bio degradation and persistence

Aerobic and Anaerobic Biodegradation Potentials of Microorganisms

Walter Reineke

Bergische Universität – Gesamthochschule Wuppertal, Chemische Mikrobiologie,Fachbereich 9, Gaußstraße 20, D-42097 Wuppertal, GermanyE-mail: [email protected]

Microorganisms represent essential components of the global carbon cycle. In addition, it ap-pears that most xenobiotic industrial chemicals can be degraded by microorganisms, either bya combination of cometabolic steps, often yielding partial degradation, or by serving asgrowth substrate which is accompanied by mineralization of at least part of the molecule.Using a number of examples, including aromatic, chloroaromatic, aliphatic, and chloroali-phatic compounds, I have presented some principles on the degradation. The great influenceof some environmental conditions on the degradation, such as the presence or absence ofoxygen, the availability of other electron acceptors such as nitrate or sulfate, has been discussedwith special emphasis on the type of reactions and the rates of degradation that occur.

While aerobic microorganisms use oxidative reactions, the degradation by anaerobic bac-teria takes place by reductive types of reactions. The oxidative sequences of aromatic and chlo-roaromatic compounds in aerobic bacteria yield central intermediates with a diphenolic struc-ture. These compounds are then cleaved by enzymes that use molecular oxygen. In contrast,the anaerobes degrade aromatic compounds by reductive conversions and the central inter-mediates ready for hydrolytic ring cleavage bear a 1,3-dioxo structure.

Aerobic bacteria and fungi, especially ligninolytic ones, were shown to use mechanisticallydifferent catabolic pathways and enzymes. The ligninolytic fungi convert oxygen to hydrogenperoxide which is then used for the formation of an aryl cation radical undergoing spon-taneous rearrangements and degradation.

The broad variety of mechanisms which brings about dechlorination is another importantpart of this work.Although the diversity of the compounds discussed is very large, the strategyof the organisms used in the degradation includes various analogous reactions.

Another important aspect of discussion is the different degree of degradation. While mostresearch is done on organisms that are able to use the respective compound as the growth sub-strate, i.e., carbon dioxide and biomass result, the cometabolic potential of microorganismsshould not be neglected. Cometabolism takes place very much in nature and brings aboutsome modification of a target compound.

In general, anoxic microbial degradation seems to be of greater relevance in nature than ear-lier expected. It is remarkable that some chlorinated compounds such as chlorobenzoates, chlo-rophenols, or tetrachloroethene may function as a physiologically functional electron acceptorin a type of anaerobic respiration, which leads to non-chlorinated or lower chlorinated products.

Since various compounds will not be degraded totally by one type of organism the com-plementary potential of anaerobic and aerobic populations in combination is thought to be amethod to bring about complete mineralization.

Finally, the possibility of enhancing the degradative potential of aerobic organisms in thelaboratory, i.e., artificial evolution of enzymes and pathways, by different genetic approaches,is discussed.

Keywords. Aromatic, chloroaromatic, aliphatic, and chloroaliphatic compounds, Aerobic andanaerobic bacteria, Ligninolytic fungi, Cometabolism vs productive mineralization, Com-pounds as carbon and energy sources, Degradative pathways with oxidative sequences, De-gradative pathways with reductive sequences, Dechlorination mechanisms, Fermentations,

CHAPTER 1

The Handbook of Environmental Chemistry Vol. 2 Part KBiodegradation and Persistence(ed. by B. Beek)© Springer-Verlag Berlin Heidelberg 2001

Haloaliphatic and haloaromatic compounds as electron acceptors with dechlorination: deha-lorespiration, Sequential anaerobic-aerobic processes, Enhancing the degradative potentialby in vivo and in vitro techniques

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

1.1 Redox Processes and Mineralization of Organic Compounds . . . 51.2 Energy Yields . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61.3 Distribution of Electron Acceptors in the Environment

and Sequential Redox Conditions . . . . . . . . . . . . . . . . . . . 81.4 Organic Compounds as Electron Acceptors . . . . . . . . . . . . . 91.5 Limitations to Fermentation . . . . . . . . . . . . . . . . . . . . . . 121.6 Various Degrees of Degradation . . . . . . . . . . . . . . . . . . . 131.7 Résumé of Introduction . . . . . . . . . . . . . . . . . . . . . . . . 15

2 Degradation of Aromatic Compounds . . . . . . . . . . . . . . . . 15

2.1 Aerobic vs Anaerobic Degradation: Introduction . . . . . . . . . . 152.2 Aerobic Degradation of Aromatics: General Differences Between

Prokaryotic and Eukaryotic Organisms in the Initial Reactions . . 172.3 Degradation of Aromatic Compounds by Aerobic Bacteria . . . . 182.3.1 Reactions Converting Aromatic Compounds into

Ring Cleavage Substrates . . . . . . . . . . . . . . . . . . . . . . . 182.3.2 Ring Fission and Carbon Chain Fission . . . . . . . . . . . . . . . 262.3.3 Pathways as a Whole for Catechol, Protocatechuate, and Gentisate 292.4 Degradation of Aromatics by Fungi . . . . . . . . . . . . . . . . . 322.4.1 Degradation by Non-Ligninolytic Fungi . . . . . . . . . . . . . . . 322.4.2 Degradation of PAHs by Ligninolytic Fungi . . . . . . . . . . . . . 342.5 Résumé: Aerobic Degradation of Aromatic Compounds . . . . . . 392.6 Anaerobic Degradation of Aromatic Compounds . . . . . . . . . . 402.6.1 Channeling Reactions . . . . . . . . . . . . . . . . . . . . . . . . . 402.6.2 Activating Reductive Sequences and Ring Cleavage . . . . . . . . . 482.6.3 Anaerobic Degradation of Environmentally Important Aromatics

where Pathway Information is Missing or Minor . . . . . . . . . . 512.6.4 Résumé: Anaerobic Degradation of Aromatic Compounds . . . . . 522.7 Résumé: Aromatic Compounds . . . . . . . . . . . . . . . . . . . . 532.7.1 Degradation in the Presence of Oxygen . . . . . . . . . . . . . . . 532.7.2 Degradation in the Absence of Oxygen . . . . . . . . . . . . . . . . 54

3 Degradation of Chloroaromatic Compounds . . . . . . . . . . . . 55

3.1 Chloroaromatic Compounds as Growth Substrate for Aerobic Bacteria and the Dechlorination Mechanisms . . . . . 55

3.1.1 Elimination of Chlorine Substituents Prior to Ring Cleavage . . . 553.1.2 Late Eliminations of Chlorine After or Linked with Ring Cleavage 623.1.3 Degradation of Higher Chlorinated Aromatic Compounds

Needs Different Dechlorination Mechanisms . . . . . . . . . . . . 683.2 Degradation of Chloroaromatic Compounds

by Ligninolytic Fungi . . . . . . . . . . . . . . . . . . . . . . . . . 71

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3.3 Anaerobic Microbial Populations with the Potential to Dechlorinate Chloroaromatic Compounds . . . . . . . . . . . . 75

3.3.1 Potential of Environmental Materials and Undefined Enrichments 753.3.2 Pure Cultures: Chloroaromatic Compounds

as Electron Acceptors . . . . . . . . . . . . . . . . . . . . . . . . . 773.3.3 Pure Cultures: Chloroaromatic Compounds as Growth Substrate . 823.3.4 Dechlorinating Organisms, Part of a Food Web . . . . . . . . . . . 823.3.5 Phototrophic Bacteria and Chloroaromatic Compounds . . . . . . 833.4 Résumé: Chloroaromatic Compounds . . . . . . . . . . . . . . . . 84

4 Degradation of Aliphatic Hydrocarbons . . . . . . . . . . . . . . . 84

4.1 Aerobic Degradation of Aliphatic Hydrocarbons . . . . . . . . . . 844.1.1 Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 844.1.2 Branched Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . 894.1.3 Alkenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 924.2 Anaerobic Degradation of Aliphatic Hydrocarbons . . . . . . . . . 954.3 Résumé: Aliphatic Hydrocarbons . . . . . . . . . . . . . . . . . . . 96

5 Degradation of Chloroaliphatic Compounds . . . . . . . . . . . . 96

5.1 Chloroaliphatic Compounds as Growth Substrate for Aerobic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . 96

5.1.1 Hydrolytic Dehalogenation . . . . . . . . . . . . . . . . . . . . . . 995.1.2 Glutathione S-Transferase-Dependent Dehalogenation . . . . . . . 1035.1.3 Lyase-Catalyzed Dehalogenation . . . . . . . . . . . . . . . . . . . 1045.1.4 Hydratase-Catalyzed Dehalogenation . . . . . . . . . . . . . . . . 1045.1.5 Dehalogenation by Oxygenases . . . . . . . . . . . . . . . . . . . . 1045.1.6 Dehalogenation During b-Oxidation . . . . . . . . . . . . . . . . . 1055.1.7 Dehydrohalogenation . . . . . . . . . . . . . . . . . . . . . . . . . 1065.1.8 Dehalogenation by Methyltransferase/Dehydrogenase . . . . . . . 1065.2 Chloroaliphatic Compounds as Growth Substrates

for Anaerobic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . 1075.2.1 Fermentative Degradation . . . . . . . . . . . . . . . . . . . . . . . 1075.2.2 Degradation Under Denitrifying Conditions . . . . . . . . . . . . 1095.2.3 Degradation Under Methanogenic Conditions . . . . . . . . . . . 1095.3 Cometabolic Transformations . . . . . . . . . . . . . . . . . . . . . 1105.3.1 Aerobic Bacteria: Oxidative . . . . . . . . . . . . . . . . . . . . . . 1105.3.2 Ligninolytic Fungi: Reductive . . . . . . . . . . . . . . . . . . . . . 1125.3.3 Anaerobic Bacteria: Reductive . . . . . . . . . . . . . . . . . . . . . 1125.4 Chloroaliphatic Compounds as Electron Acceptors . . . . . . . . . 1135.5 Résumé: Chloroaliphatic Compounds . . . . . . . . . . . . . . . . 115

6 Sequential Anaerobic-Aerobic Processes for the Degradation of Problematic Compounds . . . . . . . . . . . . . . . . . . . . . . 116

6.1 Studies with Environmental Materials . . . . . . . . . . . . . . . . 1176.2 Studies with Undefined Enrichment Cultures . . . . . . . . . . . . 118

Aerobic and Anaerobic Biodegradation Potentials of Microorganisms 3

6.3 Studies with Undefined Enrichment Cultures Supplemented with Pure Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . 118

6.4 Studies with Pure Cultures . . . . . . . . . . . . . . . . . . . . . . . 1206.5 Résumé: Sequential Anaerobic-Aerobic Processes . . . . . . . . . . 121

7 Enhancement of the Catabolic Potential of Microbial Strains in the Laboratory . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121

7.1 Uptake of a Target Compound . . . . . . . . . . . . . . . . . . . . 1227.2 Expansion of the Effector Specificity of Transcriptional Regulators 1237.3 Alterations in Structural Genes . . . . . . . . . . . . . . . . . . . . 1247.3.1 Widening of the Substrate Range . . . . . . . . . . . . . . . . . . . 1247.3.2 Mutations in Structural Genes to Avoid the Formation

of a Toxic Metabolite . . . . . . . . . . . . . . . . . . . . . . . . . . 1267.4 Use of External Genetic Information to Expand

the Substrate Range . . . . . . . . . . . . . . . . . . . . . . . . . . 1277.4.1 Chlorobenzoate-Degraders by Conjugal Transfer . . . . . . . . . . 1277.4.2 Chloronitrophenol-Degraders by Conjugal Transfer . . . . . . . . 1287.4.3 Chlorobiphenyl-Degraders by Mating Three Strains . . . . . . . . 1297.4.4 Other Chloroaromatic-Degraders by Conjugal Transfer . . . . . . 1297.4.5 Chlorobenzoate- and Chlorosalicylate-Degraders

by Genetic Engineering Techniques . . . . . . . . . . . . . . . . . 1307.4.6 Chlorobiphenyl-Degraders by Genetic Engineering Techniques . . 1327.4.7 Trihalopropane-Degraders by Genetic Engineering Techniques . . 1327.5 Construct to Degrade TCE Without Apparent Toxic Effect . . . . . 1327.6 Creation of a Pathway for the Degradation

of Halogenated Alkanes and Alkenes . . . . . . . . . . . . . . . . . 1347.7 Creation of a Pathway for the Degradation of Mixtures of Methyl-

and Chloroaromatics by Combination of Pathway Modules . . . . 1347.8 Résumé: Enhancement of Catabolic Potential . . . . . . . . . . . . 136

8 Concluding Remarks and Outlook . . . . . . . . . . . . . . . . . . 136

9 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140

List of Abbreviations

BESA bromoethane sulfonic acidBTEX benzene, toluene, ethylbenzene, xylenes mixturecDCE or cis-1,2-DCE cis-dichloroetheneDDT 1,1,1-trichloro-bis(p-chlorophenyl)-ethaneDHB dihydrodihydroxybenzoateEDTA ethylenediaminetetraacetic acidGSH glutathione (reduced)IP ionization potentialLiP lignin peroxidaseMnP manganese-dependent peroxidase

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NTA nitrilotriacetic acidPAHs polycyclic aromatic hydrocarbonsPCBs polychlorinated biphenylsPCE tetrachloroethenePCP pentachlorophenolPQQ methoxatin (2,7,9-tricarboxy-1H-pyrrolo(2,3-f)quino-

line-4,5-dione)TCA cycle tricarboxylic acid cycle = Krebs cycleTCE trichloroetheneVC vinyl chloride2,4-D 2,4-dichlorophenoxyacetic acid

1Introduction

1.1Redox Processes and Mineralization of Organic Compounds

The biosphere presents a large diversity of different habitats within whichmicroorganisms can operate provided that an energy source and nutrients areavailable and that physical conditions are appropriate. Organic compounds innature are distributed throughout aerobic and anaerobic environments. Anoxicecosystems are created when oxygen consumption by microorganisms exceedsits supply, e.g., in soils with impeded drainage, stagnant water, municipal land-fills, sewage treatment digesters, and sediments of the oceans and other naturalbodies of water. Different groups of microorganisms are found in oxic and an-oxic situations. Although the biochemical pathways used by microorganisms todegrade organic compounds are extremely diverse, they are all directed towardsthe production of energy and carbon for growth.

Concerning energy-yielding processes, four types of microbial metabolismare recognized and they are described by the terms photometabolism, fermen-tation, aerobic respiration, and anaerobic respiration.

Fermentation is a process that does not require oxygen or the presence ofother electron acceptors such as NO3

–, Mn4+, Fe3+, SO42–, or CO2 and depends on

the ability of the microorganisms to use part of the organic molecule (often ametabolite) as an electron acceptor. During fermentation of an organic com-pound, reduced pyridine nucleotides (NADH) and adenosine triphosphate(ATP) are produced by the degradative pathway. Such an energy-yieldingsequence of reactions, which is accompanied by “substrate level phosphory-lation,” can only continue if the organism has some mechanism to regenerateoxidized pyridine nucleotides as an acceptor for the further oxidation of theorganic substrate. Oxidized pyridine nucleotides are produced by the transferof electrons to intermediates which are formed during metabolism of thegrowth substrates. The result of a typical fermentation is a mixture of productswhich are more oxidized and others which are less oxidized than the originalsubstrate (Fig. 1).


1.2Energy Yields

Metabolism of organic compounds by respiration leads to a much more effi-cient use of potential chemical energy than fermentative conversion. During re-spiration electrons in reduced pyridine nucleotides can be transferred to oxy-gen in the case of aerobic respiration or to various other acceptors such as NO3

–,Mn4+, Fe3+, SO4

2–, or CO2 in the case of anoxic respiration. The chemical energyof the redox system is used for the production of a proton gradient when trans-porting the electrons. The ATP synthetase then converts the energy of the pro-ton gradient into chemical energy (ATP) (Fig. 2). Together, this constitutes two

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Fig. 1. Schematic illustration of the general scheme of fermentation with energy generationby substrate level phosphorylation and NAD/NADH cycling

Fig. 2. Schematic illustration of the respiratory chain of energy conservation including theproton translocation steps that establish a proton motive force which is used by the ATP syn-thetase for ATP formation

to three molecules of ATP generated per electron pair that is channeled throughthe electron transport chain.

During oxidative metabolism, the organic substrate is converted to carbon di-oxide and water and part of it is assimilated into cell material. Molecular oxy-gen is usually the preferred electron acceptor; it is reduced to water. The oxida-tion of an organic substrate with oxygen or nitrate as the electron acceptorsleads to a similar high yield of ATP. The reduction of CO2 to methane and sul-fate to sulfide is carried out predominantly by strict anaerobes, whereas nitratereduction is carried out predominantly by facultative anaerobes if oxygen is notavailable. The energetics of these processes are very different. The free energychange of O2 and nitrate reduction per two electrons is about the same while thevalues are much lower for sulfate and CO2 reduction (Table 1). This explains thelower growth yields and rates on an organic substrate with sulfate-reducingbacteria and methanogens as compared to aerobic bacteria and nitrate-reduc-ing organisms.

Many complex organic compounds can be oxidized to CO2 by pure culturesof bacteria with NO3

–, Mn4+, Fe3+, or SO42– as electron acceptor, whereas metha-

nogenic metabolism usually requires mixed cultures, since most methanogenscan only ferment simple low molecular weight organic molecules such as ace-tate or methylamine. At least three physiological types of bacteria operate inmethanogenic systems: fermenters, which convert the initial substrate into or-ganic acids, such as propionate, butyrate, acetate, formate, succinate, and lactate,as well as alcohols; acetogenic proton-reducing bacteria, producing acetate; andacetate and CO2 plus hydrogen-consuming methanogens.

If light is present in anaerobic environments, photometabolism is also possi-ble. The phototrophic bacteria use light as the energy source while substratessuch as organic acids (acetate, propionate, butyrate, succinate, glutarate, benzo-ate) are usually extensively assimilated into cell material. Therefore, the oxida-tive or fermentative metabolism of a portion of the carbon source is unneces-sary.

Just recently, bacteria have been isolated and characterized that use either theoxyanions of arsenate or selenate or both as terminal electron acceptors [1].


Table 1. Free energy changes in aerobic and anaerobic respiration using hydrogen as electrondonor

DGo¢

Redox reaction (kJ/mol acceptor) (kJ/mol 2e–)

2 H2 + O2 Æ 2 H2O –474 –2372.5 H2 + NO3

– + H+ Æ 0.5 N2 + 3 H2O –560 –224H2 + Mn4+ Æ Mn2+ + 2 H+ –187 –1870.5 H2 + Fe3+ Æ Fe2+ + H+ – 30.9 – 61.84 H2 + SO4

2– + 1.5 H+ Æ 0.5 H2S + 0.5 HS– + 4 H2O –153 – 38.34 H2 + CO2 Æ CH4 + 2 H2O –135 – 33.8

1.3Distribution of Electron Acceptors in the Environment and Sequential Redox Conditions

The natural abundance of major environmental electron acceptors is summariz-ed in Table 2, including the concentrations commonly encountered in terrestrialand aquatic environments. With the exception of sulfate, which is very abundantin seawater and low but variable in freshwater, the concentrations of the other el-ectron acceptors are similar in both environments. Both in soil and freshwater, el-ectron acceptors such as oxidized nitrogen or sulfur compounds or oxidized me-tal ions such as Fe3+ or Mn4+ can serve in anaerobic respiration. In waters of neu-tral pH, solid forms of Mn and Fe will dominate. The concentration of CO2 variesdepending on the alkalinity and the organic carbon input into the ecosystem.

Oxygen is the preferred electron acceptor, in agreement with what would beexpected in view of thermodynamics. If the organic carbon influx into an envi-ronmental compartment such as groundwater is higher than that of oxygen,anaerobic conditions will occur. Then there is sequential utilization of the al-ternative electron acceptors NO3

–, MnO2(s), Fe(OH)3(s), SO42–, and CO2 in that

order, creating different redox zones (Fig. 3).The sequential use of electron acceptors is important for biodegradation,

since some compounds can only be converted by certain microorganisms, i.e.,under certain redox conditions. Consequently, a chemical present as a ground-water contaminant is likely to be degraded only in that part of the groundwaterplume where favorable redox conditions prevail. Some compounds are degrad-able under any specific redox conditions, but the degradation rate of such com-pounds may vary under different redox conditions. Table 3 summarizes datawhich roughly show the degradation kinetics occurring at various redox condi-tions for some compounds. Comparing chlorinated aromatic and chlorinatedaliphatic compounds with a low and high degree of substitution, it appears thatlower chlorinated compounds will generally be degraded rapidly under aerobic

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Table 2. Natural abundance and free energy yield of commonly used electron acceptors

Electron acceptor Natural abundance Free energy (kJ/mol glucose)

Oxygen 300 mmol/l –3190Nitrate Few mmol/l –3030Manganese, MnO2 (birnessite) <mmol/l to >mmol/l –3090Manganese, MnO2 (nsutite) <mmol/l to >mmol/l –3050Manganese, MnO2 (pyrolusite) <mmol/l to >mmol/l –2920Nitrate Few mmol/l –2750Iron, Fe2O3 (hematite) <mmol/l to >mmol/l –1410Iron, FeOOH (geothite) <mmol/l to >mmol/l –1330Sulfate <100 mmol/l (freshwater), – 380

~28 mmol/l (seawater)Carbon dioxide Variable – 350

Free energies are taken from Froelich et al. [2]. The mineralogical form of the Mn4+ and Fe3+

oxides can alter the redox potential significantly.


Fig. 3. Transport of chemicals and microbial reactions in the groundwater with respect to thepresence of electron acceptors and prevailing redox conditions (according to Bouwer andZehnder [3] and Schachtschabel et al. [4])

conditions, while the higher chlorinated analogs are preferentially dechlorina-ted under anaerobic conditions. Table 3 illustrates that a broad variety of com-pounds are subject to aerobic degradation, while compounds such as hexa-chlorobenzene are degraded in groundwater only at the most reduced part ofthe redox regime. Overall the degradative potential of anaerobic microbial com-munities is much greater than appreciated until only recently.

The existence of electron acceptor profiles in the above-mentioned order andtheir associated microbial processes in the different redox zones of the subsurfacecan also be observed in lakes and the marine environment as a function of water depth, as in the Black Sea, where oxygen reduction only occurs in the toplayer of about 50 m and alternative electron acceptors are used at greater depths.

1.4Organic Compounds as Electron Acceptors

Besides the inorganic compounds mentioned above, several organic com-pounds can also serve as electron acceptors in anaerobic respiration [5].Fumarate respiration is the most widespread type of anaerobic respiration.Fumarate is formed during the biodegradation of carbohydrates and proteinsand its reduction to succinate is linked to a respiratory chain. Dimethyl-sulphoxide can be used by many other microorganisms as an electron acceptorfor an anaerobic electron transport with dimethylsulphide as the product. Evenman-made chemicals such as tetrachloroethene and some chloroaromatic com-pounds such as chlorobenzoates and chlorophenols can function as electron ac-ceptors in a dehalorespiration (these will be discussed later in detail).

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Table 3. A general picture of microbial transformations of organic compounds at various redox conditions

Oxygen reduction Nitrate reduction Manganese Iron reduction Sulfate reduction Carbon dioxide reduction reduction

Aromatics:Benzene (very fast) Benzene Benzene (very slow) Benzene (very slow)Toluene (very fast) Toluene Toluene Toluene Toluene TolueneEthylbenzene (very fast)Xylenes (very fast) XylenesBiphenyl (very fast) Biphenyl (very slow)Naphthalene (very fast) Naphthalene (slow) Naphthalene (slow) Naphthalene (very slow)Polycyclic aromatic hydrocarbons Phenanthrene

up to 4 rings (very fast to slow) (very slow)Benzoate (very fast) Benzoate (fast) Benzoate (fast) Benzoate (fast)Phenol (very fast) Phenol (slow) Phenol (slow)Cresols (very fast) Cresols (slow)Aniline (very fast) Aniline (slow)Chloroaromatics:Chlorobenzoates (very fast)Monochlorotoluenes (very fast)Mono-, di-, tri-, pentachlorophenol Chlorophenols (slow) Chlorophenols (slow)

(very fast to fast)Chloroanilines (very fast)Mono-, dichlorobiphenyls (very fast)Mono-, di-, tri-, tetrachlorobenzene

(very fast)Aliphatics:PropaneC10–C22 n-alkanes n-Hexadecane 2,6,10,14-Tetramethylpentadecane (very slow)Alkenes

Aerobic and Anaerobic Biodegradation Potentials of Microorganism

s11

Table 3 (continued)

Oxygen reduction Nitrate reduction Manganese Iron reduction Sulfate reduction Carbon dioxide reduction reduction

Haloaliphatics:Haloalkanes Haloalkanes HaloalkanesHaloalkenes Haloalkenes HaloalkenesHaloalkanolsHaloalkenolsHaloalkanoates (very fast)Haloalkenoates

Classification of growth rate (td) of population: very fast degradation: hours; fast degradation: < 1 day; slow degradation: > 1 day, <7 days; very slowdegradation: > 3 weeks

1.5Limitations to Fermentation

Most anoxic terrestrial and subsurface environments do not receive sufficientinflux of external electron acceptors for the oxidation of the organic com-pounds that are present. This leaves fermentation as the only possible physiolo-gical process for biodegradation. However, many compounds cannot be simplyfermented by pure bacterial cultures for biochemical and energetic reasons.This is illustrated for some aromatic compounds. Fermentative degradation ofbenzoate, phenol, and monohydroxybenzoates to acetate, hydrogen, and carbondioxide is not possible in pure culture, since the reactions are endergonic understandard conditions [6–8] (Table 4). These fermentations become possible onlyif a hydrogen-utilizing methanogenic bacterium keeps the hydrogen partialpressure low (about 10–4 bar) to render the reactions exergonic (Table 5). Suchso-called “syntrophic” couplings between different metabolic types of bacteriaare widespread in methanogenic degradative processes [9]. However, the syn-trophic coupling between fermentative and methanogenic bacteria leads to verylow yields of energy compared to oxidation of benzoate with other electron acceptors (Table 6). In contrast to the non- and monohydroxylated aro-

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Table 4. Fermentation of aromatics

Overall reaction Free energy, DGo¢

C7H5O2– + 7 H2O Æ 3 CH3COO– + 3 H+ + 3 H2 + HCO3

– +70.6 kJ/mol benzoateC7H5O3

– + 6 H2O Æ 3 CH3COO– + 3 H+ + 2 H2 + HCO3– +5.4 kJ/mol p-hydroxybenzoate

C6H6O + 5 H2O Æ 3 CH3COO– + 3 H+ + 2 H2 +6.6 kJ/mol phenol

Table 5. Fermentation of aromatics coupled to methanogenesis


C7H6O2 + 4.5 H2O Æ 3.25 CO2 + 3.75 CH4 + H+ –159 kJ/mol benzoateC7H6O3 + 3.5 H2O + H+ Æ 3.25 CO2 + 3.75 CH4 –178 kJ/mol p-hydroxybenzoateC6H6O + 3.5 H2O + H+ Æ 2.25 CO2 + 3.75 CH4 –140 kJ/mol phenol

Table 6. Energetics of the oxidation of benzoate coupled to anaerobic respiration (accordingto Fuchs et al. [10])


Nitrate respiration:C7H6O2 + 15 NO3

– Æ 7 CO2 + 15 NO2– + 3 H2O –2116 kJ/mol benzoate

Sulfate respiration:C7H6O2 + 3.75 SO4

2–+7.5 H+ Æ 7 CO2 + 3.75 H2S + 3 H2O –327 kJ/mol benzoateCarbonate respiration:C7H6O2 + 4.5 H2O Æ 3.25 CO2 + 3.75 CH4 + H+ –159 kJ/mol benzoate

matics mentioned above, the fermentation of two- or threefold hydroxylatedaromatics is sufficiently exergonic to allow degradation in pure culture [11, 12](Table 7).

1.6Various Degrees of Degradation

Evidence for a microbial role in the transformation of organic chemicals bysamples from the natural environment such as soil, water, and sediments can beobtained by demonstrating that the compound is transformed in nonsterile butnot in sterilized samples. Chemicals are usually subject to a variety of microbialreactions, resulting in various types and degrees of degradation. With 4-chloro-biphenyl as a model compound, the different types of processes which can takeplace have been schematically described (Fig. 4).

Biodegradation is often a growth-linked process that brings about total(complete) degradation or mineralization. As the microorganisms convert the


Table 7. Fermentation of phenolic aromatic compounds


C7H5O4– + 5 H2O Æ 3 CH3COO– + 3 H+ + H2 + HCO3

– – 77.8 kJ/mol protocatechuateC7H5O5

– + 4 H2O Æ 3 CH3COO– + 3 H+ + HCO3– –160.0 kJ/mol gallate

C6H6O2 + 4 H2O Æ 3 CH3COO– + 3 H+ + H2 – 78.1 kJ/mol catecholC6H6O3 + 3 H2O Æ 3 CH3COO– + 3 H+ –158.3 kJ/mol pyrogallol

Fig. 4. Comparison of the degree of degradation of 4-chlorobiphenyl as a model compoundby cometabolism, by partial degradation, by the use of the compound as an electron accep-tor, and by total degradation processes. Compound (middle gray), chlorosubstituted metabo-lite (light gray), biphenyl (dotted line), chloride (dark gray), and cells (pointed line)

organic substrate to fermentation products or carbon dioxide, energy is released and fixed in the form of ATP, as described in the first sections.Simultaneously, the organisms convert some of the carbon in the substrate tocell constituents, using ATP and NAD(P)H produced by oxidation reactions forthese biosynthetic reactions. As a result, the populations increase in numbersand biomass. Organic chlorine is returned to the mineral state during the min-eralization process.

With many chemicals a type of microbial conversion quite different frommineralization takes place. The phenomenon that chemicals are subject tomicrobial action and yet do not sustain growth of the organisms has been term-ed cometabolism, or sometimes cooxidation or fortuitous oxidation [13–16].The responsible organisms are presumably growing on another substrate whileperforming cometabolic transformation reactions. The microorganisms in-volved in cometabolic transformation do not increase in numbers or biomassas a result of the degradation of the chemical of interest. This lack of growth isa reflection of the inability of the organisms to use the chemical for energy gen-eration or biosynthetic purposes, and it is in marked contrast to the increase inpopulation size or biomass when a mineralizable substrate is introduced intothe same sample. Because populations are usually small, a compound subject tocometabolism is modified slowly, and the rate of degradation does not increasewith time. The product of cometabolism will often be used by other micro-organisms.

In some cases the conversion of organic chemicals may support microbialgrowth even though no complete mineralization occurs. This was observed,for example, during the degradation of 4-chlorobiphenyl. Population increaseand chemical disappearance were observed when aerobic biphenyl-degradingpopulations were provided with 4-chlorobiphenyl. In contrast to a substratethat is mineralized, the bacteria could only use part of the molecule (thenonsubstituted ring) as a carbon and energy source for growth. The bacterialpopulation size and the rate of decline in concentration of the chemical in-creased over time. However, elimination of the chlorine substituent, whichwould be typical for mineralization, did not take place and the chlorine-con-taining metabolite 4-chlorobenzoate accumulated. This process is termed par-tial degradation.

Microbial populations present in anaerobic sediments are often able to dealwith chlorinated chemicals in a quite different way. Compounds such as chloro-aromatics and chloroaliphatics can function as electron acceptors in a type ofanaerobic respiration [17–19]. Evidence for the occurrence of such a process innatural ecosystems is provided by the observation that reductive dechlorinationdoes occur in nonsterile but not in sterilized samples. The addition of electrondonors such as pyruvate or hydrogen stimulates the process. This type of re-ductive dechlorination of 4-chlorobiphenyl has never been studied but in ana-logy to the observation with chlorobenzoates, chlorophenols, and tetrachloro-ethene, giving the non- or lower chlorinated products, the formation of bi-phenyl may occur. Reductive dechlorination proceeds relatively easily withhighly chlorinated compounds and the use of a chlorinated chemical as anelectron acceptor is sometimes called dehalorespiration.

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1.7Résumé of Introduction

Redox processes are of key importance for microbial activity and they usuallyrequire an electron acceptor in addition to the organic substance itself, since thepossibilities of fermentation are restricted. Thus, the availability of an electronacceptor is essential. Various compounds such as aromatic and aliphatic com-pounds as well as their chlorinated analogous may be used.

After the description of the more general subjects of microbial metabolism,the degradative properties of microorganisms will be discussed in more detailfor some selected chemicals. Aromatic and chloroaromatic compounds havebeen chosen to document the differences in the degradation that are related tothe presence or absence of special substituents on the aromatic ring. In addi-tion, the degradation of aliphatic and chloroaliphatic compounds will be dis-cussed because of the occurrence of these chemicals as pollutants in the en-vironment. This will be used to illustrate the potential but also the limitationsof some microbial systems in complete degradation. The abundance of reac-tions involved in the degradative pathways, and also the widespread use of va-rious analogous reactions, will be discussed. A comparison of pathways used byaerobic and anaerobic organisms, as well as the differences between bacteriaand fungi with respect to biodegradation, are subjects of the following sections.

2Degradation of Aromatic Compounds

2.1Aerobic vs Anaerobic Degradation: Introduction

Aromatic compounds can be degraded under aerobic and anaerobic conditions.In both cases a key step is the activation of the inert aromatic ring. In the pre-sence of oxygen this is carried out by oxygen-dependent enzymes. Successiveaddition of two oxygen atoms to the ring or direct introduction of molecularoxygen leads to the formation of diphenolic compounds. These compounds arethen cleaved by enzymes that use molecular oxygen again. While bacteria usesoxygen for ring activation and cleavage, ligninolytic fungi bring about the ac-tivation by use of hydrogen peroxide. The formation of an aryl cation radical isthen followed by spontaneous rearrangements and degradation. In contrast,non-ligninolytic fungi use oxygen to activate the aromatic ring which is follow-ed by coupling to give O-conjugates. Thus, oxygen plays a vital role both as a ter-minal electron acceptor and as reagent in the biochemical activation of inertaromatic compounds under oxic conditions.

Under anoxic conditions various inorganic compounds can function as anelectron acceptor instead of oxygen, but there is apparently no equivalent re-placement for oxygen with respect to its function in activation reactions.Therefore, the anaerobes have to degrade aromatic compounds by reductiveconversions. In the anaerobic conversion central intermediates ready for hydro-lytic ring cleavage bear a 1,3-dioxo structure in the alicyclic ring when resorci-


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Fig. 5. General scheme for the metabolism of aromatic compounds by bacteria under oxic (left) and anoxic (right), and by fungiunder oxic conditions (middle)

nol or phloroglucinol are the central metabolites.When benzoyl-CoA is the cen-tral metabolite after the channeling reactions, one oxo group is in the alicyclicring and the other exocyclic in the ester group.

Thus, different degradation sequences are used under oxic and anoxic con-ditions, as schematically illustrated in Fig. 5.

2.2Aerobic Degradation of Aromatics: General Differences Between Prokaryotic and Eukaryotic Organisms in the Initial Reactions

Higher organisms, including fungi, have evolved a different enzyme system forthe oxidation of aromatic hydrocarbons from that observed in bacteria (Fig. 6).Most non-ligninolytic fungi are able to oxidize aromatic hydrocarbons by acytochrome P-450 monooxygenase. One atom of the oxygen molecule is incor-porated into the aromatic substrate, while the other oxygen atom is reduced towater. The arene oxide formed then becomes a substrate for further meta-bolism. The enzymatic hydration of the arene oxide leads to the formation of adihydrodiol with a trans configuration (Fig. 6). Another pathway involves iso-


Fig. 6. Differences in the initial reactions used by prokaryotic and eukaryotic organisms forthe oxidation of aromatic hydrocarbons. The upper reactions 1–4 are used by fungi: � cyto-chrome P-450 monooxygenase; � non-enzymatic rearrangement; � coupling reactions; �epoxide hydrolase. The lower reactions are used by bacteria: � dioxygenase; � cis-dihydro-diol dehydrogenase; � and � monooxygenase, ring fission pathways starting with a di-oxygenase

merization of the arene oxide to form a phenol that can be conjugated with sul-fate, glucuronic acid, or glucose.

The following two different routes are used by bacteria:1. An aromatic compound without a hydroxyl group is activated through a di-

oxygenase using molecular oxygen. Subsequently, the resulting cis-dihydro-diol is converted by a dehydrogenase to give a 1,2-diphenol derivative, re-gaining the aromatic state. The diphenol is the substrate for ring cleavage, areaction which requires another molecule of oxygen.

2. The use of successive monooxygenase reactions has also been described. Inthis case, the diphenol is produced in two steps, and the hydroxyl groupsneed not to be on adjacent carbon atoms. In contrast to the reaction in theeukaryotic organisms, no epoxides are formed as intermediates.Details of the further routes after ring cleavage will be discussed later.

2.3Degradation of Aromatic Compounds by Aerobic Bacteria

Enzymes occurring in pathways for the aerobic degradation of aromatic com-pounds have to fulfil various functions. An important point is to overcome thechemical stability of the aromatic ring. Besides ring activation and cleavage,further degradative steps have to achieve the formation of common interme-diates of cell metabolism by cleavage of carbon-carbon bonds and by proces-sion of functional groups or side-chains of substituted aromatics and multi-substituted aromatics arising from bi- and polycyclic aromatics. I have focusedon the chemical logic of the overall metabolic pathways to illustrate how a fewtypes of chemical mechanisms are used by the organisms in linked, sequentialreactions to meet the above requirements.

2.3.1Reactions Converting Aromatic Compounds into Ring Cleavage Substrates

Aerobic bacteria and especially pseudomonads are able to use various com-pounds as their source of carbon and energy, whose chemical structures con-tain one or more benzene rings. Ring cleavage is an important enzymatic stepfor the degradation of aromatics. With rare exceptions the introduction of twohydroxyl groups onto a benzene ring is a prerequisite for ring opening. The sub-strates for ring cleavage may be 1,2-diphenolic compounds such as catechol,protocatechuate, and catechol derivatives, which bear either substituents suchas a phenyl, a hydroxy phenyl, and a phenoxy group, or an additional aromaticring condensed to the first one (see Fig. 7). Gentisate, a 1,4-diphenolic com-pound, is a ring cleavage substrate too.

2.3.1.1Formation of Diphenols from Monocyclic Hydrocarbons

The conversion of a hydrocarbon to a catecholic ring fission substrate involvestwo important enzymatic steps: a dioxygenase catalyzes the formation of a cis-

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dihydrodiol, which is then oxidized to the corresponding catechol by a dehy-drogenase. This occurs, for example, with benzene and benzoate degradation(Fig. 8). In some cases only a dioxygenase is needed to produce a catechol, forexample during aniline degradation, where a 1,2-diphenol is formed by an ani-line dioxygenase which concomitantly eliminates NH3. In phenol and salicylatedegradation a monooxygenase adds one atom of molecular oxygen to the sub-strate to give catechol, while the other oxygen atom is reduced to water.

The degradation pathways of m-cresol, naphthalene, and phthalate convergevia protocatechuate and gentisate. Monooxygenase as well as dioxygenase reac-tions are involved in the formation of these diphenols (Fig. 9). The degradationof salicylate can proceed via catechol (Fig. 8) or gentisate as the ring cleavagesubstrate (Fig. 9) depending on the organism.

Aromatic compounds which bear alkyl substituents on the aromatic ring mayundergo oxidation of the side chain before ring activation. Toluene is a goodcompound to illustrate the wide variety of reactions which can take place in re-aching a diphenolic structure (Fig. 10). Five different routes for toluene degra-dation have been found: (1) side chain oxidation by a monooxygenase and twodehydrogenases to give the carboxylic acid, which is further degraded by the di-oxygenase pathway; (2+3) two monooxygenase reactions plus side chain oxida-tion to form protocatechuate – these monooxygenase reactions may start at C-2or C-3; (4) two subsequent monooxygenase reactions activating the aromatic ringleading to 3-methylcatechol; (5) the same methylsubstituted catechol is formedwhen a dioxygenase and a dehydrogenase are involved in ring activation.

2.3.1.2Formation of Diphenols from Bi- and Polycyclic Hydrocarbons

The presence of analogous reactions in the degradation pathways of aromaticcompounds has been shown for binuclear compounds like biphenyl, dibenzo-furan, dibenzo-p-dioxin, and naphthalene (Fig. 11). The general scheme is thefollowing: ring activation by a dioxygenase, rearomatization by a dehydrogen-ase to give a 1,2-diphenol, ring cleavage by dioxygenase, fission of the aliphaticside chain by a hydrolase to give, on the one side, a-oxo acids such as pyruvateor compounds which are metabolites of the meta pathway (will be discussed la-ter), such as 4-oxalocrotonate and 2-oxopent-4-enoate and, on the other side, sali-


Fig. 7. Diphenolic compounds that can undergo ring fission

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Fig. 8. Degradative steps for monocyclic aromatic compounds that lead to catechol [20–28].� benzene; � aniline; � salicylate; � phenol; � benzoate. Catechol, the central metabolite,is further degraded via the meta or ortho pathway

Fig. 9. Sequences for the bacterial degradation of aromatic compounds that converge via pro-tocatechuate and gentisate: m-cresol via m-hydroxybenzoate and protocatechuate or genti-sate, p-cresol via protocatechuate; naphthalene via salicylate and gentisate, phthalate via 4,5-dihydroxyphthalate [29–38]. � phthalate dioxygenase; � 4,5-dihydroxyphthalate decarboxy-lase; � p-hydroxybenzoate hydroxylase; � salicylate hydroxylase

cylaldehyde, salicylate, catechol or benzoate. During the degradation of binucleararomatics, the formation and cleavage of a 1,2-diphenol takes place twice.

In the degradative pathway of dibenzo-p-dioxin and dibenzofuran the cleav-age of the stable ether bond is an important step. Interestingly, no special en-zyme is necessary to bring about this reaction. Angular dioxygenation by thering activating enzymes, i.e., oxygenation at a pair of vicinal carbon atoms, oneof which is involved in the bridge between the two aromatic rings, forms cis-dihydrodiols, which are hemiacetals so that because of their instability the etherbond is cleaved spontaneously. Therefore, a dehydrogenase reaction, normallybringing about rearomatization, is unnecessary.

Various polycyclic aromatic hydrocarbons (PAHs) containing up to fourrings, such as pyrene and chrysene, can be mineralized by pure bacterial cul-


Fig. 10. Different routes for the aerobic bacterial mineralization of toluene via catechol, 3-me-thylcatechol, and protocatechuate. Enzymes: �, toluene 2-monooxygenase [39]; �, toluene 3-monooxygenase [40]; �, toluene dioxygenase [41]; �, toluene cis-dihydrodiol dehydrogen-ase [42, 43]; �, xylene monooxygenase [44]; �, benzylalcohol dehydrogenase [45]; �, benz-aldehyde dehydrogenase [46]; �, benzoate dioxygenase [47]; , benzoate cis-dihydrodioldehydrogenase [48]; , toluene 4-monooxygenase [49, 50]; �; 4-cresol dehydrogenase [51];�, 4-hydroxybenzaldehyde dehydrogenase [51]; , 4-hydroxybenzoate hydroxylase [51]

tures [63–74]. Because of the different degrees of solubility (Table 8), the de-gradation rates vary strongly, with the higher molecular weight PAHs being de-graded very slowly. These latter PAHs tend to resist degradation and bioaccu-mulate in biological material due to the hydrophobic character. Complete deg-radation pathways have been elucidated for anthracene and phenanthrene.Various metabolites of acenaphthene, acenaphthylene, fluorene, fluoranthene,pyrene, and benzo[a]anthracene have been identified, allowing some clues onthe catabolic pathways [67, 69, 78–82]. The degradation generally follows thesame pathway as described above for the bicyclic aromatic hydrocarbons: ringactivation, rearomatization, ring cleavage, and subsequent fission of the ali-phatic side chain to give pyruvate and an aromatic ortho-hydroxy acid after oxi-dation of the aldehyde group. Then decarboxylation yields a diphenolic deriva-tive of the remaining aromatic ring which is subject to the next cycle for the eli-mination of an aromatic ring. This cycle of reactions is illustrated for the linearand angular PAHs, anthracene and phenanthrene, respectively, in Fig. 12.

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Fig. 11. Degradation sequences of bicyclic hydrocarbons that converge at catechol. � naph-thalene [52, 53]; � dibenzofuran; � dibenzo-p-dioxin [54–59]; � biphenyl [26, 27, 60–62].Catechol, the central metabolite, is further degraded via the meta or ortho pathway. The sitesof cleavage of C-C bonds are marked

dioxygenase

dehydr ogenaseor spontaneous

meta cleavingdioxygenase

hydrolase


Table 8. Polycyclic aromatic hydrocarbons used by pure cultures of bacteria as the sole sourceof carbon and energy and important physicochemical properties

Compound Structure Aqueous Octanol/ Degrada- Ionizationsolubility water co- tive path- potential(mg/l) efficient way (eV)at 25 °C (log Kow)

Naphthalene 31.7 3.37 ++ 8.12

Biphenyl 7.0 3.9 ++

Acenaphthene 3.42 4.33 + 7.61

Acenaphthylene 3.93 4.07 +

Fluorene 1.98 4.18 ++

Anthracene 0.075 4.45 ++ 7.43

Phenanthrene 1.6 4.46 ++ 8.03

Fluoranthene 0.265 5.33 + 7.85

Pyrene 0.148 5.32 + 7.53

Chrysene 0.002 5.61 + 7.81

Benzo[a]anthracene 0.014 5.61 + 7.56

++, well established; +, metabolites identified.Data compiled from [75–77].

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Fig. 12. Degradative pathway for anthracene and phenanthrene: path a – Pseudomonas [83];path b – Aeromonas, Alcaligenes, Micrococcus, Mycobacterium, Vibrio [84–89]

Two modes of initial PAH degradation may occur, as has been shown forfluorene (Fig. 13). The first pathway, which involves dioxygenation and metacleavage of one aromatic ring, with the subsequent release of pyruvate, oxida-tion of the aldehyde group, followed by decarboxylation [73, 90–93], is analo-gous to the pathways described for phenanthrene and anthracene.An importantreaction is the biological Baeyer-Villiger oxidation of the cyclic ketone, inda-none, yielding 3,4-dihydrocoumarin, which is then subject of hydrolytic fissionto form 3-(2-hydroxyphenyl) propionate.

The second pathway is initiated by monooxygenation at C-9 to give 9-fluore-nol, which is then dehydrogenated to 9-fluorenone (Fig. 13). These reactions


Fig. 13. Degradation of fluorene. Pathway (a) – with initial dioxygenase; oxidation of cyclicketones by Baeyer-Villiger oxidation. Pathway (b) with initial monooxygenase. Dashed ar-rows indicate two or more successive reactions

precede dioxygenase attack adjacent to the carbonyl group to form the angulardiol 1,1a-hydroxy-1-hydro-9-fluorenone [54, 94]. Cleavage of the five-member-ed ring then generates a substituted biphenyl, whose further degradation byreactions similar to those of biphenyl catabolism leads to the formation ofphthalic acid [95, 96]. Phthalate can be further metabolized via 4,5-dihydroxy-phthalate, protocatechuate, and b-carboxy-cis,cis-muconate [95].

2.3.2Ring Fission and Carbon Chain Fission

2.3.2.1Ring Cleavage

There are two distinct modes of oxidative cleavage of the benzene nucleus.Cleavage of the bond between adjacent carbon atoms that carry hydroxylgroups is know as ortho or intradiol cleavage (Fig. 14). The pathway by whichthe product of such a cleavage is metabolized is termed the ortho or b-ketoadi-pate pathway. Ortho cleavage of a carbon-carbon bond is catalyzed by a catechol1,2-dioxygenase or a protocatechuate 3,4-dioxygenase.

In the second mode of fission of the benzene nucleus C-C-bond cleavage oc-curs between two carbon atoms of which one carries a hydroxyl group and theother carries another substituent or a hydrogen atom. In this case, the hydroxylgroups may be either ortho or para to one another, and the enzymes catalyzingsuch cleavage reaction are again designated by the position of the carbon bondwhich is cleaved. Thus, catechol and protocatechuate are cleaved, respectively,

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Fig. 14. Mode of ring fission

by a catechol 2,3-dioxygenase or by a protocatechuate 4,5-dioxygenase or a pro-tocatechuate 2,3-dioxygenase. Gentisate is cleaved by gentisate 1,2-dioxygenase.The 1,2-diphenols derived from biphenyl, naphthalene, dibenzofuran, and di-benzodioxin are all cleaved by meta-cleaving dioxygenases.

2.3.2.2Carbon Chain Carbon-Carbon Fission

Following ring fission, further reactions yield pyruvate and/or other a-ketoacid intermediates. A key step is carbon-carbon bond cleavage, which may in-volve stabilization of carbanionic intermediates or transition states. The carbonchain is cleaved by aldol fission, hydrolytic fission, decarboxylation, or thiolyticfission after formation of compounds that carry in a b-position to a carbonylfunction a hydroxyl, an oxo, or a carboxyl group (Fig. 15).

2.3.2.3Aldol Cleavage

Substrates for aldol cleavage have the partial structure shown in Fig. 16.Oxygen as an electronegative atom stabilizes the incipient negative charge

density during the reaction, to give the enolate as the initial product. Therefore,an aldehyde and a ketone are the fission products as has been shown with com-pounds A and B in Fig. 15.


Fig. 15. Carbon chain carbon-carbon bond fission substrates

2.3.2.4Hydrolytic Fission

The fission of fumarylpyruvate (compound C) is a good example to illustratethis type of cleavage to occur from the 1,3-dioxo structure (Fig. 17). Inspectionof the fumarylpyruvate skeleton suggests that H2O could be added nucleophili-cally to the keto group of the fumaryl moiety to form a tetrahedral adduct. Thetetrahedral adduct in equilibrium with the dicarbonyl is competent for C–Ccleavage, since it could (in a low-energy path) expel the stable enolate anion ofpyruvate and fumarate.

The same type of reaction producing a carboxylic acid and a ketone was found for compounds D and E in Fig. 15.

2.3.2.5Decarboxylation

A b-keto group facilitates decarboxylation by stabilizing the carbanionic tran-sition state as decarboxylation proceeds (Fig. 18). The ready possibility for eno-lization allows the b-carbonyl group to act as an electron sink, and ketonizationfollows. This type of reaction is plausible for compound F in Fig. 15.

2.3.2.6Vinylogy

Some cleavage substrates have no b-dicarbonyl structure. However, the inser-tion of a –CH=CH– group between two functional groups produces a vinylo-gous compound whose property is similar to the compound without the vinylicgroup. The concept is called principle of vinylogy [97]. Therefore compound F,formed in the meta pathway, is a vinylogous b-keto acid ready for decarboxyla-tion, while compounds D and E are vinylogous to 1,3-dicarbonyl compoundswhich are cleaved by a hydrolytic type of reaction (Fig. 15).

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Fig. 16. Aldol cleavage

Fig. 17. Hydrolytic cleavage of fumarylpyruvate

2.3.3Pathways as a Whole for Catechol, Protocatechuate, and Gentisate

2.3.3.1Gentisate Pathway

The first mode of preparation for fission between carbon atoms, i.e., forming acarbonyl group in a b-position to another carbonyl, is seen in the gentisatepathway (Fig. 19).After the benzene nucleus has been cleaved by a gentisate 1,2-dioxygenase, one of two reaction pathways is available depending upon the na-ture of the organism. Some organisms use a sequence in which the ring fissionproduct, maleylpyruvate, is first isomerized to fumarylpyruvate. Both com-pounds are in equilibrium with the respective 1,3-diketones which are suitablesubstrates for hydrolysis whereas a 1-keto-3-hydroxy grouping permits aldolfission to occur. The point of ring fission, together with the positioning of thehydroxyl groups in gentisate, determine that 1,3-diketone substrates will befurnished for hydrolysis. The function of the isomerase is to ensure that an in-termediate of the tricarboxylic acid cycle, fumarate, results from the next reac-tion. In other organisms this isomerase is not present: maleylpyruvate is hydro-lyzed directly to give maleate which is then hydrated to form D-malate.

2.3.3.2meta Pathway

In meta fission sequences of catechol the carbon chain is prepared for the fis-sion into pyruvate and acetaldehyde the following way (Fig. 20). After ring fis-sion by catechol 2,3-dioxygenase to give 2-hydroxymuconic semialdehyde, cate-chol is metabolized by two routes. For the particular organisms studied the oxi-dative route is quantitatively the more important for catechol. The enzymecatalyzed keto-enol change is known to precede the decarboxylation step sincea vinylogous b-keto acid is formed. Both routes converge to 2-oxopent-4-eno-ate, which is hydrated to introduce a hydroxyl group at C-4. The carbonyl groupat C-2 of the resulting 4-hydroxy-2-keto acid (4-hydroxy-2-oxovalerate) wasformed by ketonization of the hydroxyl group originally present in the ring fis-sion substrate and its product. This carbonyl also appears in the pyruvatemolecule formed from the hydroxyketo acid by aldol cleavage.


Fig. 18. Decarboxylation of b-keto acids

2-Hydroxymuconic semialdehyde is also a metabolite in the degradation ofprotocatechuate formed by decarboxylation of 2-hydroxy-4-carboxymuconicsemialdehyde.

2.3.3.3b-Ketoadipate Pathway (3-Oxoadipate)

The b-ketoadipate or ortho fission pathway (Fig. 20) is a multistep convergentmetabolic route used by many microorganisms to convert either of two com-pounds, protocatechuate and catechol, to succinyl-CoA and acetyl-CoA[105–107]. Reactions of the protocatechuate and the catechol branch both giverise to the common intermediate, b-ketoadipate enol-lactone. The reactions of

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Fig. 19. Gentisate pathway [29, 30, 32, 36, 98–104]. The following enzymes are involved: �gentisate 1,2-dioxygenase, � GSH dependent or GSH independent isomerase; � maleylpy-ruvate hydrolase; � fumarylpyruvate hydrolase, � maleate hydratase; � fumarase


Fig. 20. Meta and ortho cleavage pathways for degradation of protocatechuate and catechol.The meta cleavage pathway is illustrated on the left side, while the ortho (or b-ketoadipate)pathway is given on the right side. Divergence is determined by the nature of the dioxygen-ases which cleave each diphenolic substrate. The mode of cleavage is shown by a line of stars.The following enzymes are involved in the meta pathway [108–113]: � catechol 2,3-dioxyge-nase; � 2-hydroxymuconic semialdehyde dehydrogenase; � 4-oxalocrotonate isomerase; �4-oxalocrotonate decarboxylase; � 2-hydroxymuconic semialdehyde hydrolase; � 2-oxo-pent-4-enoate hydratase; � 4-hydroxy-2-oxovalerate aldolase; � acetaldehyde dehydrogen-ase (acylating). Enzymes of the ortho pathway are as follows [106, 109, 114–118]: a, catechol1,2-dioxygenase; a¢, protocatechuate 3,4-dioxygenase; b, muconate cycloisomerase (muco-nate lactonizing enzyme); b¢, b-carboxymuconate lactonizing enzyme; c, muconolactone iso-merase; c¢, g-carboxymuconolactone decarboxylase; d, b-ketoadipate enol-lactone hydrolase;e, b-ketoadipate succinyl CoA transferase; f, b-ketoadipyl CoA thiolase

the two branches are chemically analogous: dioxygenase-mediated cleavage be-tween the carbon atoms that carry the hydroxyl groups yields cis,cis-muconatefrom catechol and b-carboxy-cis,cis-muconate from protocatechuate. Strictlyanalogous enzyme reactions convert the two muconates to muconolactone andg-carboxymuconolactone, respectively. Decarboxylation of g-carboxymucono-lactone forces the migration of the double bond within the lactone ring to yieldb-ketoadipate enol-lactone; deprotonation of the g-carbon of muconolactonegives rise to the same product via an analogous mechanism. Hydrolytic openingof the b-ketoadipate enol-lactone results in the formation of b-ketoadipate. Insummary, lactonization followed by delactonization is the strategy to form the3-keto acid. This compound is further oxidized after formation of a coenzymeA ester analogous to the catabolism of fatty acids. The thiolytic fission of co-enzyme A esters of 3-keto acids to generate Krebs cycle intermediates repre-sents the fourth mode of fission of the carbon chain. The coenzyme A ester,which is formed by the reaction of 3-ketoadipate with succinyl-CoA catalyzedby a transferase, undergoes thiolysis by reaction with coenzyme A, yieldingacetyl-CoA and generating succinyl-CoA to be used for esterifying anothermolecule of 3-ketoadipate.

2.4Degradation of Aromatics by Fungi

Whereas the mineralization of aromatic compounds including PAHs and thedegradation pathways of bacteria have been well studied, knowledge of similaractivities in fungi is limited to a few species of soil fungi and various white-rotfungi. In contrast to prokaryotes, fungi rarely utilize aromatics as a sole sourceof carbon and energy. An Aspergillus fumigatus was found to be capable ofgrowth on phenol, p-cresol, and 4-ethylphenol [119–121]. Pathways via cate-chol, protocatechuate, and hydroquinone were proposed. Weber et al. [122] re-ported the discovery of a toluene-degrading fungus, Cladosporium sphaero-sperum. In addition to toluene, the organism is able to use styrene, ethylben-zene, and propylbenzene as the sole source of carbon and energy. There areindications that the degradation of toluene is initiated by oxidation of the me-thyl group. Just recently, a hyphomycete Scedosporium apiospermum was isola-ted which is able to grow on phenol and p-cresol with 3-oxoadipate as themetabolite [123].

2.4.1Degradation by Non-Ligninolytic Fungi

A variety of fungi have been found to transform aromatic compounds includingcomplex polycyclic aromatic hydrocarbons to metabolites that are similar tothose produced by mammalian enzymes. Only a few fungi appear to have theability to catabolize PAHs to CO2 [124].

In Cunninghamella elegans, a non-ligninolytic fungus, and several otherfungi naphthalene is metabolized via a branched pathway to naphthalene trans-1,2-dihydrodiol, 1-naphthol, 2-naphthol, 4-hydroxy-1-tetralone, 1,4-naphtho-

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quinone, and 1,2-naphthoquinone (Fig. 21). Two conjugates, 1-naphthylgluc-uronide and 1-naphthylsulfate, are also produced by C. elegans. The observa-tions are consistent with a mechanism for naphthalene trans-1,2-dihydrodiolformation in which a cytochrome P-450 monooxygenase catalyzes the forma-tion of naphthalene 1,2-oxide and then an epoxide hydrolase adds water toform the dihydrodiol. Since naphthalene 1,2-oxide is unstable in solution itrapidly rearranges to form phenols, principally 1-naphthol.

The same sequence of reactions have been shown for a variety of other poly-cyclic aromatic hydrocarbons, such as acenaphthene, anthracene, phenan-threne, benzo[a]pyrene, benzo[a]anthracene, fluoranthene, and pyrene.Cunninghamella elegans initiates the oxidation of anthracene by incorporatingone atom of molecular oxygen into the aromatic ring to form anthracene 1,2-oxide, which is hydrated to form anthracene trans-1,2-dihydrodiol (Fig. 22).Anthracene 1,2-oxide is rearranged rapidly to form 1-anthrol, which is subse-quently conjugated with sulfate.

Phenanthrene is metabolized by Cunninghamella elegans predominately at the 1,2-positions to form phenanthrene trans-1,2-dihydrodiol and a glu-coside conjugate of 1-phenanthrol (Fig. 23). The carbons at the 3,4- and 9,10-


Fig. 21. Proposed pathway for the fungal oxidation of naphthalene. � cytochrome P-450monooxygenase, � epoxide hydrolase

positions may also be oxidized to form trans-3,4-dihydrodiol and trans-9,10-dihydrodiol.

2.4.2Degradation of PAHs by Ligninolytic Fungi

2.4.2.1The Ligninolytic System

White rot fungi have the ability to degrade lignin efficiently [125–128]. This ca-pacity results from the activities of a complex system (Fig. 24) composed of ex-tracellular heme-containing peroxidases, known as lignin peroxidases [129]and manganese-dependent peroxidases [130], a H2O2-generating system, otheroxidases, and laccases [126, 131]. The enzymes are produced in response to lowlevels of sources of carbon, nitrogen, or sulfur nutrients [132, 133]. Interestingly,these fungi do not use lignin as a carbon source for growth; instead they de-grade the lignin to obtain the cellulose that is in the interior of the wood fiber[134]. Ligninolysis only occurs when other readily degradable substrates areavailable.

Extracellular hydrogen peroxide will be produced by oxidases that utilizecompounds such as glucose or glyoxal from molecular oxygen by two-electronreduction [135].

Lignin peroxidases – highly potent oxidizing agents – can abstract one elec-tron from a non-phenolic moiety of the lignin molecule, thus creating a cation

34 W. Reineke

Fig. 22. Proposed pathway for the fungal oxidation of anthracene. � cytochrome P-450monooxygenase, � epoxide hydrolase


Fig. 23. Proposed pathway for the fungal oxidation of phenanthrene. � cytochrome P-450monooxygenase, � epoxide hydrolase

Fig. 24. The extracellular ligninolytic system in ligninolytic fungi. � glyoxal oxidase, � li-gnin peroxidase, � manganese-dependent peroxidase. The structure represents one aroma-tic moiety of the complex lignin molecule

radical [136] which in turn initiates a random oxidative chemical reaction thatfinally results in the oxygenation and depolymerization of lignin including C-C-bond cleavage.

