid	R (8)	20 ppm, S, 30	20% (–4m), —	—	98
(19)	flumorph	R (7–9)	1 ppm, S, 25	N (3m), —	—	99
(20)	propham	L (7.6)	0.4 ppb, S, 29	22d, —	—	87
(21)	chloropropham	11NW	0.1 and 1 ppm, S, 2–27	6 yr	n	53
(22)	carbaryl	R (8–9)	3 ppb, S, 29	13% (1w)#, —	—	93
		E, R (8–9)	0.1 ppm, S, 21	0.3–1d, —	0.6–1.0	106
		R (7–8)	1 ppm, S, 9	22–45d, —	0.6	107
(23)	carbofuran	L (6.5)	100 ppm, S, 28	2d, ca. 1d	—	109
		Pa (N)	6.5 ppm, S, 30	1–2d, —	ca. 1	108
(25)	thiobencarb	L (9)	0.2 ppm, *S, 22	3d, —	0.3	105
(26)	mexacarbate	R (7.3)	10 ppb, *S, r.t.	85% (7d), —	—	88
(28)	fenuron	R (7.3)	10 ppb, *S, r.t.	80% (14d), —	—	88
(32)	chlorotoluron	R (7.8)	5 ppm, S, 25	22d, 55d	n	119
(34)	alachlor	L (N)	1 ppb, M, 28	10% (6w), —	n	85
(37)	permethrin	L (7.8)	15 ppb, S, 21	ca. 1w (trans), —	0.1	95
(39)	fenvalerate	S (8)	0.2 ppb, *S, 20	14–17d, —	0.4	96
(41)	parathion-methyl	E (N)	0.2 ppm, M, 18	6–80d, —	<0.6	147
		E, R (6–8)	0.5 ppm, S, 28	8–13d, —	0.3	32
(43)	fenitrothion	R (8)	1 ppm, S, 25	32d, —	—	110
		L (7.5)	10 ppm, S, 23	50d, —	—	114
(44)	chlorpyrifos	E, S (8–9)	0.1 ppm, S, 21	6–15d, —	0.3–0.7	106
(45)	diazinon	E, S (8–9)	0.1 ppm, S, 21	6–41d, —	0.1–0.8	106
(46)	fenamiphos	R (7.7)	0.15 ppm, S, 22	77% (2d), —	—	115
(48)	disulfoton	R (7.5)	<0.1 ppm, M, 4–25	7–41d, —	<0.3	14
(49)	malathion	S, E (8–9)	0.1 ppm, S, 21	0.3–2d, —	0.8	106
		S (8)	0.3 ppb, *S, 20	2.5d, —	0.5–0.7	96
		14NW	10 and 100ppb, S, 2–27	1.3d, —	n	53
		R (6.9)	100 ppm, M, 37	1d, —	ca.0.5	116
(50)	acephate	P, R (7–8)	1 ppm, S, 9	57–209d, —	0.3	107
(52)	atrazine	R (N)	5 ppb, M, 20	N (16d), —	—	118
(53)	terbutylazine	R (7.8)	5 ppm, S, 25	10d, 60d	n	119
(54)	bentazone	R, E (8.7)	10 ppb, M, 25	<20% (26d), —	n	20
(56)	pyrimethanil	R (8)	10 ppm, S, 30	N (112d), —	—	117

1) Natural water. E, estuary; L, lake; M, marsh; P_o, pond; P_a, paddy field; R, river or stream; S, sea. N, no information on pH. 2) In darkness; *, under diffuse light. Concentration; S (static), M (gently mixing by swirling or shaking); temperature (°C). r.t., room temperature. f, field. 3) Degradation half-life or % of degradation (^#, % of mineralization) after the specified period in the parentheses (d, day; w, week; m, month). N, no or insignificant degradation or mineralization. L, lag period. “—” means no lag for degradation. 4) Ratio of degradation half-life, non-sterile/sterile (Eq. 5). n, no degradation in sterile water. —, no information under sterile conditions.