Manganese-dependent peroxidases function by oxidizing Mn2+ to Mn3+.Mn3+ behaves as a low-molecular-weight mediator that can initiate the oxida-tion process [137].

Interesting features are the catalytic cycles of lignin and manganese peroxi-dase (LiP and MnP). The resting lignin peroxidase (ferric state) is oxidized by atwo-electron transfer to H2O2 to form compound I (LiPI, a ferryl (Fe•4+) p-por-phyrin cation radical) [138]. LiPI oxidizes a substrate molecule (aryl, Ar) by oneelectron, forming compound II (LiPII) and a free radical product (aryl cationradical, Ar•+). LiPII reacts with another substrate molecule, forming back thenative enzyme and a free radical. The free radicals then undergo nonenzymaticreactions to form the final products.

LiP + H2O2 Æ LiPI + H2O

LiPI + Ar Æ LiPII + Ar•+

LiPII + Ar Æ LiP + Ar•+ + H2O

As shown in the next equation, the primary reducing substrate in the manga-nese peroxidase cycle is Mn2+, which efficiently reduces both compound I(MnPI) and compound II (MnPII), generating Mn3+, which subsequently oxi-dizes the organic substrate.

MnP + H2O2 Æ MnPI + H2O

MnPI + Mn2+ Æ MnPII + Mn3+

MnPII + Mn2+ Æ MnP + Mn3+ + H2O

MnPI + Ar Æ MnPII + Ar•+

Mn3+ + Ar Æ Mn2+ + Ar•+

Compared to most other peroxidases (Table 9), the redox potential of lignin andmanganese peroxidases are more positive. The redox potentials of the ferric/ferrous couple (Fe3+/Fe2+) is about –140 mV for lignin peroxidase and –90 mVfor manganese peroxidases, values that are considerably higher than the values

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Table 9. Redox potential of peroxidases

Peroxidases: isoenzymes Em7 (mV) References

Horseradish –278 [139]Cytochrome c –194 [140]Lignin peroxidase: H1, H8, H2, H10 –142, –137, –135, –127 [141]Manganese peroxidase: H4, H3 –93, –88 [141]

Em7, midpoint potential at pH 7.

for horseradish peroxidase (–270 mV) or cytochrome c peroxidase (–195 mV).The higher redox potential suggests that the lignin and manganese peroxidasecompound I and II intermediates are more electron-deficient and hence havehigher oxidation-reduction potential. The higher oxidation-reduction potentialof the active intermediates of lignin and manganese peroxidase extent thenumber of chemicals that can be oxidized to those of higher redox potential.

In some white-rot fungi, laccases (low-specificity enzymes which act on o-and p-quinols) are also present. But the role of laccase in ligninolysis by ligni-nolytic fungi is not clear, since lignin can be rapidly degraded by Phanerochatechrysosporium, an organism which misses laccase. However, this conclusioncannot be extended to eliminate the involvement of laccase in ligninolysis inthose fungi that do secrete this enzyme. It is known that laccases oxidize non-phenolic aromatic compounds as well as Mn2+ in the presence of other oxidiz-able substrates [142]. Substrate oxidation by laccase is a one-electron reactiongenerating a free radical [143]. The initial product, the carbon-centered cationradical formed by removing one electron from an aromatic nucleus [144], istypically unstable and may undergo a second enzyme-catalyzed oxidation (con-verting phenol to quinone with many substrates). In addition, the radical mayundergo non-enzymic reactions such as hydration or disproportionation.Although laccase can (directly or indirectly) cleave a significant proportion ofsubstrates found in lignin, the role of laccase in ligninolysis remains unresol-ved, but the widespread occurrence of this enzyme in wood-rotting fungi is un-likely to be coincidental.

2.4.2.2Ligninolytic Systems and PAHs

The polyaromatic structure of both lignin and polycyclic aromatic hydrocar-bons (PAHs) prompted the suggestion that the same fungi might be able to de-grade these ubiquitous pollutants [145].

White-rot fungi are able to degrade PAHs and in some cases to mineralizethem. Most of the work was done with Phanerochaete chrysosporium, demon-strating its ability to degrade non-specifically a wide range of aromatic pol-lutants [146–161]. P. chrysosporium can metabolize a variety of PAHs, includingbenzo[a]pyrene, under ligninolytic (N-limited conditions) as well as non-ligni-nolytic conditions (non-N-limited) [151, 158], i.e., in the presence and absenceof peroxidases. Other white-rot fungi such as Trametes versicolor, Bjerkanderasp., and Pleurotus ostreatus are thought to be more promising than P. chrysos-porium in their ability to mineralize PAHs to CO2 [157, 162, 163]. In addition,Crinipellis stipitaria has been reported to metabolize pyrene [164, 165]. Sackand Günther [157] showed that P. ostreatus is quite efficient in the degradationof phenanthrene and fluorene, less efficient with fluoranthene, while pyrenewas not degraded to any significant extent. Vyas et al. [163] showed that P.ostreatus is able to degrade anthracene. In general, notable differences havebeen demonstrated among Phanerochate chrysosporium, other Phanerochaetespecies, and members of other genera with regard to the extent of PAH miner-alization and transformation ability.


2.4.2.3Influence of Ionization Potential on Oxidation of PAHs

Lignin peroxidase from Phanerochaete chrysosporium directly catalyzes one-electron oxidations of aromatic substrates [166, 167]. The resultant aryl cationradicals then undergo spontaneous rearrangements and degradation. Hammelet al. [154] observed transformation by lignin peroxidase of those PAHs withionization potential (IP) values of < 7.55 eV (see values of ionization potentialof PAHs in Table 8). Isolated lignin peroxidase was unable to degrade com-pounds with an IP above the threshold value of 7.55 eV. Therefore, the meta-bolism of PAHs with high IPs (>7.65 eV) such as triphenylene, phenanthrene,fluoranthene, chrysene, benzo[b]fluoranthene, and benzo[e]pyrene observedwith whole cell cultures could not be explained by the direct action of the ligninperoxidase. Moen and Hammel [155] reported data that support a role of man-ganese-dependent peroxidase-mediated lipid peroxidation in phenanthreneoxidation by P. chrysosporium. Bogan and Lamar [168] gave evidence that thedegradation of three- to six-ring PAHs with IPs between 7.2 eV and 8.1 eV is IP-dependent during in vivo and in vitro lipid peroxidation. This implies that theparticipation of a one-electron oxidant stronger than lignin peroxidase or Mn3+

is involved. The data presented showed that compounds with up to six ringswere degraded in vitro during manganese peroxidase-dependent lipid peroxi-dation reactions.

Hammel et al. [152] emphasized that PAHs which are lignin peroxidase sub-strates are more susceptible to mineralization than PAHs which are not, so thatbenzo[a]pyrene, pyrene, and anthracene are all rapidly depleted from N-limit-ed cultures.

2.4.2.4Metabolites Formed During Degradation

The degradative pathways for anthracene and phenanthrene by Phanerochatechrysosporium are shown in Figs. 25 and 26. The formation of a quinone to pre-pare the aromatic ring for cleavage is an unusual biodegradative strategy, and itappears to be of general importance in P. chrysosporium. While the formationsof 9,10-anthraquinone and phthalate were found to be rapid processes, thefurther conversion to carbon dioxide is slow. The 2,2¢-dicarboxylic biphenyl wasfound to be the major product in the degradation of phenanthrene.

Yadav and Reddy [169] presented data indicating that P. chrysosporiummineralizes all BTEX components (benzene, toluene, ethylbenzene, xylenes)

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Fig. 25. Proposed pathways for anthracene degradation in ligninolytic P. chrysosporium [153]

either individually or as a mixture to CO2 under non-ligninolytic conditions,i.e., when no lignin and manganese peroxidases are produced.

P. ostreatus differs from P. chrysosporium in its lignin degradation mecha-nism in that it does not involve lignin peroxidase activity [126]. Instead, itslignin degradation ability is assumed to be due to laccase activity [127, 128, 131,170]. Laccase was found to be nonspecific as to its reducing substrate as well asable to oxidize a variety of substrates including polyphenols, methoxy-substi-tuted phenols, diamines, and a range of other compounds. Bezalel et al. [171,172] reported that Pleurotus ostreatus is able to mineralize various PAHs to14CO2 such as phenanthrene, pyrene (no degradation in [157]), benzo[a]pyrene,anthracene, and fluorene, but fails to mineralize fluoranthene.

2.5Résumé: Aerobic Degradation of Aromatic Compounds

Bacteria and fungi are dealing with aromatic hydrocarbons in a different way.While bacteria are able to utilize the compounds as the sole source of carbonand energy fungi cometabolize the aromatics to hydroxylated products, some-times mineralization takes place by the fungi. In bacteria degradation of anaromatic compound is initiated by dioxygenase to give a 1,2-diphenol as thering cleavage substrate, while cytochrome P-450 catalyzed epoxide formation is the first step in the degradation by the fungi. The further degradation of thediphenol in the bacteria leading to intermediates of the metabolism has beendescribed to occur via meta, ortho or gentisate pathways.

Since the polycyclic aromatic hydrocarbons are ubiquitous pollutants thebiodegradation is currently of increasing interest. As the molecular weight ofPAHs increases, their water solubility decreases. The increasing hydrophobicityusually correlates with decreasing biodegradability and with increasing poten-tial for bioaccumulation. Because of the difficulty of isolating bacteria whichcan effectively degrade high-molecular-weight PAHs with four or more fusedaromatic rings, the ability of white-rot fungi to degrade the abundant naturally-occurring polymer lignin has made these fungi appropriate candidates for PAHdegradation. Since white-rot fungi produce extracellular enzymes, the PAH bio-availability is maximized and toxicity problems for the fungi are avoided.Considering the fact that the rate of PAH degradation often does not correlate


Fig. 26. Proposed pathway for phenanthrene degradation in ligninolytic P. chrysosporium[152]. Major reactions Æ and minor ones are shown

with lignin peroxidase activity, several other enzymatic mechanisms arethought to be used by the white-rot fungi to degrade PAH.

2.6Anaerobic Degradation of Aromatic Compounds

Contrary to the well-known pathways of the aerobic degradation of aromaticsthe mesomeric stabilized aromatic ring will be attacked under anaerobic con-ditions by reductive rather than oxidative reactions. Since the concept of thedestabilization of the aromatic nucleus in the absence of oxygen has been devel-oped by Evans [173], several aromatic compounds were examined and found tobe degraded by various types of anaerobic bacteria according to that principle.

As far as is known, the pathways of substrate activation and cleavage are ba-sically the same, regardless of whether nitrate, ferric ion, sulfate, or carbonateare used as electron acceptor, or whether light can provide an additional energysource. In addition, degradation of aromatic compounds is possible under fer-mentative conditions. Most information on the degradative pathways used foraromatic compounds has been obtained with denitrifying organisms. Differentaromatics are converted through channeling pathways into a few central reac-tive intermediates: benzoyl-CoA, resorcinol, phloroglucinol, and possiblyothers. After these activation reactions, the central compounds are reduced toform an intermediate with a 1,3-dioxo structure which is no longer an aroma-tic compound. This structure allows a nucleophilic attack on a ring carbonylgroup and subsequent ring fission. A b-oxidation pathway that produces threeacetyl moieties follows the ring cleavage. Acetyl-CoA may then further be oxi-dized to CO2.

2.6.1Channeling Reactions

2.6.1.1Channeling Reactions to Benzoyl-CoA

Benzoyl-CoA is formed from a large variety of different compounds, such asphenol, 2-hydroxybenzoate, 4-hydroxybenzoate, p-cresol, phenylacetate, 4-hy-droxyphenylacetate, mandelate, hydroxymandelate, toluene, 2-aminobenzoate,4-aminobenzoate, aniline, and many others. The pathways are summarized inFig. 27 indicating that some analogous reactions take place with these quite dif-ferent compounds.

Hydroxybenzoates. The reductive dehydroxylation of aromatic hydroxyl func-tions, notably with compounds having hydroxyl functions para to a carboxylgroup, is an important reaction in the metabolism of phenol, p-cresol, and 4-hy-droxyphenylacetate. The case that has been studied in detail is the reductive de-hydroxylation of 4-hydroxybenzoyl-CoA by 4-hydroxybenzoyl-CoA reductase(dehydroxylating), indicating the requirement of coenzyme A thioester forma-tion of the aromatic acids prior to dehydroxylation [177, 182]. The metabolism

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Fig. 27. Channeling sequences for various aromatic compounds leading to benzoyl-CoA asthe central metabolite. Analogous reactions such as carboxylation, formation of CoA ester,and reductive elimination of substituents are marked by different gray boxes. The informa-tion is compiled from [174–194]

of salicylate proceeds in a similar reaction sequence via 2-hydroxybenzoyl-CoAto give benzoyl-CoA [180].

Phenol. Phenol lacks a carboxylic group and therefore cannot be activated bycoenzyme A, a prerequisite which turned out to be essential for complete ringreduction in the benzoate pathway. A phenol-degrading nitrate reducer was de-pendent on CO2 as cosubstrate for phenol oxidation. Carboxylation of phenol to4-hydroxybenzoate, analogous to a Kolbe-Schmitt reaction [195], was suggestedto be the initial reaction in phenol degradation, followed by coenzyme A ac-tivation and reductive elimination of the hydroxy substituent to form benzoyl-CoA [185]. There are indications that phenylphosphate is the physiological in-termediate used by the para-specific carboxylase [190]. The use of phenylphos-phonate instead of phenol renders the carboxylation reaction exergonic undernatural CO2 and phenol concentrations. In addition, phosphorylation would fa-cilitate the cellular accumulation of this toxic substrate in a non-toxic reactiveform.

Carboxylation of phenol to a benzoate derivative was also demonstrated inanaerobic enrichment cultures converting fluorophenols to benzoate [183, 184].These experiments also proved that phenol is carboxylated at the C-4 carbon,leading to 4-hydroxybenzoate as an intermediate. Although a net carboxylationof phenol to 4-hydroxybenzoate has never been demonstrated in a cell-free ex-tract, this concept acquires more and more support from experiments with sul-fate-reducing bacteria (Schnell and Schink, unpublished) and defined metha-nogenic cultures [189].

Catechol and hydroquinone. Studies on the degradation of hydroquinone by astrictly anaerobic fermenting bacterium indicate that the primary step in hydro-quinone degradation is a carboxylation to a gentisic acid derivative, followed ap-parently by subsequent reductive dehydroxylations to benzoyl-CoA [196, 197].

The degradation of catechol by a Desulfobacterium sp. proceeds via carboxy-lation to protocatechuate followed by esterification with CoA and reductive de-hydroxylation to give 3-hydroxybenzoyl-CoA [198].

Aniline and anthranilate. Aniline is degraded in anoxic environments only veryslowly. The degradation pathway has so far only been studied with a pure cul-ture of a sulfate-reducing bacterium [194]. Aniline degradation by this bacter-ium depends on CO2 as cosubstrate, and thus is reminiscent of phenol degra-dation by nitrate reducers. Aniline is first carboxylated to 4-aminobenzoate, ac-tivated to the coenzyme A derivative, and then reductively deaminated tobenzoyl-CoA. The conversion of 2-aminobenzoyl-CoA to benzoyl-CoA by 2-aminobenzoyl-CoA reductase is an additional example of a reductive deamina-tion [191].

Cresols. p-Cresol is converted by Pseudomonas strains to 4-hydroxybenzylal-cohol by an oxygen-independent reaction [199]. The alcohol oxygen is derivedfrom water which is added to a quinone methide intermediate formed by a de-hydrogenation reaction [186]. The hydroxybenzyl alcohol can easily be oxidized

42 W. Reineke

to 4-hydroxybenzoate and further degraded as mentioned above. The samepathway appears to underlie sulfate-dependent p-cresol degradation [200, 201].

It should be emphasized that the methyl hydroxylation requires an electron-attracting group in the 4- or 2-position to stabilize the quinone methide inter-mediate. Therefore, this reaction is also applicable to o-cresol, but not to m-cre-sol degradation. In methanogenic and sulfidogenic consortia m-cresol appearsto be activated by carboxylation to 2-methyl-4-hydroxybenzoate, analogous tophenol activation [202, 203]. The further degradation of 4-hydroxy-2-methyl-benzoate proceeds via 4-hydroxybenzoate and benzoate demonstrated withanother m-cresol-degrading methanogenic consortium [204]. The carboxyla-tion strategy seems to be used also for o-cresol by a consortium under metha-nogenic conditions [205].

The anaerobic degradation of m-cresol was later studied with a denitrifyingbacterium [181]. The transient accumulation of 3-hydroxybenzoate and benzo-ate when using inhibitory compounds supports conversion of m-cresol via ini-tial anaerobic methyl oxidation to 3-hydroxybenzoate, followed by reductivedehydroxylation to benzoate or benzoyl-CoA. The same oxidative pathway se-quence was observed with a sulfate-reducing organism, Desulfotomaculum sp.strain Groll [201, 206].

A Desulfobacterium cetonicum was only recently reported, which oxidized m-cresol completely with sulfate as electron acceptor [207]. 3-Hydroxybenzyl-succinate was detected as a metabolite indicating that the methyl group is ac-tivated by the addition to fumarate as in the case of anaerobic toluene metabo-lism.A further metabolite observed was 3-hydroxybenzoyl-CoA formed by CoAthioesterification and oxidation of 3-hydroxybenzylsuccinate.

Phenylacetate, phenylpropionate, mandelate. Phenylacetate is first converted to phenylacetyl-CoA by a specific CoA ligase [208]. 4-Hydroxyphenylacetateseems to be activated by a different ligase. The thioester apparently activates thea-methylene carbon enough to allow its dehydrogenation and hydroxylationwith water as the oxygen source, i.e., anaerobic a-oxidation. Phenylglyoxylateor 4-hydroxyphenylglyoxylate are the products formed by the nitrate-reducingPseudomonas strains [209] and Thauera aromatica [210]. The oxidative decar-boxylation of phenylglyoxylate to benzoyl-CoA by phenylglyoxylate: NAD+ oxi-doreductase (CoA benzoylating) [211] is not an unusual reaction, analogous tothe oxidative decarboxylation of pyruvate. Mandelate can be oxidized viaphenylglyoxylate to benzoyl-CoA [212, 213]. Phenylpropionate is easily b-oxidized to benzoate [214] and further degraded as such.

Phthalates. Degradation of phthalates requires elimination of the additionalcarboxyl group, and further degradation analogous to benzoate. However, di-rect decarboxylation is chemically quite difficult. Phthalates appear to be firstconverted to the CoA mono thioester by CoA ligases, followed by decarboxyla-tion of o-, m-, or p-phthaloyl-CoA to benzoyl-CoA [215–217].

Toluene and xylenes. Monoaromatic hydrocarbons such as toluene and xylenesare known to be degraded in the absence of molecular oxygen, i.e., under


nitrate-reducing [174, 218–227], Fe3+-reducing [228–230], Mn4+-reducing [231,232], sulfate-reducing [222, 233–239], and/or methanogenic conditions[240–243]. Just recently, an anoxygenic phototrophic bacterium, Blastochlorissulfoviridis strain ToP1, was isolated, which is able to use toluene for growth[244].

Well-characterized bacteria capable of degrading toluene under anaerobicconditions include several denitrifying species such as Thauera aromatica andAzoarcus tolulyticus [174, 219, 220, 223, 226, 238], the ferric-ion-reducingGeobacter metallireducens [230], and the sulfate-reducing species Desulfobaculatoluolica and strain PRTOL1 [234, 237].

While toluene was found to be the most readily degradable aromatic com-pound under anaerobic conditions [245], a plausible pathway has been obtainedonly recently. Diverse compounds have been detected accumulating in differentorganisms during the toluene metabolism which hardly fit into one pathwayscheme.

There was strong evidence that toluene oxidation in denitrifying bacteria oc-curs via benzoate as a common metabolite [174, 178, 226, 230, 233, 246–249].Seyfried et al. [249] reported the transient accumulation of benzaldehyde andbenzoate after toluene was added, while 3-methylbenzaldehyde and 3-methyl-benzoate were the products formed after the addition of m-xylene. Benzyl al-cohol and benzaldehyde have been detected in the anaerobic metabolism oftoluene in the denitrifying Thauera aromatica strain K172 (Pseudomonas) [174,250]. In contrast, p-cresol accumulated in a mixed methanogenic culture grownwith toluene [241, 242]. Carboxylation of the methyl carbon of toluene as sug-gested by Altenschmidt and Fuchs [174] might explain the accumulation ofphenylacetate.

Benzylsuccinate and benzylfumarate have been reported to accumulate dur-ing anaerobic degradation of toluene under denitrifying conditions by strainT1 [246], Pseudomonas sp. strain T [249], Thauera aromatica K172 [249, 251],and Azoarcus tolulyticus Tol-4 [252], as well as under sulfate-reducing condi-tions by strain PRTOL1 [234, 253]. Phenylitaconate was identified by Migaud etal. [254] during growth of Azoarcus tolulyticus Tol-4 with toluene. Other strainsof Azoarcus tolulyticus with the ability to degrade toluene anaerobically [223,255] also synthesized similar amounts of phenylitaconate during toluene meta-bolism. These data support the assumption that an alternative pathway mayfunction in these strains. In analogy to the attack of the methyl group by suc-cinyl-CoA forming benzylsuccinate and benzylfumarate, activation of tolueneby an oxidative condensation of toluene with acetyl-CoA to yield phenylpro-pionyl-CoA is proposed by Evans et al. [246], followed by conversion to benzoyl-CoA via b-oxidation.

The pathway for the initial attack on toluene has recently been elucidated incell-extracts of Thauera aromatica strain K172 [179, 256] and Azoarcus sp.strain T [257, 258]. Toluene is condensed with fumarate by benzylsuccinatesynthase to give benzylsuccinate as the first intermediate. There is strong evi-dence that benzylsuccinate formation is accomplished in nitrate-reducing bac-teria via formation of an enzyme-bound radical [256–259]. CoA-dependentconversion of benzylsuccinate to phenylitaconate or the CoA thioester [257]

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and subsequently to benzoyl-CoA follows [179, 257]. Present knowledge, shownin Fig. 28, includes activation and b-oxidation.

While anaerobic growth of pure cultures on m-xylene has often been shownwith denitrifying bacteria [219, 223, 238, 249, 261], strains that grows on o- orp-xylene have been isolated only recently [262].

Other monoaromatic hydrocarbons. Anaerobic degradation of ethylbenzene,propylbenzene, and p-cymene has been studied. Anaerobic mineralization ofethylbenzene has been reported in three denitrifying bacteria [176, 238]. Allthree isolates are closely related to each other, and are affiliated with the genusAzoarcus. A pathway for anaerobic oxidation of ethylbenzene to benzoyl-CoAhas been proposed. Benzoate was detected as a transient intermediate [176].Formation of 1-phenylethanol and acetophenone from ethylbenzene was de-monstrated [176, 193]. The proposed initial reaction of the pathway is the oxida-tion to 1-phenylethanol. The oxygen atom of the hydroxyl group is derived fromwater. 1-Phenylethanol is further oxidized to acetophenone. Both enzymes re-sponsible for the formation of these intermediates have been demonstrated [263].

Only minor evidence is available for the further reaction involved in aceto-phenone conversion to benzoyl-CoA. It is proposed that acetophenone is car-boxylated to benzoylacetate in a reaction analogous to reactions found in ae-robic and anaerobic degradation of aliphatic ketones [264, 265]. Benzoylacetateis proposed to be activated to the CoA thioester and to be cleaved thiolyticallyto acetyl-CoA and benzoyl-CoA.

Information on the pathway used for the degradation of p-cymene is rare.Harms et al. [266] observed the accumulation of p-isopropylbenzoate duringgrowth on p-cymene, indicating an initial attack on the methyl group.


Fig. 28. Anaerobic toluene pathway in denitrifying organisms according to Leutwein andHeider [260]

2.6.1.2Channeling Reactions to Resorcinol (1,3-Dihydroxybenzene) and Phloroglucinol (1,3,5-Trihydroxybenzene)

Dihydroxy- and trihydroxybenzoates. Aromatic acids with two or more hy-droxyl functions may become decarboxylated if the resulting product containsa diol system in the 1,3-position. These compounds have little aromaticity andcan easily be reduced. Examples are the decarboxylation of resorcylates (2,4-dihydroxy-, and 2,6-dihydroxybenzoates) to resorcinol [267] (Fig. 29).

Similarly, gallate is decarboxylated to pyrogallol (1,2,3-trihydroxybenzene)followed by a transhydroxylation to give phloroglucinol (Fig. 30). Phloro-glucinol results directly from phloroglucinate by decarboxylation. In general,the decarboxylation of aromatic acids with a hydroxyl function para to the car-boxyl group is a chemically favored reaction.

Trihydroxybenzenes. Anaerobically fermenting bacteria such as Eubacteriumoxidoreducens, Pelobacter acidigallici, Pelobacter massiliensis, and the homo-acetogenic Holophaga foetida degrade trihydroxybenzenes via phloroglucinol[268, 270–273].

Pyrogallol is converted to phloroglucinol [269] by an unusual reaction whichincludes 1,2,3,5-tetrahydroxybenzene as cosubstrate [268]. The hydroxyl groupis transferred from the tetrahydroxybenzene to pyrogallol, thus yielding phlo-roglucinol and a new tetrahydroxybenzene molecule [274, 275]. Since the co-substrate is cyclically regenerated in the course of the reaction, it has to be con-sidered as a cocatalyst. Its function is to donate one hydroxyl group to C-5 ofpyrogallol.

Hydroxyhydroquinone (1,2,4-trihydroxybenzene) is degraded by the fer-menting organisms through phloroglucinol using a different hydroxyl transferreaction [276]. Three hydroxyl transfers seem to be involved (Fig. 31). First, thesubstrate is disproportionated to 1,3-dihydroxy- and 1,2,4,5-tetrahydroxyben-zene. Then the tetrahydroxybenzene is isomerized to the 1,2,3,5-tetrahydroxy

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Fig. 29. Decarboxylation of dihydroxybenzoates leading to resorcinol [11, 267]


Fig. 30. Degradation of trihydroxybenzoates to phloroglucinol involving decarboxylationand transhydroxylation [268, 269]. The hydroxyl group moving within the sequence is mark-ed. Gallate: top sequence; phloroglucinate: bottom

Fig. 31. Sequential transhydroxylation of hydroxyhydroquinone to phloroglucinol in Pelo-bacter massiliensis according to Brune et al. [276]. The hydroxyl group moving within the se-quence is marked

isomer by an unknown reaction. Finally, 1,2,3,5-tetrahydroxybenzene formsphloroglucinol by transferring its 2-hydroxyl group to either hydroxyhydroqui-none or resorcinol, thus also regenerating the cosubstrate involved in earlier re-actions of the sequence [276].

Besides the strategy of isomerization of 1,2,4-trihydroxybenzene to phloro-glucinol and further degradation through the phloroglucinol pathway (see la-ter), which is used by all fermenting bacteria, recently alternative pathways for1,2,4-trihydroxybenzene have been observed with nitrate- and sulfate-reducingbacteria [277–280].

2.6.2Activating Reductive Sequences and Ring Cleavage

2.6.2.1Benzoate Pathway

Utilization of benzoate by some species of nonsulfur purple bacteria has beenknown for many years [281–285], but is restricted to a few strains of Rhodo-pseudomonas palustris [282, 286, 287], Rhodocyclus sp. [288], Rhodospirillum sp.[289], and Rhodomicrobium sp. [290]. A characteristic feature of the phototro-phic metabolism of these bacteria is that the growth substrate is usually exten-sively assimilated into cell material. This follows from their use of light asenergy source, so eliminating the need for oxidative or fermentative manipula-tion of a portion of the carbon source.

In addition, various denitrifying Pseudomonas, Alcaligenes, and Moraxellaspecies are able to use benzoate in the absence of oxygen [175, 291–294].

It is clear that the reduction of the aromatic ring is preceded by coenzyme Athioesterification of benzoate, first observed by Hutber and Ribbons [187] withcrude extract of Rhodopseudomonas palustris, enabling the cells to accumulatean otherwise permeant molecule [286]. Benzoate is activated by a benzoyl-CoAsynthetase reaction. ATP is cleaved into AMP and pyrophosphate which sug-gests the occurrence of an intermediate acyl-AMP. Pyrophosphate is thought tobecome subsequently hydrolyzed. This renders the overall reaction stronglyexergonic.

The further pathway has become clearer by studies with enzymes ofRhodopseudomonas palustris [295] and of Thauera aromatica strain K172[296–298] (Fig. 32). Buckel and Keese [299] proposed a possible mechanism forthe benzoyl-CoA reductase (for a further discussion see Buckel and Golding[300]). The reaction proceeds in two successive one-electron reactions to givecyclohex-1,5-diene-1-carboxyl-CoA, analogous to a chemical Birch reduction[301]. During the first circle of this biological Birch reduction a highly reactiveketyl radical may be generated, having its origin in the transfer of a superre-ductive activated electron to the thioester carbonyl group of benzoyl-CoA.Uptake of a proton in para-position would then neutralize the charge makingthe radical able to accept the second electron followed by the addition of the se-cond proton. The CoA thioester may play an important role in this catalytic pro-cess. Facilitation of binding of the substrate and correct positioning in the ac-

48 W. Reineke

tive center of the enzyme might be one function. In addition, the thioesterifiedcarboxyl group is an important substituent mechanistically as the entrancepoint of the electrons and thermodynamically by lowering the midpoint poten-tial of the electron transfer step and of the whole process. However, althoughthe reduction of benzoyl-CoA is facilitated by the CoA thioester, the reductaserequires input energy of two ATP molecules to overcome the considerable activation energy, one ATP for each electron introduced [296, 297]. The nitrate-reducing bacteria can recover the high energy input for substrate-activationand for dearomatization through the further breakdown of the cleavage prod-uct, which is totally oxidized to CO2. Since fermenting bacteria can recover only little energy in the further breakdown they apply a different reaction forbenzoyl-CoA dearomatization to have the reaction exergonic so that no ATP isnecessary. They introduce four electrons and protons from NAD(P)H into thering structure which directly leads to a cyclohexene derivative [302].


Fig. 32. Reaction sequences of benzoyl-CoA degradation according to [295–298, 302, 303,305, 306]. Path (a) Rhodopseudomonas palustris and Syntrophus gentianae. Path (b) Thaueraaromatica

Therefore, different pathway branches were found in the denitrifying orga-nism Thauera aromatica, the phototrophic organism Rhodopseudomonas pa-lustris, and a fermenting organism. In Thauera aromatica the next intermediatefrom the diene is 6-hydroxycyclohex-1-ene-1-carboxyl-CoA which is derived byaddition of water. Then there is a gap between 6-hydroxycyclohex-1-ene-1-car-boxyl-CoA and 3-hydroxypimelyl-CoA, the first non-cyclic intermediate isolat-ed. The easiest explanation is the addition of water to the double bond in 6-hy-droxycyclohex-1-ene-1-carboxyl-CoA, the oxidation of the resulting alcohol togive an oxo group, and the hydrolytic cleavage of the ring. In Rhodo-pseudomonas palustris the cyclic diene is further reduced to cyclohex-1-ene-1-carboxyl-CoA. Subsequent b-oxidation results in the formation of a cyclic b-oxo compound, followed by hydrolytic carbon ring opening yielding pimelyl-CoA, which is subsequently oxidized via 3-hydroxypimelyl-CoA as in Thaueraaromatica [303]. Recently, the pathway branch via cyclohex-1-ene-1-carboxyl-CoA used by the phototrophic organism has also been demonstrated inSyntrophus gentianae when fermenting benzoate [302].

The further b-oxidation of 3-hydroxypimelyl-CoA yields glutaryl-CoA plusacetyl-CoA. The oxidation of glutaryl-CoA to 2 acetyl-CoA plus CO2 proceedsvia glutaconyl-CoA and crotonyl-CoA and is catalyzed by a glutaryl-CoA dehy-drogenase, which is present at a significant level [304].

2.6.2.2Resorcinol Pathway

In the resorcinol molecule, the two meta-oriented hydroxy substituents polarizethe p-electron cloud in such a way that selective reduction by two electrons todihydroresorcinol becomes possible, thus abolishing the molecule’s aromaticcharacter. Resorcinol-degrading fermenting bacteria (Clostridium sp.) followthis degradation strategy and 1,3-dioxocyclohexane is formed as the ultimatealicyclic compound (Fig. 33). The Clostridium also channels the carboxylatedderivatives 2,4-dihydroxybenzoate and 2,6-dihydroxybenzoate (resorcylicacids) through initial decarboxylations into this pathway [11, 267]. Obviously,the C-3 atom of 1,3-dioxocyclohexane carries sufficient positive charge to allowhydrolytic cleavage to 5-oxocaproic acid.

An obligate nitrate-reducing resorcinol degrader employs a different path ofresorcinol degradation which does not involve an initial ring reduction.

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Fig. 33. Proposed degradative sequences for resorcinol (top) and phloroglucinol (bottom)

Resorcinol is hydrolytically transformed in a one-step reaction to a non-cyclicproduct, 5-oxo-hex-2-enecarboxylic acid [307].

2.6.2.3Phloroglucinol Pathway

The three hydroxy substituents of phloroglucinol polarize the p-electron cloudeven more than the two hydroxy substituents of resorcinol. As a consequence,the trioxo tautomer prevails in aqueous solution. Consequently, phloroglucinolhas no aromatic character and can easily become reduced chemically by mildreducing agents.

Anaerobic phloroglucinol-degrading bacteria, such as Rhodopseudomonasgelatinosa [285], Coprococcus sp. [308], Pelobacter acidigallici [309], orEubacterium oxidoreducens [270], first reduce phloroglucinol to dihydrophlo-roglucinol (1,3-dioxo-5-hydroxycyclohexane) in an NADPH-dependent reac-tion. The corresponding enzyme of E. oxidoreducens has been purified and cha-racterized [310].

Nucleophilic attack on one of the carbonyl groups of dihydrophloroglucinolopens the ring to form a 3-hydroxy-5-oxocaproic acid. The further degradationof the partially oxidized caproic acid residue no longer poses basic biochemicalproblems. Details have been studied with E. oxidoreducens and P. acidigallici[270, 311, 312].

2.6.3Anaerobic Degradation of Environmentally Important Aromatics where Pathway Information is Missing or Minor

Benzene. Benzene persists in most anoxic environments [313, 314]. However,partial mineralization of benzene to carbon dioxide and methane in the ab-sence of molecular oxygen has been observed in enrichment cultures [241] andmethanogenic river sediments [315].

Benzene was completely mineralized to carbon dioxide in enrichment cul-tures in which sulfate was provided as a potential electron acceptor [235].Lovley et al. [316] observed that after an adaptation period benzene was rapidlyoxidized to carbon dioxide with the reduction of sulfate in petroleum-contami-nated sediments from San Diego Bay, California. Recently, enrichment culturesfrom marine sediments were found to be able to mineralize benzene whileusing sulfate as the terminal electron acceptor [317]. However, cultures fromriver marsh failed to show the activity. Weiner and Lovley [318] showed thatsupplementing aquifer sediments with benzene-oxidizing sulfate reducers cangreatly accelerate anaerobic benzene degradation.

Major et al. [319] reported that the degradation of benzene occurred underdenitrification conditions with material from an aquifer. Benzene degradationlinked to nitrate reduction has been found recently with enrichment culturesdeveloped from soil and groundwater microcosms [320].

Anaerobic oxidation of benzene coupled to Fe3+ as an electron acceptor hasbeen documented by Lovley et al. [321]. Stoichiometric studies demonstrated


that the electrons derived from benzene oxidation to carbon dioxide weretransferred to Fe3+, when the availability of Fe3+ had been artificially modifiedwith a chelator such as nitrilotriacetic acid (NTA). Fe3+ chelated to compoundssuch as ethylenediaminetetraacetic acid (EDTA), N-methyliminodiacetic acid,ethanol diglycine, humic acids and phosphates stimulated benzene oxidationcoupled to Fe3+ reduction in anaerobic sediments from a petroleum-contami-nated aquifer as effectively as or more effectively than when being chelated toNTA [322].

A study by Kazumi et al. [323] showed that benzene degradation takes placeunder methanogenic conditions.

Information on the degradative pathway for benzene is not available. Studiesperformed with H2

18O revealed that, in enrichment cultures, the oxygen fromwater was incorporated into benzene as a hydroxyl group with the formation ofphenol [242], but these results could not be confirmed later. No pure cultures ofbenzene-degrading bacteria have been isolated to date, which, however, are im-portant for gaining a detailed understanding of the biochemistry of anaerobicbenzene degradation.

Polycyclic aromatic hydrocarbons (PAHs). Various studies [324–332] have in-dicated that PAHs are not degraded in the absence of oxygen [64]. Other studieshave suggested that some PAHs can be degraded in the absence of oxygen ifnitrate is available as an electron acceptor, but that PAHs persist under sulfate-reducing or methanogenic conditions [333–336]. Recently, Coates et al. [337]reported that (14C) naphthalene and phenanthrene were oxidized to 14CO2 with-out a detectable lag period under strictly anaerobic conditions in sedimentsfrom San Diego Bay, California, which were heavily contaminated with PAHs,but not in less contaminated sediments. When molybdate, a specific inhibitor ofsulfate reduction [338], was added to the sediments the production of 14CO2from naphthalene and phenanthrene was immediately inhibited. First informa-tion on the metabolites of PAH degradation under anaerobic conditions wereobtained by Bedessem et al. [339] and Zhang and Young [340]. Naphthalenolwas tentatively identified as a potential metabolic intermediate of naphthalenedegradation from aquifer enrichments under sulfate-reducing conditions [339].Zhang and Young [340] gave evidence using sulfidogenic consortia that car-boxylation is an initial key reaction for the anaerobic metabolism of naphtha-lene and phenanthrene forming 2-naphthoate and phenanthrenoate, respec-tively. A pure culture of a naphthalene-degrading sulfate-reducing bacteriumhas recently been described which now allow biochemical investigations in an-aerobic degradation of PAHs [341].

2.6.4Résumé: Anaerobic Degradation of Aromatic Compounds

It appears as a general pattern that different aromatic compounds are conver-ted to give one of three central aromatic intermediates: benzoyl-CoA, resorci-nol, and phloroglucinol. Subsequently, the aromatic nucleus is destabilized byreduction in all three pathways. Cleavage of the ring is possible by a nucleophi-

52 W. Reineke

lic, probably hydrolytic, attack when the 1,3-dioxo structure is reached. The 1,3-dioxo structure exists in the cyclic, non-aromatic ring when resorcinol or phlo-roglucinol are the metabolites. In contrast, when benzoyl-CoA is the central me-tabolite after the channeling reactions, one oxo group is in the cyclic ring andthe other exocyclic as part of the CoA-ester group. In all cases the ring is brokendown to three acetyl residues which are either excreted as acetate or further oxi-dized to CO2.

Whereas those compounds which enter directly one of the basic pathwaysmentioned (benzoate, resorcinol and resorcylic acids, trihydroxybenzenes) aredegraded relatively quickly, other compounds which depend on endergonicactivation reactions (e.g., phenol, m-cresol, catechol, hydroquinone, aniline,phthalates) or dehydrogenations at a comparably high redox potential (p-cre-sol, toluene) are decomposed more slowly, indicating that the modification re-actions limit the transformation rates. Enrichment for anaerobic utilizers ofthese substrates takes, in general, much longer than with the previously men-tioned substrates (benzoate, resorcinol and resorcylic acids, trihydroxyben-zenes). It is obvious that in the case of p-cresol and toluene the type of alterna-tive electron acceptor available will also influence the degradation kinetics: anitrate reducer can derive more energy from the oxidation of an aromatic com-pound than a sulfate reducer or a methanogenic association.

Summarizing, it can be concluded that all mononuclear aromatics can be de-graded anaerobically if they carry at least one carboxy, hydroxy, amino, or me-thyl substituent. Nonetheless, the degradation kinetics differ considerably:whereas fermenting bacteria degrading resorcinol have doubling times of 6–8 h,catechol or hydroquinone degrading anaerobes have doubling times of severaldays. These differences in degradation efficiency of isomeric substrates can tosome extent be explained by the basically different pathways outlined above.

It should be emphasized that non-substituted aromatic compounds such asbenzene and naphthalene are also subjects of anaerobic degradation, althoughno or minor information on the pathway used is presently available.

2.7Résumé: Aromatic Compounds

The section has attempted to summarize what is known about the metabolismof aromatic compounds by bacteria and fungi. The results presented clearlyshow that some general features have emerged which suggest that it may bepossible to predict the types of reactions that will occur with different sub-strates and different microorganisms. The availability of molecular oxygen isthe important factor which clearly determines which strategy is in use by themicroorganisms to degrade an inert aromatic compound.

2.7.1Degradation in the Presence of Oxygen

In all cases that have been examined, bacteria initiate the oxidation of unsub-stituted aromatic compounds by incorporation of molecular oxygen into the


aromatic nucleus to form cis-dihydrodiols. These are further oxidized to di-phenolic intermediates such as catechols. The dihydroxylated intermediates undergo intra- or extradiol ring cleavage by dioxygenases to form aliphatic car-boxylic acids which are further degraded to intermediates of central metabolicpathways.

In contrast, fungi oxidize unsubstituted aromatic hydrocarbons by the inser-tion of one atom of molecular oxygen into the aromatic ring by P-450-catalyzedmonooxygenase reaction. The reactive arene oxides can isomerize to phenols orcan undergo enzymatic hydration to yield trans-dihydrodiols. The phenols arecoupled to sulfate, glucose, or glucuronic acid.

The ligninolytic fungi such as Phanerochate chrysosporium can use a batteryof extracellular enzymes, cosubstrates, and molecular oxygen to degrade aro-matics. Lignin peroxidases, manganese-dependent peroxidases, and laccases,which are one-electron oxidants, produce aromatic cation radical intermediatesthat undergo spontaneous fission reactions and formation of highly reactivequinones. H2O2 needed for the peroxidase reaction is produced by an extracel-lular oxidase which oxidizes glyoxal or glucose and reduces O2.

It is important to underline the different function of the degradation of aro-matic compounds by bacteria and fungi. Bacteria appear to have evolved suitesof enzymes for the degradation of aromatic hydrocarbons to smaller moleculesthat can support growth. In contrast, fungi appear to have evolved a detoxifica-tion system for the cellular elimination of aromatic hydrocarbons.

2.7.2Degradation in the Absence of Oxygen

Because of the absence of the molecular oxygen for ring activation and cleav-age, the anaerobic bacteria used a completely different strategy to break downaromatic compounds. The general feature is that the aromatic ring is reducedand the alicyclic ring formed is cleaved hydrolytically. The anaerobic pathwaysmay be divided into the following general steps:1. Reactions channeling the variety of substrates into a few central interme-

diates such as benzoyl-CoA, phloroglucinol, or resorcinol and thereby pre-paring the substrates for ring reduction.

2. Ring reduction, formation of 1,3-dioxo structure, and hydrolytic cleavage.3. A type of b-oxidation to central metabolites (acetyl-CoA).

The easy laboratory handling of aerobic bacteria allows their isolation in pureculture much more readily than anaerobic bacteria. However, pure cultures areprerequisites for elucidation of degradative pathways. This clearly explains thehigher number of studies on the aerobic degradation in former times. In the lastfew years, because of the interest in application of organisms to the clean-up ofcontaminated aquifers, mostly lacking oxygen, the degradative potential of an-aerobic bacteria attracted much interest. In addition, the ability of the an-aerobes to carry out “unusual” biochemical reactions made them highly inter-esting candidates for research activities.

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3Degradation of Chloroaromatic Compounds

Microorganisms can degrade chloroaromatic compounds under aerobic andanaerobic conditions. First the degradation of chloroaromatic compounds,which are used by aerobic bacteria as the sole carbon and energy source, will bediscussed. Then we will deal with the degradation potential of some fungi,especially some ligninolytic species such as Phanerochaete chrysosporium to-wards chloroaromatics. The role of chloroaromatics as electron acceptors forgrowth of anaerobic populations is the final topic.

3.1Chloroaromatic Compounds as Growth Substrate for Aerobic Bacteria and the Dechlorination Mechanisms

The biodegradation of a chlorosubstituted arene can be considered completeonly when its carbon skeleton is converted into intermediary metabolites andits organic chlorine is returned to the mineral state. The crucial point is the re-moval of chlorine substituents from the organic compound. This may occur atan early stage of the degradative pathway prior to cleavage of the aromatic ring.Alternatively, degradation proceeds through chlorinated diphenols as centralmetabolites, and HCl is eliminated from aliphatic structures, which are generat-ed after ring cleavage, or is linked with ring cleavage.

Both dechlorination mechanisms, i.e., early and late eliminations, may takeplace with multiple chloroaromatics. If the chloroaromatic bears only one chlo-rine substituent, initial dechlorination reactions lead to the formation of di-phenolic ring cleavage substrates, which are further degraded in a way similarto normal aromatics. In contrast, the early dechlorination of higher chlorinatedaromatics leads to chlorocatechols, chloroprotocatechuates, or chlorohydroqui-nones. The degradation of chlorohydroquinones proceeds through the so-cal-led hydroquinone pathway, which includes elimination of chlorine substituentsfrom the aromatic structure. However, chlorocatechols are subject to degrada-tion through the so-called modified ortho pathway, where two late dechlorina-tions take place from non-aromatic structures. The degradation pathways ofchlorocatechols or chlorohydroquinones converge at the stage of (chloro)-ma-leylacetates. A step later 3-oxoadipate is the common metabolite formed in thedegradation pathways used for aromatics and chloroaromatics, if all chlorinesubstituents have been eliminated (see simplified overview in Fig. 34). Chloro-3-oxoadipates occur from the higher chlorinated compounds. Besides thisfunneling of pathways, some divergence is seen for chlorocatechols and chloro-protocatechuates leading into the meta pathway.

3.1.1Elimination of Chlorine Substituents Prior to Ring Cleavage

The mechanisms of the dechlorination prior to ring cleavage with hydrolytic,oxygenolytic or reductive elimination of chlorine from the aromatic ring areschematically summarized in Fig. 35.


3.1.1.1Replacement of Chlorine by a Hydroxyl

Hydrolytic dechlorinations have been observed in the degradation of 4-chloro-benzoate and some chlorophenols. The mechanism of a hydrolytic dechlorina-tion process was clarified for 4-chlorobenzoate initially by labeling experimentsusing 18O2 and H2

18O [342, 343]. The data indicated that the dechlorinationreaction utilizes water as the hydroxyl donor and not molecular oxygen. Thismechanism has been shown for the degradation of 4-chlorobenzoate byMicrococcus spp., Pseudomonas spp., Nocardia sp., Alcaligenes sp., and Arthro-bacter spp. to give 4-hydroxybenzoate [344–355].

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Fig. 34. Scheme of the convergence of degradative pathways for aromatic and chloroaromaticcompounds with (chloro)-maleylacetate and (chloro)-oxoadipate as the common intermedia-tes. The aromatic ring is a representative for any aromatic structure (e.g., benzene, phenol,benzoate) and the chlorinated ring for a chloroaromatic structure. � Aromatic compound de-graded via catechol and the 3-oxoadipate pathway. � Monochloroaromatic compound con-verted via catechol: chlorine elimination as part of the peripheral pathway. � Chloroaromaticcompound (such as pentachlorophenol) converted via (chloro)-hydroquinone. � Trichloro-and tetrachloroaromatic compounds converted via chlorocatechol. One chlorine eliminationstep as part of the peripheral pathway. � Chloroaromatic compound converted via chloro-catechol. � Chlorocatechols dechlorinated as part of the modified ortho pathway.� Chlorocatechol or chloroprotocatechuate dechlorinated as part of a meta pathway

The 4-chlorobenzoate dehalogenase system from Pseudomonas sp. strainCBS3 has been shown to be a three component enzyme complex [356, 357]. Therole of each component of the system dehalogenating 4-chlorobenzoate hasbeen clarified by cloning of the respective genes [358–360] and by detailedstudies with purified enzymes (Fig. 36).

Activation of the substrate to its coenzyme A derivative needs ATP and is car-ried out by a ligase [361–363]. The 4-chlorobenzoate:coenzyme A ligase sharessignificant sequence similarity with proteins, which catalyze similar chemistryin the b-oxidation pathway [364].

The activation reaction precedes dehalogenation, which is catalyzed by adehalogenase that has sequence similarity to crotonyl-CoA hydratase [364]. Thedata support a proposal that the ligase and dehalogenase evolved from a b-oxi-dation pathway.

The studies of the Pseudomonas sp. strain CBS3 4-chlorobenzoyl-CoA deha-logenase have shown that it utilizes a unique form of catalysis in which an ac-


Fig. 35. Schematic presentation of the early dechlorinations prior to ring cleavage. � hydro-lytic; � oxygenolytic; � reductive dechlorination

Fig. 36. Activation and hydrolytic dechlorination of 4-chlorobenzoate. The following en-zymes are involved: � 4-chlorobenzoate-CoA ligase, � 4-chlorobenzoyl-CoA dehalogenase,� 4-hydroxybenzoyl-CoA thioesterase

tive site carboxylate (aspartate functions as the active site nucleophile) bonds toC-4 of the benzoyl ring of the bound substrate to form a Meisenheimer-likecomplex. Expulsion of the chloride from the Meisenheimer complex with con-comitant rearomatization of the benzoyl ring generates an arylated enzyme asthe second reaction intermediate. Hydrolysis of the arylated enzyme occurs byaddition of a water molecule to the acyl carbonyl carbon to form a tetrahedralintermediate which expels the hydroxylbenzoyl group to generate the catalyticcarboxylate residue and form 4-hydroxybenzoyl-CoA (Fig. 37).

The last reaction step in the reaction to form 4-hydroxybenzoate is carriedout by the 4-hydroxybenzoate:coenzyme A thioesterase leading into the proto-

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Fig. 37. Proposed chemical pathway for the hydrolysis of 4-chlorobenzoyl-CoA by dehaloge-nase (according to [365–369]). The role of the amino acids functioning in the catalysis isshown. � substrate; � Meisenheimer intermediate; � arylated enzyme; � arylated enzyme;� tetrahedral intermediate; � product

catechuate pathway [357, 361, 370–373]. No sequence homology of the thio-esterase with other proteins was found [364].

Dehalogenation is restricted to halobenzoates substituted in the para-posi-tion [349, 374].

Hydrolytic eliminations from other chloroaromatics such as various chloro-phenols and the chlorohydroquinones have been postulated to occur in the so-called hydroquinone pathway (Fig. 38). A hydrolytic elimination brings aboutthe formation of 6-chlorohydroxyhydroquinone from 2,6-dichlorohydroqui-none in the degradation pathway of 2,4,6-trichlorophenol of Azotobacter sp.GP1 and Streptomyces rochei 303 [375–377] as well as 5-chlorohydroxyhydro-quinone from 2,5-dichlorohydroquinone in the degradation pathway of 2,4,5-trichlorophenol by Burkholderia cepacia AC1100 [378].

Hydrolytic elimination of chlorine was found to initiate the degradation of te-trachloro-p-hydroquinone – the metabolite in the pentachlorophenol mineraliza-tion – yielding trichloro-1,2,4-trihydroxybenzene in Rhodococcus chlorophenoli-cus PCP-1 [379] as well as Mycobacterium fortuitum CG2 [380] and Sphingomonaschlorophenolica (formerly Flavobacterium sp.) ATCC 39723 [381].

The following conclusion can be reached: hydrolytic dechlorination of anaromatic ring is difficult, since substituents are difficult to remove by nucleo-philic displacement from a p-electron rich system and, therefore, the ring mustbe activated by CoA or by the presence of hydroxyl or halogen substituents.

3.1.1.2Chlorine-Carbon Bond Cleavage by Use of Mono- and Dioxygenase Reaction

Ring activating dioxygenase. Dechlorination by ring activating dioxygenases isanother mechanism to remove chlorine from chloroaromatic compounds.Catechols are produced. The oxygen of the hydroxyl groups originates from


Fig. 38. Hydrolytic eliminations as part of the hydroquinone pathway

molecular oxygen. The oxygenolytic elimination has been shown for 2-chloro-,3-chloro-, 2,4-dichloro-, 2,5-dichloro-, and 3,4-dichlorobenzoate, 1,2,4,5-te-trachlorobenzene, and 4-chlorophenylacetate [382–396]. Other haloaromaticssuch as 2-fluorobenzoate, 2-bromobenzoate, and 3-fluorotoluene were also de-graded by use of this elimination mechanism [397–402].

Initial dioxygenases, which are responsible for ring activation, produce cis-dihydrodiols, which are further transformed by dehydrogenases to give cate-chols. The molecular oxygen is placed to the aromatic ring by a benzoate 1,2-di-oxygenase, a benzene 1,2-dioxygenase, a benzoate 3,4-dioxygenase, or a phenyl-acetate 3,4-dioxygenase in such a way that one of the vic-hydroxyl groups in thecis-dihydrodiol is bound to the same carbon as the chlorine substituent (seeFig. 39 with 2-chlorobenzoate). From such an unstable vic-dihydrodiol, thechlorine substituent will be eliminated to give an ortho-diphenolic compound.

For instance, in the case of the dehalogenation of 2-fluorobenzoate the na-ture of the dehalogenation seems to be a spontaneous reaction. While the mu-tant B9 of Alcaligenes eutrophus, which is defective in the dihydrodihydroxy-benzoate dehydrogenase, whose function is to form catechol in the benzoatepathway from dihydrodihydroxybenzoate, fails to grow with benzoate, 2-fluoro-benzoate can function as growth substrate although the dehydrogenase is miss-ing [397]. 2-Chlorobenzoate cannot be tested since the benzoate 1,2-dioxygen-ase is highly specific and does not tolerate the bulky chlorine substituent inortho-position. In Alcaligenes eutrophus especially, the benzoate 1,2-dioxygen-ase has its function in the degradation of benzoate and the activity with benzo-ate is higher than with the halogenated one. Therefore, the dehalogenation pro-cess seems to be a fortuitous one. In contrast, phenylacetate 3,4-dioxygenaseprefers the halogenated compound in comparison to the non-halogenated sub-strate, so that the physiological function seems to be that of a dehalogenase.

Because of the site specificity of the introduction of oxygen in positions 1and 2, a benzoate 1,2-dioxygenase can only bring about elimination of a chlo-rine substituent present at the position 2 in benzoates. The same narrow elimi-nation potential has been observed with the phenylacetate 3,4-dioxygenase.While 4-chlorophenylacetate is a substrate, the enzyme fails to use other chlori-nated phenylacetates.

Monohalosubstituted substrates such as 2-chloro- and 2-fluorobenzoate, 3-fluorotoluene, and 4-chlorophenylacetate, respectively, are oxygenolytically de-halogenated to yield catechol, methylcatechol, protocatechuate, and dihydroxy-

60 W. Reineke

Fig. 39. Oxygenolytic dechlorination of 2-chlorobenzoate by benzoate 1,2-dioxygenase. �benzoate 1,2-dioxygenase, � spontaneous, � ring cleavage

phenylacetate. The products are subject to further degradation by enzymesused for the mineralization of non-halogenated aromatic compounds. In con-trast, higher chlorinated substrates will be oxidized and partially dechlorinatedto chlorocatechols, which will be further degraded via the modified orthopathway (see later). 2,4-Dichloro- and 2,5-dichlorobenzoate are converted to 4-chlorocatechol, whereas 3,5-dichlorocatechol is the product formed from2,3,5-trichlorobenzoate. 1,2,4,5-Tetrachlorobenzene is dechlorinated to yield3,4,6-trichlorocatechol. 3,4-Dichlorobenzoate is dechlorinated to 5-chloropro-tocatechuate, which is subject of meta cleavage (see later).

Monooxygenase. A dechlorination of pentachlorophenol (PCP) by a hydrolyticdisplacement of chlorine in Rhodococcus chlorophenolicus PCP-1 was discussedfor quite a while. However, the results were questionable because of the contra-dicting results published by Apajalahti and Salkinoja-Salonen [403] and Schenket al. [404]. Uotila et al. [405] presented evidence that dechlorination proceedsby cytochrome P-450-mediated hydroxylase.

It is now well established that the initial dechlorination of pentachlorophe-nol occurs by a monooxygenase reaction [406, 407] producing tetrachloro-p-hydroquinone. The enzymes catalyzing chlorohydroquinone formation frompentachlorophenol have been studied in an Arthrobacter, Flavobacterium, andMycobacterium fortuitum [380, 408–411].

In analogy, an oxygenase-reaction is proposed to bring about the eliminationof a chlorine substituent from 2,4,6-trichlorophenol, yielding 2,6-dichlorohy-droquinone by Azotobacter sp. strain GP1 [412], Pseudomonas pickettii [413],and Streptomyces rochei 303 [375]. Similarly, the degradation of 2,4,5-trichloro-phenol by Burkholderia cepacia AC1100 proceeds through 2,5-dichlorohydro-quinone [378]. The chlorophenol 4-monooxygenase from strain AC1100 hasrecently been purified [414]. Hydroquinone was detected as the transient inter-mediate in the degradation of 4-chlorophenol by Arthrobacter ureafaciens[415].

3.1.1.3Reductive Displacement of Chlorine

A reductive dechlorination mechanism has been shown as part of the degrada-tion of chloroaromatic compounds like chlorobenzoate or pentachlorophenolby aerobic pure cultures. A corynebacterium strain NTB-1 and Corynebac-terium sepedonicum strain KZ-4 were found to degrade 2,4-dichlorobenzoatevia 4-chlorobenzoate [416, 417]. Recently, the degradation was shown to startwith the formation of 2,4-dichlorobenzoyl-CoA followed by a NADPH-depen-dent ortho dehalogenation yielding 4-chlorobenzoyl-CoA, hydrolytic removalof chlorine from the para-position to generate 4-hydroxybenzoyl-CoA, andhydrolysis to form 4-hydroxybenzoate [417].

Reductive dechlorination steps are also involved in the degradation of themetabolites of pentachlorophenol through the so-called hydroquinone path-way. In a Flavobacterium sp. and a coryneform-like strain [418, 419] reductivedechlorinations of tetrachloro-p-hydroquinone followed the initial oxidative


dechlorination of pentachlorophenol. The reductive dehalogenase has been purified by Xun et al. [420]. A chlorine is removed from tetrachloro-p-hy-droquinone through substitution with glutathione. Glutathione is then removedby displacement of the aromatic moiety by a second glutathione molecule,producing oxidized glutathione. This reaction sequence occurs a second time,leading to the formation of 2,6-dichlorohydroquinone (Fig. 40). The net re-action is equivalent to reductive dechlorination occurring in anaerobic or-ganisms.

In contrast, the pathway of pentachlorophenol degradation in Myco-bacterium fortuitum CG-2 is different [380] (Fig. 41). Tetrachloro-p-hydro-quinone, which is formed from pentachlorophenol by an oxygenase-reaction,seems to be initially ortho-hydroxylated to produce trichloro-1,2,4-trihydroxy-benzene followed by three reductive dechlorinations to give 1,2,4-trihydroxy-benzene.

62 W. Reineke

Fig. 40. Glutathione transferase catalyzed dechlorination of tetrachloro-p-hydroquinone to2,6-dichlorohydroquinone

Fig. 41. Hydrolytic and reductive dechlorinations in Mycobacterium fortuitum CG-2

3.1.2Late Eliminations of Chlorine After or Linked with Ring Cleavage

Chlorocatechols are key intermediates in the degradation of several chloro-aromatics (Fig. 42). The enzymatic reactions bringing about the formation ofchlorocatechols are similar to the peripheral sequences used for the degrada-tion of non-chlorinated aromatic compounds. This means that a number of re-actions dealing with chlorinated intermediates take place before dechlorinationsteps are reached. The late dechlorinations occur after ring cleavage from non-aromatic structures or are linked with the ring cleavage.

3.1.2.1Dechlorination as Part of the Modified Ortho Pathway

A common feature of a chlorocatechol degradative pathway (Fig. 43), used bymany organisms able to grow with chloroaromatics such as chlorinated anili-nes, benzenes, biphenyls, benzoates, naphthalenes, phenols, phenoxyacetates,salicylates, and toluenes, is the ortho cleavage of chlorocatechols by chlorocate-chol 1,2-dioxygenases with consumption of molecular oxygen to produce thecorresponding chloro-cis,cis-muconates [421–429].

The elimination of the first chlorine substituent was assumed to occur spon-taneously after 2-chloro- and 3-chloro-cis,cis-muconate have been converted bychloromuconate cycloisomerases to 5-chloro- and 4-chloromuconolactone, re-spectively [430]. Dienelactones are formed due to the anti-elimination of hy-drogen chloride and the formation of an exocyclic double bond [430, 431].While cis-dienelactone is formed from 3-chloro-cis,cis-muconate, trans-diene-lactone is the product from 2-chloro-cis,cis-muconate. 2,4-Dichloro-cis,cis-


Fig. 42. Schematic presentation of the mineralization of chloroaromatics with chlorocate-chols as key metabolites

64 W. Reineke

Fig. 43. Modified ortho pathway for the degradation of 3-chloro-, 4-chloro-, and 3,5-dichlorocatechol. � chlorocate-chol 1,2-dioxygenase, � chloromuconate cycloisomerase, �dienelactone hydrolase, � maleylacetate reductase. The bro-ken arrows indicate a spontaneous elimination of chlorinesubstituents from intermediates in parentheses

muconate, the metabolite in the 3,5-dichlorocatechol degradation, is convertedto 2-chlorodienelactone probably in the cis-configuration in analogy to theconversion of 3-chloro-cis,cis-muconate.

With respect to spontaneous elimination of chlorine during 2-chloro-cis,cis-muconate cycloisomerization, this view was challenged by the observation that(+)-5-chloromuconolactone is a stable compound when formed by muconatecycloisomerase, an enzyme of the normal 3-oxoadipate pathway [432]. The dataof Vollmer and Schlömann [433] corroborate the assumption that the chloro-muconate cycloisomerase of the 2,4-D-degrading Alcaligenes eutrophus JMP134purified by Kuhm et al. [434] or the 3-chlorobenzoate-degrading Pseudomonasputida AC866 have the ability to catalyze chlorine elimination from (+)-5-chlo-romuconolactone, the primary product of 2-chloro-cis,cis-muconate cycloiso-merization and are therefore dehalogenases. Overall, a chloromuconate cycloi-somerase brings about the conversion of 2-chloro-cis,cis-muconate, the productof the ortho cleavage of 3-chlorocatechol, to trans-dienelactone.

The dienelactones are converted into the respective maleylacetates by diene-lactone hydrolases.

The following enzyme, maleylacetate reductase, plays a major role in the de-gradation of chloroaromatic compounds either in the modified ortho pathwayor as part of the hydroquinone pathway. The original function is the reductionof the double bond by using NADH to channel maleylacetate into the 3-oxoadi-pate pathway.

In the case of maleylacetates with chlorine substituents in the 2-posi-tion such as 2-chloromaleylacetate, the intermediate in the degradation of 3,5-dichlorocatechol, the substrate is reduced to the respective chlorinated 3-oxo-adipate. This product is converted back into maleylacetate without a substi-tuent in position 2. This step, probably a spontaneous one, is accompanied bythe elimination of chloride. Therefore, two moles of NADH per mole substrateare consumed for the conversion of maleylacetates which contain a chlorinesubstituent in the 2-position [435–439]. In contrast, only 1 mol of NADH wasnecessary to convert 1 mol of those substrates without a chlorine substituent inthe 2-position as it is in maleylacetate or 3-chloro- and 5-chloromaleylacetate.

The modified ortho cleavage pathway described tolerates substitution at thearomatic ring of up to three chlorine atoms (see pathway for the degradation of1,2,4,5-tetrachlorobenzene later). Two dechlorination steps have been describ-ed up to now. Whether tetrachlorocatechol can serve as substrate for the knownchlorocatechol sequence is at present unknown.

3.1.2.2Dechlorination Linked with Ring Cleavage

For a long time the degradation of chloroaromatics has not been shown to oc-cur via the meta pathway. One reason has been found in the formation of a sui-cide product, a reactive acyl chloride, from 3-chlorocatechol by the catechol 2,3-dioxygenase of Pseudomonas putida PaW1 [440], which leads to inactivation ofthe ring cleavage enzyme (Fig. 44). In addition, 3-chlorocatechol is able to inac-tivate reversibly a catechol 2,3-dioxygenase because of its potential to chelatethe ferrous ion [441]. Some publications postulated that compounds degraded


via catechols chlorinated in the 4-position might be mineralized via the metapathway [442–448]. However, information about the way in which the productsare dechlorinated is not available.

Recently, Pseudomonas putida GJ31 was found to degrade chlorobenzene rapidly via 3-chlorocatechol and uses a meta cleavage pathway [449] (Fig. 44).In contrast to other catechol 2,3-dioxygenases, which are subject of inactiva-tion, the chlorocatechol 2,3-dioxygenase of strain GJ31 productively converts 3-chlorocatechol [450, 451]. Stoichiometric displacement of chloride occurs,leading to the production of 2-hydroxymuconate, which is further convertedthrough the meta pathway.

A productive meta cleavage without suicide effect has been known for morethan 15 years. Kersten et al. [452, 453] reported that a distal extradiol cleaving

66 W. Reineke

Fig. 44. Degradation of 3-chlorocatechol through the meta pathway. � Conversion leading tothe suicide inactivation of the 2,3-dioxygenase; � productive conversion in strain GJ31 grow-ing with chlorobenzene using a chlorocatechol 2,3-dioxygenase

protocatechuate 4,5-dioxygenase catalyzes 2-pyrone-4,6-dicarboxylic acid for-mation by nucleophilic displacement of a halide ion from protocatechuates sub-stituted with a halogen at the C-5 of the nucleus (Fig. 45). This indicates thatcyclization entailing nucleophilic displacement of halogen provides an effectivealternative to the enzyme suicide inactivation that occurs when a nucleophilicgroup of the dioxygenase undergoes acylation. An important aspect of this me-chanism is that the ring fission product remains bound to the enzyme during acomplete configuration change that precedes nucleophilic displacement.Hydrolysis of the pyrone is followed by degradation through a meta pathway.

In contrast, in the case of 3-chlorocatechol cleavage by the chlorocatechol 2,3-dioxygenase pyrone formation does not take place. Instead, the reaction of theacyl chloride with water directly leads to an intermediate of the meta pathway.

A similar type of oxygen-dependent, acyl chloride forming ring cleavage isassumed to occur in the degradation of g-hexachlorocyclohexane (lindane) andpentachlorophenol [454–456] (Fig. 46). There is evidence that 2-chlorohydro-quinone, the intermediate in the lindane degradation, is directly subject to ringcleavage by a new dioxgenase. 2,6-Dichlorohydroquinone, the metabolite in thepentachlorophenol degradation, is cleaved by an oxygen-dependent reaction


Fig. 45. Degradation of 5-chloroprotocatechuate through a meta pathway

Fig. 46. Dechlorination in the degradation of lindane and pentachlorophenol linked withoxygen-dependent ring cleavage

rather than hydrolyzed as published in former times. The ring cleavage pro-ducts of both reactions, acyl chlorides, seems to react with water to give maley-lacetate and 2-chloromaleylacetate plus HCl.

3.1.3Degradation of Higher Chlorinated Aromatic Compounds Needs Different Dechlorination Mechanisms

”Early” and “late” dechlorinations allow the degradation of tetrachlorobenzeneby a Pseudomonas sp. [394]. While the steps involved in the elimination of thefirst three chlorine substituents are well documented (Fig. 47), i.e., oxygenoly-tic dechlorination by the benzene dioxygenase, dehydrochlorination by thechloromuconate cycloisomerase, and reductive elimination by the maleylace-tate reductase, the fourth elimination step remains unclear as yet.

68 W. Reineke

Fig. 47. Degradation pathway for 1,2,4,5-tetrachlorobenzene by Pseudomonas sp. strain PS14involving early and late chlorine elimination steps according to Sander et al. [394]. The fol-lowing enzymes are involved: � benzene dioxygenase; � spontaneous; � chlorocatechol 1,2-dioxygenase; � chloromuconate cycloisomerase; � dienelactone hydrolase; � maleylacetatereductase; � probably 3-oxoadipate: succinylCoA transferase and 3-oxoadipylCoA thiolase;� unknown sequence


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The degradative pathway for 3,4-dichlorobenzoate (Fig. 48) is another exam-ple where elimination of chlorine substituents takes place due to the ring ac-tivation by benzoate 3,4-dioxygenase followed by ring cleavage by protocate-chuate 4,5-dioxygenase, i.e., oxygenolytic and nucleophilic displacements ofchloride following dioxygenase reactions.

The occurrence of a sequence of different types of chlorine eliminations, i.e.,oxygenolytic, hydrolytic, and reductive dechlorinations, can also be illustratedwith the hydroquinone pathway used for the degradation of pentachlorophenoland trichlorophenols, the different steps of which were discussed above (Fig. 49).

Ring cleavage of 6-chlorohydroxyhydroquinone or the nonchlorinated ana-logue by 6-chlorohydroxyquinol 1,2-dioxygenase brings about the formation of

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Fig. 49. Proposed hydroquinone pathway for chlorophenols involving different types of dechlorination mechanisms [375–377, 381, 410–413, 419, 420,456, 457]. The following enzymes are involved: � monooxygenase; � tetrachloro-p-hydroquinone reductive dehalogenase; � 2,6-dichloro-p-hydro-quinone chlorohydrolase; � oxygen-dependent ring cleavage followed by reaction with water; � 6-chlorohydroxyquinol 1,2-dioxygenase; � maleyl-acetate reductase

chloromaleylacetates or maleylacetate, which are also metabolites formed in themodified ortho pathway. In contrast, recent results indicate that the dichlorinat-ed hydroquinone rather than chlorohydroxyhydroquinone is the subject of di-rect ring cleavage in Sphingomonas chlorophenolica ATCC39723.

3.2Degradation of Chloroaromatic Compounds by Ligninolytic Fungi

Unlike bacteria, fungi generally do not utilize chloroaromatic compounds as asource of carbon and energy. Degradation of chloroaromatics and of manyother xenobiotic compounds is not the consequence of enzyme systems target-ed to this function. Fungal enzyme systems generally exist to serve other pur-poses such as degradation of wood components like ligninocellulose. Enzymesisolated and identified as having chloroaromatic degradative potential are thephenol oxidases, lignin peroxidases, manganese peroxidases, and laccase inligninolytic fungi. Phanerochaete chrysosporium and other white-rot fungi aresuch organisms bearing the biodegradative capabilities that encompass a broadrange of organopollutants like chlorinated anilines, benzenes, phenols, phe-noxyacetates, biphenyls, and dibenzo-p-dioxins.

Arjmand and Sandermann [458] found that chlorinated anilines are minera-lized by P. chrysosporium.

P. chrysosporium can substantially degrade and mineralize monochloro-benzene and dichlorobenzenes under nutrient-rich culture conditions, in whichthe lignin peroxidases and manganese peroxidases are not produced [459]. Thisindicates that the lignin peroxidases and manganese peroxidases are not requir-ed for degradation of the chlorobenzenes.

Identical results concerning the non-necessity of ligninolytic enzymes wereobtained with 2,4-D as the substrate. Yadav and Reddy [460] presented evidencefor mineralization of 2,4-D in nutrient-rich media by P. chrysosporium and by aperoxidase-negative mutant of this organism with about 40% of initial radioac-tivity found as 14CO2, indicating that the ligninolytic enzymes are not necessary.

The fungal degradation of PCBs has been studied in various laboratories.Eaton [461] reported that P. chrysosporium mineralized a significant fraction ofa 14C-labeled Aroclor 1254. Bumpus et al. [148] found significant rates of 14CO2evolution from radiolabeled DDT and lindane, but did not observe significantrates from two different polychlorinated biphenyls. The previous and recentstudies by Bumpus et al. [148] and Thomas et al. [462] indicated only low levelsof mineralization of 0.9–1.1% for individual PCB congeners such as 3,3¢,4,4¢-te-trachlorobiphenyl, 2,2¢,4,4¢-tetrachlorobiphenyl, and 2,2¢,4,4¢,5,5¢-hexachloro-biphenyl by P. chrysosporium. Results of Zeddel et al. [463] showed that degra-dation of a nonspecified PCB mixture by white-rot fungi Pleurotus ostreatus andTrametes versicolor was limited to mono- and dichlorinated congeners. Yadav et al. [464] presented evidence for substantial degradation of PCB mixtures byP. chrysosporium based on congener-specific gas chromatographic analysis.Degradation of Aroclor 1242, 1254, and 1260 (60%, 30%, and 18% by weight, re-spectively) was observed in both ligninolytic as well as non-ligninolytic media.Elimination of chlorine substituents was shown to be nonspecific, involving


ortho-, meta-, and para-substitution. Data from Dietrich et al. [465] providedfurther insight into the degradative activities of P. chrysosporium with the fol-lowing model PCB congeners: 4,4¢-dichlorobiphenyl, 3,3¢,4,4¢-tetrachlorobi-phenyl, and 2,2¢,4,4¢,5,5¢-hexachlorobiphenyl. Extensive degradation of 4,4¢-dichlorobiphenyl was found while negligible mineralization and metabolism of3,3¢,4,4¢-tetrachlorobiphenyl, and 2,2¢,4,4¢,5,5¢-hexachlorobiphenyl was observ-ed. 4-Chlorobenzoate and 4-chlorobenzyl alcohol were identified as metabolitesproduced from 4,4¢-[14C]-dichlorobiphenyl.

Information on the degradative sequence was obtained for 2,4-dichloro- and2,4,5-trichlorophenol [466, 467]. Extensive mineralization of 2,4-dichlorophenoloccurred only under nutrient nitrogen-limiting conditions [467], i.e., theligninolytic enzymes are essential for degradation. Valli and Gold [467] elucidat-ed a pathway for the degradation of 2,4-dichlorophenol with purified lignin pero-xidase and manganese peroxidase as well as cultures of P. chrysosporium based onisolation and characterization of metabolites formed and transformed. Thepathway involves several cycles. Oxidative dechlorination by either peroxidaseproduces a p-quinone. The p-quinone intermediate is then converted by intra-cellular enzymes and methylated to generate a peroxidase substrate. Such a cycleof oxidative dechlorination, quinone reduction, and hydroquinone methylationleads to the removal of the second chlorine atom in the second turn (Fig. 50).

P. chrysosporium rapidly mineralizes 2,4,5-trichlorophenol in nitrogen-limi-ted culture. Overall, the multistep pathway for 2,4,5-trichlorophenol resemblesthe 2,4-dichlorophenol pathway. It involves cycles of peroxidase-catalyzed oxi-dative dechlorination reactions followed by quinone reduction reactions toyield the key intermediate 1,2,4,5-tetrahydroxybenzene, which is presumablyring cleaved. The removal of all three chlorine atoms occurs before the ringcleavage followed by degradation to CO2 [466].

Mileski et al. [468] reported that P. chrysosporium also oxidizes pentachloro-phenol. In general, a negligible amount of PCP is mineralized by most fungistudied. Most of the PCP was transformed, often by O-methylation, to interme-diates such as pentachloroanisole. Phanerochaete spp. including P. sordida havealso been shown to degrade pentachlorophenol [469]. In addition, high-mole-

72 W. Reineke

Fig. 50. Proposed pathway for the degradation of 2,4-dichlorophenol by Phanerochaetechrysosporium. The compounds are converted by lignin peroxidase (LiP), manganese peroxi-dase (MnP), or whole cells


cular-weight polymers can be produced by enzymes of P. chrysosporium frompentachlorophenol [470].

Like P. chrysosporium the white-rot fungi Pleurotus ostreatus, Phellinus wei-rii, and Polyporus versicolor also mineralized DDT [471]. In general, the extendsof mineralization varied very much with significant amounts of water-solubledegradation products also observed in some cases.

Bumpus et al. [148] also reported that 2,3,7,8-tetrachlorobenzo-p-dioxin wasmineralized, but only low 14CO2 evolution was observed (2% of total), and theformation of metabolites was not elucidated. However, recently a mixture ofpolychlorinated dibenzo-p-dioxin was degraded at high yield by Phanerochaetesordida YK-624 in low-nitrogen medium [472]. 4,5-Dichlorocatechol was de-tected as metabolite from 2,3,7,8-tetrachlorodibenzo-p-dioxin, while tetrachlo-rocatechol resulted from the degradation of octachlorodibenzo-p-dioxin. Sincethe strain does not excrete lignin peroxidase, and breakdown was not mediatedby manganese peroxidase, enzymes other than these ligninolytic enzymes areresponsible for the degradation of polychlorinated dibenzo-p-dioxins.

A pathway for the model dioxin, 2,7-dichlorodibenzo-p-dioxin, was elucidat-ed by characterization of fungal metabolites generated by lignin peroxidase,manganese peroxidase, and crude intracellular cell-free extracts [473]. The mul-tistep pathway shown in Fig. 51 involves the degradation of 2,7-dichlorodibenzo-p-dioxin and subsequent intermediates by oxidation, reduction, and methylationreactions to yield the key intermediate 1,2,4-trihydroxybenzene. P. chrysospo-rium extensively degrades 2,7-dichlorodibenzo-p-dioxin only under nutrient-limiting conditions, suggesting that a lignin-degradative system is involved.

The dechlorination and cleavage of the dioxin was thought to function as fol-lows (Fig. 52). The first step is the one-electron oxidation by the oxidized en-

Fig. 51. Proposed pathway for the degradation of 2,7-dichlorodibenzo-p-dioxin by Phan-erochaete chrysosporium (LiP, lignin peroxidase, MnP, manganese peroxidase)

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Fig. 52. Proposed mechanism for the dechlorination and ring cleavage of 2,7-dichlorodibenzo-p-dioxin by lignin peroxidases of Phanerochaetechrysosporium

zyme intermediate LiPI resulting in the formation of the aryl cation radical A,which is probably short-lived. Attack of H2O at the cation would result in theloss of chloride and the formation of the carbon-centered radical intermediateB. One-electron oxidation by LiP or MnP would result in the formation of thecation intermediate C. Attack of H2O on this intermediate would lead to the firstC-O-C bond cleavage and the formation of the quinone intermediate D.Subsequent oxidation of the phenolic function would generate the phenoxy ra-dical E which is in resonance with the carbon-centered radical E¢. Oxidation byeither LiP or MnP would yield the cation F. Finally, attack of H2O on the cationwould result in the cleavage of the second C-O-C bond and generation of 4-chloro-1,2-benzoquinone and 2-hydroxy-1,4-benzoquinone.

In general, the data obtained with ligninolytic fungi concerning mineraliza-tion show an indistinct direction. Some degradation rates are very low.

3.3Anaerobic Microbial Populations with the Potential to Dechlorinate Chloroaromatic Compounds

The anaerobic biodegradation of a chloroaromatic compound was first demon-strated for pentachlorophenol in 1972 by Ide et al. [474], and later by severalother groups with both flooded soil and sewage sludge incubation systems[474–478]. However, the significance of reductive dechlorination of chlorinatedaromatic compounds by anaerobic bacteria has gained recognition only in thelast few years, beginning with the report by Suflita et al. [479].

In most cases, the microbial activities have only been shown in situ or withenvironmental material, i.e., sewage sludge, sediment, aquifer material, andwith undefined enrichments. While, in the main, the responsible microbes havenot been identified, a few studies are available now with defined consortia andwith isolated anaerobic bacteria.

In addition, phototrophic bacteria can metabolize chloroaromatics.

3.3.1Potential of Environmental Materials and Undefined Enrichments

By analyzing concentration changes, anaerobic dechlorination has been shown tooccur in anoxic materials with a large variety of chloroaromatic compounds(Table 10) under denitrification, sulfate reduction and methanogenic conditions.

Clear evidence for the role of microbial processes in dechlorination reactionswith environmental materials came from the following observations:1. No dechlorination occurs in autoclaved samples.2. Dechlorination is very specific and different in different systems: the reactions

are specific – only certain congeners were used as the substrate such as meta-substituted benzoates or meta- and para-substituted PCBs [479,492,527].Othersystems ortho-dechlorinate PCB congeners [519].Such a pronounced specificityfor a definite substitution pattern would not be expected to occur from abioticreactions. Enrichments from different sediments clearly indicate different pat-terns of dechlorination when adding the same congener mixture [527, 531].


Enrichment cultures from sewage sludge adapted to a certain monochloro-phenol show different dechlorinating activities with various chlorophenols[500]. Complete dechlorination of pentachlorophenol was obtained when amixed culture was produced from single cultures which are able to degradedifferent single chlorophenol isomers. In contrast, each single culture alonefailed to show this property towards pentachlorophenol [506]. The regio-selectivity of reductive dehalogenation in methanogenic sediments dependson the source of the microbial community and adaptation conditions[539–541].

3. The more highly chlorinated PCB congeners generally appear to be morereadily dechlorinated than the lower chlorinated congeners, which causes ac-cumulation of mono- and dichlorobiphenyls [524, 528].

4. The dechlorination of the lower substituted congeners started when thehigher chlorinated aromatics were totally converted into the lower chlorinat-ed ones [479].

5. The reductive dechlorination can be stimulated by the addition of organicelectron donors, such as lactate, acetate, pyruvate, ethanol, and glucose. H2can also function as electron donor. In some cultures the addition of an or-ganic electron donor is essential for the dechlorination.

A different degree of degradation by anoxic microbial populations has been ob-served. While some chloroaromatic compounds will be mineralized to giveCO2, others such as PCBs and chlorobenzenes are only partial dechlorinatedand lower chlorinated congeners are formed.

Dehalogenation and degradation of halogenated aromatic compounds byanaerobic bacteria populations have mostly been demonstrated under me-thanogenic conditions [542, 543]. Reductive dechlorination is followed by cleav-age of the aromatic ring, so that the chloroaromatic compound is ultimatelymineralized to CH4 and CO2. In addition, reductive dechlorination has been de-monstrated in other than methanogenic conditions. Inhibitors, such as bromo-ethanesulfonic acid (BESA) (for methanogens) or molybdate (for sulfate-reduc-ing bacteria), are used to determine which certain group of organisms is re-sponsible for the dechlorination. Often inhibition of the methanogenic activityby BESA did not show an effect on the dechlorinating activities [484], indicatingthat methanogenic organisms do not always take part in the dechlorination.

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Table 10. Chloroaromatic compounds dechlorinated by anoxic microbial populations

Compound(s) References

Chlorobenzenes [480–488]Chloroanilines [489–491]Chlorobenzoates [479, 492–498]Chlorophenols [494, 499–508]Chlorophenoxyacetates [505, 507, 509, 510]Chlorocatechols [511–513]Chlorobiphenyls (PCBs) [514–536]Chlorodibenzo-p-dioxins [537, 538]

Another interesting observation is that the presence of other electron accep-tors such as nitrate, sulfate, and other sulfur oxyanions may lead to a partial ortotal inhibition of the reductive dechlorination of aromatics by consortia andpopulations from sediments [490, 502, 515, 544–549]. However, the effect of sul-fate, for instance, might be in most instances a question of competition for elec-trons, since DeWeerd et al. [544] observed with resting-cells of Desulfomoniletiedjei, a model organism with aryl reductive dechlorination potential, that H2uptake was much faster when sulfate was available as the electron acceptor in-stead of 3-chlorobenzoate, the substrate for dechlorination. In addition, thiosul-fate and sulfite, but not sulfate, were found to be potent inhibitors and to repressinduction of the aryl dechlorination activity of Desulfomonile tiedjei [549].

However, the inhibition of the dechlorination in a consortium by electron ac-ceptors other than CO2 is not always the case, as these alternative electron ac-ceptors may indeed support anaerobic degradation of halogenated aromaticcompounds [503, 509, 550]. It has been demonstrated that chlorinated phenolsand benzoates can be degraded under sulfidogenic conditions in both freshwa-ter (Hudson and Nile Rivers) and estuarine sediments [551–553]. The depen-dency on sulfate reduction and inhibition by molybdate [552] suggests that sul-fate-reducing bacteria may be directly responsible for chlorophenol degrada-tion [554]. The reductive dechlorination as the initial step in chlorophenoldegradation by the sulfate-reducing consortium was confirmed by usingchlorofluorophenols as analogous compounds and the detection of the stoi-chiometric accumulation of fluorophenols [555].

Kazumi et al. [556] presented data indicating that Fe3+ can serve as a termi-nal electron acceptor in the microbial degradation of monochlorinated aromat-ic compounds such as phenols and benzoates in anoxic sediment enrichments.A systematic evaluation of the utilization of monochlorobenzoates under deni-trifying, Fe3+-reducing, sulfidogenic and methanogenic conditions showed thatanaerobic microbial consortia from the River Nile have the capacity to degradeall three chlorobenzoate isomers in the absence of oxygen and in the presenceof the alternative electron acceptors nitrate, ferric ion, sulfate, or carbon dioxide[553]. The degradation of chlorobenzoates was coupled stoichiometrically to NO3

–

loss, Fe2+ production, SO42– loss, or CH4 production, indicating that the chloro-

benzoates were oxidized to CO2. The loss of chlorobenzoate isomers was fastestunder denitrifying conditions when compared to the other reducing condi-tions. There was little difference in the rate of initial substrate loss among Fe-reducing, sulfidogenic, and methanogenic conditions. Degradation of mono-chlorobenzoates with the population from the River Nile was dependent notonly on the electron acceptor present but also on the position of the chlorine sub-stituent with the pattern meta>para>ortho. This relative degradability has pre-viously also been observed with cultures from the Hudson and East Rivers [551].

3.3.2Pure Cultures: Chloroaromatic Compounds as Electron Acceptors

Although a great number of dechlorinations of aromatics have been reported tooccur under anoxic conditions, only a few pure bacterial cultures have been iso-


lated until now which are able to dechlorinate an aromatic compound reduc-tively. While the physiological function of the dechlorination of 1,2,4-trichloro-benzene by the intestinal bacterium Staphyloccoccus epidermis [487, 488] is un-known, probably being a cometabolic step, the other strains use the chlorinatedaromatic compound as an electron acceptor in an anaerobic respiration(Table 11).

The first detailed data on the dechlorination mechanism were obtained fromDesulfomonile tiedjei DCB-1, which was isolated from a methanogenic 3-chlo-robenzoate-degrading consortium. The strain dechlorinates 3-chlorobenzoateto benzoate but cannot utilize the benzoate. The strain, a gram-negative, stric-tly anaerobic rod, was found to be a sulfate-reducing organism. The same wastrue for other isolates able to use 3-chlorobenzoate. However, the idea that sul-fate-reducing organisms in general have dechlorinating activities was found tobe wrong. The oxidation of formate is coupled to the reductive dechlorinationof 3-chlorobenzoate leading to energy for growth [557, 558]. Although thestrain is also able to dechlorinate chlorophenols and tetrachloroethene, thesedechlorinations seem not to be coupled to growth [559, 560]. The dechlorinat-ing activity towards 3-chlorobenzoate is inducible and co-induced with a te-trachloroethene dechlorinating activity [561]. The dehalogenating activity ofDesulfomonile tiedjei, located in the membrane, was found to be active in cell-free extracts [562]. The measurement of proton release in a cell suspension dueto the addition of 3-chlorobenzoate clearly indicates that the reductive dechlo-rination of 3-chlorobenzoate results in the formation of a proton gradientacross the cytoplasma membrane [563]. The 3-chlorobenzoate reductase hasbeen purified from the cytoplasmic membrane of Desulfomonile tiedjei DCB-1and characterized [564]. The dechlorination was found to represent a noveltype of anaerobic respiration [479, 496, 565]. Knowledge of the components in-volved in electron transfer from a donor molecule to the electron-accepting 3-chlorobenzoate is limiting. Louie et al. [566] found a unique membrane-boundcytochrome c induced in Desulfomonile tiedjei DCB-1 which is co-induced withthe 3-chlorobenzoate-dechlorinating activity. Louie and Mohn [567] demon-strated that the reductive dehalogenase is oriented towards the cytoplasm in themembrane of Desulfomonile tiedjei DCB-1, and the active site seems to be loca-ted on the cytoplasmic side of the membrane. Protons are produced in the pe-riplasm generating a proton motive force by a scalar mechanism, i.e., no pro-tons are translocated.

A model for the process is given in Fig. 53.Other pure cultures dechlorinate ortho-substituted phenols. Desulfito-

bacterium dehalogenans strain JW/IU DC1 is a gram-positive strictly anaerobicbacterium, which is able to use – besides the chloroaromatic compounds –other electron acceptors such as nitrate, fumarate, sulfite, thiosulfate, sulfur, and3-chloro-4-hydroxyphenylacetate [568, 569]. The dechlorinated products from2-chlorophenol or 3-chloro-4-hydroxyphenylacetate, phenol, and 4-hydroxy-phenylacetate, respectively, will not be used further by the strain. Desulfito-bacterium dehalogenans strain JW/IU DC1 has now been shown to grow via dehalorespiration [570]. The ortho-chlorophenol reductive dehalogenase ofstrain JW/IU DC1 has now been purified and characterized [571]. In addition,

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s79

Table 11. Properties of strains using halogenated aromatic compounds as electron acceptors

Strains E-donor E-acceptor C-source td (h) with the chloro-aromatic compound as electron acceptor

Desulfomonile tiedjei DCB-1 H2, formate, lactate, meta-Substituted chlorobenzoates, sulfate, CO2 and organic 26 (3-chlorobenzoate)pyruvate, benzoate, sulfite, thiosulfate compoundsmethoxybenzoates

Desulfitobacterium H2, formate, lactate, 3-Chloro-4-hydroxyphenylacetate, ortho- Organic compounds 3.5 (3-chloro-4-dehalogenans JW/IU-DC1 pyruvate, substituted chlorophenols, nitrate, fumarate, (yeast extract) hydroxyphenylacetate)

sulfite, thiosulfate, sulfurStrain 2CP-1 Formate, acetate ortho-Substituted chlorophenols, oxygen n.d. 89 (2-chlorophenol)

(<6%), fumarateDesulfitobacterium sp. Lactate, pyruvate, Tetrachloroethene, 2-chlorophenol, 2,4,6-tri- Lactate 23 (3-chloro-4-strain PCE1 butyrate, formate, chlorophenol, 3-chloro-4-hydroxyphenyl- hydroxyphenylacetate)

succinate, ethanol acetate, sulfite, thiosulfate, fumarateDesulfitobacterium Pyruvate Pentachlorophenol sulfite, thiosulfate nitrate n.d. n.d.frappieri PCP-1TDesulfitobacterium chloro- Formate, butyrate, 2,3-Dichlorophenol, 2,6-dichlorophenol, n.d. 23 (3-chloro-4-respirans Co23 crotonate, lactate, 2,4,6-trichlorophenol, 3-chloro-4-hydroxy- hydroxyphenylacetate)

pyruvate, H2 benzoate, 3-chloro-4-hydroxyphenylacetate,sulfite, thiosulfate, sulfur

Desulfovibrio sp. strain TBP-1 Lactate, pyruvate, H2, 2-Bromo-, 4-bromo-, 2,4-dibromo-, n.d. n.d.fumarate 2,6-dibromo-, 2,4,6-tribromophenol,

sulfate, sulfite, thiosulfate, sulfur

n.d.: no data.

the cloning of the gene coding the dehalogenase showed structural resemblancewith haloalkane reductive dehalogenases.

The strain 2CP-1 is a facultative anaerobic gram-negative rod, more closelyrelated to the myxobacteria than to D. tiedjei or other sulfidogens. It is able togrow under semi-aerobically conditions (oxygen concentration< 6%) [572]. 2-Chloro- and 2,6-dichlorophenol were substrates for dechlorination. Phenol, theproduct of the dechlorination, is excreted into the medium and supports cellgrowth. The strain was enriched from a culture supplied with 2-chlorophenol asan electron acceptor and formate and acetate as potential electron donors. Coleet al. [572] added BESA to the enrichment culture to block methanogenesis aswell as the syntrophic fermentation of the expected phenol. It is probable thatstrain 2CP-1 gains energy by using the chlorinated substrate as a respiratoryelectron acceptor. The range of preferred substrates for dehalogenation by 2CP-1 appears extremely limited. Dehalogenation activity is not constitutive inthis isolate but is induced by the presence of 2-chlorophenol, implying specificrecognition of the substrate. These data indicate that the dechlorination is nota cometabolic reaction.

It is unknown if dechlorination is coupled to growth via dehalorespiration in strain DCB-2 [573]. Strain DCB-2 is capable of rapidly catalyzing ortho de-chlorination of pentachloro-, 2,4,5- and 2,4,6-trichloro-, and 2,4-dichlorophe-

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Fig. 53. Scheme of the chemiosmotic coupling between reductive dechlorination of 3-chloro-benzoate and energy generation in Desulfomonile tiedjei (according to Louie and Mohn [567])

nol. A unique property is the meta dechlorination of 3,5-dichlorophenol. Re-cently the strain was identified as Desulfitobacterium hafniense [574]. It was reported that DCB-2 is able to dechlorinate reductively 3-chloro-4-hydroxy-phenylacetate to 4-hydroxyphenylacetate.

More recently, a tetrachloroethene-dechlorinating strain PCE1 was isolatedfrom a tetrachloroethene-dechlorinating enrichment culture [575, 576]. Thestrain was placed within the genus Desulfitobacterium and was found to use se-veral ortho-substituted phenolic compounds as electron acceptors in additionto tetrachloroethene when lactate or pyruvate were added as electron donors.2,4,6-Trichlorophenol was reductively dechlorinated via 2,4-dichloro- to 4-chlorophenol, whereas ortho dechlorination of 2-chlorophenol resulted in theformation of phenol. Reductive dechlorination of 3-chloro-4-hydroxyphenyla-cetate to 4-hydroxyphenylacetate supported growth of strain PCE1 on lactate.Other chlorinated aromatics, such as hexachlorobenzene, 2,5-dichloro-, 3,4-dichloro-, and 2,3,6-trichlorobenzoate, were not dechlorinated.

An anaerobic bacterium, PCP-1T, characterized as the new species Desulfito-bacterium frappieri, was isolated from a methanogenic consortium with penta-chlorophenol as the electron acceptor [577]. The organism dechlorinates severaldifferent chlorophenols in ortho-, meta-, and para-position to the hydroxyl group.Pentachlorophenol is dechlorinated via 2,3,4,5-tetra-, 3,4,5-trichloro-, and 3,5-dichloro- to 3-chlorophenol. The strain fails to dechlorinate 2,3-dichloro-, 2,5-dichloro-, 3,4-dichloro-, and monochlorophenols. There are indications that twodechlorination systems are involved in the dechlorination of pentachlorophenol,one for the ortho and the other for the meta and para dechlorination. Ortho- andpara-dechlorinating activities were found to be induced by different chlorophe-nols [578]. Sulfite, thiosulfate, and nitrate, but not sulfate, function as electron ac-ceptors when pyruvate and yeast extract were used for growth.

An anaerobic spore-forming microorganism, Desulfitobacterium chlorore-spirans strain Co23, was enriched on the basis of the ability to grow with 2,3-dichlorophenol as its electron acceptor [579]. Chlorines in ortho-positions wereremoved from di- and trichlorophenols such as 2,3-dichloro-, 2,6-dichloro-, and2,4,6-trichlorophenol, but not from monochlorophenols as well as 2,4-dichloro-,2,5-dichloro-, 2,3,5-trichloro-, and pentachlorophenol, when lactate was addedas electron donor. In addition to the chlorophenols, Desulfitobacterium chloro-respirans strain Co23 could dechlorinate 3-chloro-4-hydroxy-, 3,5-dichloro-4-hydroxybenzoate, and 3-chloro-4-hydroxyphenylacetate, but fails to usedchlorobenzoates as electron acceptors. Additional electron donors were pyru-vate, formate, butyrate, crotonate, and H2. Strain Co23 also used sulfite, thiosul-fate, and sulfur as electron acceptors for growth, but did not use sulfate, nitrate,or fumarate.

In contrast to most other anaerobic dechlorinating organisms, strain Co23grows rapidly and attains high cell densities on pyruvate in the presence of 3-chloro-4-hydroxybenzoate as the electron acceptor. 3-Chloro-4-hydroxybenzo-ate was shown to be the optimum electron acceptor for growing the cells, sincethe product of dechlorination, 4-hydroxybenzoate, was not inhibitory at highconcentrations, while 3-chlorophenol, the product from the dechlorination of2,3-dichlorophenol, was inhibitory to growth and dechlorination activity. The


dechlorination activity is inducible and was found to be membrane-bound andoxygen-insensitive. It exclusively dechlorinates chlorophenols in ortho-posi-tions [580]. Interestingly, cell-free membrane preparations dechlorinated somechlorophenols, such as 2,3,5-trichloro- and pentachlorophenol, that could notserve as electron acceptors supporting growth of strain Co23. Overall the de-chlorination takes place with a wide range of chlorophenols, i.e., from dichloro-to pentachlorophenol, but chlorobenzoates or tetrachloroethene were not de-chlorinated.

Boyle et al. [581] described the isolation and characterization of a Desul-fovibrio sp., designated strain TBP-1, capable of reductive dehalogenation fromestuarine sediments. The obligately anaerobic bacterium removes bromine sub-stituents in the ortho- and para-positions of brominated phenols (2-bromo-, 4-bromo-, 2,4-dibromo-, 2,6-dibromo-, and 2,4,6-tribromophenol) but not 3-bromo-or 2,3-dibromophenol or monobrominated benzoates. It does not dehalogenatechlorinated phenols. Strain TBP-1 possesses the ability to grow by coupling theoxidation of lactate (electron donor) to the reductive dehalogenation of 2,4,6-tri-bromophenol, yielding stoichiometric amounts of phenol. Pyruvate, hydrogen aswell as fumarate can function as electron donor besides lactate. Alternative elec-tron acceptors are sulfate, sulfite, sulfur, thiosulfate, but not nitrate.

A 16S rRNA sequence analysis showed a highly phylogenetic relationship be-tween chlorophenol-dechlorinating organisms. Desulfitobacterium chlorore-spirans strain Co23 is related to Desulfitobacterium dehalogenans JW/IU DC1,Desulfitobacterium sp. PCE1, and Desulfitobacterium hafniense DCB-2 with se-quence similarities of 97.2%, 96.8%, and 98.5%, respectively [579]. Desulfito-bacterium frappieri PCP-1T exhibits 95% similarity with Desulfitobacterium de-halogenans JW/IU DC1 [577].

Overall, little is known at present of the biochemical mechanism of the re-ductive dechlorination.

3.3.3Pure Cultures: Chloroaromatic Compounds as Growth Substrate

Häggblom and Young [582] recently isolated a denitrifying bacterium, Thaueraaromatica strain 3CB-1, from Hudson River sediment after enrichment on 3-chlorobenzoate under anoxic, denitrifying conditions. Other halobenzoateswith substituents in the 3-position are also degraded with stoichiometric re-lease of halide under conditions supporting anaerobic growth by denitrifica-tion. Complete oxidation of the substrates to CO2 was observed. Oxygen couldnot replace nitrate as the electron acceptor. The degradation was specific to theposition of the halogen substituent: the strain did not utilize 2- and 4-chloro-benzoate as sole carbon source.

3.3.4Dechlorinating Organisms, Part of a Food Web

Studies with Desulfomonile tiedjei DCB-1 suggest possible reasons why an-aerobes capable of reductive dechlorination of chloroaromatics are difficult to

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isolate; they are found most readily in complex communities (Fig. 54). D. tied-jei DCB-1 was isolated from a 3-chlorobenzoate-degrading methanogenic con-sortium consisting of strain DCB-1, a fermentative benzoate-degrading bacter-ium and a Methanospirillum sp., which consumes the hydrogen and CO2, pro-duced by the fermenting bacterium, to form methane. D. tiedjei DCB-1 feeds onhydrogen and acetate as sources for redox equivalents and carbon, which areproduced by the fermenting organism. 3-Chlorobenzoate is used as the electronacceptor by D. tiedjei DCB-1 and is reduced to benzoate and chloride. Possibleelectron donors for this process include hydrogen and formate.

3.3.5Phototrophic Bacteria and Chloroaromatic Compounds

Although the degradation of aromatic compounds by phototrophic bacteria hasbeen known of for a long time, it is only recently that phototrophic mineraliza-tion of chloroaromatics has gained some attention. A phototrophic enrichmentculture using acetate as carbon source partially dechlorinated 2,3,5,6-tetra-chlorobiphenyl in the presence of light [584]. Ortho chlorines were removedpreferentially. Two Rhodopseudomonas palustris strains, WS17 and DCP3, aswell as the non-classified phototrophic bacterium H45-2, are able to photo-metabolize 3-chlorobenzoate when grown with benzoate and forming stoichio-metric amounts of chloride [585, 586]. In contrast to strains WS17 and H45-2,


Fig. 54. Syntrophic relationships in a 3-chlorobenzoate-degrading consortium (based on[479, 493, 496, 544, 583])

strain DCP3 is the only photoheterotroph capable of using 3-chlorobenzoate forgrowth independently of the presence of benzoate [586]. In addition, Rhodo-pseudomonas palustris DCP3 3-chlorobenzoate-grown cells can also use 2-chloro-, 4-chloro-, and 3,5-dichlorobenzoate.

3.4Résumé: Chloroaromatic Compounds

This section has attempted to summarize data on the microbial conversion ofchloroaromatic compounds, some of which are presently known to be natural.

Many chloroaromatics serve as carbon and energy source for aerobic bac-teria. Diverse dechlorination mechanisms exist either pre or post ring cleavage,including hydrolytic, reductive, and oxygenolytic mechanisms. Various of thesesteps were found to be spontaneous in nature, i.e., the enzymes convert a chlo-rinated substrate to an unstable product, eliminating the chlorine substituent.Other dechlorination reactions were catalyzed by dehalogenases. Overall muchbiochemical data concerning the aerobic degradation of chloroaromatic com-pounds are available.

Anoxic microbial degradation of chloroaromatics was shown to take placewith electron acceptors such as nitrate, ferric ion, sulfate, or carbon dioxide. Mostof the respective populations showed their dechlorinating potential only in mixedculture or as black box material from the environment. In addition, the chloro-aromatics can also function as an alternative electron acceptor in a novel type ofanaerobic respiration, termed dehalorespiration, as has been shown with purecultures dealing with chlorobenzoates and chlorophenols. The biochemical me-chanisms involved in the anoxic dechlorination of chloroaromatic compoundsare presently unknown. However, recent years have show a rapid increase in in-formation concerning the anaerobic degradation of chloroaromatics.

Besides these activities of bacteria, fungi, especially the ligninolytic ones, canoxidatively dechlorinate chloroaromatic compounds in a cometabolic type ofprocess by using exoenzymes such as lignin and manganese peroxidases nor-mally used for cleaving lignin.

4Degradation of Aliphatic Hydrocarbons

4.1Aerobic Degradation of Aliphatic Hydrocarbons

4.1.1Alkanes

4.1.1.1Gaseous Alkanes (Short-Chain Alkanes)

Organisms termed methanotrophs, such as Methylomonas, Methylosinus,Methylococcus, etc., use methane as the sole source of carbon and energy. Thebacteria comprise a distinct group of obligatory methylotrophic organisms re-

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viewed by Anthony [587]. They grow well on methane but often rather poorlyon methanol. Methanotrophs do not grow on higher alkanes, which clearly dis-tinguishes them from other alkane-utilizing bacteria.

For the methane molecule to be assimilated for cell growth, this most highlyreduced form of carbon must be oxidized. The initial oxidation of methane iscatalyzed by methane monooxygenase (Fig. 55). Only one atom of the dioxygenmolecule is inserted into the methane molecule to produce methanol; the otheroxygen atom is reduced to water as has been shown by Higgins and Quayle[588] by using oxygen-18.

The methanol can be further oxidized to formaldehyde. This molecule caneither be used in an assimilatory way to make cell material or continue to beoxidized to CO2 in a dissimilatory fashion to produce reduced pyridine nucleo-tides for energy generation. The formaldehyde will be assimilated either via theribulose monophosphate cycle (type I methanotrophs such as Methylomonas,Methylococcus, Methylobacter) or the serine pathway (type II methanotrophssuch as Methylosinus, Methylocystis).

A remarkable feature of the methane monooxygenases is the wide variety ofsubstrates whose oxygenation they catalyze [589–591]:1. Alkanes, yielding primary and/or secondary alcohols.2. Monosubstituted aromatic compounds, yielding hydroxylated derivatives;

for example, the saturated side chain of ethylbenzene is oxidized to the pri-mary alcohol, while the aromatic ring is oxidized to give a hydroxyl group inthe para-position.

3. Alkenes, yielding epoxides, which are sometimes chemically stable.This cometabolic potential of the methane monooxygenase will be discussedlatter in the section on the degradation of chloroaliphatic compounds.

The ability of microorganisms to utilize propane as sole carbon source is welldocumented [592]. The organisms belong mainly to the Corynebacterium-Mycobacterium-Nocardia complex, a loosely defined group of gram-positive


Fig. 55. Degradation of methane. � methane monooxygenase; � methanol dehydrogenase;� formaldehyde dehydrogenase; � formate dehydrogenase. PQQ, methoxatin, (2,7,9-tricar-boxy-1H-pyrrolo(2,3-f)quinoline-4,5-dione)

bacteria containing other genera such as Rhodococcus, Brevibacterium, andArthrobacter [592, 593]. As with methane, the initial metabolic attack on pro-pane is by a monooxygenase, which produces propanol or isopropanol, i.e., byterminal or sub-terminal oxidation. The metabolism of 1-propanol will runthrough propanoate, and further the methyl malonate pathway [594, 595].

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Fig. 56. Pathways of propane metabolism [596–599]. � terminal oxidation via propanoate;�, sub-terminal oxidation via acetol and hydroxymethylacetate; � sub-terminal oxidationvia pyruvate; � sub-terminal oxidation via methyl acetate

Isopropanol degradation proceeds via acetone, which is further metabolized viaacetol (1-hydroxy-2-propanone) and hydroxymethylacetate, via pyruvate or viamethyl acetate to reach the central metabolism (Fig. 56). For an alternative de-gradative acetone pathway see Sect. 4.1.3.

4.1.1.2Long-Chain n-Alkanes

The utilization of C1–C4 alkanes is restricted to specialized species, which caneasily be enriched. n-Alkanes in the C5–C9 range are toxic to many microor-ganisms but can be biodegraded by some specific strains that have the correctset of catabolic enzymes. n-Alkanes of the C10–C22 range have been found to bereadily degradable in the environment and support growth of laboratory cul-tures [600–603]. Higher-molecular weight alkanes tend to be solid waxes andare not readily biodegraded; however, slow biodegradation of n-alkanes up toC44 has been shown [604].

In most cases – microorganisms such as Pseudomonas spp., Nocardia spp.,Mycobacterium spp., and certain yeasts such as Candida spp. and molds – theinitial metabolic attack on medium chain-length n-alkanes is by a monooxyge-nase to produce the corresponding alkane-1-ol. Attack by dioxygenase enzymeshas also been reported but is less common. In such cases the n-alkanes are con-verted to give the corresponding hydroperoxides which are reduced to yield analkane-1-ol (Fig. 57).

The sub-terminal oxidation of alkanes to yield secondary alcohols is rarer.Certain Aspergillus, Fusarium, and Bacillus strains were found to carry out sub-terminal oxidation of medium chain-length alkanes, producing alcohols with ahydroxyl group in the 4-, 5-, or 6-position. Lower quantities of 2- and 3-substi-tuted compounds were produced [605].

Subsequent metabolism of the alcohols may follow a number of pathways asillustrated in Fig. 58. The alcohol is normally oxidized to the correspondingaldehyde and fatty acid. Less commonly, w-oxidation may result in the produc-tion of a,w-dioic acids and/or w-hydroxy fatty acids [605]. The fatty acids pro-duced by all the pathways are then further metabolized by b-oxidation.Secondary alcohols produced by sub-terminal oxidation are further oxidized bya Baeyer-Villiger type of reaction to the corresponding ester and hydrolyticallycleaved to produce an acid and alcohol. The alcohol is then oxidized and thefatty acid is also subjected to b-oxidation.

The b-oxidation of fatty acids metabolism has been reviewed by Finnertyand Makula [606]. Fatty acyl-CoA synthetase activates the fatty acid. Although


Fig. 57. Dioxygenase catalyzed activation of alkanes

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Fig. 58. Pathways involved in the metabolism of n-alkanes. The three metabolic routes are:� terminal oxidation; � sub-terminal oxidation; � w-oxidation. They have been demon-strated to occur in various microorganisms, with terminal oxidation being the most common

TCA cycle

b-oxidation

this reaction is theoretically reversible, the equilibrium is shifted since AMPand pyrophosphate are generated rather than ADP and ortho-phosphate:

RCOOH + HS-CoA + ATP Æ RCO-SCoA + AMP + PPi + H2O

The pyrophosphate is readily hydrolyzed by a pyrophosphatase which ensuresirreversibility of the first reaction.

The b-oxidation cycle involves four separate reactions – fatty acyl-CoA de-hydrogenase, 2,3-enoyl-CoA hydratase (crotonase), 3-hydroxylacyl-CoA dehy-drogenase, and 3-oxoacyl-CoA thiolase (thiolase) – yielding one acetyl-CoA percycle (Fig. 59).


Fig. 59. b-Oxidation cycle. Enzyme reactions involved: � fatty acyl-CoA dehydrogenase;� 2,3-enoyl-CoA hydratase; � 3-hydroxyacyl-CoA dehydrogenase; � 3-oxoacyl-CoA thio-lase

For unsaturated acids, such as oleic acid, 18:1 (c9), the b-oxidation cycle canproceed only for three complete sequences before a metabolic block occurs[607] (Fig. 60). The product is 3-cis-dodecanoyl-CoA which has the doublebond in the wrong position and in the wrong configuration. The double bondshould be at the 2-position and the intermediate should have trans- not cis-con-figuration. Accordingly, an isomerase then converts 3-cis-dodecanoyl-CoA tothe corresponding 2-trans isomer. This then continues in the b-oxidation se-quence as before – hydratase, dehydrogenase, and thiolase to give decanoyl-CoA – which is then handled without further deviations.

4.1.2Branched Alkanes

In general branched chain alkanes are more slowly degraded than theirstraight-chain counterparts. However, it has become apparent that many of

these compounds are more degradable than had previously been thought. It isgenerally true that highly branched compounds are more recalcitrant thansimpler compounds, and particularly recalcitrant are b-branched and quater-nary compounds due to steric hindrance of oxidation enzymes [608, 609].

The ability of diverse microorganisms to grow at the expense of branched-chain hydrocarbons is variable with the indication that 2-methyl-branched al-kanes are usually good growth substrates, whereas 3-methyl-branched alkanesare attacked by very few microorganisms.

Degradation of 2,6,10,14-tetramethylpentadecane (pristane) has been parti-cularly well studied. Griffin and Cooney [610] found that 5 of 21 bacterial and11 fungal isolates obtained from freshwater environments could degrade thiscompound. The metabolic pathways responsible for pristane (Fig. 61) have beenstudied in detail in Brevibacterium sp., Corynebacterium sp., and Rhodococcussp. [611–613] and may involve b- or w-oxidation [614]. One turn, involving b-oxidation, yielding one propionyl-CoA, is followed by another one with acetyl-CoA as the product. These two turns follow each other in sequence.

Nakajima et al. [615] isolated a Rhodococcus sp. capable of degrading othercomplex branched chain alkanes such as 2,6,10,14-tetramethylhexadecane,2,6,10-trimethylpentadecane, and 2,6,10-trimethyldocedane, which were used

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Fig. 60. b-Oxidation sequence of unsaturated fatty acids: oleic acid; cis-D3-trans-D-2-enoyl-CoA isomerase


Fig. 61. Degradative pathways of 2,6,10,14-tetramethylpentadecane (pristane)

as sole sources of carbon and energy. Metabolism proceeded by oxidation ofisopropyl units to yield terminal alcohols and then fatty acids.

The problem of alkyl branching at the b-methyl position can be circumven-ted by b-alkyl removal as demonstrated in the citronellol pathway [616–621].The main features of this pathway are a cis-geranyl-CoA carboxylase and a 3-hydroxy-3-isohexenylglutaryl-CoA lyase which catalyzes the removal of the car-boxylated b-methyl group. The overall result is the conversion of the 3-methyl-acyl-CoA to a 3-ketoacyl-CoA intermediate, which can be subsequently cleavedvia b-oxidation. The citronellol pathway could provide the mechanism for oxi-dation of b-methylsubstituted alkanes or alkenes as has been shown by Fall etal. [622]. In combination with the initial monooxygenation as part of the alkanepathway the use of the citronellol pathway allowed the growth of a strain with2,6-dimethyl-2-octene for example (Fig. 62).

There is only one report on the microbial utilization of quaternary car-bon compounds because these hydrocarbons are extremely resistant tobiodegradation. An Achromobacter sp. was found to use 2,2-dimethylheptane assole carbon and energy source [623]. The organism simply attacked the unhin-dered terminus of the branched alkane and accumulated 2,2-dimethylpropionicacid.

4.1.3Alkenes

Olefins tend to be more toxic to microorganisms and, at least under aerobicconditions, are less readily utilized than the corresponding alkanes [600].Conversion of unsaturated aliphatic hydrocarbons may be initiated either viaattack on the double bond or by the same mechanisms employed in n-alkanemetabolism. It should be noted that some organisms capable of growth onshort-chain alkenes cannot grow on the corresponding alkanes since they canonly initiate metabolic attack at the double bond.

Four main patterns of initial attack can be recognized (Fig. 63): oxygenaseattack upon a terminal methyl group to produce the corresponding alkene-1-ol;sub-terminal oxygenase attack to produce an alkenol with the hydroxyl groupat a non-terminal carbon; oxidation across the double bond to give an epoxide;oxidation across the double bond to produce a diol.

The metabolism of short-chain alkenes (C6 and below) is thoroughly review-ed by Hartmans et al. [624] and Watkinson and Morgan [625].

Recently, an additional aspect of metabolism of alkenes has been obtained.The metabolism of propylene is initiated by a monooxygenase to the corre-sponding epoxide followed by a carboxylation reaction that forms acetoacetateas a product [627–629]. Further degradation in the Xanthobacter strain Py2,isolated by van Ginkel and de Bont [630], is proposed to proceed through ace-toacetyl-CoA and thiolysis to give two acetyl-CoA (Fig. 64). Carboxylation togive acetoacetate is also a significant strategy for acetone metabolism by aero-bic bacteria [631, 632].

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s93

Fig. 62. Combined action of alkane monooxygenase and citronellol pathway for the degradation of 2,6-dimethyl-2-octene, a branched alipha-tic compound substituted in the b-position. � 2,6-dimethyl-2-octene; � citronellol; � citronellic acid; � citronellyl-CoA; � cis-geranyl-CoA; � isohexenylglutaconyl-CoA; � 3-hydroxy-3-isohexenylglutaryl-CoA; � 2-methyl-6-oxo-2-octenyl-CoA

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Fig. 64. Proposed pathway for propylene in Xanthobacter strain Py2 and aerobic bacteria foracetone

Fig. 63. Metabolic pathways involved in the microbial degradation of alkenes [626]

4.2Anaerobic Degradation of Aliphatic Hydrocarbons

In contrast to the detailed studies on the aerobic degradation of aliphatic hy-drocarbons, little is known about the degradation of alkanes under anoxic con-ditions, where oxygen-initiated reactions cannot occur. Although the degrada-tion by pure cultures of Desulfovibrio under sulfate-reducing conditions hasbeen reported in the early literature [633–635], the potential degradability ofaliphatic hydrocarbons under anoxic conditions has remained a matter for de-bate. Over the past decades some studies on the alkane degradation with en-richment cultures or in microcosms were reported [636–642].

The first reliable proof of an anoxic degradation was provided for a cultureof a sulfate-reducing bacterium, strain Hxd3 [643, 644]. Growth of this culturewith hexadecane is very slow, with doubling times of more than one week un-der optimal conditions. The following stoichiometric relationship between n-hexadecane degradation and sulfate-reduction was observed:

C16H34 + 12.25 SO42– + 8.5 H+ Æ 16 HCO3

– + 12.25 H2S + H2ONo intermediates were detected and the metabolic pathway involved remainsunknown. Since that key publication by Aeckersberg et al. in 1991 [643] threenovel alkane-degrading, sulfate-reducing bacteria, strain TD3 [245, 645], strainPnd3 [646], and strain AK-01 [647] have been isolated from anoxic and hydro-carbon polluted environments, which now allow studies on the mechanism ofthe anaerobic alkane metabolism.

So and Young [648] provided evidence that alkanes are oxidized to fatty acidsby strain AK-01 under strictly anaerobic conditions. Subterminal addition of acarbon to the hydrocarbon chain as an initial reaction is demonstrated to givea carboxylic acid. This represents a novel mechanism by which alkanes can beactivated without oxygen in contrast to the well-known hydroxylation reactionmediated by monooxygenases in aerobic organisms.

Recently, three denitrifying bacteria, strains HxN1, OcN1, and HdN1, wereisolated in pure culture, which are able to utilize alkanes such as n-hexane, n-octane, or n-hexadecane anaerobically [649].

In many deep sediments, oxygen, nitrate, ferric ion, and sulfate are depleted,leaving methanogenesis as the only terminal degradation process. Zengler et al.[650] showed that under strict anoxic conditions long-chain alkanes (hexade-cane) can be transformed by enrichment cultures to methane with the follow-ing stoichiometry:

4 C16H34 + 3 H2O Æ 49 CH4 + 15 CO2

There has been one report of the anaerobic degradation of unsaturated long-chain hydrocarbons such as hexadecene and squalene by methanogenic en-richments [651]. However, the degradation was very slow. A pathway with hy-dration of the terminal double bonds to give the corresponding primary al-cohols and complete degradation via b-oxidation was proposed.

Acetylene as a highly unsaturated hydrocarbon is fermented fast comparedto acetate and ethanol by disproportionation through acetaldehyde, which isformed by a hydratase [652, 653].


Harder and Probian [654] reported that Pseudomonas citronellolis can growanaerobically on 3,7-dimethyl-1-octanol and citronellol with nitrate as an elec-tron acceptor.

4.3Résumé: Aliphatic Hydrocarbons

Oxygen as the substrate for oxidation of aliphatic hydrocarbons is necessary.The following statement about the relationship between structure and biode-gradability for alkanes and alkenes can be made: Long-chain n-alkanes are as-similated more readily than short chains. Saturated aliphatic hydrocarbons aredegraded more readily than unsaturated ones. Branched-chain compounds aredegraded less readily than straight-chain ones. In general, the degradation pro-ceeds to form fatty acids which are subject to b-oxidation.

Aliphatic hydrocarbons were thought to be more or less recalcitrant in theabsence of oxygen. However, according to recent literature the anaerobic alkanedegradation by sulfate reducers may be a more widespread phenomenon thanwas previously thought.

5Degradation of Chloroaliphatic Compounds

Chlorinated aliphatic compounds form one of the most important group of in-dustrially produced chemicals. Several of these compounds are poorly degrad-ed in the environment. This lack of biodegradation is mainly related to bio-chemical factors rather than thermodynamics. Both oxidative conversion ofchlorinated compounds with oxygen as the electron acceptor and reductive de-gradation to methane or alkanes should yield sufficient energy to supportgrowth [655].

Five types of action of microorganisms on chloroaliphatic compounds areknown:1. The compound serves as a sole carbon and energy source for the growth of a

pure culture of aerobic bacteria (see Table 12).2. The compound serves as growth substrate for organisms that use an electron

acceptor other than oxygen (nitrate respiration).3. The compound serves as a growth substrate in an acetogenic fermentation.4. The compound serves as a substrate for some enzymes in aerobic and an-

aerobic bacteria, while the microorganism grows with another compound,i.e., cometabolism (see Table 13).

5. The compound serves as an electron acceptor under anaerobic conditions(see Table 14).

5.1Chloroaliphatic Compounds as Growth Substrate for Aerobic Bacteria

The degree of recalcitrance of chlorinated aliphatic compounds to aerobic degradation generally increases with an increasing degree of chlorine substi-

96 W. Reineke

tution. Attempts to isolate organisms that grow on compounds such as tri- and tetrachloroethene, chloroform, and 1,1,1-trichloroethane have been unsuc-cessful.

A significant amount of work on the degradation of chloroaliphatics hasbeen carried out with cultures that grow on hydroxylated or carboxylated chlo-roaliphatics. This includes organisms growing on 2-chlorocarboxylic acids[666, 669, 670, 682, 747–752], 2-chloroethanol [677, 678], chloroallyl alcohols[675, 686], epichlorohydrin [753], 3-chloroacrylic acid [686, 687], and variouschloropropanols [675, 679, 753].

Organisms capable of degrading 2-chlorocarboxylic acids are easily isolatedfrom soil [754] and use hydrolytic dehalogenases. Oxidative conversion may oc-cur in organisms that grow on chlorinated alkanes [670], but has been poorlystudied. Figure 65 schematically shows the enzymatic reactions responsible forthe dehalogenation.