2.1 Organochlorines

Lindane (1) and chlordane (6) were resistant to microbial degradation in river and sea waters under microbial growth by 3 orders of magnitude, and the apparent dissipation of (6) was due to volatilization.⁹²⁾ Eichelberger and Lichtenberg⁸⁸⁾ have reported abiotic processes dominant for the dissipation of cyclodiene insecticides in river water. DDT (8) was slowly degraded in marsh water, and microbial involvement was suspected by detection of its dehydrochlorinated derivative.⁸⁹⁾ Pentachlorophenol (2) showed moderate degradation in river water but after a long lag period.⁴⁹⁾ These results suggest the low bioavailability of organochlorines to microbes in surface water.

2.2 Carboxylic acids and esters

Microbial degradation of aryloxyalkanoate herbicides has been extensively examined. 2,4-d (10) labeled with ¹⁴C at the methylene carbon was mineralized in river water with a high concentration dependence.⁹³⁾ Moderate degradation of (10) by riverine microbes after a lag time was observed without any clear relationship between the half-life and the standard plate count of colonies.⁹⁴⁾ This type of herbicide, such as (10), is generally susceptible to microbial degradation with 98% contribution in degradation, as estimated from Eq. 5.²⁰⁾ Pyrethroids with low water solubility are mostly resistant to abiotic hydrolysis at neutral pH,¹⁷⁾ but their dissolved fractions at a ppb level are subjected to microbial hydrolysis possibly due to esterases,⁴⁾ as demonstrated for permethrin (37)⁹⁵⁾ and fenvalerate (39).⁹⁶⁾ In the case of (37), the trans-isomer was much more susceptible to microbial degradation than the cis-one.

2.3 Amides, anilides, and ureas

Propanil (16) was quickly degraded to form 3,4-dichloroaniline in various surface waters after a lag period of 3–7 days,^51,97) possibly by the action of microbial arylacylamidases.⁴⁾ In contrast, either lack of microbes having relevant enzymes or steric hindrance posed by a bulky cyclohexyl moiety resulted in insignificant degradation of fenhexamid (18).⁹⁸⁾ The different amide reactivity by conjugation with the bulky alkenyl moiety might cause insignificant biodegradation of flumorph (19).⁹⁹⁾ Alkylation of the anilide nitrogen greatly reduced the bacterial availability, as reported for propachlor (33),^51,85) alachlor (34)⁸⁵⁾ and metolachlor (36).¹⁰⁰⁾ However, white rot fungus transformed (36) via demethylation, hydroxylation and hydrolytic dechlorination. In the case of deet (17), moderate biodegradation in river water was observed with formation of various products via oxidation, showing the involvement of a different mechanism from (16).¹⁰¹⁾

A wide range of biodegradation was reported for urea herbicides, depending on their chemical structure, concentration, and surface water used. Fenuron (28) and monuron (29) at 10 ppb were completely degraded in river water for 8 weeks without any detection of the corresponding anilines,⁸⁸⁾ while those at 1 ppm gave half-lives of 5 months and 2 years, respectively.⁵¹⁾ Microbial processes to form N-demethylated derivatives were observed for diuron (30), but they were of minor importance.¹⁰²⁾ Similar oxidative reactions were reported for isouron (31) through incubation in river water.¹⁰³⁾ Incidentally, Steen and Collette¹⁰⁴⁾ measured the second-order rate constants (k_b) of seven amides and anilides by amending pond water with inorganic nutrients. The authors reported a good correlation between log k_b and the wavenumber of each carbonyl group in infra-red spectroscopy, indicating that cleavage of the amide bond is a key step in their biodegradation.

2.4 Carbamates

N-Alkyl(thio)carbamates, susceptible to abiotic hydrolysis,¹⁷⁾ were quickly biodegraded in river water.^88,105) In contrast, the contribution of microbial processes was more pronounced for N-phenyl carbamates, which show greater hydrolytic stability. Moderate biodegradation of propham (20) at 0.4 ppb was observed in lake water with mineralization, but the half-life increased by a factor of 7 at 1 ppm.⁸⁷⁾ A similar concentration effect may partially account for the insignificant degradation of chlorpropham (21) in 11 surface waters.⁵³⁾ Various contribution of biodegradation (0–40%, from Eq. 5) was reported for carbaryl (22), depending either on the pesticide concentration or the type of water.^90,106,107) The type of surface water was found to be critical for the biodegradation of carbofuran (23). Insignificant biodegradation but rapid hydrolysis were observed for (23) in paddy water.¹⁰⁸⁾ The rapid biodegradation of (23) in oligotrophic lake water (pH 6.5) contrasted with its insignificant dissipation in more alkaline eutrophic one (pH 7.4),¹⁰⁹⁾ indicating the effect of the trophic level.

2.5 Organophosphorus pesticides (OPs)

Through extensive degradation studies of OPs in surface waters, the relative contribution of physical, chemical, and biological processes was found to be highly dependent either on its chemical structure or water chemistry.^{34,35,110–113)} Filtration of river water slightly enhanced their degradation in most cases, indicating less bioavailability due to adsorption to suspended particles.^34,113) LC-MS identification of products formed via oxidative desulfuration and S-oxidation showed the involvement of microbial processes.¹¹²⁾ As listed in Table 2, phosphorothioates are moderately degraded in natural waters with 20–90% contribution of biological processes. The detection of the amino derivative of fenitrothion (42) supported the idea of microbial degradation.¹¹⁴⁾ By comparing the degradation rates in four estuarine and sea waters, Bondarenko et al.¹⁰⁶⁾ showed much less biological activity in seawater due to higher salinity with low nutrients for microbial growth. Phosphorodithiolate and phosphoramidate pesticides are also biodegraded in surface waters with their extent being dependent on temperature.^14,107,115) Malathion (49) has been most extensively examined and shown to be rapidly degraded with half-lives around 1 day in various waters.⁵³⁾ The contribution of microbial processes widely ranged from a negligible level^32,90,110) to around 50%.^96,106,116)

2.6 Miscellaneous

Bentazone (54) has an amide linkage in the ring, but microbial processes were of much less importance than photolysis.²⁰⁾ The anilinopyrimidine fungicide pyrimethanil (56) is also resistant to biodegradation in river waters.¹¹⁷⁾ In the presence of 0.1% peptone, chlornitrofen (57) was rapidly biodegraded in river water via nitro reduction.⁴⁴⁾ Triazine herbicides were mostly resistant to biodegradation in river waters,^{49,90,113,118)} but some of them were degraded at moderate rates after a long lag period.¹¹⁹⁾ By using ¹⁴C-labeled atrazine (52), Rice et al.¹²⁰⁾ have reported about 10% contribution of biotic processes in its degradation via N-dealkylation and hydrolytic dechlorination.

3. Transformation by enriched community or isolated microbes

Bacterial communities in surface waters were concentrated in an appropriate medium by using an enrichment culture technique, and some species were identified by the analysis of 16S ribosomal RNA sequences. The biodegradation of pesticides by isolated bacteria is summarized in Table 3. Organochlorine pesticides underwent dechlorination¹²¹⁾ and glutathione-mediated reactions⁴²⁾ by the bacterial community. Lindane (1) was completely degraded by Streptomyces sp. pre-cultured in the presence of (1) with an increase of biomass and its uptake was an energy-dependent process.¹²²⁾ Biotic hydrolysis is a general metabolic pathway for organophosphorus pesticides^123,124) but the resultant phenols seem to be further degraded by different bacterial species.¹²³⁾ In addition to hydrolysis, fenitrothion (42) was degraded via nitro reduction by Bacillus subtilis.¹²⁵⁾ A similar reduction was reported for the degradation of dichloran (5)⁵⁹⁾ and mesotrione (58).⁸⁶⁾ Although metabolites were not identified, (42) and disulfoton (48) were biodegraded by many kinds of bacteria isolated from ponds.⁴³⁾ Amides, ureas and carbamates are either hydrolyzed^62,126,127) or oxidized^128,129) during their microbial degradation. Phytoplanktonic bacteria in lake water degraded carbendazim (24) more on average than benthic ones isolated from the bottom sediment.⁶²⁾ Correia et al.¹²⁸⁾ reported temperature as the most important factor to control both bacterial growth and the degradation of molinate (27). Hydrolytic dechlorination was reported for atrazine (52) by bacterial communities.¹³⁰⁾ Satsuma¹³¹⁾ recently isolated Nocardioides sp., Gram-positive and short-rod shape, from a river water microcosm including sediment, and identified two genes regulating dechlorination and N-dealkylation of (52).