Table 12. Halogenated aliphatic as growth substrates for aerobic bacteria

Substrate Reference

HaloalkanesMethyl chloride CH3Cl [656–658]Dichloromethane CH2Cl2 [659–664] Ethyl chloride CH3CH2Cl [665]1,2-Dichloroethane CH2ClCH2Cl [666–669] 1-Chloropropane CH3CH2CH2Cl [667] 1-Bromooctane CH3(CH2)6CH2Br [670, 671]1-Chlorooctane CH3(CH2)6CH2Cl [665]1,3-Dichloropropane CH2ClCH2CH2Cl [667]1,9-Dichlorononane CH2Cl(CH2)7CH2Cl [672]HaloalkenesVinylchloride CH2=CHCl [673, 674] 1,3-Dichloropropene CH2ClCH=CHCl [675, 676]Haloalkanols2-Chloroethanol CH2CH2OH [677, 678]2,3-Dichloro-1-propanol CH2ClCHClCH2OH [679]1,3-Dichloro-2-propanol CH2ClCHOHCH2Cl [680]Haloalkenols2-Chloroallyl alcohol CH2=CClCH2OH [681]3-Chloroallyl alcohol CHCl=CHCH2OH [675]HaloalkanoatesChloroacetate CH2ClCOOH [682]Dichloroacetate CHCl2COOH [683]Trichloroacetate CCl3COOH [684]2,2,-Dichloropropionate CH3CCl2COOH [685]Haloalkenoates3-Chloroacrylate CHCl=CHCOOH [686, 687]3-Chlorocrotonate CH3CCl=CHCOOH [688]

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Table 13. Examples of halogenated aliphatic compounds as substrates for cometabolic processes

Substrate Growth substrate Organism(s) Reference

a) Transformation aerobic/oxidativeTrichloroethene Methane Mixed culture [689–691]

Methanotroph strain 46–1 [692]Methylosinus trichosporium [693]Methylosinus trichosporium OB3b [694–696]Methylocystis sp. [697]Methylomonas methanica 68–1 [698]

Methane and propane mixed culture [699]Propane Mycobacterium vaccae JOB5 [700]

Rhodococcus ssp. [701]Phenol Burkholderia cepacia G4 [702]

Alcaligenes eutrophus JMP134 [703]Toluene Pseudomonas putida F1; [704, 705]

toluene 2,3-dioxygenase pathwayBurkholderia cepacia G4; [704,toluene 2-monooxygenase pathway 706–709]Burkholderia pickettii PKO1;toluene 3-monooxygenase pathway [704]Pseudomonas mendocina KR1; [50, 51,toluene 4-monooxygenase pathway 704]

p-Cymene Rhodococcus erythropolis [710]Propene Xanthobacter sp. [711, 712]Isoprene Alcaligenes denitrificans JE75 [713]Ammonia Nitrosomonas europaea [714, 715]

Chlorinated Methane Mixed culture [690]ethenesChloroform Methane Mixed culture [689]

Toluene Pseudomonas sp. ENVBF1; [716]toluene 2-monooxygenase pathwayPseudomonas mendocina KR1; [716]toluene 4-monooxygenase pathway

Vinylchloride Propane Actinomycetales [717]Rhodococcus ssp. [701]

Halomethanes, Methane Methylococcus capsulatus (Bath) [718, 719]ethanes, ethenes

b) Transformation by pure cultures anaerobic/reductiveTetrachloro- Trichloromethane, Methanobacterium [720, 721]methane dichloromethane, CO2 thermoautotrophicum

Trichloromethane, Methanosarcina barkeri [722]dichloromethaneTrichloromethane, Desulfobacterium autotrophicum [720, 721,dichloromethane, CO2 723]Trichloromethane, Acetobacterium woodii [720, 723]dichloromethane,chloromethane, CO2


Table 13 (continued)

Substrate Product(s) detected Organism(s) Reference

Tetrachloro- Trichloromethane, Clostridium thermoaceticum [723]methane dichloromethane,

chloromethane, CO2

Trichloromethane, Clostridium sp. [724]dichloromethane,unidentifiedTrichloromethane Escherichia coli [725]Formate, CO2 Pseudomonas sp. strain KC [726, 727]Trichloromethane Shewanella putrefaciens 200 [728]Trichloromethane Shewanella putrefaciens MR-1 [729]

Trichloromethane Dichloromethane, Methanosarcina spp. [730]chloromethane

1,2-Dichloro- Chloroethane, ethene Methanogens [721, 731,ethane 732]

1,1,1-Trichloro- 1,1-Dichloroethane Methanobacterium [721]ethane thermoautotrophicum

1,1-Dichloroethane Desulfobacterium autotrophicum [721]1,1-Dichloroethane Acetobacterium woodii [720]1,1-Dichloroethane, Clostridium sp. [724]acetate, unidentified

Bromoethane Ethane Methanogens [731]

1,2-Dibromo- Ethene Methanogens [731]ethane

Tetrachloro- Trichloroethene Methanogens [561, 721,ethane 733]

Trichloroethene Desulfomonile tiedjei [561]Trichloroethene Methanosarcina sp. strain DCM [734]Trichloroethene Acetobacterium woodii [723]

1,2-Dibromo- Acetylene Methanogens [731]ethene

Trichlorofluoro- CHFCl2, CH2FCl, CO, Methanosarcina barkeri [735]methane fluoride(Freon 11)

5.1.1Hydrolytic Dehalogenation

Hydrolytic dehalogenases catalyze a nucleophilic displacement reaction withwater as the sole cosubstrate. The hydrolytic cleavage has been found in micro-organisms that grow with chlorocarboxylic acids and haloalkanes. The removalof the halogens from chlorocarboxylic acids results in the formation of hy-droxyalkanoic acids from monosubstituted compounds and oxoalkanoic acidsfrom disubstituted compounds.

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Table 14. Properties of bacteria reductively dechlorinating tetrachloroethene

Strain E-donor E-acceptor Product td (h) Referencesof PCE dechlorination

Dehalospirillum multivorans H2, formate, lactate, PCE, TCE, fumarate, nitrate cis-1,2-DCE 2.5 [736]pyruvate, ethanol, glycerol

Dehalobacter restrictus strain H2 PCE, TCE cis-1,2-DCE 19 [737]PER-K23Desulfitobacterium sp. strain Formate, lactate, pyruvate, PCE, ortho-chlorinated phenolics, TCE 58 [576]PCE-1 ethanol, butyrate, succinate fumarate, sulfite, thiosulfateDesulfitobacterium frappieri H2, acetate, pyruvate PCE, TCE, fumarate cis-1,2-DCE n.d. [738, 739] strain PCE-SDehalococcoides ethenogenes H2 PCE, TCE Ethene 19 [740, 741]strain 195Isolate TEA H2 PCE, TCE cis-1,2-DCE n.d. [742]Desulfuromonas chloroethenica Acetate, pyruvate PCE, TCE, fumarate, Fe(III)NTA cis-1,2-DCE 48–96 [743, 744]strain TT4BDesulfitobacterium frappieri H2, formate, lactate, ethanol, PCE, TCE, fumarate, nitrate, sulfite, cis-1,2-DCE 9 [745]strain TCE1 butyrate, crotonate thiosulfateIsolate MS-1 Polymers, carbohydrates, O2, nitrate cis-1,2-DCE n.d. [746]

esters, carboxylic acids,amides, aromatics, alcohols

n.d., no data.


Fig. 65. Mechanisms for the cleavage of carbon-chlorine bonds in microorganisms that growon chlorinated aliphatic compounds. � hydrolytic cleavage as occurring with chlorocar-boxylic acids and chloroalkanes; � haloalcohol dehalogenases catalyze a lyase reaction pro-ducing an epoxide from vic-haloalcohols; � hydratase-like mechanism as occurring withchloroacrylic acids; � dehalogenation catalyzed by a glutathione transferase; � monooxyge-nation of a chloro-substituted carbon atom produces a gem-chloroalcohol that decomposesto an aldehyde; � dehydrohalogenation from highly substituted chlorinated compounds;� dehalogenation by methyltransferase (X = unknown)

�

�

�

�

�

�

�

At least four different classes of hydrolytic 2-halo acid dehalogenases havebeen grouped according to their substrate specificity and stereospecific actionon 2-chloropropionic acid. Two classes of enzymes are active with either the L- or D-substrate, yielding products with inversion of configuration at the chiralC-2 carbon atom. The two other classes act on both stereo-isomers, one with inversion of configuration, the other with retention (Fig. 66). In general, highamino acid sequence identities are observed among the dehalogenases withinany of the separate classes. Mostly no homology is evident between the 2-halo-acid dehalogenases from different classes.1. About ten members of the class of hydrolytic dehalogenases, which is speci-

fic for the L-isomer of 2-chloropropionate and dehalogenates the substratewith inversion of configuration at the chiral carbon atom to produce the cor-responding D-2-hydroxy acid, are known and a catalytic mechanism has beenproposed [755–763].

2. The dehalogenase of Pseudomonas putida strain AJ1 belongs to the class,which is highly specific for the D-isomer of 2-chloropropionate and bringsabout inversion to form L-hydroxy acids [764].

3. DL-2-haloacid dehalogenases such as from Pseudomonas sp. strain 113 cat-alyze the hydrolytic dehalogenation of both enantiomers of 2-haloalkanoicacids, producing the corresponding 2-hydroxyalkanoic acids with inversionof the C-2 configuration [765].The enzyme of Pseudomonas sp. strain 113 showed significant sequence sim-ilarity with D-2-haloacid dehalogenases, but little with L-2-haloacid dehalo-genases [766].

4. DL-2-haloacid dehalogenases (retention type) dehalogenate both enantio-mers of 2-haloalkanoic acids to the corresponding D- and L-2-hydroxyalka-noic acids respectively [767].

Another type of dehalogenase is the enzyme from Moraxella sp. B that convertsfluoroacetate, and is mechanistically and structurally similar to haloalkane de-halogenase of Xanthobacter autotrophicus GJ10 [768].

A strain growing on trichloroacetate [769] dehalogenates chloroacetate ordichloroacetate to glycolate and glyoxylate. No oxalate was formed from tri-

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Fig. 66. Inversion and retention during hydrolytic elimination of chloride from enantiomersof 2-chloropropionate to the corresponding 2-hydroxypropionate (broken arrow: retention;full arrow: inversion)


Fig. 67. Catabolic pathway of 1,2-dichloroethane in strains of Xanthobacter autotrophicus andAncylobacter aquaticus [667, 669]

chloroacetate; instead carbon-carbon bond cleavage was observed, indicating adifferent mechanism of dehalogenation.

Hydrolysis of 2-bromobutane was accompanied by inversion of configura-tion [770].

Two hydrolytic dehalogenating enzymes have been shown in Xanthobac-ter autotrophicus and Ancylobacter aquaticus functioning in the degradation of 1,2-dichloroethane. The compound is degraded via 2-chloroethanol, chloro-acetaldehyde, and chloroacetate to glycolate in four successive reactions beforeit enters the central metabolic routes (Fig. 67). The dehalogenase (DhlA) con-verting 1,2-dichloroethane belongs to the a/b-hydrolase fold group [771].A modified aldehyde dehydrogenase is then essential for the conversion of thetoxic aldehyde [772]. The conversion of chloroacetate proceeds by a chloro-acetate dehalogenase (DhlB) that belongs to the L-specific haloacid dehaloge-nases.

A haloalkane dehalogenase hydrolyzing 1,3-dichloropropene to 3-chloro-allyl alcohol has recently been characterized from Pseudomonas cichorii 170 [676].The whole pathway, including the second dechlorination step, is given in Fig. 70.

5.1.2Glutathione S-Transferase-Dependent Dehalogenation

The role of glutathione transferases in degradation of halogenated compoundsis the nucleophilic displacement of a halogen substituent and further spon-taneous decomposition of the resulting glutathione adduct (Fig. 68). The firstevidence for the involvement of a glutathione transferase in bacterial dehaloge-nation was obtained with facultative and obligate methylotrophs of the generaMethylobacterium and Hyphomicrobium that can utilize dichloromethane as acarbon source for growth [661].

The role of these enzymes and their mechanistic aspects were recently re-viewed [773, 774]. Dichloromethane dehalogenases are atypical enzymesamong glutathione S-transferases. In contrast to most enzymes of the gluta-thione S-transferase superfamily, their substrate range is very narrow andrestricted to dihalomethanes. Halomethanes and dihaloethanes are not con-verted.

5.1.3Lyase-Catalyzed Dehalogenation

Haloalcohol lyases catalyze the intramolecular nucleophilic displacement ofvic-haloalcohols to their respective epoxides. Both the enzymes fromCorynebacterium sp. strain N-1074 [776] and from Arthrobacter sp. strain AD2[680] are more active with bromoalcohols than with chloroalcohols. The en-zyme reaction (Fig. 69) is reversible, allowing formation of epoxides from vic-haloalcohols, conversion of the epoxides to haloalcohols, and trans-halogena-tion reactions of haloalcohols. The enzymes use 2-propanols with halogen sub-stituents in the 1- or 3-position.

A lyase reaction has recently been detected to be involved in the degradationof 1,2-dibromoethane by Mycobacterium sp. strain GP1 [777]. The first step inthe degradation is catalyzed by a hydrolytic haloalkane dehalogenase. The re-sulting vic-haloalcohol, 2-bromoethanol, is then rapidly converted by the lyaseto ethene oxide. The further metabolism is unclear.

5.1.4Hydratase-Catalyzed Dehalogenation

A hydration type of dehalogenation reaction has been proposed for the conver-sion of aliphatic acrylates (Fig. 70) [686, 687]. The degradation of cis- and trans-3-chloroacrylate by coryneform bacteria is catalyzed by dehalogenases that arespecific for the cis and trans isomers. Two 3-chloroacrylate dehalogenases havealso been found as part of the degradation pathway of 1,3-dichloropropene ofPseudomonas cichorii 170 [676].

5.1.5Dehalogenation by Oxygenases

Little biochemical information is available on the role of oxygenases in dehalo-genation of halogenated aliphatic compounds. Oxygenation can lead to dehalo-

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Fig. 69. Microbial dehalogenation of chloropropanols via chloroepoxides. (a) haloalcohol dehalogenase, (b) epoxide hydrolase

Fig. 68. Degradation of dichloromethane [775]

genation if a saturated carbon atom to which a chlorine is bound is hydroxyla-ted to yield a gem-chloroalcohol. These compounds are chemically unstable anddecompose to produce aldehydes. A monooxygenase may play a role in the uti-lization of methyl chloride by Hyphomicrobium sp. strain MCl [657] and in growth of Pseudomonas sp. strain DE2 on 1,2-dichloroethane [668].

Just recently, a monooxygenase reaction was shown to initiate the degradationof 1,2-dichloroethane by Pseudomonas sp. strain DCA1, an organism exhibitinghigh affinity to the substrate (Fig. 71) [778]. Oxidation results in the formation ofan unstable intermediate, 1,2-dichloroethanol, which spontaneously releaseschloride yielding chloroacetaldehyde. The further degradation proceeds throughthe pathway described for X. autotrophicus and Ancylobacter aquaticus [667, 669].

5.1.6Dehalogenation During b-Oxidation

There is evidence that dehalogenation of chlorinated carboxylic acids may oc-cur after formation of CoA-derivatives (Fig. 72). Kohler-Staub and Kohler [688]


Fig. 70. Pathway of cis-1,3-dichloropropene bringing about the formation of malonic semial-dehyde. � haloalkane dehalogenase; � alcohol dehydrogenase; � aldehyde dehydrogenase;� 3-chloroacrylate dehalogenase. The same pathway functions for the trans isomer [676]

Fig. 71. Catabolic pathway of 1,2-dichloroethane in Pseudomonas sp. strain DCA1 [778]

showed that dechlorination of trans-3-chlorocrotonate and 3-chlorobutyrate,used as a sole carbon and energy source by Alcaligenes sp. strain CC1, occurs af-ter activation of the acids to their coenzyme A derivatives. The dechlorinationreaction is not yet understood. The same could be the case in the organism iso-lated with chloroallyl alcohols as the growth substrates [681].

5.1.7Dehydrohalogenation

The first step in the metabolism of lindane (g-hexachlorocyclohexane) is an eli-mination by a dehydrochlorinase (Fig. 73). The initial product is g-pentachlo-rocyclohexene, which is further converted by the same enzyme to 1,3,4,6-te-trachloro-1,4-cyclohexadiene. Two hydrolytic dechlorinating steps follow yield-ing 2,5-dichloro-2,5-cyclohexadiene-1,4-diol which is then metabolized by adehydrogenase to 2,5-dichlorohydroquinone [779–782].

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Fig. 72. Proposed degradation sequence used for trans-3-chlorocrotonate [688]

Fig. 73. Pathway for lindane (g-hexachlorocyclohexane) including dehydrohalogenation andhydrolytic dechlorination

5.1.8Dehalogenation by Methyltransferase/Dehydrogenase

Aerobic methylotrophic bacteria of the genera Hyphomicrobium and Methylo-bacterium have been characterized, which are able to utilize chloromethane as a growth substrate [656, 657]. Physiological and genetic studies suggest that Methylobacterium sp. strain CM4 metabolizes chloromethane by initial de-halogenation via a methyl transfer reaction (Fig. 74), followed by a series ofdehydrogenase-based steps which are different from those involved in the downstream steps of methanol metabolism in the same organism [658].Dichloro- and trichloromethane are neither growth substrates nor are they de-chlorinated by strain CM4 cells, indicating a narrow specificity of the dechlori-nation step.

5.2Chloroaliphatic Compounds as Growth Substrates for Anaerobic Bacteria

5.2.1Fermentative Degradation

It is conceivable that chlorinated C1 hydrocarbons are utilized as carbon andenergy sources by anaerobic bacteria, a process that has been observed for chlo-romethane [783, 784] and dichloromethane [785–788].

Traunecker et al. [784] reported the isolation of a strictly anaerobic, homo-acetogenic bacterium, fermenting methyl chloride as the sole energy sourcewith a doubling time of near 30 h. Strain MC does not grow with dichlorome-thane as an energy source. Cells formed about 3 mol of acetate per 4 mol ofCH3Cl consumed:

4 CH3Cl(g) + 2 CO2(g) + 2 H2O Æ 3 CH3COO– + 7 H+ + 4 Cl–

DGo¢ = – 419.9 kJ/reaction

More recently, Meßmer et al. [789] reported the development of an enzyme as-say to determine the activity of the methyl chloride dehalogenase of the homo-acetogen strain MC. They showed that the dehalogenase activity was dependenton the presence of substoichiometric amounts of ATP. The pathway for chloro-methane degradation is presented in Fig. 75.

A strictly anaerobic two component culture able to grow with a doublingtime of 20 h on a medium containing dichloromethane as the carbon andenergy source was established by Mägli et al. [790]. The strain DMC was able togrow acetogenically with dichloromethane when it was associated with Aceto-bacterium woodii, Methanospirillum hungatei, or Desulfovibrio sp. strain DMB.Strain DMC contained CO dehydrogenase activity and is responsible for boththe dehalogenation of dichloromethane and the acetogenesis. The obligatorydependence on a partner for growth might be due to the need for a growth fac-tor provided by the associated organism. The partial mass balance for growthwith dichloromethane is compatible with the following fermentation balance:

2 CH2Cl2 + 2 H2O Æ CH3COO– + 5 H+ + 4 Cl– DGo¢ = –492.7 kJ/reaction

and is thus thermodynamically favorable. A major portion of this free energy,however, is associated with the dehalogenation step:

2 CH2Cl2 + 2 H2O Æ 2 HCHO + 4 H+ + 4 Cl– DGo¢ = –344 kJ/reaction

Later strain DMC was isolated from the dichloromethane-fermenting, two com-ponent mixed culture and characterized as Dehalobacterium formicoaceticum


Fig. 74. Chloromethane metabolism involving methyltransferase (X = unknown acceptor, notglutathione)

[791]. The organism converted in a mineral medium with vitamins 5 mmol/ldichloromethane within 7 days to formate plus acetate in a molar ratio of 2:1and to biomass. Only dichloromethane supported growth, while other com-pounds including chloromethane (50 potential substrates were tested) were notused by the organism.

Dichloromethane is converted to methylene tetrahydrofolate (Fig. 76), ofwhich two-thirds are oxidized to formate while one-third gives rise to acetate byincorporation of CO2 in the acetyl coenzyme A synthetase reaction. The reduc-ing equivalents generated by the oxidation through the acetyl CoA pathway areused by the methylene tetrahydrofolate reductase in the formation of acetatethrough the CO dehydrogenase pathway. The dehalogenating activity was foundto be dependent on the presence of ATP, methyl viologen, and hydrogen [787].Co(I) corrinoid seems to be involved in this anoxic dehalogenation of dichlo-romethane. The presence of a sodium-independent ATP synthetase suggests achemiosmotic mechanism of energy conservation.

108 W. Reineke

Fig. 75. Metabolism of chloromethane by the homoacetogenic bacterium MC according toMeßmer et al. [789]. The enzymes are the following: � methyl chloride dehalogenase;� methylene tetrahydrofolate reductase; � methylene tetrahydrofolate dehydrogenase;� CH ∫ FH4 cyclohydrolase; � formyl tetrahydrofolate synthetase; � formate dehydro-genase; � CO dehydrogenase; � phosphotransacetylase; acetate kinase; methyl trans-ferase. CH3-Co-E = corrinoid enzyme

5.2.2Degradation Under Denitrifying Conditions

The facultative methylotroph Hyphomicrobium sp. DM2 has been shown to becapable of growth with dichloromethane in the absence of oxygen using nitrateas a terminal electron acceptor [792].

5.2.3Degradation Under Methanogenic Conditions

The anaerobic degradation of chloroacetates has only been reported by Egli etal. [793]. A methanogenic stable mixed culture degraded chloroacetate accord-ing the following overall stoichiometry:

4 ClCH2COO– + 7 H2O Æ 5 HCO3– + 3 CH4 + 4 Cl– + 5 H+

The data suggest that the anaerobic chloroacetate degradation proceeds via gly-colate formed by hydrolytic dehalogenation. Glycolate is then cleaved to carbondioxide and hydrogen, the substrates of the carboxidotrophic methanogens.


Fig. 76. Proposed pathway for the metabolism of dichloromethane by D. formicoaceticumstrain DMC [787]. The enzymes are the following: � dichloromethane dehalogenase; � me-thylene tetrahydrofolate dehydrogenase; � CH ∫ FH4 cyclohydrolase; � formyl tetrahydro-folate synthetase; � methylene tetrahydrofolate reductase; � methyl transferase; � CO de-hydrogenase; � phosphotransacetylase; acetate kinase. CH3-Co-E = corrinoid enzyme

5.3Cometabolic Transformations

Cometabolism is the process whereby enzymes or cofactors that have evolved todegrade other substrates fortuitously convert another compound.

5.3.1Aerobic Bacteria: Oxidative

The potential of bacteria for oxidative cometabolic degradation has been stud-ied with chloroethenes and chloromethanes. Only the monochlorinated conge-ner of the chloroethenes, vinyl chloride, is known to serve as growth substratefor aerobic microorganisms. The di- and trichlorinated congeners are meta-bolized exclusively co-metabolically, and tetrachloroethene is not degraded atall by microorganisms under aerobic conditions.

The presence of C=C bonds renders vinylic halogens rather resistant tonucleophilic substitution reactions. With these compounds, dehalogenation byoxidative mechanisms involving the addition of oxygen atoms to the doublebond is possible. The enzymes responsible for the fortuitous dehalogenation aremono- or dioxygenases: toluene 2-monooxygenase, toluene 3-monooxygenase,toluene 4-monooxygenase, toluene 2,3-dioxygenase, methane monooxygenase,isoprene monooxygenase, propane monooxygenase, ammonia monooxygenase(Table 13). They are present in bacteria growing on e.g. methane [693, 695], tol-uene [704, 705, 707, 708, 794, 795], phenol [702, 703], isoprene [713], p-cymene[710], propene [711], propane [700], or ammonia [714].

Since the chlorinated ethenes such as trichloroethene (TCE) are convertedonly cometabolically, the natural substrate (e.g., toluene or phenol, etc.) isnormally required for induction of the degradative enzyme. Recently, orga-nisms have been found in which the TCE-degrading activity (toluene 2,3-di-oxygenase) can be induced by TCE [796, 797].

Cometabolic oxidative degradation results in the formation of products,some of which can be very toxic to the organisms that produce them and inac-tivate the enzyme responsible for the conversion. The toxic intermediate in thecase of trichloroethene is thought to be trichloroethene epoxide. Thus, the con-version of the epoxides is a critical step in the degradation route of chlorinatedethenes. The selection of organisms, that are able to convert the potentiallytoxic intermediates fast enough to preclude any harm being done by these intermediates, seems likely to lead to better degradation of trichloroethene.Isoprene-degrading organisms may contain epoxide-converting enzymes thatalso efficiently convert chlorooxiranes. Toxic effects of trichloroethene con-version by the isoprene degraders were indeed not observed [713].

The results presented by Fox et al. [694] obtained with purified me-thane monooxygenase from the type II methanotroph Methylosinus tricho-sporium OB3b strongly imply that neither TCE nor the immediate enzyme-catalyzed oxidation products, TCE epoxide and chloral, are responsible for theinactivation reaction. A diffusible intermediate derived from the non-enzyme-catalyzed hydrolysis or isomerization of TCE epoxide, such as glyoxyl chloride,

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formyl chloride, dichloroacetyl chloride, or dichlorocarbene, is thought to bethe modifying agent. The methane monooxygenase oxidizes TCE predo-minantly to TCE epoxide, while 2,2,2-trichloroacetaldehyde (chloral) is pro-duced at 6% of the total product yield [694]. The various products arising dur-ing conversion of TCE by the methane monooxygenase are summarized inFig. 77.

The toluene 2,3-dioxygenase oxidizes TCE to formate and glyoxylate in a 2 :1ratio [798].

In contrast to the methane monooxygenase and the toluene 2,3-dioxygenase,the toluene 2-monooxygenase shows that Burkholderia cepacia G4 oxidizes TCEwith little apparent inactivation [702, 799]. The stable products of TCE oxida-tion by toluene 2-monooxygenase were shown to be carbon monoxide, formate,and glyoxylate [800].

Besides the dechlorination of TCE chloroform was tested with differenttoluene-oxidizing bacterial strains harboring mono- and dioxygenases towardselimination of chlorine substituents [716]. Pseudomonas mendocina KR1, whichinduces a toluene 4-monooxygenase, as well as Pseudomonas sp. ENVBF1, whichappears to produce toluene 2-monooxygenase, oxidize chloroform at respect-able rate. The same is true for Escherichia coli harboring the respective clonedgenes of the monooxygenases. Degradation of 14C-chloroform and ion analysisrevealed that a great part was mineralized to carbon dioxide (approximately30–57% of the total products). Chloride was approximately 75 % of the ex-pected yield.

However, Burkholderia cepacia G4 producing toluene 2-monooxygenase,Pseudomonas putida producing toluene 2,3-dioxygenase, and Burkholderiapicketti PKO1 producing toluene 3-monooxygenase fail to show oxidation ofchloroform. So overall the picture of the potential of the oxidizing enzymes to-ward chloroform is not totally clear.


Fig. 77. Oxidation of trichloroethene by methane monooxygenase of Methylosinus trichospo-rium OB3b and following reactions according to Fox et al. [694]. � main reaction of methanemonooxygenase; � side reaction of methane monooxygenase; � spontaneous; � reduction;� oxidation

5.3.2Ligninolytic Fungi: Reductive

The enzyme system of the ligninolytic fungi is known to oxidize a number of che-micals including polycyclic hydrocarbons. Chloroaliphatics are highly electrondeficient and therefore cannot be oxidized by the lignin peroxidase. However, a li-gnin peroxidase-dependent reductive pathway seems to be possible (Fig. 78)[801]. The veratryl alcohol cation formed by the lignin peroxidase is an oxidizingspecies which can effectively oxidize certain chemicals by one electron. EDTA wasshown to be a good reductant for the veratryl alcohol cation radical and will beoxidized to the organic acid radical. The EDTA anion radical reduces other chem-icals such as tetrachloromethane, resulting in the dechlorination and formationof trichloromethyl radical and the decarboxylation of EDTA. In summary, CCl4,which is neither a substrate for lignin peroxidase nor a good reductant, is degrad-ed via free radicals generated by lignin peroxidase under reducing conditions.

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Fig. 78. Proposed scheme for the reduction of CCl4 by lignin peroxidase (LiP)

5.3.3Anaerobic Bacteria: Reductive

Considerable evidence has accrued in studies of anaerobic microcosms and cul-tures for reductive dechlorination of tetra- to trichloroethene (PCE, TCE) [560],to dichloroethene isomers [802–804], or to vinyl chloride (VC) [805, 806]. Moreimportant, complete dechlorination to ethene [805, 807] or ethane [808] hasbeen reported.

Freedman and Gossett [805] demonstrated the reductive dechlorination ofPCE and TCE to ethene by anoxic aquifer material when methanol, glucose, H2,and other electron donors were added. No conversion to CO2 and CH4 was ob-served. The authors propose a pathway for the reductive dechlorination that fol-lows the course PCE, TCE, cis-1,2-dichloroethene (cDCE), vinyl chloride, ethene(Fig. 79). The dechlorination process was inhibited by 2-bromoethane sulfonicacid, a methanogen inhibitor, indicating that these organisms may play a keyrole in the anaerobic biotransformation of PCE and TCE.

It is unclear why some anaerobic systems only partially dechlorinate PCEwhile others effect complete dechlorination. Reductive dechlorination of TCEand PCE under methanogenic conditions can proceed to VC [805–807], where-as cDCE has tended to accumulate under sulfate-reducing conditions [802, 809].

A clearer picture of the microorganisms and enzymes involved in reductivedechlorination emerges with studies using pure cultures. Reductive dehaloge-nation of molecules such as dibromo- and dichloroethane has been demon-strated with pure cultures of methanogens [731]. The anaerobic dechlorinationof chlorinated ethenes has also been demonstrated with corrinoids (Co(II)-containing) and nickel porphyrins, which play a role in the normal anaerobicmetabolism. Thus, transition metal-containing cofactors or enzymes are likelyto be responsible for the reductive dechlorination in the anaerobes.

5.4Chloroaliphatic Compounds as Electron Acceptors

The above-mentioned dechlorinating anaerobic strains cometabolically trans-form the chlorinated compounds and thus do not benefit from the exergonic re-action which they catalyze. Halogenated aliphatic compounds are quite strongoxidants: hexachloroethane is a stronger electron acceptor than is oxygen, andseveral halogenated compounds, such as tetrachloromethane, tetrachloro-ethene, and trichloromethane, are stronger acceptors than nitrate [810]. There-fore, they can in principle serve as terminal electron acceptors in an anaerobicrespiration.

Although there have been many indications that reductive dechlorination re-actions are catalyzed by bacteria that couple this reduction to growth, the isola-tion of these bacteria appears to be very difficult. To date, eight pure cultureshave been obtained that dechlorinate tetrachloroethene at high rates and utilizethe chlorinated compound in an anaerobic respiration as an electron acceptor(Table 14). In another pure strain, isolate MS-1, which is able to dechlorinatePCE, the respiratory growth with PCE has not yet been shown. With respect tothe origin of the organisms, both contaminated material as well as materialwithout known history of contamination with chloroethenes has been the


Fig. 79. Proposed degradative pathway for PCE

source. Dehalobacter restrictus strain PER-K23 was isolated from an anaerobicfixed-bed column fed with lactate and PCE [737, 803]. However, while PCE hadbeen completely dechlorinated to ethane at high rate by the sediment fromRhine river, which has functioned as the source for strain PER-K23 [808], strainPER-K23 brings about only partial dechlorination to cis-dichloroethene.Dehalospirillum multivorans was isolated from activated sludge on a mediumcontaining pyruvate plus PCE [811], while Desulfitobacterium frappieri strainPCE-S originating from PCE contaminated soil was enriched with hydrogen,acetate, yeast extract, and PCE [738, 739]. Desulfitobacterium frappieri strainTCE1 was isolated from a PCE-dechlorinating chemostat mixed culture thathad been enriched by using soil obtained from a chloroethene-polluted loca-tion. Formate plus glucose were used as the electron donor [745].

The tetrachloroethene-dechlorinating bacteria isolated so far belong to fivephylogenetically different groups of bacteria. Physiologically they range fromfacultative anaerobes, nitrate reducers, and sulfoxy anion reducers to strict tetrachloroethene reducers. Five isolates (Desulfitobacterium sp. strain PCE1,Desulfitobacterium frappieri strains PCE-S and TCE1, isolate TEA, and PER-K23) relate phylogenetically to the gram-positive bacteria with a low DNA G+Ccontent. Strain MS-1, a facultative anaerobe, belongs to the Enterobacteriaceae.Dehalospirillum multivorans is a member of the e-subdivision of the Proteo-bacteria and Desulfuromonas chloroethenica strain TT4B of the d-subdivision,while the classification of Dehalococcoides ethenogenes strain 195 is unclear.

Strain MS-1 is able to oxidize a broad spectrum of substrates. Dehalobacter re-strictus strain PER-K23, isolate TEA, and Dehalococcoides ethenogenes strain 195,on the other hand, can only utilize hydrogen as the electron donor. The other te-trachloroethene dechlorinators can use two to six different electron donors.

Desulfitobacterium sp. strain PCE1 dechlorinates tetrachloroethene mainlyto trichloroethene and is physiologically much more versatile than isolate TEAand Dehalobacter restrictus. On the basis of electron donors and electron ac-ceptors utilized, strain PCE1 has much more in common with Dehalospirillummultivorans.

Dehalospirillum multivorans and Dehalobacter restrictus show similarities re-garding the tetrachloroethene dechlorination. Both organisms dechlorinate te-trachloroethene to cis-1,2-dichloroethene and can couple this reaction to hy-drogen oxidation, and both contain b-type cytochromes and menaquinones thatare possibly involved in electron transfer [812]. The purified tetrachloroethenereductive dehalogenase of Dehalospirillum multivorans and the tetrachloro-ethene reductase of Dehalobacter restrictus are corrinoid-containing enzymes[813, 814]. A major difference between the two organisms has been found in thelocalization of the tetrachloroethene-reducing enzyme. While the tetrachloro-ethene reductive dehalogenase of Dehalospirillum multivorans is localized in thecytoplasmic fraction [811, 815], the tetrachloroethene reductase of Dehalobacterrestrictus is membrane-bound [812]. The reductase of D. multivorans has beenpurified and characterized and its gene has been cloned [816, 817].

The ability of both strains to grow on mineral medium with H2 and PCE assole energy source gave evidence that the reductive dechlorination of PCE inthese organisms is coupled to the synthesis of ATP. Since neither the oxidation

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of H2 nor the reductive dechlorination of PCE is mechanistically coupled tosubstrate level phosphorylation, ATP synthesis must proceed via chemiosmoticmechanism. The two enzymes, hydrogenase and dehalogenase, catalyze thefollowing overall reaction:

PCE + 2 H2 Æ cis-1,2-DCE + 2 H+ + 2 Cl– DGo¢ = –376 kJ/mol

During the oxidation of H2, protons are produced at the outside of the cyto-plasmic membrane. PCE-reduction in the cytoplasm uses protons. Only due tothe consumption of protons inside and the production outside an electro-chemical proton potential will be created, which will be used for the conserva-tion of energy by the proton-translocating ATP synthetase. The participation ofa proton pump from the inside to the outside seems to be non-essential in thistype of respiration. A simplified scheme of the process is given in Fig. 80.

5.5Résumé: Chloroaliphatic Compounds

Chloroaliphatic compounds are the subject of degradation by microorganismsin different ways. Some of them can serve as growth substrates for aerobic bac-teria. A set of different dechlorination mechanisms is known to be involved.Since for highly chlorinated compounds such as tetrachloro- and trichloro-ethene, both are well known pollutants, no aerobic populations are available, thecometabolic potential of various aerobic systems has been studied especially forthe application in the cleanup processes. Besides degradation in the presence ofoxygen, under anaerobic conditions mineralization has been shown.Fermentation as well as degradation under methanogenic conditions is known.


Fig. 80. Simplified scheme of the energy conservation during the reductive dechlorination oftetrachloroethene with H2 as the electron donor (modified with information from [17, 818])

In a couple of recently isolated bacteria a chloroaliphatic compound such astetrachloroethene can serve as an electron acceptor in a respiration leading toreductive elimination of chlorine substituents.

6Sequential Anaerobic-Aerobic Processes for the Degradation of Problematic Compounds

Aerobic biological processes are widely employed as cost-effective, reliable sys-tems for the removal of organic material from municipal wastewater. In addi-tion, the use of sequential anaerobic and aerobic processes allows the removalof nitrogen and phosphorus from the wastewater. The question arises whethersequential environments might also be a possibility for the mineralization ofsome man-made organic compounds mentioned in the former paragraphs andfound to be recalcitrant. Considerable interest exists to develop systems to over-come the recalcitrance.

Separate aerobic and anaerobic environments each have limitations in theirbiodegrading abilities, but they often complement each other when they arecombined. One limitation of aerobic processes involves the recalcitrance of highlychlorinated chemicals, such as hexachlorobenzene, tetrachloroethene, and carbontetrachloride. Quite the opposite is true for the reductive dechlorination reactionscatalyzed by some anaerobes. While under aerobic conditions the persistence ofchlorinated compounds generally increases with increasing chlorine substitutionand this may enhance anaerobic degradation. However, in many cases the reduc-tive dechlorination by anaerobic bacteria is incomplete but yields less-halogenat-ed compounds, which may be efficiently degraded by aerobic bacteria.

Theoretically, sequential anaerobic-aerobic systems can be developed for bothhighly chlorinated aliphatic and aromatic compounds. The first candidates forthese systems are tetrachloroethene and carbon tetrachloride. Initial anaerobicstages may accomplish reductive dechlorination, producing trichloroethene andchloroform. Subsequent aerobic, methanotrophic stages may convert trichloro-ethene and chloroform to carbon dioxide and water. Alternatively, anaerobic re-ductive dechlorination may produce vinyl chloride and chloromethane, whichmay degrade in conventional aerobic processes if volatilization losses are mini-mized.Another example of sequential anaerobic-aerobic processes is the minera-lization of chlorinated aromatic compounds such as hexachlorobenzene andPCBs. Reductive dechlorination may occur in anaerobic stages, producing lesschlorinated congeners, which may be degraded under aerobic conditions.

Indeed, the studies of anaerobic-aerobic systems performed thus far havefocused on the degradation of chlorinated compounds, and anaerobic reductivedechlorination followed by aerobic degradation of the less chlorinated productshas been realized. Studies with non-classified indigenous microflora of environ-mental materials were performed. Besides biostimulation, i.e., the addition ofsubstrates to help the native organisms, bioaugmentation has been studied,which involves the supplementation of known bacterial populations to the indi-genous populations. In addition, an artificial system consisting of two bacterialstrains has been investigated towards sequential anaerobic-aerobic degradation.

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6.1Studies with Environmental Materials

Tetrachloroethene, chloroform, and hexachlorobenzene have been degraded ina two-stage biofilm reactor consisting of an anaerobic column followed by aconventional aerobic one [819]. Reductive dechlorination occurred in the an-aerobic column, and trichlorinated and dichlorinated products were formed. Inthe aerobic column the lesser chlorinated intermediates were substantiallytransformed into carbon dioxide and nonvolatile products. The two-stage pro-cess resulted in 61%, 49%, and 23% mineralization of chloroform, tetrachlo-roethene, and hexachlorobenzene, respectively. Dechlorination was most exten-sive when acetate served as the primary substrate, but it occurred to a lesser ex-tent also when glucose and methanol served as primary substrates.

Gerritse et al. [576] investigated the degradation of tetrachloroethene by com-bining the abilities of anaerobic bacteria, capable of reductive dechlorination oftetrachloroethene, with those of aerobic methanotrophic bacteria, capable of co-metabolic degradation of the less-chlorinated ethenes formed by the reductivedechlorination of tetrachloroethene. It was demonstrated that complete degra-dation of tetrachloroethene was possible by combining two columns in series.An anaerobic community reductively dechlorinating tetra-, trichloro-, and di-chloroethenes was used for inoculating an anoxic fixed-bed upflow column thatsubsequently converted tetrachloroethene mainly to cis-dichloroethene. Theoxic fixed-bed downflow column contained aerobic methanotrophic bacteriathat grew with methane and cometabolized the less-chlorinated ethenes formedin the anoxic column. The sensitivity of the methanotrophic bacteria to chlori-nated intermediates represented the bottleneck in the sequential anoxic-oxic de-gradation process of tetrachloroethene. In a similar approach, the second oxicstep, bringing about the dechlorination of the products of the partially anoxicdechlorination of tetrachloroethene, is substituted by the cometabolic reactionsof aerobic phenol-grown bacteria [820]. The maximum capacity for chloroethe-nes degradation was significantly higher than reported thus far.

In a simpler approach the simultaneous anaerobic and aerobic degradationof tri- and tetrachloroethene has been demonstrated by Enzien et al. [821].Under aerobic conditions, a column containing three sediments from differenthorizons from the Savannah River site was run with groundwater of an uncon-taminated well supplemented with methane, oxygen, methanol, and the chloro-aliphatic compounds. About 90% removal of tri- and tetrachloroethene was ob-served when comparing the feed and the exit water. Enumerations of the micro-bial populations in the column indicated the presence of both aerobic andanaerobic populations throughout the experiment of more than one year.Methanotrophs were detected at low numbers. The presence of methanogenssuggested that anaerobic zones or microsites existed, allowing the simultaneouspresence of both aerobic and even strict anaerobic microorganisms. These re-sults may have important implications for in situ and on-site tri- and tetrachlo-roethene bioremediation projects.

Other compounds have been successfully mineralized under sequential an-aerobic-aerobic conditions. C14-labeled 2,2,2-trichloro-1,1-bis(p-methoxy-


phenyl) ethane (methoxychlor) was degraded by bacteria that were initially in-cubated for three months under anaerobic conditions and subsequently incu-bated under aerobic conditions [822]. Cultures exposed to the sequence of en-vironments showed a 10- to 70-fold increase in labeled carbon dioxide produc-tion compared with cultures maintained under aerobic conditions only. At allconcentrations of methoxychlor studied, the anaerobic-aerobic system pro-duced significantly more labeled carbon dioxide than did the purely aerobicsystem.

Results have been obtained from investigations describing sequential an-aerobic-aerobic processes for the destruction of PCBs in river sediment. Thebiotransformations that occurred when a mixture of PCB congeners (Aroclor1242) in sediment is incubated under anaerobic conditions and then underaerobic conditions have been described [823]. Methanol was added as a primarysubstrate. During the anaerobic period, reductive dechlorination occurred, andthe mass of tri-, tetra-, penta-, and hexachlorobiphenyl decreased, whereasmono- and dichlorobiphenyl concentrations increased. Under subsequentaerobic conditions, significant degradation of all mono- and dichlorobiphenylcongeners occurred. Still, 43% of the 300 mg of PCBs/kg of soil initially addedremained after treatment. The authors also describe a conceptual model inwhich the in situ mineralization of PCBs may be accomplished by injectingmethanol or other primary substrates into river sediment, monitoring the ex-tent of dechlorination, and finally injecting hydrogen peroxide and methanol tostimulate aerobic mineralization of less chlorinated homologues.

6.2Studies with Undefined Enrichment Cultures

A single microbial population enriched from anaerobic sludge allowed the de-gradation of 2,4,6-trichlorophenol by cycling between anaerobic and aerobicconditions [824, 825]. In the first step, 4-chlorophenol was the most significantproduct formed from the target compound during incubation with diluted di-gester fluid. The anaerobic population was subsequently transferred to aerobicconditions, resulting in a very slow mineralization. The results indicate thatthere is no need for a second microbial population to achieve the goal of min-eralization of the target compound. Instead the addition of oxygen to the an-aerobic population to shift facultative organisms to aerobic degradation path-ways is sufficient to bring about mineralization. An increasing rate of dehaloge-nation was observed at high pH under anaerobic conditions, while neutral pHwas essential for the aerobic step. Therefore the sequential anaerobic-aerobicprocess has to involve a pH adjustment. The population was found to be robustand resistant to fluctuations in pH.

6.3Studies with Undefined Enrichment Cultures Supplemented with Pure Cultures

The sequential degradation of 2,3,6-trichlorobenzoate using anaerobic andaerobic organisms was reported by Gerritse and Gottschal [826] (Fig. 81). A

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2,3,6-trichlorobenzoate dechlorinating methanogenic enrichment culturegrowing with medium containing yeast extract, peptone, benzoate, and a fattyacid mixture (acetate, propionate, butyrate, 2-methylbutyrate, isobutyrate, vale-rate, and isovalerate) was cultivated anaerobically in a chemostat in which anylon bag filled with particles of the clay mineral vermiculite was placed to pre-vent wash-out of bacteria. 2,5-Dichlorobenzoate was produced under these con-ditions. Aeration of the reactor neither brought about further degradation norkilled the methanogenic culture in the bag. The addition of a culture ofPseudomonas aeruginosa JB2, which is able to grow aerobically with 2,5-dichlo-robenzoate [387], resulted in total degradation of 2,3,6-trichlorobenzoate. Thefact that reductive dechlorination of 2,3,6-trichlorobenzoate and oxidative min-eralization of 2,5-dichlorobenzoate occurred simultaneously in the aerated re-actor and resulted in almost complete mineralization of 2,3,6-trichlorobenzoateshows that the strictly anaerobic and aerobic bacteria can successfully be com-bined at low oxygen tensions. O2 concentrations in the nylon bag with vermi-culite were much lower than in the liquid phase, thus ensuring suitable growth


Fig. 81. Scheme of chlorobenzoate degradation in the mixed culture reactor under aeratedconditions. 2,3,6-Trichlorobenzoate enters the vermiculite where reductive dechlorinationtakes place. The anaerobic dechlorination product 2,5-dichlorobenzoate is degraded furtherby Pseudomonas aeruginosa once it has reached the microaerobic liquid phase

conditions for strict anaerobes. A biological purification system harboring ha-bitats for both anaerobic and aerobic bacteria is thought to have advantagesover systems which are only (or sequentially) aerobic or anaerobic: inactivationor death of aerobes or anaerobes due to the periodic absence or presence of O2,respectively, is prevented.

In a soil slurry microcosm study, Evans et al. [827] investigated the degrada-tion of weathered Aroclor 1248, i.e., decreased levels of trichlorophenyls com-pared to the original congener mixture, in historically contaminated soil with alow organic carbon content. The PCB concentration was approximately 100 mg/kg dry soil. Three systems were studied. The sandy soil was inoculated withPCB-dechlorinated microorganisms from Hudson River sediment. In a secondincubation strain LB400 (Burkholderia cepacia LB400), an organism with a highpotential for the aerobic transformation of complex Aroclor mixtures, was ad-ded plus the supplementation of biphenyl as the growth substrate. The effi-ciency of a sequential anaerobic-aerobic scheme was tested in a third incuba-tion. The aerobic treatment alone proved quite effective in reducing the totalPCB concentration by 67% after 19 weeks, leaving mainly tetra- and pentachlo-robiphenyls. The sequential anaerobic-aerobic incubation after 19 weeks show-ed a reduction of about 70% of the total PCB concentration. A further dechlori-nation was not shown for the next 60 weeks. The higher efficiency in compari-son to the study by Anid et al. [823] was thought to be due to the greater abilityof strain LB400 to degrade trichlorobiphenyls.

Shannon et al. [828] reported that more than 80% of the PCBs from a 1240-ppm Aroclor 1248-contaminated sediment was biodegraded using a two-stageanaerobic/aerobic microbial system. However, few details of the procedure werepresented. Comparing the differences in the Shannon and Evans studies, thegreater degradation noted by Shannon et al. may be contributed to the presenceof easily degraded trichlorobiphenyls which are absent in the weathered soil in-vestigated by Evans et al. [827].

6.4Studies with Pure Cultures

Beunink and Rehm [829] developed an anaerobic-aerobic process by immobi-lizing two strains in calcium-alginate beads to degrade 4-chloro-2-nitrophenol(Fig. 82). The conversion of 4-chloro-2-nitrophenol by Enterobacter cloacae,growing with glucose as the substrate, led to the formation of 4-chloro-2-ami-nophenol plus minor amounts of 4-chloro-2-acetaminophenol. The main reac-tion product was further mineralized under aerobic conditions by anAlcaligenes sp. strain TK-2. Whereas both degradative steps excluded one an-other in homogenous systems with free cells, a coupled reductive and oxidativedegradation took place in an aerated reactor system with alginate beads. Theouter bead region was an aerobic environment, while the inner bead regionswere anaerobic because of the slow diffusion of oxygen in the beads and theconsumption in the outer bead region.

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6.5Résumé: Sequential Anaerobic-Aerobic Processes

The examples presented clearly show the potential of sequential anaerobic-aerobic microbial processes in the degradation of xenobiotic compounds. Thesemay be carried out in two-stage reactor systems or synchronously in a single re-actor where both anaerobic and aerobic sites occur. The idea of sequential an-aerobic-aerobic microbial degradation can also be applied to groundwatercleanup by manipulating the environmental conditions, additions of substratesto stimulate special bacteria of the indigenous populations, or the injection ofeffective populations.

However, much work remains to be done before sequential conversions areemployed to their fullest potential.

7Enhancement of the Catabolic Potential of Microbial Strains in the Laboratory

In a number of cases, a great deal is known about the molecular changes involv-ed in the alteration of existing metabolic capabilities, resulting in the selectionof mutant strains with the ability to grow on novel growth substrates.

I will describe with a few examples why a particular pollutant may not support growth of a single microbial species and the methods used to eliminatethe respective bottleneck. Mutagenesis, transfer of genetic information, andgenetic engineering techniques will be discussed. The compounds of interest,


Fig. 82. Model of the reductive and oxidative degradation steps in a Ca-alginate bead for im-mobilization of a mixed culture to mineralize 4-chloro-2-nitrophenol

where data are available, include aromatic, haloaromatic, and chloroaliphaticcompounds and their intermediates.

Figure 83 schematically illustrates different steps which might form a bottle-neck for a target compound.

7.1Uptake of a Target Compound

In principle, the cell membrane, consisting of a bimolecular phospholipid layer,is impermeable to hydrophilic substances, and therefore bacteria have a varietyof specific systems by which hydrophilic compounds of biological importance,such as carbohydrates or amino acids, are transported into the cell. Lipophiliccompounds, however, can pass the membrane by simple diffusion. Halogenatedaromatic compounds are generally more apolar, depending on the number ofhalogen substituents, than the unsubstituted analogues. Therefore, if the halo-gen-free compound is acceptable as a growth substrate entering the cell by dif-fusion, one can assume that the halogen compound will permeate as well.

Besides permeation by a simple diffusion process, specific transport systemshave been implicated for aromatic compounds and metabolites. Evidence for aninducible active transport of mandelate and benzoate has been reported [830,831]. The aromatic compounds enter the cells by a facilitated diffusion process[832]. Later the transport of benzoate, 4-hydroxybenzoate, and protocatechuatevia an inducible permease was reported [833–837]. Recently, the transport oflow concentrations of 2,4-D was shown to be dependent on the presence of atransport protein [838].

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Fig. 83. Scheme summarizing the steps which might be responsible for recalcitrance present-ed for a chloroaromatic compound. � Uptake into the cell, � induction (effector specificity),� conversion by enzymes (enzyme specificity), � formation of toxic products

The necessity of permeability mutants and transport systems for the use ofthe polar metabolites 3-oxoadipate, cis,cis-muconate, and g-carboxy-cis,cis-mu-conate as the growth substrate has been documented [839–843].

In general, information concerning the mechanisms for the production oftransport mutants is rare. Overall, the transport into the cell seems to be abottleneck only for ionized compounds.

7.2Expansion of the Effector Specificity of Transcriptional Regulators

Biological activities that mineralize pollutants generally consist of multisteppathways. Degradation of simple compounds such as toluene can involve ten ormore enzymatic reactions. The expression of the enzymes is carefully control-led by multicomponent regulatory networks.

One mechanism for enhancing the range of inducers is the alteration of tran-scription circuits of catabolic operons. The genes of catabolic pathways aretypically organized in operons that assure coordinated synthesis of the com-ponent enzymes of the pathways. Transcription of such operons from theoperon promoters is generally controlled by positively acting regulatory pro-teins that are activated by pathway substrates or metabolites, i.e., the effectors.

Among the genes for degradative pathways the xyl genes, laying on the TOLplasmid pWW0, are the best characterized ones. These genes, which are essen-tial for the total degradation of toluene and xylenes, form two functional clus-ters, the so-called upper and the meta operon. A simplified model is given inFig. 84.

The “upper” operon xylCMABN encodes three enzymes which oxidize tol-uene and xylenes to benzoate and toluates, respectively. The promoter of theupper pathway genes, Pu, is positively regulated by the regulatory gene xylR.This gene encodes a protein which enhances transcription after the binding ofan inducing molecule (toluene, m-xylene, and the respective benzoates).Subsequently, benzoate and toluate are transformed to the respective catecholsand mineralized by the enzymes of the meta pathway, encoded by the genes ofthe meta operon. The meta operon contains the genes xylXYZ and xylL, which


Fig. 84. Simplified model of the regulation of xyl gene operons (xyl, xylene). The upperoperon (xylCMABN) codes xylene monooxygenase, benzylalcohol and benzaldehyde de-hydrogenase, while 13 enzymes are encoded by the meta operon (xylXYZLTEGFJQKIH),including toluate 1,2-dioxygenase, toluate dihydrodiol dehydrogenase and catechol 2,3-di-oxygenase. The regulator genes xylS and xylR are shown in gray. The promoter regions aremarked by small boxes. The arrows indicate induction by XylR and XylS regulatory protein inconcert with the respective aromatic effectors. Data from [844–850]

encode the enzymes toluate 1,2-dioxygenase and toluate dihydrodiol dehydro-genase. Not less than nine enzymes are co-ordinately expressed in the polycis-tronic meta operon from the Pm promoter. For efficient expression of the metaoperon by the Pm promoter, the product of the xylS gene is needed. Inducingcompounds like benzoate and toluate bind to XylS, bringing it to an activatedform.

Studies on mutants with expanded substrate range and induction patternshave shown that the xylS gene is the target for adaptive mutations. Some ben-zoate analogues that can be metabolized by the enzymes of the pathway, such astoluate dioxygenase and toluate dihydrodiol dehydrogenase, fail to induce syn-thesis of these enzymes. Instead, these non-effector benzoate analogues compe-titively inhibited effector-mediated activation of the regulator protein. Thesecompounds clearly interacted with the effector-binding site of the regulator butfailed to establish the productive contacts needed to induce the conformationalchange leading to activation of the protein. It was relatively easy to isolate mu-tant bacteria producing regulators that were activated by such benzoate analo-gues [849]. A mutant of the XylS regulatory protein can mediate three- to eighttimes higher levels of transcription than the wild type regulator [851].

Subsequent studies showed that selective elevation of XylS expression and anincrease in the intracellular level of XylS can be obtained either by replacing therelatively weak native promoter with a stronger one or by increasing xylS genedosage. This results in a substantial increase in Pm activation [852, 853].

Selective changes in the –10 region of the Pm promoter can also increasetranscription of the meta pathway enzymes in a fully regulated manner [854].

Other mechanisms which have been shown to expand the substrate range ofcatabolic pathways include activation by insertion elements [855, 856]. van derPloeg et al. [855] showed that the substrate range of a chloroacetate-utilizingstrain of Xanthobacter autotrophicus can be expanded to include bromoacetateby spontaneous insertion of an insertion element, which copies itself from an-other position on the chromosome to a site closely in front of the dehalogenasegene. This leads to a five- to tenfold increase in expression.

By judicious manipulation of regulatory proteins and their levels of expres-sion and of the structure of the cognate promoters of the regulators, one canachieve very high levels of expression of catabolic operons and create effectivedegradative organisms. It has been shown that the substrate range can be ex-panded. However, from a practical point of view it should be noted that most ofthe “new” substrates such as 4-ethylbenzoate can also be degraded by naturallyoccurring bacteria that can be enriched from polluted environmental samples.

7.3Alterations in Structural Genes

7.3.1Widening of the Substrate Range

An excellent example of mutations in a structural gene that alter enzyme speci-ficity to allow a strain to grow with a novel substrate is given by Pries et al. [857].

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They expressed the haloalkane dehalogenase (DhlA) of Xanthobacter autotro-phicus that hydrolyses short-chain (C2–C4) 1-chloro-n-alkanes to the corre-sponding alcohols in a strain of Pseudomonas that grows on long-chain alco-hols. Different spontaneous mutants were selected able to use 1-chlorohexaneas the growth substrate. All the mutants showed alterations, deletions, pointmutations, or tandem repeats, in a distinct region of the dehalogenase, the so-called cap domain. The kinetic constants indicate that the mutants are betteradapted to use the novel substrate than the wild type but lost some efficiencyagainst the original substrate 1,2-dibromoethane. Pries et al. [857] found thatthe generation of duplication was a common event during adaptation to 1-chlorohexane. Since duplications were also present in the cap domain of wildtype dehalogenase they suggest that the wild type enzyme has undergone re-cent adaptive mutations leading to utilization of 1,2-dichloroethane.

Besides these natural processes, site-directed mutagenesis allows the con-struction of enzyme variants that exhibit broader substrate specificities.Biphenyl dioxygenases catalyze the first step in the aerobic degradation of chlo-rinated biphenyls. The nucleotide and amino acid sequences of the biphenyl di-oxygenases from Pseudomonas sp. strain LB400 and Pseudomonas pseudoalca-ligenes KF707 were found to be nearly identical, yet these enzymes exhibiteddramatically different substrate specificities for chlorinated biphenyls (LB400broad substrate spectrum, KF707 narrow substrate spectrum). Site-directedmutagenesis of the LB400 bphA gene, encoding the large subunit of the termi-nal dioxygenase, resulted in an enzyme combining the broad congener specifi-city of LB400 with an increased activity against several congeners, includingdouble para-substituted ones, which is characteristic for KF707 dioxygenase[858]. The mutagenesis procedure altered a region of the LB400 bphA gene en-coding a block of four amino acids, which differ from those of strain KF707. Theregion of amino acids 335 to 341 in LB400 BphA (TFNNIRI) was converted tothe corresponding KF707 sequence (AINTIRT) at the underlined positions.This multi-amino acid modification brought about the greatest improvement in activity. The novel dioxygenase combined the broad congener specificity ofLB400 with the increased activity against several congeners characteristic ofKF707.

Later, Mondello et al. [859] examined the BphA sequences from a variety ofbacteria whose PCB substrate specificities differ from that of strains LB400 andKF707 but which contain related bphA genes. With that database four regionswere identified in which specific sequences were associated with either broad ornarrow PCB substrate specificity. The most important one was region III whichhas been modified in the Erickson and Mondello study [858]. Based on these as-sociations, site-directed mutagenesis was used to alter specific regions of theLB400 bphA nucleotide sequence and the effects of these mutations on PCBsubstrate specificity were determined. The most important region again was re-gion III, but also a modification of one amino acid in region IV can contributeto the broader substrate specificity.

Similarly, Kimura et al. [860] produced chimeric enzymes from the LB400and KF707 dioxygenases combining the substrate range of both parental en-zymes by exchanging restriction fragments.


Besides the rational, site-directed approaches, which allow the exploration ofonly very limited sequence space at a time,“irrational” approaches such as DNAshuffling, a technique using PCR [861], can be alternatives for the production ofenzymes with altered substrate specificity. In two independent studies [862,863] the substrate range of biphenyl dioxygenases towards PCB congeners havebeen successfully extended using the random cross-breeding of the genes fromstrain LB400 and KF707 by DNA shuffling. One major advantage of the in vitroDNA shuffling of enzymes over the structural remodeling by site-directed mu-tagenesis is that only a minimum of prior information is required.

7.3.2Mutations in Structural Genes to Avoid the Formation of a Toxic Metabolite

Pseudomonas sp. strain B13, which is able to use 3-chlorobenzoate as the growthsubstrate but fails to use 2-fluorobenzoate, was adapted to growth with the lat-ter compound over a period of six months in a chemostat in which 3-chloro-benzoate was stepwise replaced by 2-fluorobenzoate [397]. The resulting strainB13-1 was then cultivated in sequential batch cultures for 250 generations with2-fluorobenzoate as the sole carbon and energy source. The adapted strain B13-2, growing at high rate with the new substrate, degrades 2-fluorobenzoatefor 95% via pathway a, with oxygenolytic elimination of fluoride and catechol(Fig. 85), while pathway b is not much used. Instead, strain B13 degrades 2-fluo-robenzoate for 22% via pathway b, which yields production of high amounts of3-fluorocatechol, which accumulates and negatively affects the cells so that nogrowth occurs with 2-fluorobenzoate. The improved strain B13-2 is able to grow

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Fig. 85. Alternative pathways for the degradation of 2-fluorobenzoate due to the attachmentof the substrate to the dioxygenase with substituents in the 2- or 6-position

in the presence of 2-fluorobenzoate due to a change in selectivity of the initialbenzoate 1,2-dioxygenase towards degradation via 2-fluorodihydrodihydroxy-benzoate. Therefore, only about 5% of the 2-fluorobenzoate will be converted instrain B13-2 to give the toxic metabolite 3-fluorocatechol.

A quite different strategy in avoiding the formation of a toxic metabolite wasshown to allow a mutant of Alcaligenes eutrophus to grow with 2-fluorobenzo-ate. The wild type organism uses benzoate as the sole source of carbon andenergy. 2-Fluorobenzoate will be converted via pathway a (20%), so that greatamounts of the toxic metabolite 3-fluorocatechol accumulate. The accumula-tion is avoided in the mutant B9 by a defect of the dihydrodihydroxybenzoatedehydrogenase. 6-Fluorodihydrodihydroxybenzoate will not be convertedfurther to the toxic 3-fluorocatechol. Growth resulted because of the minera-lization of catechol formed after spontaneous elimination of fluoride.

These examples show how modification of enzyme selectivity prevents mis-routing to give toxic metabolites.

7.4Use of External Genetic Information to Expand the Substrate Range

7.4.1Chlorobenzoate-Degraders by Conjugal Transfer

The first report of the development of a catabolic pathway for the mineraliza-tion of chlorinated aromatics using external genetic information for the acqui-sition of a novel phenotype described work with Pseudomonas sp. strain B13and Pseudomonas putida PaW1 and the novel growth substrates 4-chloro- and3,5-dichlorobenzoate. Strain B13 was isolated by enrichment culture with 3-chlorobenzoate. It oxidizes 3-chlorobenzoate to 3- and 4-chlorocatechol anduses the modified ortho pathway for further breakdown. Strain B13 is unable toutilize 4-chloro- and 3,5-dichlorobenzoate, since the benzoate 1,2-dioxygenasehas a very narrow specificity and will not accept 4-chloro- and 3,5-dichloro-benzoate as substrates. However, strain B13 can oxidize 4-chloro- and 3,5-di-chlorocatechol, the expected metabolites in the degradation of 4-chloro- and3,5-dichlorobenzoate. The toluate 1,2-dioxygenase in Pseudomonas putidaPaW1, which is isofunctional to the benzoate 1,2-dioxygenase, encoded by theTOL plasmid, has a broader range than the B13 enzyme and can accept 4-chloro- and 3,5-dichlorobenzoate as a substrate.

In one enrichment experiment the two organisms were grown in a chemostatinitially with 3-chlorobenzoate (substrate for strain B13) and 4-methylbenzoate(substrate for strain PaW1). After four weeks of operation, 4-chlorobenzoatewas added as an additional carbon source since it could not support the growthof either organism. After another four-week period, the culture was switched toonly 4-chlorobenzoate and during the next twelve weeks 3,5-dichlorobenzoatewas added and colonies able to grow on 3,5-dichlorobenzoate were isolated.Eventually, Pseudomonas sp. strain WR912 was isolated and shown to be cap-able of growth on 3-chloro-, 4-chloro-, and 3,5-dichlorobenzoate [864]. Thecomplexity of this experiment, with the prolonged selection period, made it dif-


ficult to interpret, but one of the predictions was that strain B13 would acquirethe toluate 1,2-dioxygenase coded by the TOL plasmid of Pseudomonas putidaPaW1. That a novel catabolic pathway could be developed in this way was laterconfirmed by direct transfer experiments, but the results were unexpected[865–867]. Transconjugants from a mating between Pseudomonas sp. strain B13and Pseudomonas putida PaW1 were obtained on plates containing 4-methyl-benzoate to select for TOL plasmid transfer and streptomycin to contraselectagainst the Pseudomonas putida parent. Strain WR211 had the phenotype ofstrain B13 with the additional ability to grow on 3- and 4-methylbenzoate, butwas unable to grow on 4-chlorobenzoate. Strain WR211 was plated on 4-chloro-benzoate and gave rise to strains, such as WR216, which had gained the abilityto utilize 4-chlorobenzoate but had lost the ability to utilize the methylbenzoa-tes. 4-Chlorobenzoate utilizers were derived directly from other plate matings.Plasmid transfer by itself was clearly inadequate for the development of thenovel pathway.

The TOL plasmid of Pseudomonas putida PaW1 is known to undergo variousrearrangements of its DNA. In the development of the 4-chlorobenzoate utilizerit was suggested that the events are as follows [868]:1. Transfer of TOL plasmid into strain B13.2. Integration of a 56-kb segment of TOL DNA into the chromosome.3. Deletion of a 39-kb segment from TOL.4. Insertion of a DNA segment of about 3 kb into the xylE gene, the gene encod-

ing the catechol 2,3-dioxygenase.

For the selection of the 4-chlorobenzoate derivatives of WR211 it was essentialthat the meta pathway is inactivated. The catechol 2,3-dioxygenase has a highaffinity for 4-chlorocatechol, which is channeled into the meta cleavage path-way, resulting in the production of dead-end metabolites. Strain WR216 has no catechol 2,3-dioxygenase activity and 4-chlorocatechol is catabolized via theortho cleavage pathway.

In addition to the bottlenecks concerning the turnover of substrates, some-times the induction of a pathway by the novel compound does not take place.One example for this fact is 3,5-dichlorobenzoate. While 3,5-dichlorobenzoatefails to induce the toluate 1,2-dioxygenase in the strains WR211 and WR216, thecompound is an effector in the strains which have occurred on solid mediacontaining 3,5-dichlorobenzoate from the respective origins.

7.4.2Chloronitrophenol-Degraders by Conjugal Transfer

The chlorocatechol degradative genes of strain B13 and JMP134 have also beenused to obtain strains able to grow with 4-chloro-2-nitrophenol (Fig. 86).Pseudomonas sp. N31, isolated with 3-nitrophenol and succinate as sole sourceof nitrogen and carbon, respectively, expresses a nitrophenol oxygenase elimi-nating nitrite from 4-chloro-2-nitrophenol to produce 4-chlorocatechol.Conjugal transfer of the genes coding the modified ortho pathway from B13 orJMP134 into strain N31 allowed the isolation of hybrid strains such as N31-1

128 W. Reineke

able to use 4-chloro-2-nitrophenol as sole source of carbon, nitrogen, andenergy [869].

7.4.3Chlorobiphenyl-Degraders by Mating Three Strains

Some chlorobiphenyls fail to be growth substrates because the peripheral en-zymes do not convert them to the chlorocatechols. In these cases a “novel” path-way has to be constructed in one organism by segments from at least three or-ganisms. Such a mating is summarized schematically in Fig. 87 for various chlo-robiphenyls. However, the nature of some events which have to take placeduring the development is presently unknown.

7.4.4Other Chloroaromatic-Degraders by Conjugal Transfer

The development of hybrid pathways by DNA transfer with whole degradativeplasmids has been demonstrated for various other mono- and dichlorosub-stituted aromatics (Fig. 88). The procedure is superior due to its technical sim-plicity and effective positive selection. Hybrid strains can be developed by thewell-aimed mating within weeks, since the organisms with the suitable pathwaysegments are inoculated together on solid media, making a gene transfer easy.However, the selection steps have to be done in the right order. It is importantto establish the chlorocatechol degradative sequence at the beginning of the de-velopment of the hybrid strains, since otherwise the accumulated chlorocate-chols will harm the cells.

Various chloroaromatic-degrading strains have been isolated by enrichmenttechniques, such as chlorobenzenes, chlorophenols, and chlorophenoxyacetatesdegraders. An inspection of the pathway in some strains indicated that they arealso a product of the patchwork assembly described above.

In principle, similar hybrid strains can also be constructed using geneticengineering techniques. The major disadvantage is the huge amount of researchnecessary prior to the in vitro construction experiment leading to the hybridstrains. Cloning of structural as well as regulatory genes has to be done follow-ed by establishment in a suitable host.


Fig. 86. Pathway for 4-chloro-2-nitrophenol in the hybrid strain N31–1

7.4.5Chlorobenzoate- and Chlorosalicylate-Degraders by Genetic Engineering Techniques

Using genetic engineering techniques, chlorobenzoate-degrading strains wereprepared. A DNA module carrying the genes of toluate dioxygenase (xylXYZ)and of the subsequent enzyme of the pathway, toluate dihydrodiol dehydro-

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Fig. 87 a– c. Hybrid pathways for the mineralization of chlorobiphenyls in hybrid strains:a BN210, 3-chlorobiphenyl+; b KE210, 3-chlorobiphenyl+, 4-chlorobiphenyl+; c JHR22, 2-chlo-robiphenyl+, 3-chlorobiphenyl+, 4-chlorobiphenyl+, 2,4-dichlorobiphenyl+, 3,5-dichlorobi-phenyl+. The color or the hatch on the right side of the pathway characterizes the origin of therespective pathway segment in the hybrid strains: �, biphenyl degrading strains JHR (c) orBN10 (a, b); , p-toluate degrading strain PaW1; , o-toluate degrading strain WR401,� , modified ortho pathway of strain B13, and � , late 3-oxoadipate pathway segment ofPseudomonas putida strain BN10 (a, b) or Burkholderia cepacia strain WR401 (c).Information compiled from [870, 871, K. Engelberts and W. Reineke, unpublished results]

genase (xylL), plus the Pm promoter and the xylS regulatory gene, was clonedinto a broad-host-range plasmid vector and introduced into Pseudomonas sp.strain B13 [886]. The B13 derivative could grow on 3-chloro- and 4-chloro-benzoate, and synthesis of all catabolic enzymes involved in their metabolismwas fully regulated. The B13 derivative did not grow on 3,5-dichlorobenzoate,even though the catabolic enzymes present in 4-chlorobenzoate-grown bacte-ria can degrade this compound, because of the inability of the XylS regulatorto be activated by 3,5-dichlorobenzoate. However, 3,5-dichlorobenzoate be-came a substrate for the hybrid pathway when a XylS mutant regulator that isactivated by this compound was also recruited by Pseudomonas sp. strain B13[886].

Central pathways, such as the chlorocatechol ortho cleavage pathway, can beused as a base upon which to assemble additional enzymatic steps to permit the catabolism of more complex compounds. Construction of a derivative cap-able of degrading chlorosalicylates represents a simple example of verticalexpansion of the chlorocatechol pathway of Pseudomonas sp. B13. Strain B13cannot degrade either salicylate or chlorosalicylates, and such bacteria are notreadily isolated from soil. Plasmid NAH7 specifies a pathway for the catabolismof naphthalene via salicylate and catechol. Salicylate hydroxylase encoded byNAH7 exhibits a relaxed substrate specificity and oxidizes salicylate andmethyl- and chlorosalicylates to the corresponding catechols. A DNA fragmentof the NAH7 plasmid containing the gene encoding salicylate hydroxylase, itspromoter, and the regulator gene was introduced into Pseudomonas sp. B13,which thereby acquired the ability to grow on 3-, 4-, and 5-chlorosalicylates[886].


Fig. 88. Hybrid pathways for the mineralization of chloroaromatics developed by in vivo con-struction using peripheral pathway segments plus the modified ortho pathway (data com-piled from [865, 867, 870–885])

7.4.6Chlorobiphenyl-Degraders by Genetic Engineering Techniques

Hrywna et al. [887] reported that the use of sequenced and well-characterizedchlorobenzoate dehalogenase genes is an effective strategy for the constructionof hybrid organisms able to grow on ortho- and para-substituted chlorobi-phenyls. Two plasmids were introduced separately by transformation into thebiphenyl-growing, chlorinated biphenyls-cometabolizing Comamonas testo-steroni VP44. Plasmid pE43 carries the cloned gene coding for the terminal oxy-genase of the ortho-halobenzoate 1,2-dioxygenase [888], which performed oxy-genolytic ortho dehalogenation of 2-chlorobenzoate, a dead-end product in the2-chlorobiphenyl transformation by strain VP44. Hydrolytic dechlorination of4-chlorobenzoate is catalyzed by 4-chlorobenzoate dehalogenase coded bycloned genes on plasmid pPC3 [889]. The resulting hybrid strains harboringone of these plasmids expressed a dehalogenating enzyme, grew rapidly on, andcompletely dechlorinate high concentrations of 2-chloro- or 4-chlorobiphenyl,respectively (Fig. 89).

This is an example where the addition of one enzymatic reaction, missing inthe host organism, directly brings about the formation of a “normal” metaboliteof the aromatic degradation so that growth with the new compound can occur.

7.4.7Trihalopropane-Degraders by Genetic Engineering Techniques

The rational combination of catabolic segments from different organisms inone recipient strain creating a complete metabolic route has been shown for tri-halopropanes, for which mineralization has not yet been described [890]. Broadhost-range plasmids were constructed, which contained the gene coding forhaloalkane dehalogenase from Rhodococcus sp. strain M15-3, an enzyme cap-able of efficient transformation of trihalopropanes to dihalopropanols, underthe control of different heterologous promoters. Recombinant organisms wereobtained, which are able to grow on trihalopropanes, by introduction of theseplasmids into Agrobacterium radiobacter AD1, capable of utilizing dihalogenat-ed propanols for growth.

7.5Construct to Degrade TCE Without Apparent Toxic Effect

Cometabolic conversion of trichloroethene by mono- and dioxygenases of wildtype cells resulted in toxic effects which drastically reduced the conversion rate.To overcome this problem, Winter et al. [891] have introduced the gene of a to-luene monooxygenase in Escherichia coli under the control of the Trp-Lac(tac)promoter. This construct, in contrast to the situation in the wild type, was thenable to degrade TCE without apparent toxic effects.

132 W. Reineke


s133

Fig. 89. Hybrid pathways for growth with chlorobiphenyls by introduction of dehalogenase genes (for details of dehalogenating reac-tions see Figs. 36 and 39)

7.6Creation of a Pathway for the Degradation of Halogenated Alkanes and Alkenes

The design of a new pathway to carry out sequential reductive and oxidative re-actions in an organism is another example demonstrating the value of using theknowledge of catabolic enzymes and recombinant DNA technology. Hur et al.[892] and Wackett et al. [893] reported the construction of a useful metabolicpathway for the degradation of fluoro, chloro, bromo, and chlorofluoro alkanesand alkenes. Three genes coding cytochrome P-450CAM monooxygenase fromPseudomonas putida, which is able to dehalogenate reductively polyhalo-genated compounds [894], were combined with four genes coding the toluenedioxygenase of Pseudomonas putida F1 to construct the cometabolic dehalo-genation sequence of consecutive reductive and oxidative reactions. The con-version of pentachloroethane by the recombinant bacterium is given in Fig. 90.

7.7Creation of a Pathway for the Degradation of Mixtures of Methyl- and Chloroaromatics by Combination of Pathway Modules

The existence of both ortho and meta cleavage enzymes produces problems forthe conversion of a mixture of methyl and chloroaromatic compounds.Catechol and chlorocatechols are generally subjected to ortho fission, whereasmethylcatechols suffer meta fission. Although both pathways may exist in indi-vidual microorganisms, only one is usually functional at any given moment de-pending on the available substrate. However, when both chloro- and methyl-catechols are formed from mixtures of chloro- and methylaromatics, both path-way types are functional, and the catechols are subjected to both types ofcleavage. Although ortho cleavage of chlorocatechols leads to their productivemetabolism, ortho cleavage of methylcatechols leads to the formation of dead-end products. Similarly, whereas the meta cleavage of methylcatechols leads totheir productive metabolism, the meta cleavage of chlorocatechols leads to theformation of either dead-end products or products that inactivate catechol 2,3-dioxygenase, the ring cleavage enzyme. The misrouting of catechol cleavageproducts during simultaneous metabolism of chloro- and methylsubstitutedaromatics creates a sort of biochemical anarchy and eventually perturbs theproductive metabolism of aromatics [895].

Before designing a catabolic route in Pseudomonas sp. B13 for chloro- andmethylaromatics (Fig. 91) that employs only one type of catechol-ring fissionmode, the peripheral pathway was expanded [895]. Pseudomonas sp. B13 pos-

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Fig. 90. Hybrid pathway for the degradation of pentachloroethane

Fig. 91 A – C. Construction of 4-methylbenzoate degraders using the ortho cleavage pathway for the mineralization of mixtures of chloro- and methyl-substituted aromatics by combining pathway segments: A pathway segments involved in the construction: white: from strain B13, gray: segments co-ded by xylXYZL, i.e., toluate 1,2-dioxygenase and toluate dihydrodiol dehydrogenase of strain PaW1; dark-gray: 4-methyl-2-enelactone isomerase fromA. eutrophus. Full arrows indicate pathways used for growth, broken arrows those pathways able to convert compounds; B hybrid pathway for the de-gradation of 4-methylbenzoate in strain FR1(pFRC20P)-1; C genealogy of the strains. Abbreviations: 3CB, 3-chlorobenzoate; 4CB, 4-chlorobenzoate;4MB, p-toluate; 4CP, 4-chlorophenol; 4MP, 4-methylphenol

(A)

(B)

(C)

sesses only an ortho cleavage route for catechols; it can grow on 3-chloroben-zoate and acquires the ability to grow on 4-chlorobenzoate when it recruits a re-laxed substrate-specificity toluate dioxygenase through transfer of the xylXYZgenes of the TOL plasmid. To create a stable B13 derivative that can degrade 4-chlorobenzoate and partially metabolize 4-methylbenzoate, the cloned TOLgene module (xylXYZL including their promoter Pm, and xylS) was inserted intotransposon Tn5; the hybrid transposon was then transposed into the B13 chro-mosome. The derivative strain, FR1, grew on 3- and 4-chlorobenzoate and co-metabolized 3- and 4-methylbenzoate via ortho cleavage to the dead-end pro-ducts 2- and 4-methyl-2-enelactone, respectively.

Strain FR1 – like B13 – grew on 3-methyl-2-enelactone as a sole source ofcarbon and energy. Therefore, the degradation of 4-methylbenzoate by strainFR1 in principle requires recruitment only of an isomerase that converts 4-me-thyl-2-enelactone to 3-methyl-2-enelactone. The recruitment of 4-methyl-2-enelactone isomerase from Alcaligenes eutrophus which is able to transform 4-methyl- into 3-methyl-2-enelactone [896] resulted in the transformation of 4-methylbenzoate to 3-methyl-2-enelactone, which was then mineralized by B13enzymes.

This pathway was further expanded through mutational activation of thepreviously cryptic phenol hydroxylase of strain B13 to produce catechols whichallowed metabolism of 4-methylphenol exclusively via ortho cleavage. 3-Me-thylbenzoate and 3-methylphenol are no substrates, since they are mainly co-metabolized to 2-methyl-2-enelactone [897, 898], which is not further metabo-lized by B13 enzymes nor by the 4-methyl-2-enelactone isomerase cloned fromA. eutrophus.

Thus, a novel ortho cleavage pathway for the degradation of mixtures of 3-and 4-chloro- and 4-methylbenzoates and 4-chloro- and 4-methylphenol hasbeen constructed by the rational assembly of pathway modules from three dif-ferent bacteria: P. putida PaW1, Pseudomonas sp. B13, and A. eutrophus JMP134.

7.8Résumé: Enhancement of Catabolic Potential

The identification of pathways and genes have allowed an understanding of thebiochemical causes of recalcitrance and degradability of chlorinated aromaticand aliphatic compounds. Blocks have been identified and enzyme sequenceswith broader specificity have been obtained. Based on this knowledge, newpathways have been constructed. Some of them seem to be copies of thosefound in strains enriched from environmental samples. A drawback for the ra-tional design of novel microorganisms is that a lot of biochemical knowledge isneeded, which is not available for many compounds.

8Concluding Remarks and Outlook

In this chapter I have been concerned with drawing together information froma variety of sources to illustrate the current state of knowledge of microbial de-

136 W. Reineke

gradation of some natural and man-made molecules, and aliphatic and aromat-ic compounds and their chlorinated counterparts were chosen. In former timesthe chlorinated hydrocarbons were classified as being solely anthropogenic andtherefore should be attributed to be xenobiotics. However, today it is knownthat more than 2400 organohalogen chemicals are produced by living orga-nisms and over half of these contain chlorine as part of their innate molecularstructure [899]. The difference between the anthropogenic and natural organo-halogen compounds is mostly the difference in the amounts produced. Whilethe chemical production is in the range of 106 tonnes per year world-wide themajority of the natural organohalogens can be termed to be niche products.However, in contrast, some natural halogenated compounds exceed the anthro-pogenic production drastically, e.g. natural generation of chloromethane wasestimated to be 28 ¥ 106 tonnes per year [900] compared to an estimated annualindustrial world production of about 0.7 ¥ 106 tonnes [901].

There was and is one compelling reason among several why attention shouldbe focused upon the special area of microbial metabolism of haloorganics. Alarge number of these compounds either deliberately or unintentionally havefound their way into our environment as a consequence of the activity ofmodern industry and agriculture, and by persisting there for various periods oftime have caused concern to society.“We need to understand why some chemi-cals persist while others disappear” [902].

While in those days simpler answers were given, such as it is the structuralelement chlorine substituent on an aromatic ring which is responsible for therecalcitrance of these compounds, today‘s answers show a more complex pic-ture.

Biochemical reactions have to follow chemical principles: the negative in-ductive effect of halogen substituents decreased the nucleophilic character ofan aromatic ring and thus impedes the electrophilic attack of dioxygenases.However, this simple view of a chemist, when trying to explain recalcitrance ofchloroaromatic compounds, is misleading. Steric effects more drastically con-tribute to the slow down of dioxygenase reactions. In one organism a dioxyge-nase reaction might be a bottleneck with chlorosubstituted substrate analo-gues, but not a bottleneck in another one. It is hard to give predictions on thestereospecificity of an enzyme. The only simple argument which can be given isthat if an enzyme tolerates substrates with a methylsubstituent, it will also beable to deal with a chlorosubstituted substrate because of the identical size ofboth substituents. In contrast, the substitution of a hydrogen in the natural sub-strate by a chlorine substituent results in dramatic effects because of the greatdifference in size, while a fluorine substituent will be tolerated from this pointof view.

Another observation along the same lines was that a high number of chlorinesubstituents on an aromatic ring strongly reduced the electron density and hin-dered an electrophilic reaction. This implies that an aerobic degradation ofcompounds such as tetrachlorobenzene or pentachlorophenol cannot takeplace. However, this simplifying view of a chemist does not fit in with reality.Tetrachlorobenzene and pentachlorophenol are the subjects of degradation byaerobic pure cultures.


In reference to the argument that anaerobic degradation has little impact incomparison to aerobic oxidation, it should be noted that there are numerousnatural anaerobic environments. The great influence of some environmentalconditions on the degradation, such as the presence or absence of oxygen, or theavailability of electron acceptors such as nitrate or sulfate, is evident. Differenttypes of reactions occur and the rates of degradation are different under oxicand anoxic conditions. In general, anoxic microbial degradation of aromaticsseems to be of greater relevance in nature than earlier expected.

A very important explanation for the persistence of a chemical is the fact thatthe physico-chemical behavior might bring about low bioavailability as shownfor PAHs, with the result that microbial degradation cannot take place or pro-ceeds slowly. A possible way to enhance the bioavailability and the biodegrada-tion is the application of (bio)surfactants. However, recent observations showthat single surfactants can have contrasting effects on the degradation of or-ganic pollutants by different bacteria [903]. Surfactants were found to enhancethe oxidation of phenanthrene by a Pseudomonas but inhibited its oxidationand growth on various aromatic compounds by a Sphingomonas strain.Therefore there is a need for design of optimal (bio)surfactants.

Reaction sequences must make chemical sense, a view which has been ad-dressed by Dagley [902]: “Catabolic pathways are usually economical and directto the point of elegance, if that is not the case it will probably be incorrect.”Attempts to discuss common aspects of the degradative pathways kept thisstatement in mind.

The book of Gibson from 1984 [904] has elegantly summarized the state ofthe art on the subject of microbial degradation available at the beginning of the1980s. Since then considerable progress has been made in understanding themineralization of chlorinated compounds. In contrast, only smaller pieces offundamental knowledge in the field of non-substituted aliphatic and aromaticcompounds were collected such as ether cleavage of dibenzofuran and dibenzo-p-dioxin. While most of the information on the aerobic degradation was avail-able in 1984, astonishing “unexpected” results have been obtained in recentyears concerning anoxic degradation.

Degradation of aliphatic and aromatic compounds is possible without molec-ular oxygen contrary to opinion in former times. The degradation of toluene isan example of the substantial increase in knowledge in the field of the anoxicdegradation of aromatic compounds. However, also for the alkanes, which wereconsidered to be non-degradable without molecular oxygen, pure cultures haverecently been enriched which now allow biochemical investigations to elucidatethe strategy used by the organisms to activate alkanes without oxygen.

Dechlorination reactions involved in the aerobic degradation of chloroaro-matic compounds via the modified ortho pathway are now known in detail. Thehydroquinone pathway as an alternative to the modified ortho pathway hasbeen elucidated more recently.

In addition, some results which formerly brought about some generalizationhave now to be assessed more carefully: It was assumed for a long time to be im-possible to metabolize 3-chlorocatechols via the meta pathway, because the re-action product would inactivate the extradiol dioxygenase. However, a novel

138 W. Reineke

chlorocatechol 2,3-dioxygenase that can effectively cleave 3-chlorocatechol,leading to simultaneous ring cleavage and dechlorination, allows a pseudomo-nad to degrade chlorobenzene rapidly via a meta cleavage pathway.

The problems with generalizing rules and statements can be illustrated byanother example. Anaerobic bacteria have to degrade aromatic compounds byreductive conversions. Gallus and Schink [905] and Philipp and Schink [277]presented evidence that the reductive strategy for ring destabilization is not theonly one used in anaerobic degradation of aromatic compounds. Instead, an-aerobic bacteria were isolated which use oxidation rather than reduction toovercome stability of an aromatic ring.

Optimistic microbiologists of the early 1980s believed that some man-madecompounds which were considered to be persistent at that time will turn out tobe biodegradable in the future when a few microorganisms acquire the neces-sary catabolic expertise and then transmit it to others through the agency ofplasmids. I think they were right. Some new properties for instance were foundto be the result of patchwork assembly of preexisting pathway segments whichin combination function as hybrid pathways [906, 907]. Besides the naturaldevelopment and the construction in the laboratory by use of conjugation, ra-tional design of pathways for novel compounds has been shown by use ofgenetic engineering techniques.

In recent years a big change in emphasis of interest in the field of degrada-tion has taken place. So information pertaining to the genetic basis of the de-gradative pathway is widely available today, but mostly omitted in this chapter.Instead, emphasis has been placed here on the biochemical activities of micro-organisms with respect to the degradation or transformation. So most of thechapter is written with the view of a biochemical microbiologist working withpure cultures and single compounds rather than of a molecular biologist or anenvironmentalist.

I believe that traditional microbiological approaches such as the enrichmentof new organisms give the chance to recruit presently unexpected, useful reac-tion sequences, for instance in the field of anaerobic degradation. The naturalpool harbors a great diversity of reactions formed during nature’s evolution,which should be used and not neglected.

Some authors have felt that laboratory studies with pure cultures and singlecompounds have nothing to contribute to the solution of environmental prob-lems. Our understanding of microbial action in the environment is still in itsinfancy; there can be no doubt that these fundamental contributions from thelaboratory, including the genetic tools, are respectable and necessary startingpoints to understand the fate, survival, and activities of microorganisms in theenvironment and during bioremediation processes. Syntrophic microbial popu-lations, whose partners cannot be isolated in pure culture but which seem to beresponsible for degradation in various anoxic environments, can becharacterized with these genetic tools.

So there is much work remaining for microbiologists, biochemists, molecularbiologists, ecologists, environmental chemists, and chemical engineers toanswer the above-mentioned questions and to help in solving problems withcurrent and future pollutants.


Acknowledgements. I am grateful to Dick B. Janssen for critical reading of the first draft ver-sion of the manuscript and highly productive comments. In addition, I thank Bernd Beek forpatience and fruitful comments.

I thank all my collaborators for their enthusiastic participation in the research in my la-boratory. I wish to acknowledge support provided by the European Community and theDeutsche Forschungsgemeinschaft.

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Biodegradation of Xenobiotics in Environment and Technosphere

Tom N.P. Bosma, Hauke Harms, Alexander J.B. Zehnder

Swiss Federal Institute of Environmental Science & Technology, Ueberlandstrasse 133,CH-8600 Dübendorf, Switzerland, E-mail [email protected]

Microorganisms play an important role in the removal of synthetic organic compounds fromthe environment. This chapter gives an overview of the evolution of biodegradation pathwaysand describes the strategies that microorganisms have evolved to transform important molec-ular structures. The actual effectiveness of biodegradation in the environment is determinedby the bioavailability of the compounds.As a general rule, one could state that the release ratesof synthetic compounds should not exceed the environment’s ability to degrade them.

Keywords. Biodegradation, Xenobiotics, Pathways, Bioavailability, Technosphere, Remedia-tion, Evolution, Recalcitrance

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

1.1 Behavior of Chemicals in the Biosphere . . . . . . . . . . . . . . . 1641.2 Behavior of Chemicals in the Physical Environment . . . . . . . . 1661.3 Decontamination by Microorganisms . . . . . . . . . . . . . . . . 166

2 Biodegradation Pathways . . . . . . . . . . . . . . . . . . . . . . . 168

2.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1682.1.1 Evolution of Metabolic Pathways . . . . . . . . . . . . . . . . . . . 1692.1.1.1 Events in Vertical Evolution . . . . . . . . . . . . . . . . . . . . . . 1692.1.1.2 Events in Horizontal Evolution . . . . . . . . . . . . . . . . . . . . 1722.1.1.3 Generation of Novel Degradative Pathways . . . . . . . . . . . . . 1722.2 Fate of Substituents . . . . . . . . . . . . . . . . . . . . . . . . . . . 1742.2.1 Removal of Halogen . . . . . . . . . . . . . . . . . . . . . . . . . . 1742.2.1.1 Nucleophilic Substitution of Halogen . . . . . . . . . . . . . . . . 1772.2.1.2 Dehydrodehalogenation . . . . . . . . . . . . . . . . . . . . . . . . 1782.2.1.3 Oxidative Dehalogenation . . . . . . . . . . . . . . . . . . . . . . . 1782.2.1.4 Reductive Dehalogenation . . . . . . . . . . . . . . . . . . . . . . . 1792.2.2 Nitro Groups . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1812.2.2.1 Nitroaromatic Compounds . . . . . . . . . . . . . . . . . . . . . . 1812.2.2.2 Nitrate Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1832.2.3 Sulfonic Acid Groups . . . . . . . . . . . . . . . . . . . . . . . . . . 1832.2.4 Organophosphonates . . . . . . . . . . . . . . . . . . . . . . . . . . 1842.3 Degradation of the Carbon Backbone . . . . . . . . . . . . . . . . 1842.3.1 Aliphatic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . 1842.3.1.1 Aerobic Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1852.3.1.2 Anaerobic Hydrocarbon Degradation . . . . . . . . . . . . . . . . 185

CHAPTER 2


2.3.2 Aromatic Rings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1872.3.2.1 Aerobic Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1872.3.2.2 Anaerobic Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . 1912.3.2.3 Fungal Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . 1922.3.3 Ethers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1922.3.4 Chiral Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . 1942.3.5 Complexing Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . 1942.4 Recalcitrance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195

3 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . 196

4 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197

1Introduction

Due to extensive production and use in almost all human activities, syntheticorganic compounds are widely distributed in the environment. They enter theenvironment via waste disposal, accidental spills, application as pesticides, andvia losses during transport, storage, and use. It has been recognized since theearly 1960s that environmental pollution with low concentrations of organicchemicals is world-wide. In the last two decades, the danger of heavily pollutedsites to nature and mankind has received increased attention.

The objective of the current chapter is to present the state-of-the-art knowl-edge on the biodegradation of xenobiotics in the environment and the techno-sphere on the basis of biodegradation mechanisms that have evolved in the pastdecades. The introduction (1) will discuss the behavior of man-made chemicalsin the biosphere and the response of ecosystems on their presence. Then theevolution of new biodegradation pathways and the transformation mechanismsthemselves will be discussed (2). It appears that microorganisms use convergentstrategies to transform chemicals so that they can be channeled into existing me-tabolic pathways. The mechanisms of enzymatic conversions are discussed fromthe point of view of molecule constituents (for example, halogen or nitro groups)instead of discussing the biodegradation of certain groups of compounds (forexample, halogenated vs non-halogenated substances). The impact of environ-mental conditions on the possible biodegradation and the implications for theengineering of bioremediation are discussed, together with the factors govern-ing exposure of microorganisms to contaminants in soil and groundwater.

1.1Behavior of Chemicals in the Biosphere

The response of the biosphere to the appearance of man-made compounds canbe understood by using the concepts developed in systems ecology. The bio-sphere is characterized by the cycling of matter through the expenditure ofenergy flowing through the system and is thus able to maintain a status of highentropy. Upon their release in the environment, man-made chemicals enter the

164 T.N.P. Bosma et al.

biosphere, and may be taken up by organisms or become part of the pool ofnon-living matter (the contaminant pool, Fig. 1). Biota take up contaminants di-rectly from the contaminant pool, e.g., via leaves or the skin, or they ingest themby feeding on a lower trophic level. Organisms have systems at their disposal toexcrete or detoxify contaminants. Excretion brings contaminants back to thecontaminant pool, while detoxification results in a decontamination as indicat-ed in Fig. 1.

Plants and animals are not always able to detoxify or excrete contaminantsafter uptake. The inability of organisms to deal with xenobiotic compoundsmay have several causes. One example is the absence of appropriate enzymes totransform the compounds, another the accumulation in (animal) fat tissue be-fore excretion or enzymatic transformation has taken place. Contaminants willthen accumulate in the food chain. Accumulation is indicated by the use of dif-ferent gray shades in Fig. 1.

The population of “decomposers” (Fig. 1) is specialized in the uptake andconversion of all kinds of dead organic material, like for instance dead animalsand plant debris. Decomposers are crucial for the functioning of ecosystems be-cause they recycle nutrients back to the nutrient pool. Contaminants which areaccumulated in the tissue of organisms are recycled back to the contaminant

Biodegradation of Xenobiotics in Environment and Technosphere 165

Fig. 1. Cycling of hydrophobic organic contaminants in ecosystems. Input to the system oc-curs as a result of human activities. Output occurs via chemical or biological pathways result-ing in the formation of harmless products or in complete mineralization. Microorganismsthat are able to transform and mineralize hazardous organic compounds may be viewed as“decontaminators” and belong to the ecological group of the decomposers

pool together with the recycling of nutrients back to the nutrient pool. Somebacteria and fungi are able to detoxify and mineralize man-made organic com-pounds like chlorinated benzenes and polyaromatic hydrocarbons. Thus, theyprevent the accumulation of such chemicals in the environment. These micro-organisms may therefore be viewed as the “decontaminators” of ecosystems andthe biosphere (Fig. 1).

1.2Behavior of Chemicals in the Physical Environment

Global exchange of organic chemicals mainly occurs via trade, the atmosphere,the oceans, and biota [1–6]. The world-wide character of pollution is illus-trated by the presence of man-made organics in Arctic snow and in the air ofthe Northern and Southern hemispheres [1, 7]. The spread of chemicals is caus-ed by high production and release rates combined with their stability againstbiotic and abiotic transformation and their relative mobilities in air, water, soil,and biota [8].

PCBs, dioxins, and PAH are released directly to the atmosphere in combus-tion processes and can exist there both unbound and bound to particles [2, 4,6]. Wet and dry deposition then lead to soil and water pollution [2, 4, 6].Subsurface contamination results from infiltration of contaminated surfacewater into river borders, from deposition with settling particles onto the sedi-ment in sedimentation areas of rivers and large water bodies, and from seepagefrom the top-soil to deeper layers [9–12]. Thus, the atmosphere and subsurfaceare two major sinks where hydrophobic organic contaminants accumulate(Fig. 2). Volatile compounds accumulate more in the atmosphere while lessvolatile compounds accumulate more in the subsurface. Mineralization of theseotherwise recalcitrant compounds in one of these sinks is the only pathwayremoving these compounds from our environment.

Transformation reactions in the atmosphere are almost exclusively(photo)chemical and may interfere with other atmospheric compounds.Volatile CFCs (chlorofluorocarbons), for instance, can survive unchanged formore than 100 years in the atmosphere due to their physical and chemical inert-ness. They diffuse eventually upward into the stratosphere where chlorine radi-cals are released by short-wavelength UV-radiation [13]. This reaction initiatesthe well-known breakdown of the stratospheric ozone-layer and is the only re-moval mechanism of CFCs known to occur in the atmosphere [13].

1.3Decontamination by Microorganisms

Photochemical reactions are not possible in the subsurface, where microbiallymediated transformations constitute the dominant removal mechanism of per-sistent organic compounds. Many microorganisms live in soil and groundwaterwhere hazardous compounds may accumulate. It was recognized early in thiscentury that bacteria are able to oxidize rapidly complex, chemically stable or-ganic structures originating from petroleum, paraffin, and benzine [14]. Their



167

Fig. 2. Global exchange of hydrophobic contaminants. They tend to volatilize or to bind to soil due to their physical-chemical properties. As a conse-quence, the subsurface and the atmosphere are major sinks where these contaminants accumulate. Exchange between the northern and southern he-mispheres is minor and occurs via human intervention (transport by air and sea) and biota (animals traveling over the equator)

capacity to detoxify anthropogenic chemicals under similar environmentalconditions is variable among various habitats. This may be related to previousexposure of the microorganisms to the compound under consideration. An ad-apted microflora capable of converting and mineralizing new compounds mayevolve after a long exposure time. The microflora in a not pre-exposed en-vironment may not be able to detoxify the same compound. Dichloropropeneand 2,4-D (dichlorophenoxy acetic acid) are examples of pesticides that micro-organisms have “learned” to transform. Degradation of these compounds in thefield can be so rapid nowadays that their effectiveness as pesticides is stronglyreduced. As a result, farmers have to apply considerably larger amounts of thesepesticides than was necessary when they were first being used.

2Biodegradation Pathways

2.1Introduction

For organisms to grow, electron donors and acceptors, a carbon source, andnutrients need to be present. In addition to the naturally occurring organic substrates, many anthropogenic compounds can fulfill the growth requirementsof microorganisms [15]. Many aliphatic and aromatic contaminants serve as electron donors, thereby undergoing substantial transformation or even min-eralization to inorganic end products such as carbon dioxide, water, and in-organic ions. Intermediates of the degradation may be assimilated as the car-bon source for the formation of biomass, and functional groups of the con-taminants such as amino, nitro, and sulfonate groups may be used as nutrients.The electron acceptor for contaminant degradation may be molecular oxygenor, under anoxic conditions, the oxidized inorganic compounds nitrate, metalions, sulfate, and carbon dioxide. It appears that not all of these electron ac-ceptors are functional with any organic contaminant. The fate of many organiccontaminants therefore relies on the presence of the appropriate electron ac-ceptors, or even the absence of other electron acceptors. Oxygen, for instance,cannot be replaced by other compounds for its function as a direct reactant[16], whereas the absence of oxygen, i.e., a reducing environment, may berequired to allow reductions which are involved in the catabolism of certaincontaminants.

Several important pollutants, such as polychlorinated biphenyls (PCBs) [17],polychlorinated dibenzofurans, and dibenzo-p-dioxins (PCDF/D) [18], anddichlorodiphenyltrichloroethane (DDT) [19] do not support growth of themicroorganisms, which achieve their primary degradation. Such fortuitous orco-metabolism of pollutants may be catalyzed by enzymes which are needed forthe metabolism of the growth-supporting substrate, but also exert activity onthese co-substrates due to relaxed substrate specificities. The strategy of manyfungi is to non-specifically oxidize the organic matter in their neighbourhood,such as lignin. The fungi seem to benefit not only directly from the oxidationproducts but also indirectly from the fertilization of their habitat by growing on


other organisms or on excretion products of those organisms. The effectivenessand the low substrate specificity of the fungal exo-enzymes leads to the co-oxi-dation of many pollutants.

2.1.1Evolution of Metabolic Pathways

Microorganisms have evolved a broad range of biochemical pathways in orderto be able to utilize all naturally occurring organic compounds [20–23].Microbial communities which are exposed to xenobiotic compounds, have of-ten been found to adapt to the utilization of these chemicals [24–26]. Such ad-aptation events may be the result of induction of specific enzymes in membersof the community or the growth of subpopulations capable of metabolizing thexenobiotic. However, there is a wealth of information suggesting that micro-organisms use genetic mechanisms to evolve metabolic pathways for the degra-dation of xenobiotic compounds. The possibility was considered that microor-ganisms may eventually be able to degrade any kind of molecule [21, 27]. Thefact that evolution of metabolic activities has occurred becomes evident fromthe similarities between individual genes and enzymes involved in the degra-dation of different xenobiotics and natural substrates and the similarities be-tween the arrangements of genes encoding for different metabolic pathways.Direct evidence comes from studies on experimental evolution. Figure 3 showsschematically the two principle groups of mechanisms involved in the geneticadaptation to xenobiotic compounds. Genetic changes can be subdivided inthose acting on the level of the cell, like point mutations and DNA rearrange-ments which are transmitted to the descendants (vertical evolution), and thoseinvolving the transfer of genes between related or non-related organisms (hori-zontal evolution) [22].

2.1.1.1Events in Vertical Evolution

Point mutations as well as changes affecting larger DNA sequences occur at lowfrequencies as a result of errors in DNA replication or repair. It has been shownthat point mutations, i.e., changes only affecting a single base pair of the DNA,can alter the specificities of enzymes and regulatory proteins or affect the re-cognition of promoter sequences. An example is the extension of the specificityof the catechol 2,3-dioxygenase encoded by TOL plasmid pWW0 to 4-ethylcate-chol by an exchange of a single amino acid [28]. There are indications that en-vironmental stresses may lead to increases in the frequency of point mutations,thereby accelerating vertical DNA evolution [29]. Increased mutation rates wereobserved in non-growing bacterial cultures in the presence of substrates, whichpotentially could provide energy or carbon for growth [22]. However, it is stillunclear whether external factors may also control the direction of changes, forinstance, by specifically promoting ‘useful’ point mutations. Divergence of DNAsequences may also result from erroneous repair of defective DNA or duringDNA replication. Such mutations are particularly frequent in DNA regions


where sequence repetitions or palindromic sequences facilitate false hybridiza-tion of DNA strands or the shift of the template DNA. A number of recent re-views and articles summarize the current knowledge of such mutational events[30–35]. Mobile DNA sequences, so-called insertion elements (IS-elements)which are present in high number in bacterial genomes [21, 36] are often sus-pected to be involved in DNA rearrangements. The replication of such elementsand integration into other regions of the DNA may, however, disturb the func-tionality of genes. On the other hand IS-elements may contain transcriptionsignals, such as promoter sequences which, after insertion, activate formerlyunrecognized genetic information (silent genes) in their neighborhood. MobileDNA elements may also co-mobilize genes for degradation of pollutants.A well-documented example is the tcbAB element encoding for the chlorobenzene di-oxygenase of Pseudomonas sp. P51 which is flanked by two IS-elements of thesame type. The composite transposable element, referred to as a transposon,was shown to be capable of inserting at random into the bacterial genome. It isobvious that transposons may translocate DNA sequences containing largeamounts of genetic information, thereby substantially rearranging bacterial ge-nomes [37].


Fig. 3. Scheme of the two principle mechanisms in molecular evolution. Vertical evolutioncomprises genetic events such as replication errors leading to single base-pair substitutions(1) or deletions (2). Horizontal evolution refers to the mobilization of DNA fragments ormolecules from one organism to another

Most gene mutations will result in deactivation of the enzyme they encodeand in case of the destruction of an essential function will not propagate.However, when mutation occurs in combination with a preceding gene duplica-tion, one of the gene copies may not be subject to selective pressure and maytherefore act as a playground for mutational changes [38]. The two isoforms ofthe 6-aminohexanoate dimer hydrolase in Flavobacterium sp. strain K172, forexample, differed by two orders of magnitude in their activity [39]. A detailedsummary of the mechanisms known to alter the DNA and examples for the ef-fects of these alterations in degradation of aromatic compounds is given by vander Meer et al. [21].

Comparison of the genetic information for degradation of xenobiotic com-pounds revealed that the operational DNA regions (operons) and gene clustersencoding different pathways have genes in common. The abundance of similar-ities of gene functions between pathways suggests that the assembly and re-combination of existing genetic material is the most important mechanism forevolving or expanding metabolic pathways [21, 22]. The TOL plasmid pWW0contains two separate operons, one encoding the enzymes oxidizing toluenes tobenzoates, the other one encoding the “meta cleavage pathway” enzymes de-grading methylbenzoates to pyruvic acid and acetaldehyde (Fig. 4) [40, 41]. Themeta cleavage pathway encoded by the NAH7 plasmid is homologous to that ofthe TOL plasmid but contains the gene encoding for the salicylate 1-hydro-xylase required to channel salicylic acid into the catechol meta cleavage path-way instead of the multicomponent aromatic ring dioxygenase necessary forbenzoate dioxygenation [42].


Fig. 4. Organization of the TOL plasmid-encoded pathway for the degradation of alkylben-zenes. The genetic information for the catabolic enzymes is organized in two regulatedoperons. The XylR protein stimulates transcription of the upper operon when activated by,e.g., toluene. The XylS regulator protein stimulates transcription of the meta operon when activated by, e.g., benzoate (redrawn from [45])

2.1.1.2Events in Horizontal Evolution

Horizontal evolution comprises those genetic events which involve the transferof genes between organisms, and are believed to have resulted in the obviousdistribution of common genetic information in the microbial community. Theobservation that slightly different homologous operons encoding for degrada-tion pathways have been frequently found in phylogenetically distant orga-nisms suggests the occurrence of extensive horizontal gene transfer. Examplesfor highly mobile DNA are self-transmissible plasmids such as the homologousTOL, NAH, and SAL plasmids [43]. These plasmids not only encode for the de-gradation pathways of toluene, naphthalene, and salicylate, respectively, but alsocontain the instructions for their transfer into other organisms. As a conse-quence, these plasmids are found in a wide range of bacterial species. In micro-cosm experiments, such catabolic plasmids could be transferred from labora-tory-derived organisms to indigenous recipient bacteria [44]. Alternative me-chanisms of gene transfer known to occur in the environment are transduction,i.e., gene transfer mediated by bacteriophages, and transformation, i.e., theuptake of free DNA. Novel metabolic activities of bacteria have been created inmany studies on experimental evolution. There are examples for both the suc-cessful expansion of existing pathways by addition of peripheral enzymes andthe broadening of substrate ranges by exchange of narrow substrate range en-zymes with enzymes having low substrate specificity.

2.1.1.3Generation of Novel Degradative Pathways

Most of the information on the evolution of metabolic pathways reviewed so farhas been deduced from comparative biochemical and molecular analysis ofcurrently existing microorganisms. The direct investigation of the underlyingnatural evolutionary events is nearly impossible because of their low frequencyand randomness [21]. Therefore, many studies on experimental evolution in thelaboratory were conducted which aimed at the change, extension, or de novoconstruction of metabolic pathways. Different approaches to the laboratoryevolution of metabolic pathways were used [45]:

1. Long-term selection may involve the progressive replacement of a utilizablesubstrate by the recalcitrant target substrate and the use of mutagens. An ex-ample for such an approach is the successful adaptation of naphthalene de-grading microorganisms to the degradation of naphthalenesulfonic acid [46].

2. In vivo construction of pathways, i.e., the recruiting of genes of one organisminto another, is achieved by a stimulation of the natural genetic transfer pro-cesses, such as transduction, transformation, and conjugation. By means of astimulation of conjugation, the degradation range of 3-chlorobenzoate de-grading Pseudomonas sp. strain B13 could be expanded to chlorosalicylates,chlorobiphenyls, chloroanilines, and chloronitrophenols [47–50].

3. In vitro construction of pathways involves the directed and selective combina-tion of well-characterized genes by molecular techniques [45]. The construc-


tion of a degradative pathway can be very straightforward when the recal-citrant target compound exhibits substantial structural analogy to a compoundwhich is readily degradable by a known pathway [51]. It involves the identifica-tion of the steps in the known pathway which are not permissive for the targetcompound and their modification into permissive ones. Such a modificationcan be the broadening of the substrate or effector specificity of a certain en-zyme or regulator protein, respectively, or the upward extension of the knownpathway. Timmis achieved the modification of the toluene degradation path-way of Pseudomonas putida encoded by the TOL plasmid pWW0 [45] (Fig. 5).


Fig. 5. Schematic representation of the successful modification of the TOL plasmid-encodedpathway for the degradation of alkylbenzenes leading to the broadening of its substrate rangeto 4-ethylbenzoate. Unlike 4-methylbenzoate (1), 4-ethylbenzoate (3) is neither an effectormolecule for the XylS regulator protein (see Fig. 2) nor a substrate for the meta pathway.Mutation of the xyl S gene led to a XylS* regulator protein that accepted 4-ethylbenzoate andactivated the meta cleavage pathway. This resulted in the conversion of 4-ethylbenzoate to 4-ethylcatechol (4), which inactivated the ring cleavage protein. Mutation of the xyl E gene ledto an inactivation resistant ring cleavage enzyme, permitting complete 4-ethylbenzoate de-gradation (redrawn from [45])

A sequence of changes was required to restructure the toluene pathway in sucha way that eventually the formerly recalcitrant analogue 4-ethyltoluene wasutilized [28, 52, 53]. The effector specificity of the XylS regulatory protein con-trolling the expression of the meta cleavage pathway turned out to be too nar-row to accept 4-ethylbenzoate as an inductor. XylS mutants with a relaxedeffector specificity could be obtained by chemical mutagenesis. One of themutants degraded 4-ethylbenzoate to 4-ethylcatechol. This product, however,was not further metabolized since it appeared to be a suicide substrate for thecatechol 2,3-dioxygenase. Clones with a catechol 2,3-dioxygenase being resis-tant to 4-ethylcatechol were also obtained by chemical mutagenesis. These or-ganisms mineralized 4-ethylbenzoate, but not 4-ethyltoluene.An analysis of thesubstrate specificity of the three enzymes leading from toluene to benzoate andthe effector specificity of the XylR protein controlling the expression of this up-per pathway revealed that only the toluene oxidase was not permissive for 4-ethyltoluene. Chemical mutagenesis resulted in mutants with relaxed substratespecificity of the toluene oxidase. These mutants completely mineralized 4-ethyltoluene via the meta cleavage pathway.

An alternative approach to new degradation pathways is the patchwork assem-bly of enzymes from different organisms [54]. One of the problems associatedwith the degradation of mixtures of aromatic pollutants arises when these che-micals simultaneously activate both the meta and the ortho cleavage pathwaysof catechols. The ortho route productively degrades chlorinated catechols,whereas alkylated catechols are converted to dead-end metabolites (Fig. 6). Themeta cleaving catechol 2,3-dioxygenase is appropriate for alkylated catechols,but converts 3-chlorocatechol and 4-chlorocatechol to metabolites which ra-pidly inactivate the enzyme [55]. Pseudomonas sp. B13, a strain degrading 3-chlorobenzoate via the ortho pathway, served as the basis for the constructionof a derivative which simultaneously metabolized chlorinated and methylatedphenols and benzoates via the ortho cleavage pathway (Fig. 7). The transforma-tion of 4-chlorobenzoate, 3-, and 4-methylbenzoate to the corresponding cate-chols was achieved by adding a broad substrate range toluate dioxygenaseencoded by the TOL plasmid pWW0 [49]. The production of dead-end meta-bolites from 3- and 4-methylbenzoate was overcome by adding a 4-methyl-2-enelactone isomerase from Ralstonia eutropha (i.e., the former Alcaligenes eu-trophus) [54].

2.2Fate of Substituents

2.2.1Removal of Halogen

Although many halogenated organic molecules occur in nature [56], industri-ally produced halo-organics constitute a large group of environmental pol-lutants and are often considered the quantitatively most important compoundsof xenobiotic character. Examples for the widespread industrial and agricul-



175

Fig. 6. Differential degradation routes for halogenated and alkylated catechols. Productive further degradation of 3-chlorocatechol A and 4-chlorocatechol B requires ortho-cleavage, whereas degradation of methylcatechol degradation requires initiation by meta-cleavage C

tural application of synthetic halogenated compounds are their use as solvents,intermediates, hydraulic and heat transfer fluids, plastics, pesticides, and inter-mediates for chemical syntheses. In contrast to natural halogenated com-pounds, which readily degrade [56], many man-made haloorganics were foundto persist in the environment [57].


Fig. 7. Patchwork assembly of a catabolic pathway for the simultaneous degradation ofchloro- and methylaromatics. The modified ortho pathway for the degradation of chlorocate-chol from Pseudomonas sp. B13 was extended by gene functions for the conversion of alkyl-benzoates from the TOL plasmid pWW0 (see also Fig. 2). This expanded the degrada-tion range to include 4-chlorobenzoate and methylbenzoates. Extension by gene functionsfrom Ralstonia eutropha (the former Alcaligenes eutrophus) for the conversion of 4-methyl-2-enelactone to 3-methyl-2-enelactone allowed the complete metabolization of alkylben-zoates. Methylphenols and chlorophenols became subject to degradation upon mutationalactivation of the expression of the phenol hydroxylase of Pseudomonas sp. B13 (redrawn from [45])

The crucial reaction in the microbial degradation of halogenated compoundsis the removal of the halogen substituent. This is reflected by the large numberof review articles dealing with bacterial dehalogenation [58–81]. Carbon-halo-gen bonds are either cleaved enzymatically or by chemical dehalogenation.Four widespread enzymatic dehalogenation reaction types will be discussedhere:

1. In nucleophilic substitution reactions, the halogen is displaced by an othernucleophile, such as the hydroxyl ion OH– in the course of hydrolytic deha-logenation.

2. During dehydrodehalogenation a halogen is removed together with an adja-cent hydrogen resulting in the formation of a double-bond.

3. Oxidative dehalogenation is often achieved by monooxygenase or dioxygen-ase activity replacing the halogen by a hydroxyl group, the oxygen of whichoriginates from molecular oxygen.

4. Reductive dehalogenation comprises hydrogenolytic dehalogenation, dihalo-elimination, coupling of halogenated molecules, and hydrolytic reduction[82–84].

2.2.1.1Nucleophilic Substitution of Halogen

In hydrolytic dehalogenation reactions, the halogen substituent is displacedthrough a hydroxyl group which is derived from water. The oxidation state ofthe reacting molecule remains unchanged (Fig. 8A). Hydrolytic dehalogenationof halo-benzoates seems to be very common, and has been shown to act on flu-orinated, chlorinated, brominated, and iodinated compounds. Under anaerobicconditions halobenzoates are often activated with coenzyme A (CoA) prior tohydrolytic dehalogenation [62]. Dechlorination of dichloroethane by Xantho-bacter autotrophicus GJ10 was achieved by two hydrolytic reactions, the first


Fig. 8 A , B. Nucleophilic substitution of halogen from: A 1,1-dichloroethane by hydroxyl [85];B dichloromethane by reduced glutathion and hydroxyl [86, 87]

forming 2-chloroethanol and the second dechlorinating chloroacetic acid toglyoxylate [85]. Dechlorination of dichloromethane by Hyphomicrobium sp.strain DM2 involved the replacement of the first chlorine atom through reduc-ed glutathion (thiolytic dehalogenation) followed by hydrolytic removal of thesecond chlorine atom (Fig. 8B) [86, 87]. Subsequently, the reduced glutathionwas recovered and formaldehyde was formed. An example of a hydrolytic de-chlorination not involving cofactors is the enzymatic removal of chlorine fromS-triazine deethylsimazine by Rhodococcus corallinus [88]. Chloride is non-en-zymatically eliminated from intermediates of the ortho cleavage pathway ofchlorocatechol degradation (see also Fig. 6). Muconate cycloisomerase II con-verts 2-chloro-cis,cis-muconate and 3-chloro-cis,cis-muconate, respectively, tothe corresponding 4-carboxychloromethylbut-2-en-4-olides which spontane-ously release HCl to give the trans and cis isomers of 4-carboxymethylbut-2-en-4-olide, respectively (for overview see [78]). The overall reaction is an intra-molecular substitution of halogen by hydroxyl.

2.2.1.2Dehydrodehalogenation

In dehydrodehalogenation halogen is removed together with a hydrogen atomfrom an adjacent carbon resulting in the formation of a double bond (Fig. 9).Pseudomonas paucimobilis UT26 dehydrodechlorinated g-HCH via penta-chlorocyclohexene and, possibly, via the unstable metabolite 1,3,4,6-tetra-chloro-1,4-cyclohexadiene to 1,2,4-trichlorobenzene [89, 90]. Eukaryotic orga-nisms were shown to dehydrodehalogenate DDT to DDE [91].

2.2.1.3Oxidative Dehalogenation

None of the previously described reaction mechanisms involved molecular oxy-gen. Hydroxylation of pentachlorophenol to tetrachloro-p-hydroquinone byArthrobacter sp. and Flavobacterium sp. depended on NADPH and oxygen(Fig. 10A). Labeling experiments revealed that the oxygen of the hydroxyl ori-ginated from molecular oxygen [92]. The broad substrate range of the pentach-lorophenol 4-monooxygenase allowed the removal of halogen, nitro, amino,and cyano groups. Oxygenolytic dehalogenation was also found in the course ofthe dioxygenation of halogenated aromatic compounds. In the degradation of1,2,4,5-tetrachlorobenzene by Pseudomonas sp. strain PS14, initial dioxygena-


Fig. 9. Stepwise dehydrodehalogenation of g-hexachlorocyclohexane to 1,2,4-trichloroben-zene (redrawn from [89, 90])

tion at C5 and C6 is followed by rearomatization due to the release of hydro-chloric acid yielding 3,4,6-trichlorocatechol (Fig. 10B) [93]. The oxidation oftrans-1,2-dichloroethylene by a culture of methanotrophic bacteria involvedthe spontaneous release of chloride from the intermediately formed epoxide[94] (Fig. 10C).

2.2.1.4Reductive Dehalogenation

Four different mechanisms of reductive dehalogenation have been distinguish-ed (Fig. 11) [82]. First, in hydrogenolytic dehalogenation, the halogen substi-tuent is replaced by hydrogen at the expense of reduction equivalents [74]. Thisreaction is therefore mainly found under anaerobic conditions, although hy-drogenolytic dehalogenation by facultatively anaerobic microorganisms hasbeen reported [95]. Aromatic and aliphatic compounds can be subject to hy-drogenolytic dehalogenation. Hydrogenolytic dechlorination of aromatic com-pounds has been proposed for various chlorophenols including pentachloro-phenol, 2,4,5-trichlorophenoxyacetic acid, polychlorinated benzenes, polychlo-rinated biphenyls, and polychlorinated dibenzo-p-dioxins [17, 18, 26, 96–98].The redox potentials of chlorinated aliphatic or aromatic hydrocarbon couplesrange between E0¢ = 0.3 V and 0.5 V. Thus, from the energetic point of view, the


Fig. 10 A – C. Different types of oxidative dechlorination: A monooxygenation of pentachloro-phenol yielding tetrachloro-p-hydrochinone [93]); B dioxygenation of 1,2,4,5-tetrachloro-benzene yielding 3,4,6-trichlorocatechol [93]; C epoxide formation by monooxygenation oftrans-1,2-dichloroethylene followed by spontaneous dechlorination [94]

utilization of chlorinated hydrocarbons as electron acceptors is possible [99,100]. Accordingly, hydrogenolytic dechlorination of perchloroethylene by thestrictly anaerobic bacterial isolates Dehalobacter restrictus and Desulfitobac-terium sp. strain PCE1 and of 3-chlorobenzoate by the sulfidogenic bacteriumDesulfomonile tiedjei DCB-1 was found to be coupled to chemiosmotic energyconservation [65, 99–101]. The halogenated compounds are used as the termi-nal electron acceptor for the growth of these organisms. In the course of the me-tabolism of 2,4-dichlorophenoxyacetic acid by the aerobic Alcaligenes eutro-phus JMP134, the NADH dependent maleylacetate reductase was shown to hy-drogenolytically dehalogenate 2-chloromaleylacetate. Although dehalogenationgenerally decreases the xenobiotic character of a chemical, in some cases it mayform more hazardous chemicals than the educts [102]. The hydrogenolytic de-


Fig. 11. Different types of reductive dehalogenation (redrawn from [82])

chlorination of tetrachloroethylene to vinylchloride is an example of such anundesired reaction [103]. Second, in dihalo-elimination, halide is removed andan additional carbon to carbon bond is formed. Dichloro-elimination fromdichloroethane by methanogenic bacteria yielded ethene. g-Hexachlorocyclo-hexane (lindane) was dihalodechlorinated to g-tetrachlorocyclohexene [104,105]. Third, coupling of two chloromethane molecules resulted in reductive de-halogenation and the concomitant formation of ethane [82]. Fourth, the oxida-tions of tetrachloromethane and 1,1,1-trichoroethane to CO2 and acetate, re-spectively, were found to be initiated by two electron reductions, followed byhydrolysis [106].

2.2.2Nitro Groups

2.2.2.1Nitroaromatic Compounds

The use of nitroaromatic compounds as explosives, pesticides, and dyes, and asintermediates in many chemical syntheses led to their entrance in all majorcompartments of the environment. Since organic nitro compounds of naturalorigin are very rare, virtually all nitroaromatic compounds nowadays found inaqueous systems [107, 108], terrestrial systems [109, 110], and the atmosphere[111, 112] were formed by human activities. Nitroaromatic compounds may betoxic and/or mutagenic to microorganisms, plants, animals, and humans. Theexplosive 2,4,6,trinitrotoluene (TNT), one of the most abundantly producednitro compounds, has been shown to cause anemia in humans [113]. The muta-genicity of airborne 3-nitrodibenzofuran was shown to be three times strongerthan that of benzo(a)pyrene [114].