Table 3. Aerobic degradation of pesticide by bacteria in natural water

No.	Pesticide	Bacteria1)	T, Deg/Miner, L2)	Path3)	Ref.
(1)	lindane	L: Streptomyces sp.	30, ca. 50% (4d), N	n	122
(3)	PCNB	R: bacterial community	30, 10–100% (1w), N	R, C	42
(5)	dicloran	NW: bacterial community	20, 11–21d*, N	R	59
(9)	methoxychlor	NW: 5 bacteria	25, 5–10d*, 0–3d	X	121
(16)	propanil	L: bacterial community	25, 100% (ca. 7d), 3d	H	127
(24)	carbendazim	L: bacterial community	20, 45d, N	H	62
(27)	molinate	NW: bacterial community	20, 100% (2d), N	O	128
(30)	diuron	R: Pseudomonas sp.	28, 25–60% (2d), N	n	126
		Po: bacterial community	30, —, N	O	129
(42)	fenitrothion	Pa: Flavobacterium sp.	r.t., 100% (2d), N	H	123
		L: Bacillus subtilis	r.t., 7% (10d)*, N	R	125
(48)	disulfoton	Po: Alcaligenes, Acinetobacter sp.	25, 28–49% (4d)*, N	n	43
(51)	methamidophos	R: bacterial community	35, 66–74% (1w), N	H	124
(52)	atrazine	R: bacterial community	r.t., —, N	X, O	130
		R: Nocardioides sp.	25, 1d*, N	X, O	131
(58)	mesotrione	R, L: Pantoea ananatis	30, 100% (18h), N	R, O	86

1) Origin: NW, natural waters; E, estuary; L, lake; M, marsh; P_o, pond; P_a, paddy field; R, river or stream; S, sea. 2) T, temperature of incubation (°C; r.t., room temperature); degradation half-life or % of degradation with (*) or without organic nutrients after the specified period in the parentheses (d, day; w, week; m, month); L, lag period. N, no lag for degradation. 3) Metabolic pathway. C, conjugation; H, hydrolysis; O, oxidation or hydroxylation; R, reduction; X, dehalogenation. n, no detection of any metabolite.

MacRae³⁹⁾ has pointed out some concerns regarding metabolism studies using an isolated pure microbial culture from the viewpoint of complex interactions among different microbes in a consortium. Enhanced degradation by microbial mixtures was reported for diuron (30) and deltamethrin (38) relative to that by a single strain isolated from a river,¹²⁶⁾ a surface microlayer and epidermis of common reed in a lake.¹³²⁾ A similar trend was observed for carbendazim (24) by benthic bacteria, while the biodegradation decreased by mixing several planktonic strains.⁶²⁾ Retarded degradation by mixing strains was also reported for propanil (16).¹²⁷⁾ Therefore, pesticide biodegradation in surface water by a bacterial community cannot be simply deduced from a batch culture study using isolated strains. Furthermore, the toxicity of metabolite(s) produced by one strain on another should also be considered for degradation by a community.³⁹⁾ Through the biodegradation of 2,4-d (10) and 2,4,5-T (11) mixture by Pseudomonas and Alcaligenes sp., Haugland et al.¹³³⁾ reported that chlorohydroquinone from (10) inhibited the hydrolytic dechlorination of the phenols of both herbicides and (11) inhibited the formation of the phenol from (10).

The population of fungi in surface water and their metabolic activity seem to be much less than those of bacteria, and less information on the fungal biodegradation of pesticides is available. The population of phenol-mineralizing fungi in a pond and a lake was smaller than those of bacteria by two orders of magnitude, with a ratio of mineralization rate (bacteria/fungi) of 2–6.¹³⁴⁾ Aspergillus sp. isolated from the pond metabolized malathion (49)⁵⁸⁾ and N-phenyl carbamates²¹⁾ with a much lower rate than that of bacteria by 2–3 orders of magnitude. Intra- or extra-cellular laccases²⁹⁾ most likely participate in the degradation of pesticide, as reported for pentachlorophenol (2).^135,136) Through a biodegradation study of nonylphenols with a mitosporic fungal strain and aquatic hypomycete, Junghanns et al.¹³⁷⁾ have identified 2 and 3 laccases that gave high-molecular-weight products due to oxidative coupling. Removal of fungi by filtration of an enriched microbial community retarded the degradation of diuon (30), and the addition of glucose did not improve the reduction, indicating the importance of fungal degradation.¹²⁹⁾

Factors Controlling Aerobic Transformation

1. Substrate concentration and microbial adaptation

Most of the pesticide concentration in surface water is extremely low due to various reduction processes; however, microbial degradation studies are sometimes conducted at a higher concentration due to analytical limitations. The rates of uptake and mineralization in three lake waters were found to be proportional to the initial concentration of phenol at ca. 1 ppt to the ppm level,^78,138) and the maximum specific growth rate of degrading microbes increased with the concentration.⁵⁶⁾ However, Rubin and Alexander¹³⁹⁾ found a smaller mineralization rate at <1 ppt than expected from the relationship. These authors pointed out the participation of different microbes depending on the trophic level of water, and they concluded that the slower energy gain at a lower concentration may not meet the demand by the small population of degrading microbes. A similar concentration effect on mineralization was reported for 2,4-d (10) and carbaryl (22) in stream water,⁹³⁾ while (10) and propham (20) were more efficiently mineralized in lake water at lower concentrations.^87,140) In the case of isouron (31), no clear concentration effect was observed on the degradation and mineralization at 10 ppb–1 ppm.¹⁰³⁾

Incidentally, there sometimes appears to be a lag period due to microbial adaptation (acclimation) between the application of a substrate and the initiation of its degradation (or mineralization); moreover, the period decreases with its repeated application. Two triazine herbicides in river water started to degrade 40–60 days after the first application, and the repeated application greatly decreased this lag time by a factor of 2–3 but with less effect on their half-lives.¹¹⁹⁾ Using Bacillus cereus pre-exposed to anilines, El-Dib and Aly¹⁴¹⁾ clearly demonstrated the shorter lag periods and half-lives of four phenylamide herbicides by their repeated application. Microbial adaptation has been extensively examined for 2,4-d (10) by using lacustrine microbes.¹⁴²⁾ The lag period before degradation was affected by neither protozoa nor fungi and greatly shortened by pre-exposure to (10). The population of (10)-degrading microbes estimated by the MPN technique increased long before the initiation of degradation. Therefore, the lag phase represents the time for a small population of a degrader to reach a sufficient size to form a product to be detected. A series of similar studies on p-nitrophenol supported this mechanism.¹⁴³⁾ The induction of catabolic gene expression was another mechanism of acclimation, as demonstrated by the differences in dynamics of tfdA transcript abundance and tfdA loci in two microbial communities in ponds when exposed to (10).¹⁴⁴⁾

2. Spatial and seasonal variations

Spatial variation, generally by less than 1–2 orders of magnitude, has been reported for the first-order microbial degradation rate of pesticide in surface waters.^{42,45,49,145)} When the second-order rate constant k_b (Eq. 1) was used for comparison, the spatial variance was less pronounced⁹⁷⁾ or statistically disappeared.⁵³⁾ The different population of pesticide degraders in water partially accounts for the variation,^49,146,147) supported by either no correlation between the degradation rate and standard plate counts¹⁴⁵⁾ or insignificant differences in the number of total aerobic heterotrophs.⁶⁶⁾ Microbial adaptation by pre-exposure to pesticides is one possible mechanism for the spatial variation of microbial degradation in urban and agricultural areas.^42,45,148) In addition to the geographical features controlling the hydraulic retention time,¹⁴⁸⁾ water chemistry also plays a role in the variation. The different pH value and temperature affect the hydrolysis of pesticides, and a higher salinity reduces microbial activities.¹⁰⁶⁾ A multivariate analysis on the biotransformation of 4 pesticides in 11 types of water showed that three components can explain 84% of the total variance in their degradation rates; microbial activity, macro-/micro-nutrients and phosphorus content in water.¹⁴⁹⁾