In spite of their xenobiotic character, microorganisms have developed en-zymatic mechanisms to degrade nitro compounds, i.e., to convert or release thenitro group prior to further metabolism. A number of recent reviews deal withthe degradation of nitroorganic compounds [115–121]. In the following, thefate of the nitro group will be used for a classification of degradation mecha-nisms.

Oxidative Release of the Nitro Group. The degradation of a number of nitro-benzenes, nitrotoluenes, and nitrophenols is initiated by an NADPH- or FAD-dependent monooxygenase attack of the aromatic ring that replaces the nitrogroup by hydroxyl and results in the release of nitrite (Fig. 12A) [122, 123].Dioxygenolytic attack liberated nitrite from compounds such as 2,4,5-trichlo-ronitrobenzene, 3-nitrobenzoic acid, and nitrobenzene, and resulted in the for-mation of the corresponding catechols [93, 124, 125].

Reductive Removal of the Nitro Group. Nitrite can also be reductively removedfrom the aromatic ring (Fig. 12B). The nucleophilic attack of picric acid result-ed in the transient formation of a hydride-Meisenheimer complex, which re-aromatized by releasing nitrite [126].


Reduction of the Nitro Group. The nitro groups are reduced prior to the clea-vage of the carbon-nitrogen bonds of nitrobenzoic acids, nitrotoluenes, andnitrophenols (for an overview see [117, 119]). The reduction is catalyzed bynitroreductases and proceeds via a nitroso and a hydroxylamino to an aminogroup (Fig. 12C). The oxygenolytical removal of the amino group yields the cor-responding catechol as the substrate for further metabolism. Recently, in thecourse of the degradation of 4-nitrobenzoate, nitrobenzene, and nitrotoluene,the hydrolytic removal of the hydroxylamino group from the aromatic nucleusyielding the corresponding catechol has been described [127–129]. The generalobservation that the ability to reduce the nitro group of nitroaromatic compo-unds is much more common than the utilization of the products has been at-tributed to the broad substrate ranges of many nitroreductases. The reductionof various substituted nitrobenzenes to the corresponding amino compoundsin anaerobic aquifer material occurred by a reaction with surface bound ironspecies, which served as mediators for the transfer of electrons originating fromthe oxidation of organic material by iron-reducing bacteria [130].

Aromatic amino compounds tend to form complexes with the humus frac-tion of soils. There is an ongoing discussion as to whether such complexes maybe regarded as stable and, as a consequence, desirable deposits of amino aro-matic compounds or represent reservoirs of remobilizable hazardous metabo-lites [131].


Fig. 12 A – C. Mechanisms of nitrogen removal from nitroaromatic compounds: A oxidative re-lease of the nitro group from 4-nitrophenol yielding p-hydroquinone [122]; B reductive re-moval of the nitro group from picric acid involves the formation of a hydride-Meisenheimercomplex [126]; C reduction of the nitro group prior to oxidative deamination (redrawn from[119])

2.2.2.2Nitrate Esters

Nitrate esters are abundantly used in ammunition and for the therapy of car-diovascular diseases. Up to now, no naturally occurring nitrate esters areknown. Despite the strict xenobiotic character, the hydrolytic microbial cleav-age of the nitrate ester bonds of the explosives glycerol trinitrate, ethylene gly-col dinitrate, pentaerythritol tetranitrate, cellulose nitrate, and the pharmaceu-tical isosorbide 2,5-dinitrate, and the concomitant liberation of nitrite, havebeen shown [118, 121].

2.2.3Sulfonic Acid Groups

Sulfonated organic contaminants are very rare in nature. Large amounts ofthese compounds arise as lignosulfonates in the course of the industrial pro-duction of paper from wood, as surface active alkylarylsulfonates, and asamino- and hydroxynaphthalenesulfonic acids which serve as building blocksin the synthesis of azo dyes. It appears that the removal of the bulky sulfonategroup is the first step in the degradation of many sulfonated compounds.Methanesulfonic acid degradation by marine microorganisms was initiated byNADH-dependent monooxygenase reaction yielding formaldehyde and sulfite[132]. The degradation of naphthalene-2-sulfonic acid and naphthalene-2,6-di-sulfonic acid was found to be initiated by the dioxygenolytic removal of a sul-fonate group as sulfite (Fig. 13) [46]. In the latter case, the resulting sulfonateddihydroxynaphthalene is channeled into the naphthalene metabolism. The re-maining substituent is removed later in the metabolism by hydroxylation of 5-sulfosalicylic acid yielding gentisic acid [133]. Several bacteria are able to usearylsulfonates as the sole sources of sulfur. Pseudomonas putida S-313 [134]converted a number of structurally diverse sulfonated aromates, including sur-factants and azo dyes, to the corresponding phenols. The resulting molar yieldof biomass was as high as with sulfate.


Fig. 13. Oxygenolytic desulfonation of naphthalene-2-sulfonic acid [46]

2.2.4Organophosphonates

Organophosphonates are a class of compounds possessing one or more carbon-phosphorus (C-P) bonds. C-P bonds are chemically stable and withstand hy-drolysis, thermal decomposition, and photolysis [135]. Organophosphonatesare chemicals of environmental concern. Although their toxicity has long beenknown, they are widely used as herbicides, insecticides, lubricant and polymeradditives, corrosion inhibitors, and antibiotics [136]. Organophosphonates arereadily catabolized and utilized as source of carbon, phosphorus, or nitrogen bya large number of bacterial and fungal isolates [138–140]. There are a numberof reports on the partial degradation of the organophosphonate molecule priorto the cleavage of the C-P bond [141, 142]. In a recent report, Bujacz et al. [140]distinguish three enzymatic mechanisms of C-P bond cleavage (Fig. 14): (i) thehydrolysis of phosphonoacetaldehyde to acetaldehyde and inorganic phosphateby means of a phosphonatase [141] and (ii) the metal cation-assisted hydroly-sis of phosphonoacetate to acetate and inorganic phosphate by means of thephosphonoacetate hydrolase [137]; (iii) the hydrolysis of a broad spectrum oforganophosphonates by the ”so-called” C-P lyases in fact comprises reductiveand oxidative pathways [143–145].

2.3Degradation of the Carbon Backbone

2.3.1Aliphatic Compounds

Aliphatic compounds comprise a great number of straight-chain and branchedmolecules including gases, liquids, and long-chain molecules which are solid atphysiological temperatures [146]. Many of these molecules serve as growth sub-


Fig. 14. Enzymatic reactions involved in the cleavage of carbon-phosphorus bonds (redrawnfrom [140])

strates for a very wide variety of microorganisms [146–152]. The low water so-lubility of most aliphatic hydrocarbons is the major obstacle for the uptake ofthese compounds. Microorganisms have therefore developed various mecha-nisms to overcome substrate limitation. Some yeasts are known to take up hy-drocarbon droplets [153], whereas many bacteria excrete emulsifying agents orpossess highly hydrophobic cell surfaces, mediating their permanent associa-tion with hydrocarbon droplets [154–158].

2.3.1.1Aerobic Pathways

Aliphatic compounds from C1 up to C44 are subject to microbial degradation[159]. The most readily degraded hydrocarbons are the straight chain alkanesfrom C10 to C18 [151]. The principle strategy of the metabolism of aliphatic com-pounds is to convert the alkane chains into fatty acids (Fig. 15). Under aerobicconditions, the alkane is attacked by an NADH2-dependent monooxygenase (al-kane-hydroxylase) system consisting of the terminal oxidase and two or threeelectron transfer components. The monooxygenase forms a fatty alcohol bytransferring one oxygen atom of molecular oxygen onto the terminal C-atom ofthe molecule.Another effective multicomponent enzyme known to catalyze thisreaction is the methane monooxygenase of methylotrophic bacteria [151].Alcohol dehydrogenase further oxidizes the fatty alcohol to the correspondingaldehyde, which is then oxidized to a carboxylic acid by means of an aldehydedehydrogenase. The carboxylic acid is channeled into the central metabolismfor further oxidation by b-oxidation. b-Oxidation is initiated by the formationof a CoA-thioester of the fatty acid by means of the activity of ATP-dependentacyl-CoA synthetase. The products of each b-oxidation cycle are reductionequivalents and acetyl-CoA. b-Oxidation of C-odd fatty acids yields propionyl-CoA in the last oxidation cycle which is carboxylated to methylmalonyl-CoAand can enter the central metabolism after being isomerized to succinyl-CoA.The aerobic degradation of alkenes can be initiated in various ways. Besides in-itial terminal or subterminal oxygenation, the reactive double bond can be sub-ject to epoxidation or hydroxylation [147]. Straight-chain aliphatic compoundsare generally more rapidly degraded than branched compounds, although com-plete metabolism of a number of branched compounds has been reported.Examples are the utilization of the isoprenoid pristane (2,6,10,14-tetramethyl-pentadecane) and of 2,2,4,4,6,8,8-heptamethylnonane by various bacteria [160,161]. The metabolism of alkynes, such as acetylene, by both aerobic and anaer-obic bacteria is initiated by hydration of the triple bond [148]. For informationon the microbial degradation of cyclic aliphates, the reader is referred to re-views by Trudgill [152] and Perry [162].

2.3.1.2Anaerobic Hydrocarbon Degradation

Under anoxic conditions, oxygen is unavailable for the initial oxidation of al-kanes and for a long time it was believed that alkanes (with the exception of



Fig. 15. Aerobic and anaerobic mechanisms of aliphatic hydrocarbon degradation (redrawnfrom [84])

methane) are anaerobically recalcitrant [146, 163]. Recently, nitrate-reducingand sulfate-reducing bacteria which were isolated from an oil production plantwere shown to grow anaerobically with straight chain alkanes as the sole car-bon sources. Strains exhibited different specificities with respect to the range ofalkanes that could be utilized. A sulfate-reducing bacterium was found to growwith short-chain alkanes (C6 to C13), whereas nitrate-reducing strains were en-riched under anoxic conditions with hexadecane as the substrate [164, 165].One of the sulfate reducing isolates did not utilize alcohols, indicating that thereare no intermediates in alkane degradation. The pattern of total cell fatty acidssuggested that the alkane oxidation occurred via an elongation of the carbonchain by a C1-unit [166]. The double bonds of alkenes can be hydrated anaerobi-cally to form an alcohol (Fig. 15) [167]. The further conversion to a fatty acid isthen performed by oxygen-independent enzymes as in the aerobic pathway.

2.3.2Aromatic Rings

Next to the glucosyl residues, the benzene ring, as one of the main constituentof wood, is the most widely distributed structural unit in nature [168].Understandably, many microorganisms have evolved enzymatic pathways tomake use of the gigantic source of carbon and energy provided by aromaticcompounds. The recycling of carbon bound in aromatic rings actually relies al-most entirely on microbial activities. Nowadays, there is much concern aboutanthropogenic aromatic bulk chemicals such as benzene, toluene, and xylene,which are widely used as solvents [169] and polyaromatic hydrocarbons (PAH),which were formed and unintentionally entered soils and groundwater duringmanufacturing of combustible gases [170]. The thermodynamic stability ofbenzene, substituted benzenes, and condensed aromatic rings caused by theirpossession of a negative resonance energy, required the development of specialdegradation mechanisms.

2.3.2.1Aerobic Pathways

The main strategy of microorganisms to degrade aromatic pollutants aerobi-cally is to use a range of peripheral enzymes which convert the substances to akey intermediate, which is most often a (substituted) catechol (Fig. 16). Thisstrategy appears to be very economic, since it allows one to channel a large va-riety of substrates into the same central catabolic pathway. Different mecha-nisms of catechol formation exist in eukaryotes and prokaryotes (Fig. 17).Eukaryotic organisms produce catechols from aromatic compounds by insert-ing one atom of molecular oxygen by means of a monooxygenase yielding epox-ides. Subsequent addition of water leading to trans-dihydrodiols is followed bydehydrogenation. Prokaryotes introduce an entire oxygen molecule by a dioxy-genase reaction forming a cis-dihydrodiol which is then subject to dehydroge-nation. Aromatic hydrocarbons, such as benzene, naphthalene, higher condens-ed polyaromatic compounds, and biphenyl are attacked in this way. Aliphatic


substituent groups on the benzene ring offer the possibility of alternativemodes of biodegradation; either side-chain attack or ring attack. It seems thatalkylbenzenes with chains of up to 7 C-atoms are preferably attacked at thearomatic rings, whereas the initial oxidation of the alkyl chain is the preferredattack when chain lengths exceed 7 carbon atoms [169]. However, the degrada-tion of toluene is a well-examined example for the realization of both degrada-tion pathways, the initial attack being either ring dioxygenation (Fig. 18A) orthe stepwise oxidation of the methyl group to benzoic acid as the substrate forring dioxygenation (Fig. 18B). In eukaryotes and in prokaryotes, catechol andvarious substituted catechols are opened by dioxygenase reactions by eitherortho- (intradiol-) or meta- (extradiol-) cleavage.

The ortho-cleavage of catechol by catechol 1,2-dioxygenase activity yieldscis,cis-muconic acid (Fig. 19). Further breakdown via muconolactone and 3-oxoadipate enol-lactone leads to 3-oxoadipate which enters the central metabo-lism as succinate and acetyl-CoA. Halogenated aromatic compounds are mostoften degraded in this manner, although recently a meta cleavage pathway for4-chlorobenzoate has been reported [171]. The chlorines are eliminated afterring cleavage depending on their position in the molecule during the cycloiso-merization or even later in the breakdown (see also Fig. 6). The enzymes involv-ed in the breakdown of chlorocatechols have broader substrate specificitiesthan the ordinary ortho cleavage pathway enzymes. An example is the modified


Fig. 16. Converging pathways of aromatic hydrocarbon degradation (redrawn from [84])

ortho pathway involved in the degradation of 2,4-dichlorophenoxy acid byRalstonia eutropha JMP134 (formerly Alcaligenes eutrophus) [172].

The meta-cleavage of catechol (Fig. 19) by catechol 2,3-dioxygenase bringsabout 2-hydroxymuconic semialdehyde, which after decarboxylation to 2-oxo-penta-4-enoate is cleaved into pyruvate and acetaldehyde. Alkylated and phenylsubstituted catechols are most often subject to meta cleavage [68, 169].

Alternative substrates for ring cleavage are the para-dihydroxylated aromat-ic acids gentisic acid (Fig. 20) and homogentisic acid, which are cleaved be-tween the carboxy group and the adjacent hydroxy group [173]. Homogentisicacid is known as key intermediate in the catabolism of the aromatic aminoacids phenylalanin and tyrosin. The ring cleavage products of homogentisateand gentisate enter the central metabolism as fumarate and acetoacetate, andfumarate and pyruvate, respectively. Interestingly, none of the reactions follow-ing ortho or meta cleavage of catechol or the cleavage of homogentisic and gen-tisic acid requires oxygen. Further information on the aerobic biodegradation


Fig. 17. Different ways of catechol formation from mononuclear aromatic compounds by eu-karyotes and prokaryotes (redrawn from [84])

Fig. 18 A, B. Toluene biodegradation initiated by: A dioxygenation; B successive oxidation ofthe methyl group (redrawn from [169])

190T.N

.P.Bosma et al.

Fig. 19. The ortho-(intradiol-) pathway and the meta-(extradiol-) pathway of catechol degradation

Fig. 20. The gentisate pathway of aromatic ring-cleavage

of aromatic compounds can be obtained from several of reviews and mono-graphs [57, 168, 169, 173–181].

2.3.2.2Anaerobic Pathways

Anaerobic degradation of aromatic compounds is carried out by phototrophicbacteria, by fermenting, manganese-reducing, iron-reducing, nitrate-reducing,and sulfate-reducing bacteria, and by methanogenic consortia. The destabiliza-tion of the aromatic ring prior to cleavage is achieved by reduction (Fig. 21).The key intermediate in this pathway is cyclohexanone or one of its derivatives.Ring opening proceeds through hydration of the cyclohexanone. As early as1968 the intermediates of the anaerobic biodegradation of benzoic acid byRhodopseudomonas palustris were identified and a degradative pathway wasproposed [182]. Benzoic acid was found to be activated by with co-enzyme Afollowed by complete reduction of the aromatic ring. The consecutive sequenceof dehydrogenation forming a double-bond in the a,b-position, hydration, de-hydrogenation, and thiolysis resulting in ring cleavage follows the scheme of theb-oxidation of fatty acids. The first linear product was pimelyldi-CoA. Phenolreduction by nitrate-reducing enrichment cultures was observed by Bakker[183]. Here caproate was the product of ring-cleavage. Nitrate-reducingPseudomonas strains exhibited phenol carboxylase activity forming 4-hydroxy-benzoate, which after dehydroxylation to benzoate was channeled into the re-


Fig. 21. Reductive pathways of anaerobic aromatic hydrocarbon degradation (redrawn from[84])

ductive pathway [184]. Catechol degradation by methanogenic consortia pro-ceeded via dehydroxylation to phenol followed by reduction and ring cleavageto give caproate or, alternatively, adipate. An oxidative initiation of the an-aerobic degradation of toluene, phenol, and p-cresol was proposed by Lovleyand Lonergan [185]. They could show that the oxidation of the methyl group of(substituted) toluene to the carboxy group and the further degradation of ben-zoate was coupled to the reduction of Fe (III). The anaerobic biodegradation ofaromatic compounds has been dealt with in more detail in several reviewarticles [186–191].

2.3.2.3Fungal Metabolism

White-rot basidiomycetes exhibit the ability to transform a wide range of ali-phatic and aromatic compounds including many environmental pollutants.Most of the degradative mechanisms involve the activity of lignin peroxidaseand manganese peroxidase, the major components of the lignin-degradativesystem [192, 193]. The non-specific nature of the peroxidases, which use hydro-gen peroxide to achieve a one-electron oxidation of chemicals to free radicals,allows them to attack complex mixtures of pollutants. However, primary growthsubstrates such as cellulose and glucose are required for the fungal co-metabo-lism of pollutants. The fact that peroxidases are generally excreted, allows thetransformation of insoluble and polymeric compounds, such as polycyclic aro-matic hydrocarbons or lignin. Since the products of co-oxidation are often notmetabolized further by the fungi, mixed populations of fungi and bacteria areusually required to achieve the complete mineralization of organic contami-nants. Fungi might prove valuable should molecules which are not easily trans-ported into bacterial cells or metabolized by bacteria have to be oxidized.Detailed information on fungal metabolism is available in several articles, re-views, and monographs [192–201].

2.3.3Ethers

The ether bond is a structural feature present in many natural compounds, suchas lignin and its degradation products. Synthetic ether compounds are used asagrochemicals, such as the phenoxyalkanoate herbicide 2,4 dichlorophenoxy-acetic acid (2,4-D), as additives, such as the antifreeze agent polyethylene glycol(PEG), and as detergents. Ether compounds of considerable concern are thehighly toxic polychlorinated dibenzofurans (PCDF) and dibenzo-p-dioxins(PCDD) which are unintentionally formed in various incineration processes[202]. Due to its chemical stability, the ether linkage represents a major chal-lenge to enzymatic attack. Microorganisms have, nevertheless, evolved a num-ber of mechanisms resulting in ether cleavage. A common mechanism ofaerobic ether cleavage involves the insertion of oxygen into one of the carbonatoms next to the ether bond by either monooxygenase [203, 204] or dioxygen-ase activity [205]. This yields unstable hemiacetal structures which sponta-


neously dismutate into alcohol and aldehyde (Fig. 22A). Hydroxylation of theadjacent C-atom can also be achieved by the addition of water to a C=C doublebond next to the ether linkage (Fig. 22B). Such a mechanism is involved in theaerobic and the anaerobic metabolism of polyethylene glycol [16, 206]. A furth-er common mechanism of ether cleavage is the oxidation of one of the ethercarbons to a keto group yielding an ester, which is subject to enzymatic hydro-lysis (for an overview see [207]). Whereas all mechanisms discussed thus farachieve ether cleavage by an initial destabilizing reaction, followed by a sponta-neous C-O fission or ester hydrolysis, carbon-oxygen lyases represent a meansof direct ether cleavage by a b-elimination reaction as shown in Fig. 22C [208].


Fig. 22 A – C. Different mechanisms of ether cleavage: A destabilization of 4-chlorodiphenylether by angular dioxygenation is followed by spontaneous cleavage [205]; B destabilizationof the ether bond by addition of water to an adjacent double bond [206]; C direct ether cleav-age by a carbon-oxygen lyase [208]

2.3.4Chiral Compounds

Many of the chemical compounds in use are chiral. Chirality refers to the sym-metry properties of the molecules. A chiral molecule is not superimposable(i.e., not identical) with its mirror image. In the most simple case of a moleculecontaining one asymmetric carbon atom (a carbon atom connected to four dif-ferent residues) there are two stereo-isomers, called enantiomers. Many chem-icals in agricultural and medical use are marketed as racemates, i.e., mixturesof equal amounts of both enantiomers. A review by Ariëns gives a good ac-count of the manifold occurrence of racemates among pesticides and drugs[209]. Other chiral compounds of environmental relevance are the linear alkyl-benzene sulfonates (LAS) which are constituents of commercial detergents,with an annual production of 1–2 million tons [210]. Enantiomers have iden-tical chemical and physical properties only in a symmetrical environment.Enzymes, however, are generally asymmetrical, due to their composition ofchiral amino acids. Compared with the quantitative importance of racemates,relatively few studies considered the environmental degradation of chiral com-pounds [211]. There are many examples found in the literature of the enantio-selective degradation of racemates, i.e., the preferred usage of one of the enan-tiomers [212–216]. A recent investigation of the complete microbial degrada-tion of the racemate of the herbicide mecoprop [(RS)-2-(4-chloro-2-methylphenoxy) propionic acid] revealed, that both enantiomers were degrad-ed by different mechanisms [217]. These reports clearly indicate the necessity totreat enantiomers separately with respect to environmental degradation studiesand environmental regulations.

2.3.5Complexing Agents

Complexing aminopolycarboxylates such as nitrilotriacetic acid (NTA) andethylenediaminetetraacetic acid (EDTA) form water-soluble metal complexes.They are used in household detergents to inhibit the formation of insolubleCa2+ and Mg2+ salts from tensides or carbonate. Nowadays, these compoundssubstitute phosphate-based complexing agents which had been found to causethe world-wide eutrophication of rivers and lakes [218]. NTA is readily degrad-ed in laboratory cultures by several taxonomically different bacteria as well asduring waste water treatment [219]. Aerobic primary biodegradation involvesthe cleavage of NTA by the oxygen and NADH-dependent monooxygenase giv-ing iminodiacetic acid and glyoxylate (Fig. 23). Interestingly, the same productsare formed under denitrifying conditions by the NTA dehydrogenase [219].EDTA, in contrast, has been found to persist in wastewater treatment plants, ri-vers, and in aerobic groundwater infiltration zones [220–222]. The recal-citrance of EDTA in natural environments is of considerable concern since theeffective metal-binding properties are suspected to have undesirable environ-mental consequences such as the remobilization of heavy metals from river se-diments [223].


2.4Recalcitrance

Chemicals that resist biodegradation are known as recalcitrant molecules.Alexander listed several examples of anthropogenic compounds which per-sisted in soils for between 3 and more than 20 years [57]. Recalcitrance may bethe result of adverse environmental factors preventing the degradation of a che-mical. This implies that changes of the environment may turn formerly recal-citrant chemicals readily biodegradable. Polychlorinated biphenyls (PCBs) andpolychlorinated benzenes carrying more than seven and four chlorine atoms,respectively, are recalcitrant in the presence of oxygen. However, under an-aerobic conditions they are readily dechlorinated to biphenyl and benzenewhich, in turn, persist under anaerobic conditions, but readily degrade underoxic conditions. Recalcitrance of highly chlorinated aromates may therefore beovercome either by environmental conditions changing between anoxic andoxic or by the transport of intermediates from anoxic to oxic zones.

Recalcitrance in the strict sense refers to inherently non-biodegradable che-mical structures. Several reasons for the intrinsic recalcitrance of moleculeshave been suggested [57]: (i) Enzymes able to attack chemicals of strong xeno-biotic character may not exist. This may for instance explain the considerableresistance of highly branched molecules and molecules containing bulky sub-stituents, which may mask potential sites for enzymatic cleavage. (ii) Uptakesystems allowing the intracellular degradation of the chemicals may not exist.


Fig. 23. Reaction scheme of the degradation of nitrilotriacetate (NTA) (adapted from [230])

The considerable recalcitrance of most polymers can be at least partly attribut-ed to the impracticability of the uptake in combination with the non-existenceof extracellular enzymes. The observed recalcitrance of polyesters and nylon incontrast to the ready degradability of the respective oligomers is a strong indi-cation for such a mechanism of recalcitrance [39]. (iii) Regulatory proteins ableto recognize the chemicals and induce the enzyme synthesis may not exist. Thisis understandable in terms of the necessity to reconcile the evolution of degra-dative enzymes with the parallel evolution of appropriate regulator proteins[21]. An example of the lack of such a parallel evolution is the aerobic metabo-lism of PCBs which requires the presence of biphenyl or monochlorobiphenylsas an inductor of the degradation pathway [68, 224]. (iv) The intrinsic toxicityof the molecule may, in case of an unspecific mechanism of action, affect allmicroorganisms with degradative potential. The recalcitrance of tetralin may atleast partly be attributed to its accumulation in cell membranes, causing mem-brane expansion and adversely affecting membrane function [57, 225]. Bacteriawere found to overcome such mechanisms of recalcitrance by specific adapta-tion. For instance, the solvent tolerances of Pseudomonas putida strains weremediated by changed membrane compositions and modified lipopolysacchari-des [226]. In some cases not the anthropogenic pollutants but metabolites resistbiodegradation. The poor effectiveness of the cometabolic degradation of PCBsin the environment has recently be attributed to the fortuitous formation of thebroad spectrum antibiotic protoanemonin from 4-chlorocatechol by the natur-al microflora via the widespread ortho cleavage pathway of catechol [227].

3Concluding Remarks

Microbial transformation is required to achieve detoxification of organic con-taminants that accumulate in soil and groundwater. Microorganisms can there-fore be viewed as a sub-population of the decomposers with a special function,namely detoxification of the environment. Microbial processes have been usedin the fields of waste water technology, the clean-up of polluted air streams inbiofilters and in soil and groundwater remediation. The effectiveness of micro-bial processes in soil and groundwater can be reduced by the relative immobi-lity of organic compounds in the soil matrix where microorganisms live.Pollutants which are immobilized in the soil due to sorption are considered tobe unavailable for biodegradation. Other reasons for a limited bioavailability insoil can be the presence of the compounds as pure solids or liquids as is oftenthe case at former gas manufacturing plants [170] or the formation of bound re-sidues due to chemical binding to organic matter [228].

The relative availability of a compound can be expressed in a bioavailabilitynumber Bn which simply is the ratio of the rate of mass transfer to the degrad-ing organisms to the rate of biodegradation [229]. Bn expresses control of bio-degradation by mass transfer at values less than unity and control by microbialactivity at values greater than unity [229].A lot of effort in the past has been putinto resolving the limitations to biodegradation caused by mass transfer fromsorbed phases or pure solids or liquids. There has been a trend recently to ad-


apt the strategy in bioremediation technologies to the limited availability ofpollutants rather than try to speed up the remediation to faster than the masstransfer rates. This approach has been facilitated by the acceptance that classi-cal clean-up goals (for instance, the Dutch list) can only be reached throughcostly – in terms of money and energy – ex situ treatment technologies of pol-luted soil. Thus, it became possible to design approaches that are adapted to thelimitations set by mass transfer. An excellent example of such a remediationstrategy is intrinsic bioremediation or natural attenuation. This strategy makesuse of intrinsic degradation processes for the remediation of polluted soil andgroundwater. The attenuation of the pollutants is the overall result of physical,chemical, and biological removal processes. An important clue to natural at-tenuation is that the technology is adapted to the limited availability of pol-lutants for biodegradation. Thus, the realm of biodegradation pathways as de-scribed in the previous pages becomes available for the solution of the prob-lems arising from environmental pollution with man-made chemicals, especiallyin soil and groundwater.

From an ecological stand-point, it can be argued that production and releaserates of toxicants have to be smaller than in situ biotransformation rates to keepenvironmental pollution within acceptable limits. Treatment as close to thesource as possible during the manufacturing and use of chemicals will be animportant strategy to reach such a goal. The use of pesticides should be regula-ted such that the amount applied in a growth season is completely transformedin situ in the same season.

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Dordrecht


Protozoa in Wastewater Treatment:Function and Importance

Wilfried Pauli 1, Kurt Jax 2, Sandra Berger1

1 Institut für Biochemie und Ökotoxikologie, Freie Universität Berlin, Ehrenbergstr. 26–28,D-14195 Berlin, Germany, E-mail: [email protected]

2 Zentrum für Ethik in den Wissenschaften, Universität Tübingen, Keplerstrasse 17,D-72074 Tübingen, Germany

Protozoa constitute a major link between the highly productive and nutrient retaining micro-bial loop and the metazoans of the classical food web. Protozoa are efficient at gatheringmicrobes as food, and they are sufficiently small to have generation times that are similar tothose of the food particles on which they feed. They are, in quantitative terms, the most im-portant grazers of microbes in aquatic environments, balancing bacterio-plankton produc-tion. Protozoa not only play an important ecological role in the self-purification and mattercycling of natural ecosystems, but also in the artificial system of sewage treatment plants. Inconventional plants ciliates usually dominate over other protozoa, not only in number of spe-cies but also in total count and biomass. It is generally accepted that their feeding on bacteriaimprove the treatment, resulting in a lower organic load in the output water of the treatedwastes. Due to their biodegradation potential some attempts have been made to use ciliatesspecifically in environmental biotechnology. As biosensors they could provide valuable infor-mation regarding adverse effects of environmental chemicals on this part of the biocoenosisessential for the effective operation of biological waste-water treatment processes.

Keywords. Protozoa, Ciliates, Ecology, Sewage treatment, Environmental biotechnology

1 Ecological Role of Aquatic Protozoa with Special Regard to Ciliates Within the Microbial Food Web . . . . . . . . . . . . . 205

1.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2051.2 Traditional Food Webs and Microbial Food Webs . . . . . . . . . . 2051.3 The Role of Protozoa in Aquatic Food Webs . . . . . . . . . . . . . 2081.4 Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211

2 Protozoa in Wastewater Treatment . . . . . . . . . . . . . . . . . . 212

2.1 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2122.1.1 Wastewater . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2122.1.2 Biological Treatment Processes . . . . . . . . . . . . . . . . . . . . 2142.1.3 Bacterial Biofilms . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2162.1.4 Activated Sludge . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2162.2 Protozoa in Biological Wastewater Treatment Plants . . . . . . . . 2172.2.1 Occurrence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2172.2.2 Species Composition . . . . . . . . . . . . . . . . . . . . . . . . . . 2182.2.3 Plant Specific Basic Communities . . . . . . . . . . . . . . . . . . . 2202.2.4 Biomass . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2212.2.5 Ecological Framework . . . . . . . . . . . . . . . . . . . . . . . . . 2212.2.5.1 Sludge Loading . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 223

CHAPTER 3


2.2.5.2 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2232.2.5.3 pH-Value . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2232.2.5.4 O2-Content . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2242.3 Significance of Protozoa for Wastewater Treatment . . . . . . . . . 2252.3.1 Nutrition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2252.3.2 Reduction and Elimination of Suspended Particles and Bacteria . 2272.3.2.1 Clearing Rate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2272.3.2.2 Experimental Findings . . . . . . . . . . . . . . . . . . . . . . . . . 2282.3.2.3 “Field”-Observations . . . . . . . . . . . . . . . . . . . . . . . . . . 2312.3.3 Elimination of Dissolved Substances . . . . . . . . . . . . . . . . . 2322.3.4 Flocculation and Composition of the Bacterial Community . . . . 2322.3.5 Reduction of the Total Biomass . . . . . . . . . . . . . . . . . . . . 2352.3.6 Influence of Protozoa on Bacterial Metabolism . . . . . . . . . . . 2372.3.7 Filamentous Bacteria and Protozoa . . . . . . . . . . . . . . . . . . 239

3 Impairments of Protozoa: Consequences for Water Purification . 241

4 Environmental Biotechnological Aspects . . . . . . . . . . . . . . 243

4.1 Biodegradation Potentials of Ciliates . . . . . . . . . . . . . . . . . 2434.2 Ciliates as Biosensors . . . . . . . . . . . . . . . . . . . . . . . . . . 245

5 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246

List of Abbreviations

BOD(5) biological oxygen demand (index: within 5 days)COD chemical oxygen demanddw dry weightfm sludge loading [g BOD (g MLSS day)–1 or g BOD (g MLVSS day)–1],

also known as “food to micro-organism (F/M) ratio”F/M-ratio see fmMLSS Mixed-liquor suspended solids, sludge solids (g m–3; concentration

of the suspended solids in an aeration tank including inorganic mat-ter)

MLVSS Mixed-liquor volatile suspended solids (g m–3; corresponds to the or-ganic, i.e., combustible content of the sludge, which amounts to ca.70% of the sludge solids: 0.7 MLSS≈MLVSS; this parameter is oftenused as indicator of microbial concentration, although it does notdistinguish between biochemically active material and inert or deadmaterial in the sludge)

EC/LC50 50% effective and lethal concentration, respectively

204 W. Pauli et al.

1Ecological Role of Aquatic Protozoa with Special Regard to Ciliates Within the Microbial Food Web

1.1Introduction

There is hardly any place on earth in which protozoa cannot be found. They areabundant in terrestrial as well as in aquatic systems. In the latter they are pre-sent in high numbers of species and individuals both in the oceans and in fresh-water habitats. Some taxa live attached to solid substrates or within the sedi-ment, some as part of the plankton. An overview of the data about the abun-dance of protozoa in aquatic habitats gives a first indication that these organismsare not negligible in aquatic environments – although in fact they are still oftenneglected. In the plankton of highly productive lakes, densities of small flagel-lates (< 20 mm body size) of more than 106 cells per ml were reported [1] and instudies on the periphyton of small bodies of waters maximum values of morethan 1350 cells per cm2 of the much larger testate amoebae specimens were en-countered [2]. However, these numbers do not make any statements about theecological interactions in which the species are involved and the role they playwithin those processes which mostly are seen as the essence of ecosystem dy-namics, namely the fluxes of energy and material. It is the objective of this pa-per to provide a short introduction to the current knowledge of these roles asregards aquatic environments.

1.2Traditional Food Webs and Microbial Food Webs

Traditionally, food webs in aquatic systems were illustrated as in Fig. 1. Goingback to the limnologist August Thienemann, the different species within a bodyof water were characterized by the categories of producers, consumers of differ-ent order (primary consumers, secondary consumers and so on) and decom-posers [3]. The latter live on the dead organic matter and mineralize the organ-ic compounds to inorganic nutrients, e.g., phosphorus, nitrogen, etc. These ca-tegories were also the basis on which Raymond Lindeman [4] built his famoustrophic dynamic concept of ecology which was the first implementation ofArthur Tansley‘s ecosystem concept [5]. Energy enters the system as light and isprocessed as organic matter along the food chain or food web until most of theenergy is dissipated by respiration.

In aquatic habitats these functional categories – trophic levels in Lindeman’sparlance – were commonly attributed to phytoplankton (producers), zooplank-ton (primary consumers), and different kinds of vertebrates on the highertrophic levels. Protozoa and particularly bacteria were seen as decomposers,mainly restricted to sediments and other surfaces, but of minor importance inthe pelagic food web.

This association of bacteria and protozoa with decaying matter was recog-nized and used for applied purposes rather early. Protozoa were used as bioin-

Protozoa in Wastewater Treatment: Function and Importance 205

dicators for the saprobic states of natural and manmade freshwaters as early as1908 (e.g., [7, 8]). Their dynamics in the process of decomposition of organicsubstances were clarified by the middle of the century. Meanwhile, classical stu-dies on this topic were made by Bick and co-workers (e.g., [9, 10]) who investi-gated the succession of micro-organisms, in particular ciliated protozoa, in thecourse of the “self-purification” of water enriched with sewage and other organ-ic substances.

However, during the last two decades there have been some new insightswhich have broadened and fundamentally changed our way of looking at thewater of lakes and oceans and which affect the role protozoa and other micro-organisms are supposed to play within aquatic systems. These insights were in-itiated by the appearance of some new actors on the stage of the ecological thea-ter which also radically changed the roles in which protozoa were perceived. In1974 Pomeroy [11] presented a paper in which he developed new ideas aboutthe interactions of the pelagic organisms. Although these ideas were first devel-oped in connection with marine systems they were soon transferred to fresh-water habitats. The main point made is that, besides and connected with theclassical “macroscopic” food web, there exists a microbial food web. The reasonwhy these microbial food webs were discovered so late can, to a high degree, beattributed to the development of new methods in aquatic ecology.

206 W. Pauli et al.

Fig. 1. Diagram of the “classical” food web in lakes. Modified, according to [6]

By the early 1970s it was recognized that an important part of the pelagic or-ganisms had been neglected as a result both of the methods used and of thetheories regarding interactions in the water. Using direct counts of bacteria withepifluorescence methods instead of plate counts, it turned out that the abun-dance of bacteria in the open water had been underestimated by orders of mag-nitude. Only 0.1–1% of the actual abundance had been counted [12]. Fur-thermore, most investigations of marine and freshwater plankton used plank-ton nets with a mesh size of 20 mm or even 60 mm, while all smaller organismswere thought to be of minor importance. Finally, the methods of conservingplanktonic protozoa were inadequate and even larger protozoa were neglectedor underestimated as components of the pelagic species assemblages [13].What was collected and counted were those fractions of the plankton which wenow call the micro- and macroplankton, i.e., organisms bigger than 20 mm(Table 1).

Thus, not only all smaller organisms, the pico- and nanoplankton – consist-ing of bacteria, Cyanobacteria, small protozoa, and small eukaryotic algae [14]– but also many larger protozoans were to a large extent excluded from thequantitative sampling. However, it turned out that especially this small sizedfraction of the plankton is of extreme importance in terms of energy- and ma-terial fluxes. New measurements revealed that the major part of the metabolicactivity in plankton was displayed by the size fraction below 10 mm [15]. Themost productive component of the pelagic food webs was not, as thought ear-lier, the planktonic eukaryotic algae of the microplankton, but the tinyCyanobacteria, mostly of the genus Synechococcus, and some small eukaryoticalgae. The percentage of primary production in terms of carbon varies between1% and 90% in marine waters – with higher ratios in more oligotrophic condi-tions – and 16–70% in fresh waters [16]. For oligotrophic lakes 50–70% are do-cumented, while the autotrophic picoplankton amounts to 10–45% of the totalphytoplankton biomass (standing stock, measured as chlorophyll) [17]. Data formarine habitats give estimates of 20–80% [18]. Similarly, the abundance of he-terotrophic picoplankton, i.e., heterotrophic bacteria, is much higher than pre-viously thought and can approach 109 cells in highly eutrophic fresh waters [1].However, the new theory incorporates some new links rather than just addingpicoplankton to the classical food web. Figure 2 presents a very simple diagramof what a microbial food web might look like, given the current status of knowl-edge.


Table 1. The size classes of planktonic organisms

Picoplankton Nanoplankton Microplankton Macroplankton

0.2–2 mm 2–20 mm 20–200 mm > 200 mmBacteria Algae Algae CiliatesCyanobacteria Flagellates Rhizopods RotatoriaAlgae Rhizopods Ciliates CrustaceansFlagellates Ciliates Rotatoria Fish larvaeCiliates Nauplii

The earlier food chain from algae via macrozooplankton to fish still existsbut is supplemented by a new section which is commonly called the microbial“loop.” This consists of the picoplankton (“algae,” i.e., Cyanobacteria and he-terotrophic bacteria), protozoa, and a compartment of non-living material, i.e.,dissolved organic matter (DOM). DOM is lost and excreted in substantial amounts by both algae and Cyanobacteria and constitutes the energy source forthe heterotrophic bacteria. The rate of fixed carbon lost by phytoplankton cellsmay vary between 10% and 40% depending on the physiological status of thecells [13]. The picoplankton is grazed by protozoa which themselves are preyedupon by the metazoan zooplankton, thus coupling the microbial loop to the tra-ditional parts of the food web. As cells with a size of up to 2 mm hardly get lostthrough sedimentation, the microbial loop not only adds some new links to theclassical food web but keeps the nutrients (DOM and inorganic nutrients) with-in the water body and minimizes losses to the deeper, non-productive regionsof the waters or even the sediment. This seems to be particularly important dur-ing the summer stratification of oligotrophic lakes, in which the epilimnion, theupper and photosynthetically active region of the lake – the euphotic zone – istemporarily cut off from the richer nutrient supply of the deeper waters [17].

1.3The Role of Protozoa in Aquatic Food Webs

From this scheme the new role of protozoa within the food webs of aquatic sys-tems seems obvious. They are not only – in the same way as bacteria – decom-posers associated with the decay of organic material, but they are a link between

208 W. Pauli et al.

Fig. 2. The food web of the lake plankton. The classical food chain (open circles) is supple-mented by the elements of the microbial loop (filled ovals and square). DOM: dissolved or-ganic matter

the highly productive and nutrient retaining microbial loop and the metazoansof the classical food web. Most microplankton organisms are unable to utilizeparticles smaller than 5 mm directly [18]. Protozoa “repack” the organic mate-rial into edible portions and thus make it available to crustaceans, rotatoria, andother metazoans. There is empirical evidence that planktonic protozoa graze ef-fectively on picoplankton and also that protozoa constitute a valuable diet forcrustaceans [19]. Thus both necessary links between picoplankton and metazoahave been established.

The details of the microbial webs, however, are still the subject of researchand discussion. The specific pathways and the number of steps over whichenergy and nutrients are transferred are subject to much variation. There istemporal variation, e.g., seasonally, [20] and there is spatial variation both with-in lakes and even more if different lakes are compared.

The compartment of protozoa can be divided in several ecological relevantways. Not only is there a taxonomic division between flagellates and ciliates, butalso a physiological one, relating to the nutritional mode (autotroph, he-terotroph, mixotroph, etc.) which does not correspond with the classic taxonom-ic or “trophic level” boundaries [21]. Furthermore the body sizes of the differ-ent taxa are important features for their position within the food webs.

In many cases bacteria are grazed upon mainly by small heterotrophic fla-gellates, the heterotrophic nanoplankton (HNAN), which in most cases turnedout to be the most efficient predators of bacteria that were able to control thebacterial populations even during their highest productivity (e.g., [1, 22]).Berninger et al. [1] found a clear correlation between the abundance of bacteriaand HNAN in comparing samples from more than hundred freshwater sites ofdifferent trophic states. The numbers of the two groups of organisms differedby two or three orders of magnitude, with maxima of more than 106 specimenof HNAN and 109 specimens of bacteria per ml. They inferred predator-prey re-lationships between these groups.

HNAN are sometimes grazed upon directly by metazoa, while in other bo-dies of water ciliates constitute the main predators [17, 23]. Heterotrophic fla-gellates, possessing high turnover rates, inhabit a central position in the trans-fer of organic carbon in most microbial food webs.

But what about the ecological roles of ciliates? In some cases, especially in pro-ductive waters, ciliates can also graze effectively on picoplankton and can even bethe most important bacterivores, taking a key position for the transfer of matterto the metazoan links [23]. However, smaller bacterivorous ciliates with highgrazing efficiencies need a threshold abundance of bacteria to persist on this diet.Beaver and Crisman [24] gave an estimate that small ciliates (20–30 mm) were“largely excluded from lakes having <5 ¥ 106– 5 ¥ 108 bacteria ml–1 – a concen-tration normally found only in more productive systems.” Large ciliates(> 50 mm), being mainly phytophagous and grazing on nanoplanktic algae, do-minate the ciliate assemblages in oligotrophic lakes, with low bacterial abun-dance. Mixotrophic ciliates with endosymbiotic algae can even contribute sub-stantially to pelagic autotrophic biomass in some lakes (15% of annual total [25]).

The overall number of planktonic ciliates in lakes is correlated with the tro-phic state of the water bodies.While under oligotrophic conditions abundancies


of 3–10 cells ml–1 were recorded, 90–215 cells ml–1 were recorded in hypereu-trophic waters [25].

The length of the food chain originating from bacteria and Cyanobacteriaand the identity of links involved is important to the still unresolved questionas to whether the microbial loop is acting as a link or a sink for organic mate-rial. Adherents of the latter position argue that a microbial food chain with foursteps will be unlikely to transfer any substantial amount of organic carbon tothe metazoan part of the web [15, 26, 27]. The answer to this question is depen-dent on several variables. Besides the trophic states of the waterbodies, otherabiotic variables such as temperature and acidity are relevant for the specificpatterns of the microbial web [25] and also the species composition of thewhole food web [28].

In some cases organic material is transferred from picoplankton via he-terotrophic flagellates to larger ciliates and then to crustaceans or other meta-zoans. In other cases crustaceans may directly feed on nanoplankton, while ci-liates are of minor importance [29]. Even though most metazoans cannot feedeffectively on small particles of the order of few mm, some freshwater species, inparticular cladocera of the genus Daphnia, can effectively control bacterialabundance (although they may not persist on bacteria alone), thus shortcuttingthe microbial loop [17, 28]. The presence or absence of a single species can thuschange the pathways completely, deciding the coupling or decoupling of themicrobial loop from the metazoan web. The proportion to which differentgroups of organisms contribute to different nutritional types in a lake is alsoseasonally variable [17, 20, 28].

In this regard, the scheme displayed in Fig. 3 comes closer to the perceivedprocesses than many other representations, in that a multitude of pathways ispossible which may be more or less important at different times.

210 W. Pauli et al.

Fig. 3. Diagram of the food web in lake plankton. In contrast to the scheme in Fig. 2, the com-partment of protozoa has been differentiated. Note that not all pathways are realized at anyone time. See also text. DOM: dissolved organic matter

As mentioned above, the microbial loop is not only important for the trans-fer of energy in the form of organic carbon, but also for the cycling and reten-tion of nutrients. This is especially important in oligotrophic situations, wherenutrients like phosphorus and nitrogen are scarce – at least during certaintimes of the year. The phosphorus dynamics of the pelagic zone seem to bestrongly determined by the interactions of algae, bacteria, and protozoan gra-zers. Algae and bacteria compete for P, with bacteria being more efficient in theuptake of P. Bacterial grazing by protozoa was demonstrated to enhance phos-phorus turnover and mineralization [30]. As grazed bacteria populations growfaster their excretion of P also becomes stronger. Furthermore, protozoan gra-zers increase the amount of organic P by excretion, which seems to be of spe-cial importance for phytoplankton [31]. Although this compound is also excret-ed by micro- and macrozooplankton, the high metabolic rate of protozoa leadsto higher excretion rate of this group of organisms. Buechler and Dillon [32]estimated that if ciliates only contribute 1% to the biomass of a zooplankton as-semblage, they should be able to contribute 50% to the release of dissolved P.

A similar situation exists with regard to nitrogen in cases where nitrogen isa limiting factor for the growth of algae and bacteria. Bacteria can also out-compete phytoplankton for N and thus serve as a sink for nitrogen within thefood web. However, as has been demonstrated experimentally, the presence ofbacterivorous protozoan grazers leads to a partial remineralization of N and al-lows an increase in algal biomass [33]. The degree to which this process is of im-portance depends on the carbon available for the bacteria. As Caron et al. [33]concluded: “the role of bacterivorous protozoa as mineralizers of a growth-limiting nutrient is maximal in situations where the carbon:nutrient ratio of thebacterial substrate is high”.

1.4Outlook

Most of the interactions described above were investigated in the pelagic partof aquatic habitats. However, as mentioned above, many protozoa are closely re-lated to surfaces within the water bodies, be they sediments, plants, and stones,or even microscopic aggregates within the pelagic zone. In lakes or oceans themain metabolic activity is certainly associated with the pelagic zone. Regardingstreams or small water bodies, the surface-related biota gain in importance forthe fluxes of energy and materials. In streams, a true plankton only exists in theslow flowing lower reaches of large rivers. Thus, most organismic activities arefound in and on the benthic parts. Many of the aspects discussed above will alsobe valid in these environments. However, there will surely be differences.Although some data is available on the numbers and production of protozoa inthese microhabitats [34–36], our understanding of the complex web of interre-lations is much less than for the open water. To a considerable degree this seemsto be a consequence of the methodical difficulties. Benthic assemblages arehighly heterogeneous in space and time and this heterogeneity, i.e., the smallscale spatial arrangement of the different components, is by itself of importancefor the nature of the interactions between protozoa and the other parts of these


assemblages. Thus we are only just beginning to delve deeper into the compli-cated patterns and dynamics of those biofilms. There is now important evi-dence that these biofilms are also highly productive but also very retentive inregard to nutrients [37]. Nutrient pulses are retained much longer within theperiphyton assemblages of streams than would be expected on the basis of acontinuous water flow.

There are certainly many other important ways in which protozoa are in-volved in the ecology of aquatic systems. For example, little is known aboutinformational relations between protozoa and other members of the speciesassemblages, although there may be indications in this direction (e.g., [38]).Also, our view of microbial food webs may change during the next years withthe new awareness that even the pelagic zone of lakes is not as homogenous asit seems at first sight. In addition to rather macroscopic stratifications of abio-tic factors and the related stratifications of organisms, the role of tiny and – inthe realm of human time-scales – fleeting aggregates of small detritus particles,bacteria, protozoa and algae come into prominence, the so called “lake snow.”These aggregates may turn out to be hot spots of microbial activity, and especi-ally for the grazing activities of protozoa. There are data that indicate that ciliatebacterivory is especially high in lakes with high amounts of suspended organicmatter [39]. Similar to biofilms on solid substrates, the microenvironment on,in, and around these aggregates can be chemically strangely different from theaverage water column data. It remains to be seen, what these new insights willbring about for the understanding of the ecological processes in freshwaterhabitats.

2Protozoa in Wastewater Treatment

2.1Background

2.1.1Wastewater

Wastewater includes municipal, industrial, and agricultural wastewater as wellas rainwater. The relative proportions of wastewater for West Germany (1980)were 32% municipal, 47% industrial, and 1% agricultural wastewater, plus 20%rainwater run-off in areas with main drainage. All wastewater produced intowns and communities is termed municipal sewage. This expression coversdomestic wastewater (50%), extraneous water (leachates 14%), and wastewaterfrom industry and commerce (36%) [40].

Municipal sewage is treated as follows:

– Initial mechanical purification or sedimentation– Biological purification or clarification – Further purification, e.g., elimination or reduction of the nitrogen, sulfur, or

phosphate content, polishing, filtration– The treated wastewater is then discharged into the receiving stream (Fig. 4)

212 W. Pauli et al.

Protozoa in Wastew

ater Treatment:Function and Im

portance213

Fig. 4 a– c. Types of common sewage treatment plants – flowdiagram of: a activated sludge plants; b, c biofilm processes(trickling filter and Rotating Biological Contactor, RBC, re-spectively). In the activated sludge process (a) the wastewateris exposed to a mixed microbial population in the form of aflocculent suspension. In fixed medium systems the waste-water is brought into contact with a film of microbial slime (b)on the surfaces of the packing medium, (the wastewatertrickles through the bed, most commonly consisting ofstacked stones), or (c) on a partly submerged support mediumwhich rotates slowly on a horizontal axis in a tank throughwhich the wastewater flows

All substances present in sewage are classified according to their significancefor wastewater treatment plants. Organic content is of particular importance fordegradation processes. It is quoted in terms of the chemical or biochemical oxy-gen demand (COD, BOD) of the organic substances. Furthermore, a differentia-tion is made between suspended and dissolved wastewater components.

Approximately two-thirds of the total load (organic and inorganic) of muni-cipal sewage is in solution. With regard to the organic load almost half is in so-lution, the rest consists of colloidal material (25%) or is bound to particleswhich sediment (75%). Similarly, about half of the oxygen demand of biochem-ically degradable organic compounds is attributed to the dissolved fraction, ofthe other half one third to floating and two thirds to particulate matter. After a 2 h sedimentation period, two-thirds of the total organic load remains in thesupernatant (also two-thirds of the total BOD). About 25% of the dissolved or-ganic load is bound to colloids and particles which do not sediment (Table 2).Carbohydrates are not usually present in municipal wastewater plants. They aremetabolized on route in the sewage. Proteins are also hydrolyzed in the sewers.The main task of the wastewater treatment plant is then to eliminate fatty acidsand the amino acids formed by protein hydrolysis.

Municipal sewage averages an organic load of 300 mg BOD5 l–1 (ca. 450 mg l–1

organic content). Activated sludge plants aim for effluent values < 20 mgBOD5 l–1, i.e., a reduction in the organic content of more than 90% [41]. For in-dustrial – as opposed to municipal – wastewater, no generalizations can bemade regarding type and amount of load. Diverse organic and inorganic loadsare produced by different industrial sectors. Even within a sector values varyaccording to the production methods and environmental requirements.Wastewater from the chemical industry often exhibits toxic or inhibitory ef-fects.

2.1.2Biological Treatment Processes

It is well known that a microbial degradation of organic substances takes placein natural flowing waters. This natural, self-purifying capacity of water becameovertaxed by the increase in population and industrialization. Attempts werethen made to pre-treat partially or fully sewage by mechano-biological pro-cesses, before discharging it into the surface water.

214 W. Pauli et al.

Table 2. Average contribution of settleable (sedimentation within 2 h) and non-settleablematter and their respective biochemical oxygen demand (BOD5) to the total organic load ofmunicipal sewage, according to [157]

Organic load (in Æ Settleable: 33% (w/v) or –total ca. 450 mg/l) 150 mg/l, 33% (BOD)

Æ Non-settleable: 67% (w/v) Æ Dissolved: 83% (w/v)or 300 mg/l, 67% (BOD) or 250 mg/l, 75% (BOD)

Æ Suspended: 17% (w/v) or 50 mg/l, 25% (BOD)

A conscious use of biological degradation began after bacteria were discov-ered in the nineteenth century. Two principles were implemented: activated andfixed-bed processes. The latter have been in use since 1882 and utilize the slimegrowth of organisms in the receiving stream. The activated sludge process,which takes advantage of the self-purification properties of the suspended or-ganisms in the receiving water body, was developed in 1913, and the firstGerman plant was operational in 1926 [42]. Both methods are still in use today.In Germany the activated sludge technique has taken precedence, due to itshigher performance capacity, particularly for extended wastewater treatmentincluding nutrient elimination. However fixed-bed reactors in combinationwith activated sludge techniques are finding increased application today. Assubmerged aerators they increase the active biomass and the age of the sludgein activated sludge plants, making a positive contribution to the purification ef-ficiency [43].

The underlying principle of biological wastewater treatment is to transformthe majority of dissolved and suspended substances into biomass which canthen be removed either by sedimentation (activated sludge) or by fixing (sub-merged aerator contactors). In this way, a nutrient concentration exceeding thedegradation capacity of local surface waters, resulting in disruption or even de-struction of natural biological systems, can be avoided: Direct discharge of sub-stances would result in anaerobic or aerobic burdening of the sediment of sur-face waters; high oxygen consuming, organic content (BOD5) in the effluent canovertax the oxygen household of the water, through its rapid conversion by he-terotrophic organisms; direct discharge of plant nutrients, particularly nitrogencompounds and phosphates, encourages algal growth, with negative effects onthe water (larger pH- and O2-fluctuations, sludge formation). At the same time,however, the discharge of bacteria – used for the fixation of wastewater sub-stances – should be kept to a minimum.

All biological processes have in common that they involve sectors of naturalmetabolic cycles. In wastewater treatment plants, the only difference from na-tural processes is that part of the reaction chain is technically controlled. Theperformance is dependent not on one specific species with a high degradationcapacity, but on the interaction of a wide range of different organisms. Over thelast 20 years the traditional model of a vertical material and energy flow, start-ing from nutrients through to decomposers and primary producers and bothprimary and secondary consumers, has been replaced by a more complex eco-logical web, which takes into account the network of microbial systems andtheir significance for turnover of matter (see Sect. 1.2).

In treatment plants, due to the high organic content of the wastewater, a bio-coenosis of organisms forms, primarily made up of members of the group ofdecomposers, i.e., saprophytic bacteria. The majority of the bacteria degradedead organic matter, in the presence of oxygen, to carbon dioxide and water.Nitrogen is released in the form of ammonia. Bacteria are significant in waste-water treatment due to their large surface area in relation to their body volumeand their associated high metabolic and reproductive rates. Apart from theseprokaryotic forms of life, protozoa (unicellular, animal organisms) are the nextmost important group of organisms in the wastewater biocoenosis. Together


with bacteria they form a closely related microbial system which forms the ba-sis of the so-called natural self-purification process.

2.1.3Bacterial Biofilms

In both fixed-bed and activated sludge processes, microbial biofilms – either asslime growth or flocs – are fundamental for the turnover of organic waste. Thecolonization of surfaces by bacteria is a widespread process in the environment.In natural biotopes, bacteria favor the colonization of suspended particles andsediment. By far the majority (99%) of all bacteria in the environment adhereto surfaces such as stones, sediment, and soil. Important physico-chemical pro-cesses, forming the basis for the biomass layer, precede the attachment of a bio-film. Dissolved organic molecules (polysaccharides, proteins, humic acids) ac-cumulate spontaneously on the surface of very different materials forming a“conditioning film,” on which bacteria colonization follows. The cells are im-mobilized and produce extra-cellular polymeric substances which anchor theorganisms to the surface and to each other. Embedded in this matrix, microbialcommunities of complex composition are built up, usually in several layers.Biofilms are not static systems, rather a dynamic equilibrium exists betweenfreely suspended bacteria and those adhering to particles. From the moment abacterial biofilm forms, a detachment of cells or cell-aggregates takes place [44],dependent on the prevailing conditions. Several bacteria species, dependent ontheir nutrient supply, can exist either freely suspended or mainly aggregated inboth pure and mixed cultures [45].

2.1.4Activated Sludge

Existing literature regarding protozoa and wastewater treatment deals mainlywith aerobic processes, with the focus on activated sludge technology. This is due to the significance of this technology for wastewater treatment on the one hand and that suspended activated sludge is more easily accessible for bio-logical investigations than slime-growth areas of fixed-bed reactors on theother.

Activated sludge processes operate with typical sludge concentrations be-tween 2–3 g l–1 [46]. About 70% of the activated sludge is organic content and30% inorganic (clay: Si; Al; Fe; ferric oxide; calcium phosphate) [47]. Non- – ornot easily – oxidizable organic matter makes up 20–25% of the sludge [41].

In a conventional activated sludge tank flocculate suspended material con-tains about 6 ¥ 109 bacteria ml–1, i.e., 1–3 ¥ 1012 bacteria g–1 dry weight [48].They represent about 90% of the total biomass of the activated sludge. The pro-portion of living or metabolically active bacteria found in the flocs varies con-siderably, depending on the method of analysis. Estimates based on glucose,stearate and acetate uptake rates imply active proportions of 8–13%, 14–28%,and 5–10% of the total biomass, respectively [48]. More recently, direct mea-surements by fluorescence-microscopy indicate a proportion of 35–40% (de-

216 W. Pauli et al.

hydrogenase activity [49]) and 70% (rRNA directed oligonucleotide probes,[50]), whereby a similar level of activity was assumed for all zones of the floc[51].

2.2Protozoa in Biological Wastewater Treatment Plants

2.2.1Occurrence

Systematic investigations at a large number of wastewater treatment plants re-veal protozoa as typical components of the biocoenosis (Table 3). Thus, for ex-ample, in all ten South African activated sludge plants studied by Bux and Kasan[52] “basic communities” of protozoa, typical for sewage plants were found.Similarly, Curds and Cockburn [53] found protozoa biocoenoses in 53 of 56British activated sludge plants and all 52 biological percolation filter plantsstudied. In New Jersey, Chung and Strom [54] found protozoa in all the rotatingdisc contactors and according to Madoni and Ghetti [55], typical ciliate com-munities were detected in 38 of 39 activated sludge plants and 47 of 49 rotatingdisc contactors in the Emilia region of Italy. The presence of protozoa is closelyassociated with biofilms and restricted mainly to aerobic processes and there-fore to certain areas of the wastewater treatment plant; only a few specialistsamong the protozoa take part in anaerobic processes. Thus protozoan commu-nities can be typically encountered in activated sludge tanks as well as in the se-dimentation tanks, whereas no protozoa are found in sludge digestion or in thesupernatant of the sedimentation tank (effluent), with the exception of malfunc-tions [56].


Table 3. A survey of the protozoan fauna in sewage treatment plants (only microfaunistic in-vestigations based on ten and more plants are taken into consideration), according to [52–55]

Type of plant No. of plants Occurrence of Typical Protozoainvestigated typical protozoan protozoan absent(country) communities communities

absent

Activated sludge 56 (Great Britain) Within 53 plants 2 plants a 1 plant39 (Italy) Within 38 plantsb 1 plant b ? 10 (South Africa) Within all 10 plants – –

Trickling filter 52 (Great Britain) Within all 52 plants – –Rotating biological 49 (Italy) Within 47 plants b 2 plants b ? contactor

10 (USA) Within all 10 plants – –

a No ciliates, but flagellates present.b Only ciliates investigated, no comments on other protozoan groups such as flagellates and

amoebae.