Seasonal changes in water discharge, contents of nutrients and temperature strongly affect both the population and distribution of microbes in natural waters.^27,61,67) As compared with the spatial variation, less seasonal variation by less than an order of magnitude has frequently been reported for pesticide biodegradation.^{42,45,104,145)} It is noteworthy that the activation energy in microbial degradation depends on the season and site of water sampling.^90,113) Microbial degradation of 2,4-d (10) in river water showed the peaks in May and August and correlated with the amounts of DOM or suspended matters that are closely related to a total phosphorus content in water, indicating the importance of a nutrient level.^73,94) The maximum degradation of (10) was observed in June to July when the load of suspended sediment in the rivers increased by flooding.¹⁵⁰⁾ The number of aerobic heterotrophs dominantly consisting of fluorescent Pseudomonads in U.S. lakes varied around an order of magnitude and peaked in summer.⁶⁶⁾ Similar but larger variation was observed for the microbial activity in hydrolyzing fluorecein diacetate, and Pseudomonads degraded several pesticides. Seasonal variation in the biodegradability of several pesticides was reported for ponds, together with a marked change in the dominant population of microbes.⁴³⁾

3. Sediment

The effect of sediment on the microbial degradation of pesticides has been examined by typically adding 0.5–1 g L⁻¹ of sediment, considering the occurrence of suspended matters in surface waters.¹⁵¹⁾ The presence of sediment, suspended or precipitated, affects the dissipation profiles of pesticides mainly by two mechanisms. Adsorption to sediment results in an apparent reduction of pesticide in water, and its tight binding leads to less bioavailability with longer persistency, a mechanism that is more observable in hydrophobic pesticides.^{46,95,96,108,152,153)} Reduced degradation by adsorption to sediments or colloids has been shown by kinetic analysis on the biodegradation of endosulfan (7) in a river water-sediment system.¹⁵⁴⁾ Alternatively, benthic microbes frequently enhance the biodegradation of pesticides by an order of magnitude.¹⁵⁵⁾ The contribution of two mechanisms can be conveniently examined by sterilizing sediment.^{46,107,120,145,156)} The catalytic hydrolysis by clay minerals and metal cations from sediments is assumed, but it is considered limited to some types of pesticides.¹⁷⁾ An insignificant sediment effect for an ionic pesticide that is slowly biodegraded was also observed.¹⁵⁷⁾

The participation of benthic microbes was supported by approximately 5-fold acceleration in the biodegradation of fenthion (43) and p-chlorophenol with a 10-fold increase of a sediment content in estuarine waters.^47,158) In the sediment-enhanced biodegradation of parathion-methyl (41), there was no correlation between the degradation rate and c.f.u. in water.¹⁴⁵⁾ In the biodegradation study of phenols, the number of degrading microbes markedly increased in the presence of sediment by greater than two orders of magnitude.¹⁴⁷⁾ Furthermore, not only sterile sediment but also glass beads and sand enhanced the mineralization of p-chlorophenol with a marked increase of the degraders.¹⁵⁸⁾ These results indicate the importance of a substratum for microbial growth. Recently, a sediment-water interface has been shown as a biodegradation site for polycyclic aromatic hydrocarbons (PAH) by various combination of PAH, isolated PAH-degraders, and river sediment in a reaction vessel separated by a 0.22-µm pore membrane.¹⁵⁹⁾ The co-existence of sediment and PAH degraders that were not separated by membrane increased the zero-order degradation rates of PAHs by a factor of 3 with a 3–4-fold larger maximum specific growth rate. Furthermore, the faster degradation in the presence of fine silt, adsorbing less PAHs but showing an almost equal bacterial density to that in clay, strongly suggested that the biodegradation mainly proceeds at the sediment-water interface.

Some pesticide-degrading microbes have been isolated and identified from sediments. Although detailed species identification was not conducted, several microbes degrading parathion (40) or malathion (49) were isolated from the estuarine sediment and each of them could metabolize both pesticides but to different extents.¹⁶⁰⁾ The Gram-negative bacterial strain RA8 collected from the river sediment was identified as Variovorax sp., which specifically metabolized ureas having the N-methoxy-N-methyl moiety.¹⁶¹⁾ From the channel sediment treated with pyrethroids, several genera of culturable bacteria in the Proteobacteria division specific to each pyrethroid were isolated, but the strong sorption of pyrethroids to sediment reduced the bioavailability.¹⁶²⁾

4. Macrophytes and biofilm

In artificial channels where gravels and macrophyte-planted sediments were alternatively placed in the bottom, 26–46% of pentachlorophenol (2) was degraded by microbial processes.⁵³⁾ Greater bacterial numbers on the macrophyte (epiphytic, ca. 10⁹ g⁻¹ dry weight) and gravel (ca. 10⁸ cm⁻²) surfaces in comparison with those in water (ca. 10⁷ mL⁻¹) showed the importance of these bacteria in the degradation of (2), also supported by the higher metabolic activities of epiphytic and benthic microbes toward sugars and carboxylic acids.²⁵⁾ The presence of the whole plant or roots of Spartina alterniflora resulted in the 100-fold accelerated degradation of fenthion (43), with the enhancement stepwise decreasing by one order of magnitude when the outside and inside portions of its leaves were used, indicating the high activity of epiphytic bacteria.⁴⁷⁾ Many epiphytic bacteria isolated from the surface of common reeds were found to efficiently degrade deltamethrin (38).¹³²⁾ Furthermore, pesticide adsorption and uptake by phototrophic macrophytes and algae are known to be important for pesticide dissipation.²⁴⁾ Recently, 2–20-fold accelerated dissipation was reported for six pesticides applied to a water-sediment system (3 : 1, v/w) when Elodea canadensis or an algal community was added under fluorescent light at >400 nm.¹⁶³⁾

The adsorption kinetics of eight pesticides having a wide range of log K_ow (−1–7) to river biofilms were investigated by using a flow-through reactor.⁶⁰⁾ Linearity between log K_d (linear adsorption coefficient) and log K_ow was observed, and a much narrower range of K_d by a factor of 50 was explained by the hydrophilic nature of EPS in the biofilms. To determine the adsorption and degradation profiles of pesticides in biofilm, the distribution of diclofop-methyl (15) and EPS glycol-conjugates was extensively assessed by using a lectin binding assay, confocal laser scanning microscopy, and LC-MS/MS analysis in combination. The distribution of fucose-binding lectin and (15) overlapped in the EPS present at bacterial surfaces in the biofilms.^164,165) Furthermore, either the partial recovery of fluorescence in the presence of KCN from the photo-bleached biofilm or the detection of the acid derivative of (15) indicated that bacteria were actively involved in the uptake and degradation of (15).^166,167) The bacterial processes in the degradation of parathion-methyl (41) were confirmed by the addition of candicidin as an inhibitor to the river biofilm.⁵⁵⁾ The biodegradation of acetochlor (35) in the lake biofilm to the ethansulfonic acid derivative resulted in less accumulation of (35) with a factor of 28 than expected from its hydrophobicity.¹⁶⁸⁾ A large variation in a degradation rate depending on the collection site of biofilms was observed for 2,4-d (10) esters and (41), indicating the contribution from a wide variety of microbes.¹⁶⁹⁾ Biodegradation of diazinon (45) by using microbes collected from the surface of submerged ceramic disks in the river varied with the seasonal change of the bacterial mass.¹⁷⁰⁾ Diuron (30) was oxidatively demethylated by microbes collected from the surface of river cobbles, and multivariate analysis showed the modified structure of a bacterial community by exposure to (30).¹⁷¹⁾ By conducting the biodegradation of 2,4-d (10) butoxyethyl ester in a reactor containing periphytons on Teflon strips, the first-order rate constant was found to be proportional to a mixing speed, indicating the mass transport as a rate-limiting step.¹⁷²⁾ Incidentally, Lewis and Gattie¹⁷³⁾ attempted to formulate the contribution of microbes in biofilms by using the biodegradation rates of 2,4-d (10) esters in the presence and absence of biofilms. The authors found the linearity in the plot of the first-order degradation rate normalized by “(surface area of biofilms)/(water volume)” of natural water vs. water flow velocity and proposed the following empirical relationship for the contributing fraction of a microbial community in the biofilm (f_B); f_B=U/(U+2D), where U and D are velocity (m h⁻¹) and depth (m) of water, respectively. This equation clearly shows the importance of biofilms for pesticide degradation in streams and shallow rivers.