2.2.2Species Composition

The majority of microfaunal investigations confirms that all of the three maingroups of protozoa – flagellates, ciliates, and amoebae (naked and shell) – canbe found in wastewater treatment plants, whereby ciliates form the largest pro-portion with regard to biomass and number of species, both in activated sludge[53, 57–62] and in fixed-bed processes (percolation filters: [53, 59]; rotating disccontactors: [63–65]), compare Table 4.

It should be noted, however, that the composition of the protozoan biocoe-nosis, as well as that of the total biomass involved in the purification process,is mainly dependent on the composition of the wastewater, together with phy-sical conditions and factors arising from the process technology used. In thecase of malfunctions, or in the initial stage of a plant, very different composi-tions can be encountered. Sydenham [57] observed 2 municipal activated sludgeplants over a period of 12 months and identified amoebae as the dominantgroup with regard to biomass. In sludge with a high organic load, Curds andCockburn [66] and Mudrack and Kunst [67] report high population densities offlagellates. The age of the sludge also has an effect on the composition of theprotozoan community. Kinner and Curds [63] quote 6–12 months as the lengthof time required to establish a steady-state community of protozoa in a pilotrotating disc contactor plant supplied with domestic effluent. Bacteria were vi-sible on the disc surfaces within one day of startup followed within a few daysby flagellates and small amoebae. Free-swimming bacterivorous ciliates appear-ed within 8–10 days. Subsequently, sessile peritrichous forms accompanied bycarnivorous ciliates, rotatoria, and large amoebae make up the stable commu-nity. Parallel to sludge aging, a typical chronological succession of dominantprotozoa populations can also be observed in activated sludge plants. After theinitial phase of 1–2 weeks where flagellates, naked amoebae, and free-swim-

218 W. Pauli et al.

Table 4. Structure of the protozoan community in three urban activated-sludge plants, oper-ating at different organic loading rates and dissolved oxygen concentrations (observationover a one year period), according to [62]. Biomass calculation is based on data, given by [61]

Plant 1 Plant 2 Plant 3

Organic load a 0.23–0.38 0.21–0.35 0.5–0.8O2-conc. (mg O2/l) 3.6–5.2 1.8–3.0 1.0–1.3Densities and biomass ind./ml mg/l ind./ml mg/l ind./ml mg/lCiliates 3000– 18–43 8600– 50–99 4500– 26–93

7400 17 000 16000Flagellates (< 20 mm) 43 000– 2.2–5.2 89000– 4.6–51 38000– 20–83

600 000 980 000 1 600 000Naked Amoebae (<50 mm) 4000– 0.21–5.3 800– 0.04– 77000– 4.1–5.4

100 000 130 000 6.9 101 000

a kg BOD5/(kg MLVSS) day.

ming ciliates predominate, more and more crawling and sessile forms appear,which remain dominant throughout the stabilization phase and can be regard-ed as typical representatives of mature sludge [62, 65, 68–70]; see also Fig. 5.Unlike the free-swimming forms, which arrive at the plant with the sewage andare flushed out at the end of the process, the existence of sessile and crawlingforms is closely associated with the development of slime growth or sludge


Fig. 5. Composition of the bacterivorous ciliate community during the establishment of amature sludge. Stabilization, i.e., steady-state occurs after about 50 (activated sludge, above fi-gure) and ca. 80 days (RBC, below figure), respectively. Bars lower than 100% indicate the ad-ditional presence of carnivorous and omnivorous ciliates, after [65]

flocs. Bound to biofilms as fixed slime growths (fixed-bed) or as sedimentablesludge, they are retained in the treatment plant and can thus build up a stablecommunity with the bacterial flora. Whereas characteristic population succes-sion takes place in both plant types, in percolation filters, due to the unequaldistribution of the organic load, a physical separation of the organisms is ob-served, dependent on the filter depth [68].

Figure 5a, b shows results from studies on the colonization behavior of cili-ates in a pilot rotating disc contactor plant as well as in an operational activatedsludge plant [65]. Both plants were fed with domestic wastewater. Whereas inthe initial stage of the activated sludge plant ciliates make up between 0.17%and 0.44% of the total biomass, in the stabilizing phase they account for morethan 9% of the sludge biomass. In the initial phase free-swimming forms fromthe wastewater dominate. After 10–15 days their numbers drop markedly andcrawling (Aspidisca cicada, A. lynceus, Euplotes affinis, Chilodonella uncinata)as well as sessile (Vorticella convallaria, V. microstoma, Epistylis plicatilis,Opercularia coarctata) ciliates characterize the protozoan fauna. Similarly, in the rotating disc contactor plant, ciliates makes up only 4–5% of the slime bio-mass in the colonization phase, as opposed to 12–19% under steady-state con-ditions. Here, too, essential changes take place during the colonization of thesubmerged contact aerator and the typical ciliate biocoenosis develops in theplant itself. In the initial phase, free-swimming ciliates such as Parameciumputrinum and Uronema nigricans are present; in the stable phase sessile formssuch as Opercularia coarctata and Vorticella convallaria dominate. Investiga-tions by Madoni [64, 65] make it clear that in both types of plants (submergedcontact aerator and activated sludge) a significant positive correlation exists between the increase of the sludge, biofilm and ciliate biomass (r2 = 0.927 andr2 = 0.853). This implies a close relationship between the size of the ciliatepopulation and the bacterial biomass.

2.2.3Plant Specific Basic Communities

The relative abundance of an organism in a particular habitat can be consider-ed as a measure for its significance within the ecological structure of the bio-logical system concerned. Alongside amoebae and flagellates, Curds andCockburn [53] identified 67 and 53 ciliate species in 56 activated sludge plantsand 52 percolation filter plants in Britain, respectively. Madoni and Ghetti [55]detected 45 and 47 ciliate species in 39 activated sludge plants and 49 percola-tion filter plants in Northern Italy. Of note is that the British and Italian activat-ed sludge plants revealed very similar ciliate fauna [55]. Nevertheless, not allspecies in the individual samples can be regarded as typical, as to their presenceand population density, for the respective wastewater treatment process. Themajority of the species are found only sporadically in a few samples and usuallywith a low population density. The overall picture of the ciliate population is de-termined by a few, primarily sessile (peritrichous) and crawling (hypotrichous),species most of which are bacterivorous (compare with Fig. 6). With cell countsof, on average, more than 104 ml–1, ciliate densities are 100–1000 times higher

220 W. Pauli et al.

here, than in the plankton of oligotrophic (10 ml–1) and eutrophic (100 ml–1)waters [24]. Table 5 summarizes the dominant ciliate species in the “basis com-munity” of each plant type identified by Curds and Cockburn [53] and Madoniand Ghetti [55]. The specific biocoenosis differs according to plant type and tothe current operating conditions [55]: in areas with a high organic load an in-crease in free swimming species is observed [61, 66] along with a decrease in thediversity of species [71, 72]; with these limitations, the community forms givenin Table 5 can be considered average for municipal plants.

2.2.4Biomass

In activated sludge plants a high proportion of the eukaryotic biomass is com-prised of protozoa. Investigations carried out by Sydenham [57] revealed thatprotozoa made up over 90% of the total eukaryotic biomass of two municipalwastewater treatment plants. According to Aescht and Foissner [61], protozoamade up 99–100% of the eukaryotes in a pharmaceutical plant with a bacterialnutrient load. The average proportion of protozoa in relation to total solids(dw) is 5% [59, 73]. Ciliates alone make up 10% of the total biomass (pro- andeukaryotic dry weight). Even higher numbers of ciliates are encountered in mu-nicipal rotating disc contactors where proportions of about 20% of the totalbiomass of the slime-growth can be observed [64, 65].

2.2.5Ecological Framework

The biocoenosis in wastewater treatment plants should not be regarded as acommunity with a rigid composition and constant characteristics but rather as


Fig. 6. Examples of free swimming (holotrichous), crawling (hypotrichous), and sessile (pe-ritrichous) ciliates in waste water treatment plants

222 W. Pauli et al.

Table 5. Ciliate species dominating and occurring with a high frequency in sludge samples ofBritish (GB) and North Italian (I) sewage treatment plants, respectively, after [53, 55].“Dominating” refers to the relative cell density, whereas “present” indicates the number ofsamples, in which the respective species – independent of its individual numbers – could beobserved

Dominant Present Life form Nutrition a

(%) (%) Ecological typeGB I GB I

Activated sludge (GB and I)Aspidisca costata a 35 85 69 90 Crawling BacterivorousVorticella convallaria a 19 77 58 84 Sessile BacterivorousTrachelophyllum pusillum a 15 30 64 58 Free swimming CarnivorousOpercularia coarctata a 12 23 54 25 Sessile BacterivorousCarchesium polypinum a 11 26 25 28 Sessile BacterivorousVorticella alba (GB) 11 – 38 – Sessile BacterivorousVorticella microstoma (GB) 10 10 75 10 Sessile BacterivorousEuplotes moebiusi (GB) 5 5 35 7 Crawling BacterivorousVorticella fromenteli (GB) 4 – 31 – Sessile BacterivorousEuplotes affinis (I) – 59 11 69 Crawling BacterivorousZoothamnium pygmaeum (I) – 33 – 33 Sessile BacterivorousTrochilia minuta (I) 2 23 12 25 Crawling Filamentous b

Trickling filter (GB)Opercularia micodiscum 44 81 Sessile BacterivorousCarchesium polypinum 15 62 Sessile BacterivorousVorticella convallaria 10 83 Sessile BacterivorousChilodonella uncinata 4 90 Crawling Filamentous b

Opercularia phryganeae 4 90 Sessile BacterivorousOpercularia coarctata 2 56 Sessile BacterivorousVorticella striata 2 52 Sessile BacterivorousAspidisca costata – 56 Crawling BacterivorousCinetochilum margaritaceum – 54 Crawling Bacterivorous

Rotating biological contactor (I)Euplotes moebiusi 53 79 Crawling BacterivorousParamecium caudatum 46 79 Free swimming BacterivorousTrachelophyllum pusillum 41 59 Free swimming Carnivorous Vorticella convallaria 53 57 Sessile BacterivorousOpercularia microdiscum 41 45 Sessile BacterivorousOpercularia coarctata 33 37 Sessile BacterivorousParamecium trichium 27 43 Free swimming BacterivorousCinetochilum margaritaceum 23 37 Crawling BacterivorousChilodonella cucullulus 18 41 Crawling Filamentous b

a Dominant both in British and Italian plants.b Filamentous: ciliates with a specialized oral apparatus, enabling the ingestion of rod-shap-

ed, filamentous bacteria.– not present.

an artificial but biological segment of natural self-purification processes, thecomposition of which is influenced by ecological conditions and physico-chem-ical factors, thus differing from plant to plant and even within a plant over time.

2.2.5.1Sludge Loading

Sludge loads with fm-values between 0.2 and 0.6 [g BOD (g MLSS · day)–1] areconsidered optimal for the purification sequence at conventional municipal ac-tivated sludge plants (e.g., [47, 67]). Ciliate densities of 6000–30,000 ml–1 arefound in sludge with these loads [71, 74]. However, similar concentrations ofciliates are also encountered in sludge with both higher and lower loads:Salvado and Gracia [71] observed a constant ciliate population density in a mu-nicipal plant with fm-values varying from 0.03 to 0.4. Experiments by Lee et al.[74] confirm only slight changes in ciliate counts at sludge loadings between0.1–1.4 [g BOD (g MLVSS day)–1]. Only under very heavy loads [1.8–2.4 g BOD(g MLVSS day)–1], was a reduction in cell density observed.

Although the population density remains constant over a wide range, theorganic load influences the number of species and the composition of domi-nant ciliates in the basis community. The number of species present sinks withincreasing organic content of the wastewater [66, 71, 72]. According to Curdsand Cockburn [66], activated sludge with a relatively low organic load[fm= 0.1–0.3 g BOD (g MLSS day)–1] shows the greatest species diversification,whereby all three groups of ciliates – peritrichs (sessile), hypotrichs (crawling),and holotrichs (free swimming) – are represented with approximately the samenumber of species. In the medium load range of fm= 0.3–0.6, peritrichous spe-cies dominate and by high organic loads of fm= 0.6–0.9 equal portions of peri-trichs and holotrichs are present (Fig. 7).

2.2.5.2Temperature

Temperatures in municipal plants are generally slightly above the outside tem-perature in winter and slightly below in summer. Performance is optimal be-tween 10 °C and 25 °C [41]. No negative effects on ciliate fauna are found up to30 °C; experimental activated sludge investigations reveal a decline in ciliates attemperatures above 30 °C and their disappearance above 40 °C [74]. The authorsdiscuss the concomitant deterioration of the settling properties of the sludge aspossibly resulting from the collapse of the ciliate population.

2.2.5.3pH-Value

Activated sludge has a relatively high buffer capacity. If no strongly acidic or al-kaline effluents are introduced, mainly from industrial processes, pH-valuesgenerally fluctuate between 6.5 and 8 [41, 67, 75]. Therefore not only the tem-


perature, but also the pH-values of municipal plants are in a favorable range forprotozoan growth [75].

2.2.5.4O2-Content

Conventional processes of biological wastewater treatment utilize the meta-bolism of the organic load, which is faster, more thorough, and easier to controlunder aerobic conditions. Aerobic conditions are also a prerequisite for a highincidence of protozoa. Few specialists can survive strictly anaerobic conditionsand little knowledge is available regarding their distribution or function in an-aerobic degradation processes. The number of facultative anaerobic protozoa isslightly higher, but almost all species seem to be able to survive low oxygen con-centrations or even the absence of oxygen, at least for a short period [75]. Apartfrom plant malfunctions (e.g., breakdown of the aeration), this ability is alsoimportant in the normal cycle of activated sludge processes, where the organ-isms are constantly alternating between the aerobic activated sludge tanks andthe sedimentation tanks, in which anaerobic conditions arise for short periods

224 W. Pauli et al.

Fig. 7. Composition and species number of ciliates in activated sludge plants operated at dif-ferent sludge loadings [food to micro-organism (F/M) ratio]. Results from an investigation of52 British plants made by Curds and Cockburn [53]. Peritrichous, hypotrichous, and holo-trichous ciliates represent sessile, crawling, and free swimming ciliates, respectively. In con-ventional municipal plants, treating domestic wastewater, a sludge loading between 0.2 and0.6 is regarded to be optimum for the functioning of the sewage treatment process

in the deeper layers of the settling sludge (less than 4 h [41]). Only longer andrepeated oxygen deprivation over several hours (continual alternation between6 h aerated and 24 h without aeration) leads to a marked decline of the ses-sile ciliates Vorticella convallaria and Opercularia coarctata, typically found inwastewater treatment plants [76].

2.3Significance of Protozoa for Wastewater Treatment

As already described (Sect. 2.2.2), the majority of protozoa in aerobic biologicalpurification systems are sessile or crawling ciliates. Whereas free-swimming ci-liates are flushed out with the clarified water, crawling and especially sessileforms are bound to bacterial biofilms (flocs and slime growths) [59]. In the caseof fixed-bed plants they remain bound to the biofilms in the plant; in activatedsludge processes they sediment with the sludge and are retained in the plantdue to continual sludge recycling.

To understand the role of protozoa and classify their position in the artificialsystem of biological wastewater treatment, the following characteristics have tobe considered: type of motion (free swimming, crawling, or sessile); form ofnutrition (e.g., filter-feeders, browsers); sources of nutrition (abiotic colloidsand particles, bacteria, algae, other protozoa). From their form of nutrition andtheir trophic level, functional aspects important for wastewater treatment be-come apparent. New understanding of natural systems as well as experimentalresults on the physiology, energy budget, and nutrient cycling of both aquaticand terrestrial protozoa provide extensive information regarding the ecologicalrole of this group of organisms, which, although quantitatively less significantthan bacteria, make a considerable contribution to wastewater treatment.

2.3.1Nutrition

Several possibilities are open to ciliates for nutrient-uptake. On the one hand,similar to bacteria, substances can be transferred directly through the plasmamembrane into the interior of the cell. Active and passive, carrier-mediateduptake mechanisms through the plasma membrane have been described forTetrahymena for amino-acids [77–79], di-peptides [80], acetate, glucose [81,82], and even for such complex nutrient solutions as proteose-peptone-yeast ex-tract (PPY) medium [83]. Another method of nutrient uptake is pinocytosis[84, 85]. It describes the active transport of dissolved substances in sub-micro-scopic, particle-free vacuoles or vesicles from the plasma membrane to the cellinterior, where they undergo normal lysosomal digestion processes. Finally,ciliates have a highly specialized oral apparatus for taking up particulate mat-ter by phagocytosis. The particles are not simply ingested with the surroundingsolution but rather undergo a highly efficient filtration process, facilitating theconcentration of particulate matter from a large volume of liquid, prior to theirintake in food vacuoles [85]. This process involves the production of a watercurrent by cilia (Fig. 8) and the extraction of particles from the flowing water


with the aid of a ciliary sieve, which retains – in the case of bacterivorous spe-cies – particles sized between 0.3 mm and 5 mm [85–87]. The particles, thus con-centrated, are subsequently ingested. Apart from food, abiotic and even indige-stible matter of the size of bacteria are efficiently ingested [86–89]. Parameciaconcentrate food particles in this manner in their oral cavity up to 1000-fold[90]. A similarly high concentration capacity can be assumed for Tetrahymena:Whereas a volume of 50–80 nl is cleared of particles per hour and cell [87, 91],a more than 1000 times lower water volume of 36 pl h–1 and cell is actually in-gested by the food vacuoles [92]. The efficiency of this form of nutrition is un-derlined by investigations comparing the growth kinetics of Tetrahymena pyri-formis with particulate and dissolved substances as nutrient source, respec-tively [93]. While under monoxenic conditions with particulate bacterialsubstrate the half maximum growth rate is already attained with a bacteria con-tent of 12 mg l–1 Klebsiella aerogenes (5.5 mg carbon l–1), 200 times that con-centration of organic matter is required in case of dissolved nutrients (2.4 g l–1

proteose-peptone-yeast medium = 1.3 g carbon l–1).

226 W. Pauli et al.

Fig. 8. Mechanisms of filter-feeding (ambiguously often referred to as ‘grazing’) used by pro-tozoa.Water currents are created by flagella or the coordinated activity of cilia, that bring sus-pended food to the mouth region of the cell

2.3.2Reduction and Elimination of Suspended Particles and Bacteria

2.3.2.1Clearing Rate

The volume of water cleared per individual and hour depends on cell size. Smallprotozoa with cell diameters of less than 5 mm, such as flagellates, filter less than1 nl h–1 at temperatures between 9°C and 17°C [20]. Higher filtration rates areobserved for larger ciliates. Sanders et al. [20] quote a yearly fluctuation rangeof 12–156 nl h–1 for the filtration performance of planktonic ciliates. In labora-tory experiments with the ciliates Halteria grandinella (diameter: 25 mm) andStrombidium sp. (size: 15 ¥ 21 mm) filtration rates of 80–90 nl h–1 at 9°C and120–140 nl h–1 at 17°C were determined. In the case of Vorticella microstoma(average cell dimensions: 60 ¥ 30 mm), a ciliate frequently present in wastewa-ter treatment plants, filtration rates as high as 156 nl h–1 at bacteria densities of106 ml–1 are reported. Tetrahymena (cell dimensions: 40 ¥ 20 mm), a speciespresent but not dominant in wastewater treatment plants, has a filtration per-formance of 80 nl h–1 [91]. Fenchel [87] observed filtration rates of 50 nl h–1 andcell at 20–22°C for Tetrahymena pyriformis and 200–1000 nl h–1 for larger(100–200 mm) representatives of crawling and free-swimming ciliates such as Euplotes, Paramecium, or Blepharisma. Assuming average filtration rates of100 nl h–1 and cell and ciliate densities of 10,000 ml–1 and above [61, 94–97], thisimplies that the entire liquid of an activated sludge plant can be filtered in lessthan 1 h. The enormous predator and selection pressure exerted on the bacteriais illustrated by the following examples.

Many heterotrophic bacteria in activated sludge have the ability to divideevery 20–40 min under optimal laboratory conditions [41, 48]. Under “field”conditions, such as those prevailing in wastewater treatment plants, theirgrowth is generally much slower due to sub-optimal physical (temperature) and physiological (nutrients, pH-values) parameters. The actual bacterial divi-sion rates under constant operating conditions and good nutrient availabilitycan be estimated from the ratio of the surplus (drawn off) sludge to the totalsludge in the activated sludge plant [98]. For low to high organic loads(fm= 0.05–0.6 g BOD per g MLSS and day), growth rates can vary from 4–50%per day [41] or, expressed in other terms, the bacteria population in the sludgedoubles every 48 h at most, i.e., in a time span by no means adequate to com-pensate for potential protozoan feeding.

Highly loaded wastewater contains ca. 106 bacteria ml–1. The majority aremedically harmless but others are pathogenic and bear health risks. Con-ventional wastewater purification involves an initial pre-clarification step of20–30 min, after which the wastewater is fed into the activated sludge tank andaerated for 4 h. In the aerated and agitated system of the activated sludge tankthe wastewater is brought into contact with a mixed microbial population in theform of a flocculent suspension.When the desired degree of treatment has beenachieved, the flocculent microbial mass, known as the “sludge”, is separated for2–4 h from the treated wastewater in a separate, specifically designed sedimen-


tation tank. The supernatant from the separation stage is the treated waste-water, and should be virtually free of sludge. Most of the settled sludge from theseparation stage is returned to the aeration stage to maintain the sludge con-centration in the aeration tank at the level needed for effective treatment and toact as a microbial inoculum. Some of the sludge is removed for disposal, and isknown as “waste” or “surplus” sludge. In both the activated sludge and the sedi-mentation tanks, the resident ciliate community has sufficient time to filter theentire wastewater several times, thus removing bacteria and abiotic particles ofsimilar size (see Sects. 2.3.1 and 2.3.2).

2.3.2.2Experimental Findings

It has long been known that protozoa are present in wastewater treatmentplants and that their species composition reflects the prevailing conditions inthe plant. However, scientific opinion was less unanimous with regard to the ac-tual contribution of protozoa to the purification process. Although Ardern andLockett [99], Pillay and Subrahmanyan [100], Pillay et al. [101] and McKinneyand Gram [102] referred to a connection between protozoa and the quality ofthe water discharged from the plant, proof of a causal relationship was lackingor inconclusive.

Curds et al. [103] succeeded in selectively removing protozoa from activatedsludge and further cultivating this protozoan-free sludge in bench-scale treat-ment plants over a long period. Through the subsequent re-introduction of ty-pical sludge ciliates they observed, under various starting conditions, positiveeffects on a series of parameters describing the success of the purification pro-cess (Table 6). The principal observation of their experiments was that in theabsence of protozoa the effluent of the plant was turbid, due to its high contentof suspended bacteria; this turbidity almost disappears after re-introduction ofthe protozoa (Fig. 9).

Similar findings are published by Sridhar and Pillai [104] and Macek [105] inprotozoan-free, pasteurized sludge and in bacteria cultures isolated from ac-tivated sludge. The addition of sessile, crawling, and free-swimming ciliates

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Table 6. Effects of ciliated protozoa on the effluent quality of bench-scale activated-sludgeplants. Results are given in mg l–1 unless otherwise noted; after [103]

Effluent analysis Without ciliates With ciliates Mean reduction

BOD 53–70 7–24 75%COD 198–250 134–142 38%Permanganate value (4 h) 83–106 62–70 30%BOD after filtration 30–35 3–9 81%COD after filtration 31–50 14–25 39%Organic nitrogen 14–21 7–10 51%Suspended solids 86–118 26–34 71%Optical density at 620 nm 0.95–1.42 0.23–0.34 76%Viable bacteria counts (106 ml–1) 160 1–9 97%

(Epistylis articulata, Vorticella microstoma, Aspidisca cicada, Chilodonella unci-nata, Stylonychia putrina, Colpidium camylum) reduces high COD values andsuspended matter content. Farrah et al. [106] confirm the causal relationshipbetween the presence of ciliates and a clear, almost bacteria-free effluent with alow organic content. Departing from a typical pro- and eukaryotic sludge bio-coenosis, the authors show that a largely selective reduction of protozoa, by theaddition of sodium fluoride (0.2 mol l–1) or sodium azide (6–20 mmol l–1) re-sults in a notably higher content of freely suspended bacteria including strep-tococci. After application of the eukaryotic cell toxins, the total count of fecalstreptococci increases about threefold and the proportion of suspended bac-teria, as compared to those bound to flocs, increases from 0.3% to 64%.Kakiichi et al. [107] made essentially the same observations. The effects of twoamphoteric detergents (orthodichlorobenzene and polyhexamethylene bigua-nide hydrochloride) with known effects on bacteria and protozoa were studiedand a causal relationship between poor quality of the outflow (increased turbi-dity and COD values) from batch cultures of activated sludge and the inhibitoryeffect (reduction in population density) on the protozoa was observed. A corre-lation between the effluent quality and the population density of protozoa isalso implied by Lee et al. [74]. Studies with a bench-scale activated sludge plant(organic load: 0.1–0.4 g BOD per g MLSS and day) show that the selective de-cline of the ciliate population density, due to running temperatures of 36°C andover (see Sect. 2.2.5.2), corresponds to a more than twofold increase in suspen-ded matter in the effluent.

Experiments with bacteria-free synthetic wastewater (e.g., [103]) exhibit thatfreely suspended bacteria, originating from the autochthonous microflora ofthe activated sludge itself, are substantially reduced in the presence of protozoa.


Fig. 9. Influence of ciliates on the bacteria content in the effluent of a bench-scale activatedsludge plant, after [103]

bacterial density

Furthermore Curds and Fey [108] observed that bacteria originating from theinfluent wastewater are also effectively removed in the presence of protozoa.After mechanical destruction of the protozoan population, by means of a ballmill, the authors determined concentrations of 6.5 ¥ 105 culturable E. coli ml–1

in the effluent of a continuously operating bench-scale activated sludge plant;after re-inoculation of the activated sludge with ciliates (Opercularia coarctata,Vorticella microstoma, Hypotrichidium conicum, Tetrahymena pyriformis) andthe establishment of a stable protozoan community, this count was reduced ten-fold to 6.3 ¥ 104 ml–1. The half life of E. coli in the activated sludge was reducedfrom 16 h to 1.8 h.

Filter-feeding ciliates in wastewater treatment plants are, in principal, notselective consumers. Along with harmless bacteria, a series of pathogenicstrains causing, for example, diphtheria, cholera, typhoid, and streptococcal in-fections are also phagocytosed (for reviews [75, 109]. Investigations by Farrahet al. [106], with activated sludge in batch cultures, illustrate the significance ofthis elimination of pathogenic bacteria from wastewater treatment plants. Afterselective reduction of the protozoan fauna by sodium fluoride (200 mmol l–1) orsodium azide (20 mmol l–1), cultures of Salmonella typhimurium and E. coli, ad-ded in densities of 105 ml–1 almost treble within 24 h (S. typhimurium and so-dium fluoride) or only decrease by ca. 50% (E. coli and sodium azide), whereasin untreated controls with protozoa, both bacteria are reduced to less than 5%of their initial density. Moreover, under conditions of aerobic sludge stabiliza-tion, the authors show that even low densities of protozoa (660 ml–1) lead to asubstantial elimination of bacteria. Figure 10 shows results with Streptococcusfaecalis.

230 W. Pauli et al.

Fig. 10. Effect of sodium azide (6 mmol l–1, selectively reducing protozoan activity) onStreptococcus fecalis in activated sludge (laboratory scale), after [106]. CFU: colony formingunits

2.3.2.3“Field”-Observations

“Field” observations leave no doubt that the results found in the laboratorymicrocosms are transferable to pilot and full-scale plants and that the presenceof a typical protozoan community is reflected by the improved quality of theplant effluent.

First, a close negative correlation is observed between the population densityof, mainly crawling and sessile, ciliate populations and the proportion of sus-pended matter in the effluent of wastewater treatment plants (domestic andmunicipal wastewater [56, 110–114] and brewery wastewater [115]). Resultsfrom a three-year investigation of three activated sludge plants with differentorganic loads in Spain [114] reveal – on average for all plants – a highly signifi-cant correlation coefficient between total ciliate population density and biolog-ical oxygen demand of r = – 0.868. In the presence of protozoa the effluent BODranges from 4 mg l–1 to 18 mg l–1, rising to values of up to 67 mg l–1 in their ab-sence. An almost identical correlation between effluent quality (COD) and thepopulation density of typical activated sludge ciliates was observed by Sudo andAiba [111] for six municipal wastewater treatment plants in Tokyo. Mean CODvalues of 10 mg l–1 were found with ciliate densities of ca. 104 ml–1; these in-crease to 40 mg l–1 when ciliate densities drop to 102 ml–1 (Fig. 11).

Second, according to Curds and Cockburn [53], plants without ciliates can berecognized by the low quality of their effluent: 3 out of 53 activated sludgeplants were selected due to the high content of suspended material in theireffluent. In one of these plants no protozoa could be found at all, in the othertwo no ciliates, only small flagellates, could be detected. The highest BOD valuesmeasured in the three plants occurred in the plant with no protozoa.


Fig. 11. Relationship between protozoan densities and effluent COD, observed in municipalactivated sludge plants of Tokyo, after [111]. Symbols represent different plants

Finally, as compared to normal activated sludge processes, the clarifying ef-fect of protozoa in activated sludge processes with submerged fixed-bed filters– a technology which creates additional surfaces for slime growth and primar-ily sessile ciliates [116–118] – improves, which is basically due to low bacteriaand suspended matter content in the fixed-bed plant effluent [117, 119].(Evidently, protozoa find optimum living conditions on the filter installed in theactivated sludge tank, an adequate oxygen supply and plenty of food, so that thedense population of mainly ciliates even crowds out attached bacterial growths.In contrast to the common activated sludge process, where ciliates contribute toabout 10% of the total, bacteria dominated biomass, an almost inverse relationof 68% protozoan and 32% bacterial biomass (dw) is found for the biofilms ofsubmerged fixed-bed filters [116].)

2.3.3Elimination of Dissolved Substances

The bulk of dissolved substances entering the wastewater treatment plant areamino-acids, products of protein hydrolysis in the sewage system, and fattyacids. Carbohydrates are usually completely degraded in the sewage beforereaching the plant.

Although many protozoa can take up organic substances [85, 89, 120, 121],their contribution to the degradation of these substances in wastewater treat-ment plants is negligible: For these substances the essential activity comes fromthe bacteria population. They dominate the biomass and possess a higher me-tabolic efficiency as a result of their high surface to volume ratio [41, 46–48, 67].An impression of the different degradation efficiencies can be gathered frommeasurements of amino-acid uptake by Escherichia coli and T. pyriformis [122].Even under the assumption that all ciliates present in wastewater treatmentplants can metabolize not only bacteria but also dissolved substances similar toT. pyriformis, the experiments reveal an 80-times higher uptake of amino-acidsby bacteria. Results from Hrudey [123] can also be well interpreted in the lightof the significantly higher degradation rate of dissolved substances by bacteria.After addition of peptone, a protein hydrolysate rich in amino-acids, an imme-diate rise in the bacterial biomass was observed, whereas ciliates were scarcelyable to convert the available peptone into their own biomass and could only re-produce substantially after the bacterial content increased considerably.

2.3.4Flocculation and Composition of the Bacterial Community

Apart from the feeding activity of protozoa, another factor is discussed as con-tributing to the reduction of the content of suspended matter and bacteria inbench and full-scale plants. In the presence of protozoa, freely suspended, singlebacteria form compact flocs, which then settle [59, 105, 106, 111, 124–128].

This is attributed, on the one hand, to polymer, particle-aggregating excre-tion products (polysaccharides) from protozoa [59, 125], which are possibly re-leased into the media to facilitate a more effective uptake of particles [24, 129].

232 W. Pauli et al.

On the other hand, this flocculation is believed to be associated with theexocytosis of indigestible, originally finely dispersed material as a digestedbundle [130, 131], which in turn could serve as a settlement surface for solitarybacteria [132–134]. However, wastewater itself contains a high proportion ofchemically complex particles of differing sizes, and bacteria themselves, domi-nant with regard to their biomass in wastewater treatment plants, produce ex-tracellular polymeric substances (polysaccharides), to which they can effec-tively adsorb [135, 136]. For these reasons, protozoa, by excretion of digested re-mains and polymers, probably play only a minor role in floc formation inwastewater treatment plants.

Bacteria feeding itself seems, not only quantitatively but also qualitatively, asignificant stimulus for complex bacterial growth forms. As a result of the pre-dator-prey relationship between protozoa and bacteria, a collapse of the bacte-ria population in the activated sludge and a reduced elimination efficiency ofthe system as a whole would be expected (see Sects. 2.1.2 and 2.3.3). Suchcollapses or phase-shifted oscillations between predator and prey can be obser-ved in model systems [111, 137–141] and in natural ecosystems [20, 142–146]and led originally to the view that protozoa are harmful for the clarificationprocess [147].

Only a few protozoa, e.g., amoebae, mostly present at low densities in waste-water treatment plants, are principally capable of taking up larger particles, dueto their ability to entrap their prey. Ciliates, typical representatives of protozoafound in wastewater treatment processes, possess a highly specialized oral ap-paratus for highly efficient filtration, which at the same time exclude particlesof several micrometers in diameter [86]. The ability of bacteria to develop larg-er forms, to grow collectively, or to merge as micro-colonies protects themagainst the predator pressure from the protozoa [148–153]. The development ofgrowth forms resistant to filter-feeding can thus be seen as an essential processin the evolution of bacterial flocs and biofilms [45, 111, 127, 148].

To what extent a qualitative selection of floc and biofilm forming bacteria ispossible [148], and what could be gained from a quantitative shift within a spe-cies to larger or aggregating phenotypes [45, 149], cannot be decided in the lightof the present literature. Güde [148] observed selection of bacteria populationswhich aggregate in pilot wastewater treatment plants. On the other handShikano et al. [149] find that, in the presence of the ciliate Cyclidium sp., phe-notypes of considerably larger dimensions appear within a bacteria species.Gurijala and Alexander [154] provide evidence of lower feeding pressure by theciliate Tetrahymena thermophila on bacteria with hydrophobic surfaces – inother words on phenotypes with water-repellent properties – which enhancetheir adhesive, i.e., aggregation, ability [155].

Many bacteria are also capable of organizing themselves spontaneously intobiofilms in the absence of protozoa, thus forming flocs [102, 135, 156].Nevertheless, the extent and persistence of the flocs seem to be influenced bythe presence of protozoa. Farrah et al. [106] show that in the absence of proto-zoa autochthonous aerobic bacteria and cultures of Salmonella typhimuriumand E. coli introduced into the sewage sludge are predominantly freely suspend-ed (43–68%). In the presence of protozoa, the proportion of freely suspended


bacteria drops significantly to 1–15%. The majority can now be found in oradhering to flocs (85–99%); compare also Fig. 10.

Experiments in model wastewater treatment plants [105] show that differentciliate species induce flocculation to different degrees. With the exception of thecrawling Aspidisca costata, which, even at low population densities, inducesgood flocculation when added to protozoan free (pasteurized: 50°C, 5 min) se-wage sludge, the tendency of bacteria to aggregate in the laboratory fermentervaries considerably for free-swimming (Colpidium campylum), crawling(Chilodonella uncinata, Stylonichiaputrina), and sessile (Vorticella microstoma)forms, essentially independent of their population density.

Ciliates feed selectively, not only – as shown for Tetrahymena – with regardto the physico-chemical surface structure of their prey [154], but also regardingthe size of the phagocytosed particles: This was shown by Fenchel [86] with fil-ter-feeding ciliates, characteristic for the ciliate fauna in wastewater treatmentplants. Each ciliate species can only filter specific size ranges of food particles,i.e., different ciliate species feed in their respective – sometimes distinct – ni-ches (Fig. 12). Dependent on the selection mechanism, different effects on thecomposition of the bacterial populations and the development of more or lessaggregated growth forms become apparent.

234 W. Pauli et al.

Fig. 12. Clearing rate (volume of water the organisms can clear of particles per unit time at low particle concentrations, here in multiples of the ciliates own volume per h) for threeciliate species as function of particle size, from [86]

2.3.5Reduction of the Total Biomass

In order not to exceed a sludge concentration favorable for the purification per-formance of the wastewater treatment plant, an amount equal to the daily pro-duction must continuously be drawn off. This excess sludge is subsequentlyconcentrated, digested, and drained and must finally be disposed of as a poten-tially pathogenic and frequently toxic waste product. Excess sludge is thereforean economic factor, even within the wastewater treatment plant itself. A reduc-tion in sludge production corresponds to savings in personnel, energy, and run-ning costs.

Since the function of the sedimentation tank is merely to separate the bio-mass from the purified water, the effluent concentration must already be attain-ed in the well-mixed activated sludge tank. The organisms therefore live in anenvironment with low nutrient concentrations, resulting in slow growth [46,67]. The average age of activated sludge (sludge residence time) for organicallyburdened municipal sewage, where the main emphasis is on the elimination ofthe carbon compounds, is 4 days [41, 46]. If nitrification is an objective, thesludge residence time increases to 8–10 days [157]. This means that the sludgebiomass doubles after 4 days, at the earliest. Generation times in this range im-ply not only stationary growth for the majority of heterotrophic bacteria butalso sub-optimal, reduced growth rates for the ciliate fauna having generationtimes of 5–15 h; see Table 7.

In principal, a lengthening of the food chain results in a reduction of the orig-inally available energy. In every heterotrophic link, part of the assimilated foodis converted into biomass. The remaining carbon compounds are used asenergy source for metabolic processes. When the chain becomes longer, lessenergy will remain locked into biomass. This means more carbon-mineraliza-tion and less biomass production.


Table 7. Doubling times of activated sludge and ciliates isolated from activated sludge plants

Doubling time (h) Temperature ( °C)

Activated sludge a 3.3–10 20Aspidisca costata b 13.6 20Aspidisca lynceus b 12.4 20Vorticella microstoma b 5.0 20Vorticella convallaria b 7.6 20Carchesium polypinum b 9.3 20Opercularia spec b 5.0 20Epistylis plicatilis b 10.2 20Colpidium campylum b 4.7 20Tetrahymena pyriformis b 4.5 20Paramecium caudatum b 12.0 20

a [158].b [111].

Protozoa assimilate about 85% of readily exploitable nutrients after uptake.They are converted into individual biomass or respired for energy purposes. Theremaining 15% are eliminated as compact digestion bundles (exocytosis) or dis-solved substances (excretion) [159]. Under optimal growth conditions, ca. 50% ofthe nutrients taken up by protozoa are converted into individual biomass, whichcorresponds to the metabolic efficiency of prokaryotes [160]. Different circum-stances are encountered under inhibited or stationary growth conditions. Herethe emphasis is on basal metabolism, not growth: The metabolic performance isreduced and energy consumption, as mineralized carbon in the form of CO2, in-creases [128, 161]. This diminished ability to utilize available nutrients for bio-mass production as a result of reduced growth rates is demonstrated by Ratsak etal. [128] with Tetrahymena pyriformis. At a high growth rate (generation time of5.5 h near the optimum of 3.4 h), 51% of phagocytosed bacterial biomass(Pseudomonas fluorescens) are converted into ciliate biomass, whereas at a lowgrowth rate with a generation time of 17 h only 39% of the prey is converted intopredator biomass. At the same time the ratio of respired mineralized carbon tothat converted into cell biomass increases from 0.65 to 1.2.

In municipal activated sludge plants ciliates are present in densities of104 ml–1 and over [61, 75, 94–97]. The number of bacteria required to maintainthis ciliate population can be estimated based on data from Macek [162]. Understeady-state conditions (20°C) and generation times of 5 days, free-swimmingciliates such as Colpidium campylum and sessile forms such as Vorticella micro-stoma at densities of 1.3 ¥ 104 ml–1 and 0.59 ¥ 104 ml–1 consume, over the 5-dayperiod, 2.5 ¥ 109 and 2.1 · 109 bacteria ml–1 (450 and 420 mg COD l–1), respec-tively. The bacterial content of sewage arriving at the plant is on average106 ml–1. With flow-through times of 2 h or more in municipal activated sludgeplants [41], no more than 0.5 ¥ 106 bacteria are available per ml and hour forthe ciliates. Based on the findings of Macek [162], however, a typical ciliate den-sity of 104 ml–1 would require more than 17 ¥ 106 bacteria ml–1 and hour (2–3¥ 105 ml–1 in 5 days). Therefore, the suspended bacterial content in the influentsewage cannot essentially contribute to the production of protozoan biomass.To supply adequately the protozoan population a 30-times higher bacterial con-tent in the influent would be required.

It is known that bacterivorous species are capable of effective filtration andingestion of abiotic particles with diameters of 0.3–5 mm [85–87; see alsoSect. 2.3.1] and exploiting them, if possible, for cell reproduction or to increaseindividual biomass. Thus in bench-scale plants, the addition of emulsified li-pids, which form suspended particulate fat droplets, leads to a rapid increase insessile ciliates, which can accumulate these lipids in their cytoplasm [123].Similarly, Tetrahymena is able to convert particulate suspended skimmed-milkfor reproduction, thereby attaining high population densities [163, 164]. It is un-clear however to what extent particulate abiotic organic materials (e.g., proteinrich colloids from feces) in municipal sewage are suitable, in terms of chemicalcomposition, size, and content, to be utilized in the biomass production of ty-pical sewage plant protozoan fauna.

The composition of the ciliate community in wastewater treatment plants isprimarily made up of bacterivorous filter-feeding organisms which efficiently

236 W. Pauli et al.

concentrate and ingest particulate matter the size of bacteria from the sur-rounding liquid (see Sects. 2.2.3 and 2.3.1). Bacteria occur both in activatedsludge and fixed-bed processes as complex, aggregated cell formations (flocsand slime growth). Firmly embedded in these structures, they are protectedagainst their protozoan predators. However, there is a dynamic equilibrium be-tween flocculation and de-flocculation (see Sects. 2.1.3 and 2.3.4) which, in thepresence of protozoa, shifts towards more complex micro-colonies and, in theirabsence, leads to high concentrations of single suspended bacteria (seeSects. 2.3.2 and 2.3.4). That ciliates indeed can exploit the micro-flora of thesludge itself as their primary source of nutrition is confirmed by experimentswith sterile synthetic wastewaters, e.g., [103, 123]. Activated sludge with an al-most exclusively bacterial biomass was supplied with sterile synthetic waste-water as nutrient source (see Sect. 2.3.3) and nonetheless, a typical protozoanbiocoenosis is developing.

The average sludge concentration at municipal plants is quoted as 2–3 g (dw)l–1 [46], which corresponds to ca. 6 ¥ 109 bacteria ml–1 [48]. In conventionalplants this bacterial mass is reproduced in 4 or more days (sludge residencetime). Referring to data from Macek [162], typical ciliate populations in activat-ed sludge consume 1.5–2.9 ¥ 109 bacteria ml–1. In other words, even at shorterretention times in a plant aimed primarily at the elimination of carbon com-pounds, a considerable proportion (25–48%) of the bacteria can be phago-cytosed by ciliates: This corresponds to a 10–19% reduction of the accumula-ted sludge, based on a mineralization of around 40% of the bacterial food [128].Observations with submerged fixed-bed filters in activated sludge plants reveala similar picture with regard to the reduction of the accumulated sludge by pro-tozoa. In the activated sludge tank (volume 756 m3) a contact aerator, whoseslime-growth makes up almost 18% of the biomass (dw) of the tank, leads to areduction of the BOD sludge accumulation of about 25% [117]. Such sub-merged fixed-beds are primarily colonized by protozoa whereby ciliates domi-nate [116, 117, 165, 166], comprising around 68% of the total biomass [116].Based on these data, an additional biomass of 12% consisting exclusively ofciliates (18% additional biomass, 68% of it ciliates) effects a sludge reduction of 25%. A transfer of these results to conventional activated sludge plants would mean that the autochthonous ciliate fauna, as the second link in the foodchain and representing 9% (dw) of the total biomass [64, 65], is in a position to reduce sludge accumulation by 19%.

2.3.6Influence of Protozoa on Bacterial Metabolism

A series of studies on degradation efficiency in bench-scale wastewater treatmentplants show that in the presence of protozoa – in spite of their antagonistic effectsas bacteria predators – the physiological performance of the bacteria is maintain-ed or even increased: In bench-scale plants, ciliates show no effects on the nitrifi-cation bound to flocs [103, 126, 127]. The degradation of nitrilotriacetic acid bybacteria is equally unaffected by the presence of ciliates; however, here a shiftfrom single suspended to complex aggregate growth forms is observed [126, 127].


Clear indications of an increase in bacterial metabolic activity were found byCurds et al. [103]: Under experimental conditions they observed, in the pre-sence of protozoa, an increased degradation (BOD, COD) of the dissolved, non-filterable portion of organic materials, attributed almost exclusively to the ac-tivated sludge flora (see Sect. 2.3.3 and Table 6). Findings by Wiggins andAlexander [167] also imply a positive influence of protozoa on bacterial degra-dation processes with regard to the organic pollutants 2,4-dichlorophenol (2,4-DCP) and 2,4-dichlorophenoxyacetic acid (2,4-D). Although protozoan feedingreduced the mixed culture of freely suspended bacteria by more than one orderof magnitude – leading to delayed degradation compared to protozoa-free cul-tures – after 15 days the environmental chemicals 2,4-DCP and 2,4-D were min-eralized in the presence of protozoa to 70% and 90%, respectively: Whereas incultures where the protozoa were inhibited by nystatin and cycloheximide, de-gradation of only 40% (2,4-DCP) and 10% (2,4-D) were observed over the sameperiod (Fig. 13).

In biocoenoses other than wastewater, i.e., in microcosms with pure andmixed cultures of typical aquatic and terrestrial bacteria, an increased bacterialmetabolism in the presence of protozoa is observed. Various explanatory at-tempts emphasize the direct influence of the protozoan metabolic activity;others attach more importance to bacteria feeding and its indirect consequen-ces on the size and composition of bacteria populations and some correlate themicro-currents, generated by the ciliates, with an improved food and oxygensupply of the bacterial flocs or multi-layer biofilms. Protozoa are capable of me-tabolizing bacterial metabolic products such as acetic-acid, butyric acid, andethanol [77, 168] and could thus avert end-product inhibition [169]. On the

238 W. Pauli et al.

Fig. 13. Effects of protozoa on the mineralization of 0.1 mg l–1 2,4-dichlorophenol and 2,4-dichlorophenoxyacetic acid (2,4-D) in sewage. Cycloheximide (250 mg l–1) and nystatin(30 mg l–1) were used to suppress protozoa; from [167]

other hand, protozoa release a series of organic substances such as amino-acids[170] and “growth factors,” having chemical structures not characterized in de-tail [22, 109, 171–175], into the surrounding medium, leading to activation ofbacterial metabolism and growth. Furthermore, protozoa have the highestexcretion rate of inorganic phosphate and nitrogen, relative to biomass, withinthe zooplankton [176]. In addition, in the presence of protozoa, an acceleratedbacterial phosphorus mineralization is observed [177]. This mutually advantag-eous interaction by nitrogen and phosphorus re-mineralization is emphasizedby many authors [22, 145, 177–183]. To what extent these additional organic andinorganic substances introduced into the wastewater cycle play a part in waste-water treatment processes is controversial, but due to the composition of thewastewater, rather unlikely [75, 184]. On the one hand, municipal sewage itselfis a complex nutrient solution with a heavy organic load; on the other hand,nitrogen and phosphorus are present in excess in wastewater treatment plants,in contrast to most limnic, marine, and terrestrial ecosystems (a BOD:N:P ra-tio of 100:5:1 is considered to be the optimal substrate composition of sewage– compared to this nutrient balance, municipal sewage with average BOD:N:Pratios of 100:17:5 [41] contains an excess of nitrogen and phosphorus).However, in the case of commercial and industrial wastewaters with high car-bon loading and comparatively low concentrations of nitrogen and phosphorus(e.g., vegetable processing businesses, fiberboard works, paper and cardboardfactories, coking plants, as well as chemical and pharmaceutical industries [41,185]) catalytic effects on bacterial metabolism by interactions with N and P setfree by protozoa are quite conceivable.

It is not self-evident that bacteria feeding and their subsequent reduction ofbacterial populations should have positive effects on bacterial metabolic turn-over. A possible cause could be the qualitative shifting of the selection condi-tions for the bacteria and therefore the composition of mixed bacteria popula-tions and their organizational forms. The success of a bacteria population is notonly dependent on its adaptation to the nutrients on offer but also on whetherit is edible for protozoa [148]. As discussed in Sect. 2.3.4, the selection of feed-ing-resistant bacterial growth forms can be viewed mainly as a result of pha-gocytic activity of protozoa: Freely suspended bacteria are succeeded by aggre-gated, sessile growth forms [136, 148, 186]. That this shift can be accompaniedby a simultaneous intensification of the microbial metabolic processes is shownby studies on marine bacteria, which as adherent cells in biofilms (“marinesnow”) display faster growth (incorporation of thymidin into DNA [187]), anincrease in electron transport (reduction of tetrazolium salts to formazan[188]), and higher hydrolytic activity [189], than as freely suspended singlecells.

2.3.7Filamentous Bacteria and Protozoa

Filamentous bacteria are present in the bacterial flora of almost all activatedsludge. Due to their large surface area, they are well-equipped for the adsorp-tion and metabolism of organic compounds. At low densities, they contribute to


the stabilization of activated sludge flocs. However processing problems arise ifmass reproduction of filamentous bacteria occurs in the activated sludge tank.The enlarged surface area of the flocs hinders the settling and thickening pro-cesses in the sedimentation tank which can, in extreme cases, due to the forma-tion of light, fluffy, poorly settling flocs, result in the discharge of sludge into na-tural waters. This phenomenon, known as “bulking sludge,” used to be causedby “high load bacteria” such as Sphaerotilus sp. or filamentous types 1863 and0961. Today, however, many of the filamentous bacteria found in sewage treat-ment are adjusted to low carbon concentrations [low F/M ( food:micro-orga-nism) bulking], e.g., types 0041, 0675, 0092, 1851, or Microthrix parvicella.Experience shows that putrid wastewater, rich in H2S or with high carbohydrateor short-chain organic acid content (i.e., wastewater from food processing, pa-per and textile industries), as well as low nitrogen, phosphorus, or oxygen con-

240 W. Pauli et al.

Fig. 14. Degeneration of bulking sludge (decrease of sludge volume index: SVI) in the aera-tion tank of an activated sludge plant and in laboratory experiment by the filamentous pre-dacious protozoan Trochilioides recta; from [194]

tent, stimulates the development of bulking sludge. Various chemical (e.g., lim-ing, chlorination, addition of H2O2, iron salts, and nitrogen and phosphoruscompounds) and physical (e.g., increased oxygen supply) methods are imple-mented to combat bulking. Sometimes even operational conditions of plantsare altered (e.g., increasing the return-flow rate, by-passing the pre-clarifica-tion, aerobic, and anaerobic selectors) [67, 185].

In principal, autochthonous ciliates appear suited to counteract abundantdevelopment of filamentous bacteria. However, only a few species capable oftaking up filamentous bacteria are present in activated sludge plants, e.g.,Trithigmostoma cucullus (Chilodonella cucullus), Trochilioides recta, Trochiliaminuta, and Chilodonella uncinata. If these ciliates attain a high populationdensity, a pronounced decline in filamentous bacteria and degeneration ofbulking sludge is observed within a few days, both in bench-scale and operatio-nal plants [190–193]; see Fig. 14. Effective cell densities for Trochilioides sp. arequoted as 1000 ml–1 [190] and for Trithigmostoma cucullus and Trochilioidesrecta as 2000 ml–1 [193].

3Impairments of Protozoa: Consequences for Water Purification

Ciliated protozoa are very numerous in all types of aerated biological treatmentsystems (compare Sects. 2.2.3 and 2.2.4). They play an important role in thepurification process removing, through predation, the major part of dispersedbacteria that cause highly turbid, i.e., low quality effluent. It has been generallyrecognized that changes in the population density and community structure ofciliates affect the food web of this artificial ecosystem, thus influencing the per-formance of plants. Excess influx of toxic wastes with detrimental effects on ci-liates would prevent clarification, thereby severely threatening the degradationprocess. A variety of chemicals can limit growth of ciliates. As with organismsfrom other taxonomic, functional, and trophic levels, the toxicological effectsinduced by organic and inorganic chemicals on ciliates vary widely, i.e., EC50-values ranking from some mg l–1 to some g l–1 (reviewed by [194, 195]). Sub-stances having toxic effects which diminish or even paralyze the purificationperformance frequently find their way into wastewater treatment plants withcommercial and industrial wastewater. Risks are particularly great from metal-finishing works with electrochemical processes and wastewater from iron andsteel pickling plants, accumulator-charging stations, stereotype, photocopy,photographic and printing works, dry-cleaning premises, industries producingpesticides, herbicides, and disinfectants, as well as tanneries, leather goodsmanufacturers and coking plants. In order to estimate the hazard potential andto lay down maximal concentrations, in addition to bacterial tests, biologicaltests with ciliates are indispensable to reflect potential risks of hazardous sub-stances on the biological system of wastewater treatment as a whole.

Tests with typical wastewater protozoa have been carried out for a number oftoxic substances. Gracia et al. [196] observed effects of copper (sulfate) in con-centrations of 1 mg Cu2+ l–1 on species diversity and population density – espe-cially of the ciliates – of natural sludge samples. Madoni et al. [197] determined


the 50% lethal effect concentrations of Cu<Hg<Cd<Pb<Cr<Zn (1 mg l–1 –50 mg l–1) on various ciliates isolated from activated sludge, whereby the au-thors report differences in species sensitivities of up to two orders of magni-tude, dependent on the heavy metal tested. Kakiichi et al. [107, 198–200] reportinhibitory effects of disinfectants and surfactants on typical activated sludgeciliates. A comparison of the effect potential of 4 disinfectants towards thewastewater bacteria Alcaligenes faecalis and the wastewater ciliate Colpoda as-pera reveals an almost 10-fold higher sensitivity of the ciliates [200]. Highersensitivities of ciliates in comparison to bacteria were also found by Yoshioka etal. [201] for 32 wastewater relevant environmental chemicals. Results from theOECD activated sludge respiration test (RI Test, [202]) – considered as an indi-cator for acute effects of chemicals on heterotrophic bacterial flora – andgrowth tests with Tetrahymena, a ciliate typical in polysaprobic surface waters,but also found in activated sludge and submerged contactor plants [53, 55, 111,112, 115, 203–209] were compared: 50% effect concentrations were, on average,10 times lower with the ciliate test. Furthermore, certain substances provedhighly toxic in the Tetrahymena test, and showed only weak effects in the respi-ration test; out of a total of 32 substances, just 6 cases had a (toxic) effect po-tential of less than 100 mg l–1. The weak correlation of r2= 0.17 confirms the dis-crepancy between the two tests (Fig. 15). Similar observations of a low correla-tion were made by Pauli and Berger [210]. Figure 16 illustrates toxic responsesof 4 ciliate species and standard tests with activated sludge towards industrialchemicals (data taken from the International Uniform ChemicaL InformationDatabase, IUCLID, including toxic data of a wide variety of industrial chemi-

242 W. Pauli et al.

Fig. 15. Acute effects of chemicals on the bacterial flora of activated sludge (OECDRespiration Inhibition Test) in comparison to those on the ciliate Tetrahymena pyriformis(growth inhibition) and on fish (OECD lethality test with Oryzias latipes); after [202]

cals). Although a generally higher sensitivity of ciliates cannot be observed forthis data set, the random distribution of points around the bisector confirmsthe dissimilarity of ciliate and activated sludge toxicities (r2< 0.01, n = 35).

Evidently ciliates are not only sensitive to pollutant induced stress, but testresults reflect a series of additional toxic interactions, not represented by testswith bacteria in activated sludge. That this different toxic profile is probably dueto the more complex cell-physiological – eukaryotic – organizational structureof the ciliates is implied by QSAR studies for heterogeneous chemical classes[211], which revealed a high correlation between the LC50 values found in thewidely accepted fish lethality test (r2= 0.78) with Tetrahymena growth, but notwith bacteria test.

4Environmental Biotechnological Aspects

4.1Biodegradation Potentials of Ciliates

Although it is well known that ciliate grazing on bacteria fulfills important tasksin the biological purification of sewage (compare Sects. 2.3.2.2 and 2.3.2.3) andthat a number of technical methods and plant operation parameters obviouslyimprove the purification efficiency by favoring ciliate growth (see Sects. 2.2.5,2.3.2.2, and 2.3.2.3); only recently some pioneering attempts have been made tospecifically use ciliates in biodegradation processes.

Generally, large amounts of biosludge are formed in biological wastewatertreatment processes and the separation, dewatering, treatment and disposal ofthis sludge represents major investment and operating costs. One of the poten-


Fig. 16. Comparison of results from standard activated sludge respiration tests and bioassayswith ciliates (data from IUCLID); from [210]

tially useful assemblies for reducing the sludge yield is the two-stage cascadeused in many experiments for the study of ciliate-bacterial interactions, e.g.,[140, 212–215]. The technique of a two-stage system enables one to manipulatethe artificial ecosystem of conventional treatment processes so that dispersedbacteria are growing in the first part of the process and being consumed by pro-tozoa in the last. Whereas in conventional treatment due to the growth of flocor film forming bacteria most of the bacterial biomass is protected against pre-dation (see Sect. 2.3.4), dispersed bacteria can be readily taken up and meta-bolized by protozoa (see Sect. 2.3.2), resulting in a lower sludge yield (seeSect. 2.3.5). Operating the first part of the treatment process as an aerated tankreactor without biomass retention and at an hydraulic retention time shortenough to prevent a significant growth of protozoa is a simple way to stimulatethis growth of dispersed bacteria. Cultivations using synthetic wastewater anddefined cultures of bacteria and ciliates in a two-stage chemostat cascade haveshown that protozoan grazing can result in a considerable biomass reduction[128]. By introducing a “predation trap” (second stage) it was possible to obtaina decrease of 12–43% in biomass yield in comparison with a system without ci-liate grazing. Studies of Lee and Welander [216, 217] confirm this potential of atwo-stage system to reduce the sludge yield. Employing synthetic wastewaterand mixed cultures of bacteria, protozoa and metazoa from activated sludgethey observed a sludge yield around 30–50% of the yields typically obtained inconventional aerobic processes [216]. If authentic instead of synthetic waste-water was used as bacterial food supply the sludge production was also con-siderably lower than in conventional treatment [217].

Cox and Deshusses [218] developed a strategy to control biomass growth inbiotrickling filters for waste air treatment by engineering predation of bacteriaby protozoa. It was shown that clogging of bench-scale biotrickling filters couldbe slowed down with the use of protozoa. Interestingly, it was found that the re-actor with protozoa had a shorter start-up time, possibly because of bacterialgrowth factors secreted by the protozoa.

For the biodegradation of whey, the ciliate Tetrahymena had been chosen byBonnet at al. [219] as a micro-organism capable of degrading and modifyingthe whey biologically in order to diminish its pollutant effect (whey is theaqueous phase that separates from the curds during cheese making or caseinproduction). Disposal of crude whey completely arrested operation of lagoonpilots serving as example of receptor media, whereas the effects of biodegradedwhey were only temporary, and normal operation was recovered within a fewdays. The authors stress that this method could be a valuable tool for smalldairy farms, being unable to use complex industrial treatment technologies toforestall pollution by waste whey.

Clearly, optimal conditions for protozoan activity need to be further evaluat-ed and pilot scale experiments have to be performed to prove the influence ofbiomass predators in real treatment systems. Nonetheless these findings areauspicious, suggesting that specific use of ciliates can be made to improve bio-degradation processes.

244 W. Pauli et al.

4.2Ciliates as Biosensors

As a constitutive group within the microbial food web, ciliates not only play animportant ecological role in the self-purification and matter cycling of naturalaquatic ecosystems, but also in the artificial system of sewage treatment plants.Their feeding on bacteria improve the treatment, resulting in higher trans-parency, i.e. lower organic loads in the output water of the treated wastes (seeSects. 1 and 2). This status of ciliates as an important functional group, improv-ing the process in municipal sewage treatment, and furthermore that‚ “valuesfrom ciliate growth inhibition tests are relevant for the risk assessment forsewage treatment plants” has been recently acknowledged by a TechnicalRecommendation of the EEC [220].

There is a broad consensus in ecotoxicology that taxonomic similarity (i.e.,close relationship, in terms of phylogeny) generally implies similar toxicologi-cal responses, e.g., [221, 222]. This is reflected in aquatic toxicology by selectingcertain fish, crustacean, and algae species to represent trophic and taxonomiclevels as a whole. A transferability of toxicological data for ciliates is also indi-cated. Although there exists an extraordinary amount of evolutionary distancebetween different genera and even between species of the same genus [223,224], comparisons between the ciliates Colpidium, Colpoda, Paramecium,Tetrahymena, Uronema, and Vorticella reveal an almost comparable toxicologi-cal susceptibility [210]. Despite the lack of standardized ciliate test protocols,only 2 substances out of 13 exert a toxic effect differing by a factor of more than100, whereas for the rest of the chemicals the deviations lie within about one or-der of magnitude (Fig. 17).