5. Illumination

Most microbial degradation studies of pesticides have been conducted in darkness to avoid not only the unfavorable effect of U.V. light on a microbial growth but also the contribution of photolysis. There are various kinds of phototrophs in surface waters represented by algae and some bacteria, and their participation in the biodegradation of pesticides cannot be assessed in darkness. Since most pesticides do not have any U.V.-vis. absorption at >400 nm, illumination by a fluorescent light for cultivation may be suitable, as prescribed in Agricultural Canada T-1–255.⁶⁾ However, photosynthetic action usually causes a large pH increase by 2–3 units¹⁶³⁾ which affects the degradation of hydrolytically unstable or dissociable pesticides. Furthermore, illumination even at >400 nm may induce indirect photolysis via reactions with excited humic substances¹¹³⁾ or photo-produced ¹O₂ and ·OH.^14,20) Therefore, sterile control under illumination is necessary to confirm the contribution of photochemical processes.

Conclusion

Many factors controlling microbial processes can be conveniently placed under the categories of pesticide properties, experimental design, or origin of surface water. Among the various processes, microbial degradation becomes more important for pesticides with a low volatility from water, no tight adsorption to suspended and bottom sediments, and hydrolytic/photochemical stability. Pesticides freely dissolved in water are bioavailable to microbes and subjected to metabolism or co-metabolism via enzymatic hydrolysis, oxidation and reduction.^4,24) The use of radio-labeled pesticides has been limited to quantify its mineralization; thus, further study of biodegradation using ¹⁴C-pesticide with the identification of metabolites is highly recommended to understand the microbial degradation processes in surface water.

The available test guidelines allow for a solid design of an experimental system based on the accumulated evidence. The concentration of pesticide is one of the most critical factors to determine its biodegradability; therefore, the most probable environmental concentration of pesticide should be selected based on field monitoring or simulation data. Temperature is another important factor and the activation energy of biodegradation should be standardized similarly to pesticide degradation in soil¹⁷⁴⁾ to describe the temperature effect. For the present, temperature close to that when surface water is sampled or more than one temperature may be favorable in a pesticide biodegradability study, as prescribed by the Canadian guidelines.⁶⁾ Concerning surface water and sediment, their history of exposure to pesticides or other anthropogenic pollutants is useful to examine if the obtained degradation profile originates from microbial adaptation. The degradation rate may be normalized by using the biomass of degraders, but either characterization of relevant microbial community or isolation of specific microbes is cumbersome and sometimes difficult. The biodegradation should be carefully examined in relation with not only the season- and site-dependent population of microbes but also the water characteristics, such as the temperature, pH, nutrient level, and contents of DOM and suspended sediments. Therefore, it is prudent to use the biodegradation in surface water as higher-tier site-/season-specific data, since its variability is considered much higher than that in aerobic soil.¹⁷⁵⁾

Microbes freely floating in water represent a part of all microbes, and those associated with suspended sediments also play a role in biodegradation of pesticides. Therefore, the effect of sediment should be examined together with its possible sink for pesticides due to adsorption, as prescribed in the guidelines.^5–7) Furthermore, there are various microbes living in biofilms on the surfaces of macrophytes and substratum which are possibly involved in the biodegradation of pesticides, and their contribution becomes more important in the shallow water body with a low flow rate. Although algal biodegradation of some pesticides has been examined, information on the contribution of phototrophs grown under illumination is still limited.

The results obtained according to the present guidelines are very useful to grasp the potential biodegradability of pesticides, but it should be kept in mind that they only represent a part of microbial processes in a surface water body. To utilize microbial aerobic transformation as one of the typical processes in the environmental assessment of pesticides, more studies should be conducted to clarify the relationship between pesticide degradation and microbes in surface water, the effects of substratum and the method for normalizing site- and season-specific data, referring to the microbial metabolism of pesticides in soil.

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