The early use of ciliates in toxicity testing was reviewed by Persoone and Dive[225]. Among the ciliates, the organism of choice in aquatic toxicity testing hasbecome the common freshwater hymenostome Tetrahymena [195, 226, 227].Many features have contributed to making Tetrahymena – particularly the speciesT. pyriformis and T. thermophila – favorite models in cell biology and facilitatedtheir modern day use as aquatic toxicity test organisms. It is worth mentioning,not only that these unicellular organisms can be grown under axenic, i.e., bac-teria-free conditions, but also that they combine important advantages from twogroups of organisms. Indeed, they belong to the higher cells, the eukaryotes, butthey can be cultured both easily and economically like the prokaryotic bacteria.

An innovative tool with the potential of a wide application has recently beenoffered by the introduction of a commercialized microtoxicity test kit withTetrahymena (Protoxkit F, Creasel Ltd., Belgium). The test is specially designedfor the use of environmental samples, thereby providing a helpful means to as-sess risks of sewage contaminants and their possible detrimental effects on theperformance of waste water treatment plants. Following the concept of ready-to-use microbiotests, with the test kit a ciliate multi-generation (growth) assaycan be conducted by non-experts without sophisticated sample preparation andexpensive equipment.

Growth impairment tests with Tetrahymena have generally reached the high-est degree of acceptance and standardization [195, 227, 228]: Based on an inter-


national pilot ring test, a growth test with the ciliate Tetrahymena is recom-mended by the German Federal Environmental Agency for ecotoxicological riskassessment [229].A final ring test to establish an internationally recognized TestGuideline has been initiated – an important step to include a traditionally un-tested, but ecologically important group of organisms in comprehensive eco-toxicity test batteries.

5References

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Predictability of Biodegradation on the Environment:Limits of Prediction from Experimental Data

Johanna B. Wesnigk 1, Maike Keskin 2, Wilfrid Jonas 3, Klaus Figge 3,Gerhard Rheinheimer 4

1 BLG Consult Bremen GmbH, Hafenstr. 55, 28217 Bremen, Germany,E-mail: [email protected]

2 proDERM, Institut für Angewandte Dermatologische Forschung, Industriestr. 1,22869 Schenefeld, E-mail: [email protected]

3 NATEC, Institut für naturwissenschaftlich-technische Dienste GmbH, Behringstr. 154,22763 Hamburg, Germany

4 Institut für Meereskunde an der Christian-Albrechts-Universität Kiel, DüsternbrookerWeg 20, 24105 Kiel, Germany

This chapter deals with the description and interpretation of different biodegradation testsdealing with the degradation of chemicals by mixed microbial communities derived from dif-ferent natural and semi-natural habitats. Prescribed standardized laboratory tests and theirlimitations are listed and compared to more complicated tests systems such as differentsimulation tests, micro- and mesocosm tests, and field studies. Where possible examples aregiven to illustrate possible outcomes of some tests, using the substances 4-nitrophenol, te-trachlorobenzene, and NTA. Tests with and without soil or sediments are described. The im-portance of environmental factors like the concentration of the chemical, size of the degrad-ing population, grazing, temperature, sorption, and others are illustrated. A focus lies on theimportance of adaptation phenomena. Interactions of the factors and their impacts on thepredictability of biodegradation are discussed and recommendations for further research aswell as management advice are given.

Keywords. Biodegradation, Predictability, Adaptation, Test systems, Environmental factors

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254

2 Typical Environmentally Relevant Substances . . . . . . . . . . . 256

2.1 4-Nitrophenol (4-Np) . . . . . . . . . . . . . . . . . . . . . . . . . 2562.2 Chlorobenzenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2562.3 Nitrilotriacetate or Acetic Acid (NTA) . . . . . . . . . . . . . . . . 256

3 Analytical Methods and Their Implications . . . . . . . . . . . . . 257

3.1 Analytical Methods Used in Screening Tests . . . . . . . . . . . . . 2573.1.1 Dissolved Organic Carbon (DOC Die-Away Test and Modified

OECD Screening Test, MOST) . . . . . . . . . . . . . . . . . . . . . 2573.1.2 Biochemical Oxygen Demand (BOD Closed Bottle Test) . . . . . . 2593.1.3 Carbon Dioxide (CO2 Evolution Test, Previously Called

Modified Sturm Test) . . . . . . . . . . . . . . . . . . . . . . . . . . 2593.2 Common Features for All Screening Tests

for Biodegradability . . . . . . . . . . . . . . . . . . . . . . . . . . 2603.3 Radioanalytical Methods . . . . . . . . . . . . . . . . . . . . . . . 2603.4 Analytical Separation and Quantitation Methods . . . . . . . . . . 2613.5 Methods for Identification of Compounds . . . . . . . . . . . . . . 262

CHAPTER 4


4 Which Test Systems are Used for Which Purpose? . . . . . . . . . 262

4.1 Screening Tests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2634.2 Simulation Tests . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2644.2.1 Degradation Studies in Water/Sediment Systems . . . . . . . . . . 2664.2.1.1 Limitations of Simulation Studies Using Soil or Sediments . . . . 2684.2.2 Possibilities to Predict from Simulation Test Results . . . . . . . . 2684.3 Mesocosms and Field Studies . . . . . . . . . . . . . . . . . . . . . 2694.3.1 Possibilities to Predict from Mesocosm and Field Test Results

and Comparison of Test Systems . . . . . . . . . . . . . . . . . . . 271

5 Potentials and Limitations for Prediction from Laboratory Results . . . . . . . . . . . . . . . . . . . . . . . . 272

5.1 Concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2725.2 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2755.3 Sorption . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2765.4 Oxygen Content . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2775.5 Sediments and Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . 2775.6 Grazing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2795.7 Interactions Between Concentration, Growth, Grazing,

and Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . 280

6 What is Persistence, and When is a Substance Biodegradable? . . 283

6.1 A Practitioner’s View . . . . . . . . . . . . . . . . . . . . . . . . . . 2836.2 An Ecologist’s View . . . . . . . . . . . . . . . . . . . . . . . . . . . 284

7 Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285

7.1 Some Research Suggestions for Better Predictability . . . . . . . . 2867.2 Some Steps Towards Sustainable Development . . . . . . . . . . . 287

8 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287

1Introduction

Of major importance for environmental management are questions relating tothe timely discovery and estimation of the potential dangers stemming fromchemical substances. To do this it is necessary to study two fundamental aspectsof pollutants in detail: their fate and effect in the ecosystem. The fate can be de-scribed by the paths of transport, the distribution, and the degradation pat-terns. The effect refers to unwanted, deleterious, toxic or other chronic or acuteeffects and their mechanisms.

The prediction of expected paths and distribution patterns of the chemicalspresent in the environment, involuntarily or intentionally, is important formanagement and therefore often attempted. A synopsis of work related to thistopic to date shows that this question has been looked at almost exclusively

254 J.B. Wesnigk et al.

from a theoretical point of view. The focus of research activities so far has beenon developing models and related calculations. They predict the behavior ofchemicals in the environment according to the underlying theory of the model.

These models are based on assumptions, several of which are specific foreach theoretical model on the nature of the different processes under naturalconditions. As examples may serve:

1. The assumed relevance of a defined balance (steady state) for the distribu-tion due to the chemical and physical properties of the substance in question.

2. The importance of several chosen transfer constants for the passage throughboundaries between the compartments.

3. The kinetic model used for the degradation of the chemicals in the separatecompartments.

In addition, the properties of the environment itself necessitate assumptionsspecific for each model, i.e., whether it can be represented by homogenous com-partments with related, amenable volumina. This chapter will try to give an an-swer to the important question as to whether degradation patterns calculatedfrom models or laboratory test results are conforming to the results of measur-ing the reaction more directly in a natural environment. Furthermore, com-monly used test procedures will be described and evaluated in this regard.

In addition to the degradation characteristics, it is necessary to evaluate thebehavior of chemicals in the environment to gain reliable data on their ecotoxi-cological effects. This evaluation is complicated by possible synergisms and an-tagonisms. A pollutant can have different effects when entering an organism indifferent ways. Certain pollutants can be present in varying physical and chem-ical states, which might change their toxicity, while others may be very specificor quite broad ranging in their effects. A recent example for a new pollutant ef-fect are the hormone mimicking substances or endocrine disruptors. They arechemically unrelated but have a structural element in common which in severalcases locks onto estrogen receptors, thus amplifying and confusing hormonalsignals in very low doses.

It has been surprisingly hard to identify and causally relate unwanted effectsin nature to a single substance. However, today’s development of modern eco-toxicological methods makes it possible to determine changes in the environ-ment and relate them to pollutants. In order to analyze effects it is not enoughto look only at segments of the ecosystem in question. To get the full picture itis necessary to study all ecosystem levels including all compartments, i.e., wa-ter, soil, air, and biota.

This leads to the following conclusions: practical experiments regardingtransport, distribution, degradation, and effect of chemical substances in ecosys-tems will continue to play an important role in preventative environmental re-search. These kinds of studies are difficult in open, natural systems. Thus stan-dardized laboratory tests or small scale ecosystem models – later named micro-or mesocosms – have to be used. If one of these systems is used, the followingquestions regarding the transfer of laboratory results to the field will arise:

– Can the results obtained in simplified microcosms reflect the reality in com-plex natural ecosystems?

Predictability of Biodegradation on the Environment: Limits of Prediction from Experimental Data 255

– Is it acceptable to use laboratory data for the calculations in theoretical eco-system models?

– To what extent can a cost-effective simplified microcosm be used as a prac-tical model to obtain results relevant for the environment?

This chapter will mainly focus on the question of how far experimental labora-tory data can predict the degradation of chemicals in nature.

2Typical Environmentally Relevant Substances

The following substances were chosen as examples. Their degradation pathwaysare presented in chapters 1 of this volume.

2.14-Nitrophenol (4-Np)

This substance was studied extensively by several authors. It shows a somewhaterratic behavior in standard tests, being sometimes mineralized and sometimesnot [1, 2], and is therefore classified as a typical gray list substance. In theUnited States 2- and 4-nitrophenol are classified as priority pollutants [3]. 4-Npis supposedly a major degradation product of the widely used insecticide para-thion and can be generated in the atmosphere from car exhausts like benzeneand NOx [4]. The degradation pathway has been identified [5] but there havebeen indications that 4-Np can be metabolized to the more persistent 4-amino-phenol instead of being mineralized [6].

2.2Chlorobenzenes

Chlorobenzenes with different chlorination grades can be found worldwide inall compartments of the environment [7, 8] and even in human blood and fattissue [9] since they are used as solvents or heat conductors and as basics for theproduction of insecticides, other biocides, and colors [10]. They have been clas-sified as priority pollutants by the US Environment Protection Agency [3].Their high persistence is mainly due to chloro-substituents which lower theelectronic density of the aromatic ring usually cleaved by electrophilic oxy-genases [11]. Therefore, highly chlorinated aromatics cannot be degraded bymost living organisms. Only for the last fifteen years several bacterial culturescapable of using chlorobenzenes as sole source of carbon and energy have beenisolated [12, 13]. This implies the evolution of new strains which have adaptedthemselves to the environmental contamination.

2.3Nitrilotriacetate or Acetic Acid (NTA)

This detergent builder and chelating agent has been intensively studied beforeand during its large scale introduction into surface waters as a phosphate sub-


stitute in Canada [14–16]. It can be degraded after an adaptation phase and itcan interact with metals [17–19]. There is still some doubt about the concen-trations and temperatures necessary for adaptation and about its degradabilityin seawater [20–22].

3Analytical Methods and Their Implications

Chemical analytical work in biodegradation studies has the following aims:

– The qualitative and quantitative determination of the test substance and ofits known metabolites at different time points and the spatial distribution inthe test system, including a balance of substance distribution and metabo-lization.

– The identification of unknown metabolites occurring during the study in or-der to develop a degradation pathway.

– The determination of the proportion of total mineralization.

These objectives can best be achieved by using radiolabeled compounds inconnection with suitable radiochemical, chemical, and physical analytical me-thods. However, before carrying out sophisticated studies using radiolabeledcompounds, it is reasonable to perform screening tests in water in order to lineout the readily biodegradable compounds and to focus resources for expensivestudies on problematic compounds and in more heterogenous media such assoils. This stepwise procedure (i.e., starting the assessment of biochemical de-gradability with a screening test) is generally required for registration of a chem-ical in Europe.

3.1Analytical Methods Used in Screening Tests

The most commonly used screening tests for testing biodegradability are listedin Table 1 [23]. One common feature in these tests is the analysis of sum para-meters as measures for the biodegradability instead of analyzing the test sub-stance directly. The most obvious advantage of analyzing sum parameters is thelow cost. Therefore these tests are the most effective way of testing a large num-ber of chemicals. In many cases, these test give satisfying answers to the ques-tion if a chemical is readily degraded. The limits of the screening tests in gen-eral will be given below. In this section, the principles and the limits of the ana-lytical methods are described.

3.1.1Dissolved Organic Carbon (DOC Die-Away Test and Modified OECD Screening Test, MOST)

The principle of this method is the oxidation of the organic matter in a sampleby combustion or by using a strong oxidant and measurement of the infrared(IR) absorption of CO2 by an IR-detector. If the sample is used directly, the to-



Table 1. Test methods for screening tests for ready and inherent biodegradability [23]

Test name Parameter Concen- Inoculum [cfua/ml] Suitable Suitablefor analysis tration for poorly for volatile

soluble solublesubstances substances

Ready:DOC Die- DOC 10–40 mg 104–105; active sludge, – –Away DOC/l sewage effluent, surface

water, soil or mixtureCO2 evolu- CO2 10–20 mg 104–105; see above + –tion DOC/lModified DOC 100 mg/l 104–105; mixture of 10 dif- – +/–MITI b (I) ferent sites, incl. Industrial

sewage effluent, acclimation in lab. 1–3 months

Closed bottle BOD 2–10 mg/l 10–103; secondary sewage +/– +effluent (domestic), and/or surface water

Modified DOC 10–40 mg 102; secondary sewage – –OECD DOC/l effluent (domestic)screeningManometric BOD 100 mg/l 104–105; like DOC + +/–respirometry Die-away testBODIS c BOD 100 104–105; supernatant of + +

ThOD/l settled activated sludge (domestic)

Special test:Biodegrada- DOC or 5–40 mg No cfu-conc. prescribed, +/– +/–tion in BOD DOC/l seawaterseawaterInherent:Modified DOC 20 mg/l > 105; activated sludge from – –SCAS d a suitable treatment plant,

adaptation permittedModified DOC 10–100 DOC (inoc.) /DOC Zahn- mg/DOC/l (test subst.) = 2.5/1 to 4/1; – –Wellens activated sludge; adapta-

tion permittedModified BOD 30 mg/l > 105; like MITI (I) + –MITI b (II)

a cfu cell forming units.b MITI Ministry of International Trade and Industry, Japan.c BODIS BOD-Test for Insoluble Substances–a modification of the Closed Bottle Test, issued

by the German Umweltbundesamt.d SCAS Semi-continuous activated sludge test.

tal carbon (TC) is determined. If the sample is filtered or centrifuged beforemeasurement, the total dissolved carbon is analyzed. For separation of organicand inorganic carbon (carbonates), the sample is acidified and the CO2 of inor-ganic origin is stripped before oxidation. This means air or inert gas is blownthrough the sample. After this pretreatment, the DOC can be analyzed. Limits ofthis method are:

– Volatile organic test substances or metabolites are not detected.– The limit of detection is approximately 2 mg DOC/l.

This means that the starting concentration needs to be a minimum of7 mg DOC/l in order to allow measurement of at least 70% degradation.

3.1.2Biochemical Oxygen Demand (BOD Closed Bottle Test)

The determination of biodegradation by means of BOD measurements is an in-direct method. The theoretical oxygen demand (ThOD) from the complete oxi-dation of a test substance is calculated on the basis of the formula of the testsubstance or by measurement of the chemical oxygen demand (COD). Inscreening tests using this parameter the oxygen depletion is determined duringthe tests using oxygen sensitive electrodes or manometers. The measured oxy-gen depletion is compared to the theoretical oxygen demand. Limits of thismethod are:

– It is only applicable for aerobic degradation.– The limit of detection is 4–5 mg ThOD/l.

This means that the starting concentration needs to be a minimum of 10 mgThOD/l in order to allow measurement of at least 60% degradation.

3.1.3Carbon Dioxide (CO2 Evolution Test, Previously Called Modified Sturm Test)

The CO2 evolution test is also an indirect method. The theoretical CO2 evolution(ThCO2) after complete oxidation of a test substance is calculated on the basisof the formula of the test substance. The CO2 produced during the tests is trap-ped in barium or sodium hydroxide and is measured by titration of the residualhydroxide, or as inorganic carbon. The CO2 evolution is compared to theThCO2. Limits of the test are:

– It is not applicable to volatile test substances or metabolites.– It is relatively laborious compared to other screening methods.

The limit of detection depends on the concentration and amount of barium hy-droxide in the traps (it corresponds to the limits given in the DOC Die-Awaytest).


3.2Common Features for All Screening Tests for Biodegradability

The following features are those all screening tests mentioned so far have incommon:

– The test media are mineral media containing prescribed concentrations ofpotassium and sodium phosphates plus ammonium chloride, magnesiumsulphate, and iron (III) chloride. For the modified OECD screening test, traceelements (Mn, B, Zn, Mo) and yeast extract are added.

– A reference compound is run in parallel to check the operation of the proce-dures.

– Pass levels for ready degradability are 70% removal of DOC and 60% ofThOD or ThCO2 production.

– For ready biodegradability tests, the inoculum may be pre-conditioned to theexperimental conditions but not pre-adapted to the test substance. For inhe-rent biodegradability tests, this pre-adaptation is allowed.

– Test duration is 28 days for ready biodegradability tests. It can be shortenedif the substance is degraded before the end of this period. It can be extendedif the plateau phase has not been reached. The test duration of the inherentbiodegradability tests depend on the individual adaption periods.

– The incubation temperature is 20°C or room temperature.

3.3Radioanalytical Methods

The most common isotope used for environmental studies is 14C, simply be-cause carbon is the most common atom in organic compounds and because ofthe relatively easy detection methods available for this carbon isotope. If thesynthesis of a [14C]-labeled compound is too complicated, it may be necessaryto use 3H. The use of this isotope, however, may cause problems in the interpre-tation of the data since 3H is known to be replaced intramolecularly. If unknowndegradation products are expected, additional labeling with 13C, 32P, 15N, 35S,36Cl, 19F, and others may be reasonable in order to facilitate the interpretation ofNMR- and mass spectrometer data.

The position of the label within the test substance is very important for thestudy design. Generally, a 14C-label will be positioned in the part of the mole-cule which is the least susceptible for biodegradation in order to be able to traceas much of the metabolites as possible. If a molecule carries a short ester group,for example, this group will normally be quickly separated by enzymatic ornon-enzymatic ester cleavage. If this group carries the radioactive isotope onewill find a quick degradation but will not know what happened to the majorpart of the compound. The most clear-cut results can be expected if the testsubstance is labeled with one atom in a hardly susceptible position such as thering of an aromatic system. Two labels in one molecule may increase the sensi-tivity but may complicate the interpretation of the results, especially if metabo-lites have to be identified.


The following equipment and radioanalytical methods are commonly used:

– Liquid scintillation counter (LSC) – suitable for counting b-decay events insamples mixed with scintillation cocktails. The number of decay events perminute is a measure for the amount of test substance present in the sample.This apparatus is absolutely necessary.

– Combustion apparatus: this apparatus is necessary for determination ofsamples bound to or present as insoluble solid compounds such as soil orsediment, plant or animal tissues.A defined amount of the sample is weighedinto a cellulose cone, this cone is transferred into the combustion coil (Pt-Rh-alloy) and combusted at 800°C in an oxygen flux. The combustion gas con-tains – besides other non-labeled compounds – the residues of the radiola-beled test substance as 14CO2. This radiolabeled carbon dioxide is collected inan automatic trapping system containing a special absorber scintillation cock-tail. The radioactivity in this cocktail is determined in an LSC (see above).

– Linear analyzer for radio thin layer chromatography (radio-TLC-scanner):the TLC-scanner enables the detection and quantitative determination ofcompounds separated by thin-layer chromatography.

– Digital autoradiograph (DAR): a detection system for samples separated bytwo-dimensional TLC or for sliced tissue samples containing a two-dimen-sional pattern of radioactive spots.

– Radio-high performance liquid chromatography (radio-HPLC): a normalHPLC-apparatus with a detector for radiolabeled compounds. There are twogeneral principles:a) The detector contains a solid scintillator.b) A scintillator liquid is mixed with the eluate of the HPLC-column.In both cases, an integrator processes the signals of the counter in thedetector and calculates the peak area resulting in a measure for quantifica-tion of each compound separated in an HPLC-run.

– Radio-gas chromatography (radio-GC): this method is only used in few caseswhen substances with high volatility are investigated.

3.4Analytical Separation and Quantitation Methods

The analytical procedures for separation and quantitation of non-radiolabeledcompounds are similar to the methods listed for radioanalytical methods: TLC,HPLC, and GC. The main difference is the detection method and the fact thatmore effort is necessary for separation of the test substance and its metabolitesfrom compounds occurring naturally in the respective matrix. All results there-fore have to be compared to blank values of the particular matrix. In addition,the samples more often need more extensive pretreatment (clean-up proce-dures) before injecting them into the chromatographic systems.

For the preparation of samples for chromatographic analysis one or more ofthe following steps may be necessary:

– Homogenization of the matrix– Liquid extraction from the matrix (e.g., in a Soxhlett apparatus)


– Separation of polar and nonpolar components by:� Liquid-liquid separation� Solid-liquid separation� Derivatization (e.g., esterification)

Commonly used GC-detection methods are:

– “Universal” detectors such as the flame ionization detector (FID): suitable forsubstances which are well combustible.

– “Selective” detectors such as the electron capture detector (ECD): suitable forsubstance with high affinity for electrons such as molecules containing halo-gens or aromatic compounds with nitro- or cyano substituents.

– “Specific” detectors such as the nitrogen/phosphorus selective detector: suit-able for substances containing several atoms of nitrogen and/or phosphorus.

– Mass spectrometer (MS): GC-MS is normally used for identification pur-poses but can also be used for the quantitation of compounds not detectablewith other GC-detectors.

Commonly used HPLC-detection methods are:

– UV/VIS-detector: suitable for substances absorbing UV- or visible light.– Polarization detector.– Mass spectrometer (MS): HPLC-MS is normally used for identification pur-

poses but can also be used for the quantitation of compounds not detectablewith other HPLC-detectors.

Besides the methods mentioned above, the eluates from the chromatographycolumns can be collected and analyzed using any suitable method not mentio-ned before (nucleo-magnetic resonance – NMR, atom-absorption spectrometer– AAS, or infrared spectrometer – IR).

3.5Methods for Identification of Compounds

Before attempting the identification of a certain compound in a mixture, thiscompound has to be separated from the other compounds present in thismixture. The separation methods are the same as mentioned above. For identi-fication, the separation method is combined with a mass spectrometer system(GC-MS or HPLC-MS) or with NMR-analysis. The function of these detectors isdescribed in the common analytical literature.

4Which Test Systems are Used for Which Purpose?

Some biodegradation tests have been used for quantitative risk assessmentwhich is mostly focused on toxicity data. For example, in the GESAMP hazardevaluation procedures (with 6 overall criteria (A–F) organized in 19 sub-co-lumns), biodegradation features only in subcolumn A2, whereas toxicity-relatedparameters are present in all other criteria (B–F) [24]. This overall process re-


views chemical, biological, and physical properties to evaluate the fate of a chem-ical in the environment. In the past it has been used mostly for new chemi-cals, for which test data have to be submitted for registration in advance, e.g.,under the Toxic Substances Control Act in the USA or the EC Directives 67/548,79/831, 91/414 and 92/32, and related national legislation in the EuropeanUnion.

4.1Screening Tests

Two extremes in biodegradability testing are screening tests for ready and in-herent biodegradation and field tests. If these different test systems are com-pared, the analytical methods have to be considered, as they and the test setupitself have a major influence on the validity of predicting biodegradation in na-tural environments. For screening tests indirect methods are mostly used, de-termining parameters like oxygen demand (BOD or COD) or the changing con-centrations of dissolved organic carbon (DOC). This is done for the sake of easydetermination and to get an indication of the extent of degradation, but it hassome drawbacks. When the different screening methods were tested againsteach other they sometimes gave the same relative order of biodegradation ofdifferent chemicals but the time frame for each of them was different [25].Other authors found widely differing results, especially for those compoundsused as examples in this chapter [1, 26].

Only the biodegradation of chemicals studied with the same test can be eas-ily compared, and therefore the name of the test should always be mentioned[27]. These authors compared the degradation of several chemicals in the Zahn-Wellens-, the MOST-, the Closed-Bottle-, and in their self-devised GSF-Test (fordetails of the tests see Table 1). For 4-nitrophenol (4-Np) the results rangedfrom 55% degradation to only 1% mineralization and 35% metabolites formed.For hexachlorobenzene only one test was appropriate, the GSF-test using 50 mg/lof radiolabeled substrate, as the solubility of this chemical was too low. It resul-ted in 1% CO2 and metabolites each. The different drawbacks of screening testslike problems with water insoluble compounds and the high initial concentra-tions needed are discussed in more detail above and in Howard et al. [25] andStruijs and van den Berg [28].

Most of the screening tests only yield qualitative results to classify substancesin groups:

1. With the potential to be (readily) biodegraded, so-called soft or white listcompounds.

2. With the potential for degradation under specific circumstances (e.g., in theZahn-Wellens Test with a very high population density), leading to gray listcompounds.

3. With no biodegradation in the test period, leading to “hard” or black listcompounds.

The results cannot be used to determine degradation rate constants, and there-fore they cannot be used for a quantitative prediction of the fate of a chemical


in a natural environment. For a prediction it needs to be known how fast and towhat extent a substance will be mineralized under natural conditions.

This should be obvious if the inocula are considered (see Table 1). For almostall screening tests sewage or sludge is or can be used, sometimes even elaborateconcoctions of the two with soil added (MITI-test). To determine whether bac-teria with the ability to degrade a certain chemical exist, this approach – a clas-sical bacteriological technique called enrichment culture – seems feasible. Butit cannot under any circumstances be used to extrapolate to degradation ratesin nature, i.e., in a surface water body, where several environmental biotic andabiotic factors are at work simultaneously. This holds true especially when littleor no waste water input is received by the surface water.

Some tests may show the potential for degradation of a chronically pollutedriver (MOST, Closed Bottle), but their results cannot be regarded as more thana potential. In reality the key controlling feature is the environment, which in-cludes the substrate [29]. Very often laboratory tests are based on environ-mental irrelevancies, using unnaturally high concentrations of chemicals andignoring important microbial interactions [30].

Even the applicability of some screening tests using sewage or sludge, e.g.,the Zahn-Wellens Test, to what is happening in real sewage treatment plants re-mains questionable, as the residence time of waste water in such a plant mayvary but is mostly short.

To sum up, screening tests can be used to rank chemicals in a specific rela-tive order of potential biodegradability if the same test and standardized refe-rence compounds are used. The results can give management advice insofar asnew substances can be grouped into white, gray, and black lists accordingly. Theblack list substances should not be registered or should be phased out as soonas possible. The gray ones should be submitted to additional testing, e.g., simu-lation or field tests using natural conditions, i.e., low concentrations, low tem-peratures, or low oxygen concentrations. This procedure may lead to differen-tiated results, advising phase outs or restrictions of use/emission in certain en-vironmental compartments. During this evaluation period the precautionaryprinciple should be enacted, i.e., no registration for new substances.

The “white” compounds which are degraded easily in all screening tests forready biodegradation may not be a problem. But depending on their mode ofapplication they may not end up in adequate sewage treatment plants. If theycan get into the environment or monitoring studies already prove they arethere, simulation tests are necessary to determine degradation rates in the rele-vant compartments and to identify problem areas as described for gray list sub-stances above.

4.2Simulation Tests

The next steps in testing are the much more specific simulation tests. They arean opportunity and a risk at the same time as much needs to be known aboutthe simulated environment. Very important are the source of the inoculum andits preconditioning, like acclimation to laboratory conditions, adaptation to the


chemical under study, or pre-adaptation to it in the environment where thesample was taken from.

Instead of using mineral salts media, natural water, sediment, or soil samplesare mostly used. These are brought to the laboratory and treated or stabilized,e.g., by filtering, acclimation in an aquarium, or other methods. Most simula-tion tests are static tests, but in some systems a flow-through mechanism ismaintained. Another important feature is the means of agitation, as the oxygenconcentration in samples with sediment or soil needs to be maintained in anenclosed system. This can be achieved by stirring, shaking, or by continuousflow of aerated water or air through the system.

Since 1955 the River Die-Away Test has been used. It has been recommendedby the US-American EPA since 1978 for biocide registration, requiring radio-labeled test compounds [25]. A major advantage of this technique is the mea-surement of mineralization of the test compound with a sensitive and specificmethod allowing low concentrations to be used. A drawback is the use of waterwithout sediment.

Therefore the core-chamber method with intact sediment cores came intouse, as described, together with other systems, by Bourquin et al. [31]. Theseeco-cores are artificial laboratory microcosms that closely mimic natural con-ditions for studying microbial interactions. They belong to the group of micro-cosms, of which an array of different systems exist. They are considered simpli-fied small-scale models for ecosystems and can consist of more than just waterand sediment components. Often they are used to quantify transport, bioaccu-mulation, and toxicity mechanisms, sometimes using flow-through systems,which limit the degradation. Every method has to be studied closely to deter-mine the advantages and drawbacks of the system, and of the environment sim-ulated.

A good example, how the size and setup of microcosms can influence results,is shown by Spain et al. [32]. In their experiments with flasks, eco-cores, and twoother kinds of larger microcosms, the rate of biodegradation of 4-Np variedwith the type of test and the adaptation status of the inoculum from initial ra-tes of 0.02–0.48 mg l–1 h–1 to rates from 2.56–17.82 mg l–1 h–1 (after 88–460 h ofadaptation).

Very often mass balancing is not done in microcosm studies, and thereforethe disappearance of the substrate has to be interpreted with care, especiallywhen the compound under study is not used in its radiolabeled form. The useof proper abiotic controls is absolutely necessary. This was shown by van Veldand Spain [33] in their experiments comparing the degradation in shake flaskswith and without sediment with that in intact sediment/water cores, in whichthey could follow the mineralization. Only in the eco-cores and in one sedimentshake flask a significant decrease of 4-Np was observed. But only 6–13% turn-ed out to be carbon dioxide, 39–65% was tightly bound to the sediment (in theabiotic control 23%), pointing to degradation products with stronger sorptioncapacities.

Shaking a sediment fully changes its properties, mainly enhancing the ac-tivity and available number of the microflora and also facilitating adsorption byexposing more surfaces. This phenomenon may be naturally occurring, e.g., in


tidal waters, but as a rule sediment and soils are more stable. Therefore un-disturbed sediment samples should be used to simulate biodegradation in mostnatural environments.

For a good simulation test the objectives and environmental conditions sim-ulated have to be clearly stated as there is no fixed definition for a simulation ora microcosm test. As the design has a large influence on the kinetics and the ex-tent of degradation, the size, the sediment/soil to water ratio, and the supplywith water and oxygen has to be appropriate for the simulated environment. Itshould be the same when the mineralization of different chemicals is compared.

4.2.1Degradation Studies in Water/Sediment Systems

As an example for a typical simulation test for the investigation of biodegrationin soil or sediments, one exemplary procedure used for water/sediment studiesis described in detail below. Other studies using soil as a matrix have differentintentions but they are similar with respect to the methodology and the limita-tions posed by the analytical methods.

Water/sediment studies are required especially for the registration of new ac-tive ingredients used in biocides (plant protectives) if the standard laboratorytests have not been positive. Since there are no international guidelines, somecountries have developed national ones. The German guideline was issued bythe Biologische Bundesanstalt (BBA) in Braunschweig [34].

Two water sediment systems are collected from an uncontaminated freshwa-ter, differing in the organic carbon content. The water is characterized by deter-mination of oxygen concentration, redox potential, pH, total N, total P, DOC,and water hardness. The sediment is characterized by determination of particlesize distribution, organic carbon, pH, total N, total P, microbial biomass, drymatter, redox potential, and cation exchange capacity.

Prior to the test, the water is separated from the sediment. Weighed amountsof the wet sediment and of the water are transferred, resulting in a sedimentlayer of 2.5 cm and a supernatant water layer of 6 cm. The flasks are placed onan orbital shaker in the dark at 20°C and are allowed to equilibrate for approxi-mately 4 weeks respectively, before the 14C-labeled substance is applied. Theamount to be applied is calculated on the basis of the maximum field applica-tion rate and, assuming a homogenous distribution, to a depth of 30 cm in thenatural system (water of creek/lake close to fields).

Through stirring, aerobic conditions are achieved in the water, whereas agradient in the redox potential is maintained in parts of the sediment (seeFig. 1). As an alternative, anaerobic conditions can be achieved if nitrogen is in-troduced into the aqueous phase in the flasks, for example by introducing asmall Teflon tube through the septum.

The experimental conditions analyzed are dissolved oxygen, redox potential,temperature and pH in the water, and redox potential in the sediment. The testsubstance concentration is measured by means of radioactivity measurementsand chromatographic methods (TLC and/or HPLC) in order to separate parentcompound from the metabolites. The radioactivity is measured separately for


water and for sediments. The compounds in the sediment can only be charac-terized after extraction of the radioactivity using organic solvents and/or basicor acidic conditions. Chromatographic separation of parent compound frommetabolites can be carried out in liquid systems only, whereas determination ofthe total radioactivity is possible by submission of sediment samples to com-bustion. Additionally the radioactivity in the traps is analyzed to determine vo-latile metabolites and CO2.

Water/sediment studies as described above simulate conditions occurring inthe environment very well. The test systems are selected sections of real eco-systems, the environmental conditions corresponding well to the conditions oc-curring in summer months in the real environment, only light effects not beingconsidered. Since the test method allows differentiation between parent com-pound and degradation products, and since several samples are collected at dif-ferent time points, the generation and further degradation of the metabolitescan be recorded – in the aqueous phase as well as in the sediment. Since a clos-ed system is used, the study is carried out with radiolabeled test substances andthe volatile compounds are also collected, a total balance of all degradation pro-ducts can be achieved.


Fig. 1. Water/sediment system as prescribed by the German BBA for the testing of plant pro-tectives. The incubation flask (volume ca. 1.0 l) is equipped with a device for slow stirring ofthe water only, with a trap for volatile components and a septum allowing the introduction ofgas. The trap contains quartz wool impregnated with paraffin oil and soda lime and is per-meable to air. Sampling is done after 0 h, 6 h, 24 h, 2 d, 7 d, 14 d, 30 d, 60 d, and more than100 d after application of the test substance. One flask is used for each sampling date

4.2.1.1Limitations of Simulation Studies Using Soil or Sediments

Studies using complex systems such as soil and sediment have their limitations.Often there are residues bound to the soil which cannot be extracted using normal chemical extraction methods. Little is known about these compo-nents. Depending on the type of test substance, the reasons for the amount and the occurrence of bound residues can be very different. One mechanism isthe irreversible adsorption of parent compound and/or metabolites to differ-ent kinds of soil particles. Other mechanisms which are discussed are the in-tegration of degradation products into humic and fulvic acids or other com-ponents of the soil. It is also possible that some parts of the test substance areused for microbial cell compounds such as structural proteins, cell wall com-ponents, or extracellular polymers which are adsorbed to surfaces of soil par-ticles. Most studies end up with a certain amount of “bound residues” and it isnot known how relevant they are for a hazard assessment of the chemical sub-stance.

Another problem is very polar degradation products which cannot be sepa-rated in normal phase TLC-systems or in reversed phase systems. In long-termstudies such as lysimeter studies, these metabolites often comprise the mainfraction of radioactivity found in leachates or in soil extracts. These substancesare regarded as a complex mixture of polar components in the soil – metabo-lites of the normal physiology of soil microorganisms and hydrolysis or oxida-tion products of these metabolites. Since chemical characterization would bevery expensive and would probably give no relevant information, leachates con-taining high concentrations of these substances (more than 0.5 mg/l test sub-stance equivalents) are used for ecotoxicity tests. If these leachates have no ef-fects, e.g., on crustaceans, they are regarded as harmless.

4.2.2Possibilities to Predict from Simulation Test Results

Simulation tests can lead to habitat specific predictions, especially when dif-ferent habitats like rivers or lakes are compared with the same test setup.Alexander and co-workers found differences in rates and extent of mineraliza-tion in lakes of different trophic status. But additionally the substrate concen-tration influenced the results, leading to a threshold for degradation or to longlag times before degradation started in some cases. Under these conditions theystrongly advise against extrapolation from laboratory conditions to naturalwaters [30, 35, 36].

Again the results do not represent the absolute truth on the biodegradabilityof a substance but they indicate in which environment and under which condi-tions major problems with a chemical might be encountered. A chemical maybe mineralized in samples from a polluted river or coastal environment but notin ocean water, or the same may happen in samples with and without sediment[37, 38].


4.3Mesocosms and Field Studies

The next steps of sophistication in measuring degradation rates close to naturalones are mesocosms and finally field studies. The exact line between micro- andmesocosms is hard to draw: mesocosms are larger and sometimes they are usedoutdoors or at least with some outdoor features like a natural light regime. Theycan be artificial ponds or canals, large plastic enclosures mounted in a lake orthe sea, open or enclosed aquatic-terrestrial systems, or wholly terrestrial me-socosms with single or different compartments. Up to six compartments havebeen analyzed for the fate of the test compound and its metabolites and the sizevaried from a half to hundreds or thousands of cubic meters in volume [39].

Table 2 explains two types of lysimeter, one of which has to be used inGermany (and Europe) for the registration of biocides if there is reason to be-lieve that the chemical itself or its degradation products can reach the ground-water in concentrations of 0.1 mg/l or 0.5 mg/l respectively. Modeling of the dis-tribution after application, e.g., with the PELMO pesticide leaching model [40]is used to estimate the potential risk. The test has a duration of 3 years.

A natural soil core is utilized in both cases. The larger closed lysimeter ismore complex as more factors can be controlled. But it has turned out to be toomuch of an effort for routine practice. Nevertheless some interesting resultshave been obtained and one example will be given below.

Both lysimeters can be used:

1. To obtain substance specific data on distribution, degradation and residuesrequired for registration.

2. To provide information on the effects of various factors, e.g., application rateor soil type on distribution and degradation.

3. To enable quantitative analysis of partitioning, accumulation, and the degra-dation process of chemicals, including the atmosphere to obtain a mass ba-lance.


Table 2. Comparison of two environmental test systems modelling terrestrial ecosystems [41]

Type of Function Number Size Surface Weather Analysis Analytical Mass-system and use of (diam. ¥ area regime method compart- balance

replicas depth) ments

Large- Legal 2 113 cm ¥ 1.0 m2 natural 14C P, L, S, Pl No,scale require- 110–120 open

ments cm systemLarge- Fate/me- 1 71 cm ¥ 0.4 m2 chosen 14C P, L, S, Pl, Yesscale tabolism 50 cm Air

P = Percolation water; L = Leachate; S = Soil; Pl = Plants.The system uses legally prescribed soil, consisting of loamy sand with low humus content, ta-ken as an undisturbed core and acclimated to the study site conditions for 2 months: pH =6.1–6.3; water capacity = 13.8–33.5; organic carbon = 0.02–1.31 (depending on depth).In all systems the temperature of the lower half of the soil and the amount of rainfall is con-trolled to stay at 8°C and no less than 800 mm of rain.

To give an example of the use of the closed lysimeter, the degradation of te-trachlorobenzene in two different setups with soil is described and compared.Figure 2 demonstrates the differences in mineralization of 1,2,4,5-tetrachloro-benzene in soil in a laboratory study and this mesocosm inoculated with a pureculture of bacterial degraders. For both studies the same soil and the samenumber of cells per gram of contaminated soil was used. The closed mesocosmor lysimeter used (see Table 2) was incubated under the climatic conditions ofan average day in June in Northern Germany regarding temperature, humidity,light conditions, and wind during a 24 h day. Only the top 5 cm (33 kg) wereremoved, homogeneously contaminated with the chemical and later on the bac-teria were added and then the contaminated material was reapplied onto thelysimeter surface; for the flasks the whole soil sample (50 g) was contaminated.The glass flasks in the laboratory were incubated at 15°C, the mean temperatureof the 24 h period of the “June day” [42].

The degradation process within the controlled mesocosm was much slowerthan under similar conditions obtained in a laboratory (see Fig. 2). This resultmight have been caused by a multitude of different factors and/or their combi-nation, possibly the periodical variation of the climate (temperature, water con-tent, etc.). One can only speculate about the exact reasons, and this exampleshows again that there is no simple correlation between data obtained in the la-boratory and mesocosm data, not to mention natural ecosystems.


Fig. 2. Comparison of mineralization of 1,2,4,5-tetrachlorobenzene by the bacterial strainAcidovorax sp. PS14 under different conditions. Unsterile soil (BBA standard soil) was usedin glass vessels containing 50 g soil (�) and in a mesocosm (soil core: 71 cm diameter and40 cm depth) (�). The tetrachlorobenzene concentration was 10 ppm in the laboratory study,the inoculum 107 cells PS14/g soil, the incubation temperature 15°C, the water content 40%of the maximal water holding capacity. The tetrachlorobenzene concentration of the meso-cosm was 19 ppm, which was only applied to the top 5 cm (= 33 kg), the inoculum 107 cellsPS14/g soil with chemical. The mesocosm was incubated under the climatic conditions of amean day in June in Northern Germany

Field studies are mostly done in soil systems, and sometimes with ponds orsmall sections of rivers [32, 43]. The mesocosms used by Kuiper and co-workers[44, 45] are the closest approach to a field study in a marine ecosystem.

A disadvantage of field studies compared to mesocosms is the necessity touse specific analytical methods to measure the disappearance of the substrate,leading to indirect evidence of biodegradation only. Besides, there is no possi-bility for abiotic controls. A mass balance in a field test and the distinction ofabiotic and biotic processes remain a formidable task and may not be possible[46].

Accompanying lab tests for mineralization are often done to obtain more in-sight into the biotic process, but they may not reflect what happens outdoors, ascould be shown by Kuiper and Hanstveit [45]. 4-Np was mineralized in bottlesincubated under almost precisely the same conditions in the laboratory, but notdegraded in the outdoor mesocosms in up to 50 days in their tests.

It seems that the best but most specific system for information on biodegrada-tion close to in situ conditions is a well modeled mesocosm in which factors at theecosystem level, which are thought to be relevant to biodegradation, are at workbut can be controlled. The size has to be chosen according to the system understudy, e.g., for an oligotrophic system the mesocosm needs to be larger [44].

Big differences remain between systems with and without sediment. The im-portance of integrating the sediment compartment is specific for the chemicalin question, e.g., it is more important for hydrophobic substances, but it shouldbe done more often, even if analyzing sediment samples is more of an effort.

4.3.1Possibilities to Predict from Mesocosm and Field Test Results and Comparison of Test Systems

It has to be kept in mind that it cannot be assumed that the physiological statusof microorganisms is unaltered after a sample of water, soil, or sediment is re-moved from the field [46]. In his review Madsen states that, despite decades ofdebate, the problems of extrapolating from laboratory results to the field havenever been solved, one reason being the impasse due to the methodologicallimitations mentioned above. It has been hard to prove degradation in situ un-equivocally. His recommendation is an elaborate stepwise strategy for deter-mining in situ biodegradation, explaining the complexities and the necessaryarray of parameters to be measured. Only if proof of biodegradation in the fieldcan be obtained, e.g., by enhanced numbers of protozoan predators or uniquemetabolites appearing, the classical microbiological methods of isolation andenumeration of biodegrading organisms and flask assays can be used as con-firmatory evidence. Adaptation alone is not conclusive evidence for in situ bio-degradation [46]. Results can be taken directly for predictions for specific habi-tats if no or slow change is documented, i.e., by monitoring data. If a fast dis-appearance is determined, additional lab tests should verify the mechanism,focusing if possible on mineralization and excluding sorption, leaching, sedi-mentation, volatilization, or bioconcentration.A promising field seems to be thestudy of combined photo- and biodegradation.


Up to this time it remains a challenge to determine in situ biodegradation ra-tes. As several competing biotransformation processes may act on the sub-strate(s) under study, the extent of mineralization and the fate of transforma-tion products need to be studied under field conditions.

A promising approach was shown by Ulrich and co-workers [47]. They com-pared measurements of the NTA concentration in the Swiss Greifensee with re-sults of a mathematical model, simulating transport and transformation pro-cesses. In the epilimnion NTA disappeared with a rate of 0.02–0.05 per daywithout any seasonal trend. They attribute this process to biodegradation.

Several habitats need to be studied, and their selection should proceed ac-cording to the application and fate of the chemical in question. An extrapola-tion from fresh water test results to what will happen in salt water should not bedone. Similarly, tests with soil or sewage cannot be used for prediction of thefate in aquatic systems or vice versa. The most sensitive and exposed habitatshould be chosen through a series of tests and calculations and the fate of thecompound tested in depth in the resulting compartment(s) to derive advice formanagement purposes.

5Potentials and Limitations for Prediction from Laboratory Results

A lot of factors have to be taken into account, which influence the predictabilityof biodegradation in nature. A main focus in this chapter will be on the inter-actions between some of these factors. These interactions can be additive, syn-ergistic, or antagonistic. A lot of studies have tried to identify and quantifysingle factors, but the identification and even more the quantification of the in-teractions is just beginning.

5.1Concentration

Starting with the limitations, it should be clear that one of the main influencingfactors is the concentration of the substance in question. It has to be seen in cor-relation with the number of bacteria capable of mineralizing the chemical[48–50]. Dilution can either bring the number of degrading bacteria down sothat no effect is measurable and/or dilute the chemical so that no growth is pos-sible anymore. Additionally it is very unlikely that concentrations below 1 mg/lcan induce enzymes, as Hanne et al. [51] determined at much higher con-centrations for an Arthrobacter and a Nocardia strain. Even if an experimentwith long incubation periods can force the enrichment/activation of degradingbacteria there still is no guarantee that this will happen in nature where theorganisms are not starved to biodegrade.

An especially interesting phenomenon, which impedes the prediction fromlaboratory results, is the threshold for growth of the microorganisms capable ofdegrading chemicals [52]. It should be assumed that the interaction between thesubstrate concentration and growth is not linear at low concentrations.Van Veldand Spain [33] noticed the following effect in fresh water sediment cores. When


a first dose of 4-Np was below 10 mg/l, they found a slow degradation rate whichwas not increased after a second addition of 10 mg/l. When the first dose was60 mg/l or higher, a second addition was degraded faster. It seemed that 10 mg/lwas a threshold for adaptation to faster degradation and was very likely relatedto the missing growth of the degrader population.

Typical graphs of mineralization after an adaptation phase cannot be model-ed with models regularly used to describe degradation kinetics. The subtractionof the lag is arbitrary, and first order kinetics do not fit. Therefore data from thiskind of degradation cannot be used as an input for most models. The multitudeof possible reasons causing an adaptation phase or, as it is often called acclima-tion phase, singly or in co-operation was intensively studied by Wiggins [53].

Even if a small population is present which can degrade a low concentrationof the chemical in question, the kinetics often differ from that at higher con-centrations, i.e., with a less steep incline (i.e., a lower rate) and lower final ex-tent of mineralization as can be seen in Fig. 3.

All samples with 4-Np concentrations ranging from 0.08 mg/l to 250 mg/lshowed no mineralization during the first 10 days of incubation at 20°C (Fig. 3).Obviously an adaptation was necessary before degradation could start. Thendifferent concentrations of 4-Np were degraded in two different patterns. Thehigher concentrations showed a steep increase after 13 days. All concentrationsbelow 10 mg/l were degraded at a much slower rate and to a lower final extent.Maximum degradation rates were 0.21%/h for 0.08 and 0.8 mg/l (for 8 no ratewas determined) whereas they were 0.62%/h and 0.77%/h respectively for80 mg/l and 250 mg/l. Therefore the increase was not proportional, the two high-


Fig. 3. Mineralization of 4-nitrophenol in different concentrations in natural seawater sam-ples from the Kiel Fjord. The incubation temperature was 20°C. Natural mixed communitiesand radiolabeled 4-Np were used. The sampling months were December for 8 mg l–l and Mayfor all other concentrations

er concentrations showing a much higher degradation rate than could be ex-pected from extrapolation alone or vice versa.

MPN counts showed that the number of bacteria capable of degrading 4-Nphad increased substantially after 16 days in both incubations with high concen-trations. At the low concentrations and in a control without 4-Np no degraderscould be measured with the MPN method till the end of the experiment after35 days (limit of detection > 3/ml).

The adaptation phenomenon is complicated as some chemicals need more orless rare enzymes to be mineralized. These enzymes will not be constitutionalin most cases but will have to be induced as is the case for 4-Np [5]. Thereforesome authors postulate a threshold for adaptation, which may be related to theformation of these non-constituent enzymes. It should be noted in this contextthat a lot of the genetic information for the degradation of xenobiotics is plas-mid coded, i.e., the ability for degradation of toluene, 2,4-D, or p-chlorobi-phenyl [54]. All enzymes involved in a plasmid coded pathway are inducible bythe primary substrate [54]. Whether enzymatic adaptation can be the onlyreason for the often observed slow or postponed degradation, or if slow or nogrowth is an additional factor, has not been resolved for degradation underclose to natural conditions. Probably both factors act together.

An interesting fact is that slow degradation remains possible at low concentra-tions. But the kinetics are totally different from those at higher concentrations. Togive an example the mineralization of 4-Np in a water sample from the Baltic Seais illustrated in Fig. 4.After 80 days of incubation 1 µg/l 4-Np was mineralized anda second addition did not show enhanced degradation (Fig. 4). There probably


Fig. 4. Mineralization of 1 mg l–1 4-nitrophenol in a water sample from the Central Baltic Sea.After 110 days a second dose of 5 µg l–1 was added (A). Or the bacteria were harvested and in-cubated in freshly prepared sterile media with a second dose of 1 mg l–1 4-Np (B – freshly ta-ken Baltic Sea water, C – aged brackish water). The incubation temperature was 20°C. Thestandard deviation is only shown when it is higher than 1.5%

was no growth and only minimal degradative capacity present in the biota of theoriginal sample. It was not enhanced by the low dose but it was still present aftera prolonged incubation. Other experiments incubating samples without 4-Np de-monstrated a loss of the degradative capability in a few weeks [55].

Examples for substances where adaptation is often observed are 4-Np, NTA,and methyl-parathion [17, 26, 56–58]. The frequent practice of just subtractingthese lag periods, sometimes without even mentioning them in the publication,does not lead to better predictability of degradation in nature. After adaptationthe test system is changed either irrevocably or at least for a prolonged period.

There are obviously different kinds of substrates, those enabling growth,those only degradable through cometabolism, and probably some in between,for which other carbon sources may enhance the degradation as they are no suf-ficient growth substrates themselves. The resulting differences in character-istics and behavior of the two types of biodegradation were summarized byTiedje [59] using NTA as a case study. Important for this chapter are the differ-ences in kinetics: exponential kinetics with growth vs first order kinetics with-out growth.

In biodegradation studies there will be a major effect if mineralization withgrowth or only cometabolism happens. If growth can happen and is necessaryfor the chemical to disappear in an adequate time, anomalous behavior due tothresholds (for induction and/or growth) is possible as low ambient concentra-tions will influence this time. If this is compared to cometabolic processes with-out induction they will also start with a very slow rate, so we may be fooled intothinking that we see acclimation. The main difference should be that a cometa-bolic rate does not increase over time, just a certain percentage of the chemicalwill be slowly transformed. Robertson and Alexander [60] found that pesticidesnot supporting growth at all, like simazine and carbofuran, were not subject toaccelerated biodegradation in soil samples, whereas 2,4-D and propham (IPC)initially showed a lag-phase, then an increase in degrader numbers and in min-eralization rate. A second addition was only mineralized faster in the last twocases. In extrapolation and decision making it is of paramount importance toknow if a substance can be mineralized with growth, because then quantitativedegradation in nature can be expected. Still a prediction of how long this willtake can be difficult. A strong influence of environmental factors on degrada-tion has to be taken into account, requiring precise rate measurements underrelevant conditions. If a substance is only cometabolized, extremely slow rateshave to be expected in any case and possibly recalcitrant metabolites.

However, the problem with low ambient concentrations not allowing anygrowth under natural conditions remains. Can there be adaptation in naturewithout growth? This still remains to be proven, thus necessitating a graded ap-proach towards prediction of degradation in different habitats.

5.2Temperature

The next factor often overlooked is temperature or to be more specific the in-fluence of naturally low ones. There is no proof that bacteria react simply as ex-


pected in a chemical reaction for which the Arrhenius equation holds true. Themicrobial ecologists have separated natural populations of bacteria into psy-chrophilic, mesophilic, and thermophilic groups because the incubation tempe-rature (or that occurring in their natural habitat) has a major influence onselecting one of these groups. Room temperature may be adequate for a simu-lation of sewage treatment, even if it is more likely to be an artifact of time pres-sure to get results fast and of restricted access to temperature controlled roomson the site of the research institution. But for natural aquatic systems in north-ern Europe a standard incubation temperature of 10°C would make much moresense as there is no general way to extrapolate downward from higher tempe-ratures. There are very few studies on the effect of temperature on adaptation ofbacteria and on the kinetics and extent of mineralization. Some authors couldshow a much bigger effect than expected by simple extrapolation [21, 55, 61].

This effect is more pronounced if low concentration and low temperature acttogether. For example when uninduced microbial communities were incubatedwith low concentrations of 4-Np at low temperature, nothing happened inmonths whereas adapted communities took only a few days longer, degradingthe chemical at 10°C rather than at 20°C, at higher concentration [55]. For theeffect of a low concentration see Fig. 7.

Hales and Ernst [21] could measure NTA mineralization at 5°C in a riverestuary. The rate depended on the salinity and the sediment content, and filter-ing the sample halved the rate. The low temperature had a stronger influencewhen concentrations in the lower microgram range were used compared withwhen 1000 mg/l were used. At 12°C they measured half-lives of 4–29 dayswhereas the mean retention time of the water body in the estuary is 17days. Palmisano et al. [20] noted than NTA was degraded not only faster but also more completely at higher temperatures in samples from different rivercompartments in a polluted river.

NTA is an interesting example for a substance which is measured in theenvironment in spite of being biodegradable in several standard tests. Perry etal. [14] stated in a review article that NTA often leaves sewage treatment plantsuntreated in substantial amounts. It can mobilize heavy metals and polluteground and drinking water and it is therefore now routinely measured in someEuropean monitoring programs, together with the even more recalcitrantEDTA. He notes that NTA seems to be degraded very slowly at temperatures un-der 7°C as concentrations up to 100 mg/l could be measured in winter whereasonly 10 mg/l or less were determined in summer.

5.3Sorption

Sorption can be caused by different mechanisms like van der Waals forces,charge transfer complexation, hydrogen bonding, and hydrophobic interactions[62]. All adsorption processes except for covalent bonds are reversible.Chemical substances covalently bound to the humus matrix are called “boundresidues.” Sorption and degradation processes are dependent on each other[63]. The limitation of degradation processes by sorption was found to differ


with different bacterial strains. This implies that the bioavailability of soil-sorb-ed substances (e.g., atrazine [64], naphthalene [65]) also depends upon the de-sorption efficiency accomplished by a bacterial strain.

Several authors noted a strong adsorption of metabolites of the pesticidemethyl parathion in sediments up to 48%. In sterile controls there was only little adsorption, and therefore a biologically mediated sorption was postulated[33, 66].

5.4Oxygen Content

Oxygen is a prerequisite for most mineralization processes. It is usually not aproblem if only water samples are tested and the concentration of substrate isnot extremely high. But in sediments and soils, oxygen can be a very importantfactor.

The water content in a soil is inversely proportional to its gas content.Therefore, the oxygen content of a soil decreases with increasing water content.Hardly any aerobic degradation of chemical substances can be found in a watersaturated soil if no oxygen is brought in by mixing (e.g., parathion [67]). Thesmaller the grains of a soil and therefore the finer its porosity, the slower will bethe oxygen exchange within the soil. No optimal water content can be defined,but in different soils an optimal degradation is achieved with different watercontents.

This is demonstrated in Fig. 5. In all soils (A, B, C) the bacterial degradationof tetrachlorobenzene was lowest at 100% water saturation. In the sandy soil Awith the largest grains and pores, no difference was found for the degradationwith 40% and 70% water saturation. Here, a fast oxygen and water exchangecould take place. Soil B with medium grain and pore size showed a better de-gradation with the lowest water content of 40%. Apparently, the oxygen contentwas not sufficient for the strictly aerobic microorganisms at the higher watercontents. In soil C with a high clay content, equaling a very small grain size,much water was bound by the clay particles and therefore inaccessible for themicroorganisms. In this case, the degradation was better with the higher watercontent of 70% [42].

5.5Sediments and Soil

There are not many examples of a transfer of experiments performed in the la-boratory to the pilot scale. On a larger scale the degradation process is often re-stricted by suboptimal water content, e.g., dryness or insufficient oxygen sup-ply accompanying a high water content, as well as deficient bioavailability of thedegradable substance or other nutrients [68]; conditions of the natural envi-ronment often do not lead to such positive results as those obtained under con-trolled laboratory conditions.

Many single factors have been investigated by the research group ofAlexander and coworkers [69, 70], but the complex interactions between these



Fig. 5 A – C. Soil types (see text). Influence of different water contents in sterile soils upon themineralization rates of 1,2,4,5-tetrachlorobenzene by the bacterial strain Acidovorax sp. PS14.Water contents: 40% (�), 70% (�), and 100% (�) respectively, of the maximal water holdingcapacity of the soil; tetrachlorobenzene concentration: 10 ppm; inoculum: 107 cells/g soil; in-cubation temperature: 25°C

factors can hardly be covered by laboratory experiments. In a concrete reme-diation case described by Steilen et al. [71], none of four remediation techniquesapplied led to the successful degradation of a PAH-contaminated soil, but theauthors had not been able to forecast this.

The toxicity of a xenobiotic substance to the autochthonous soil microflorawas shown to be higher in smaller samples than in large-scale experiments [72].This can be explained by the greater variance of the microflora found in largersamples, so different effects can be balanced. Larger samples also have a higherprobability of containing microorganisms capable of degradation. Additionally,abiotic procedures like the draining reaction of chemical substances were shownto be different in natural soil compared with disturbed soil columns [73].

A special problem arises with soil and sediments exposed to xenobiotic sub-stances for a longer period of time prior to remediation (e.g., old neglected de-posits) so that complex sorption processes have taken place. These time-depen-dent reactions can hardly be simulated in the laboratory [74]. It is often tried toforecast the behavior and degradation of chemical substances in soil eco-systems with the help of mathematical models [75, 76], but these models areonly applicable for defined chemical classes, defined soils with defined watercontents etc., and have no general validity.

An interesting experiment was performed by the research group of Short etal. [77, 78]. They created a genetically engineered microorganism (GEM) ofPseudomonas putida with a plasmid which enabled the microorganism totransform 2,4-D to 2-chloromaleyl acetate in soil. Subsequently this metabolitewas mineralized by the endogenous microorganisms of the soil [77]. After thesepositive results the GEM was introduced into another soil type. Here they foundthat 2,4-D was only metabolized to 2,4-dichlorophenol, and not any further.This metabolite accumulated in the soil. Because of the toxicity of this com-pound, which is higher than that of 2,4-D, the number of fungi decreased morethan 400-fold resulting in a reduced soil respiration rate in comparison to a soilcontaining 2,4-D. This means that the degradation pathway of the GEM led todifferent metabolites in different soils and it was not possible to transfer the re-sults from one soil type to another [78].

5.6Grazing

Another phenomenon is the influence of grazers i.e., heterotrophic nanoflagel-lates, ciliates, or other protozoa living on bacteria. Galvao [79] observed thatonly at flagellate numbers of more than 6–8 ¥ 103/ml significant grazing can beobserved. Additionally it is important to note that flagellates thrive at a temper-ature of around 10°C. Andersen and Fenchel [80] determined that a minimumof 106 bacteria/ml are necessary for grazing to start. Therefore the concentra-tion of total bacteria and of grazers is decisive for grazing to start. If part of thepopulation grows only slowly they may not survive the grazing pressure andcan be severely decimated or perhaps even eliminated [69, 81]. If this part in amixed microbial community happens to be the one degrading the substance inquestion, the mineralization may stop or never start. The slow growth rate may


of course be caused by low substrate concentrations and/or low temperatures.Furthermore the influence of grazing on the adaptation process has beenshown to prolong the lag phase [82].

An example for the influence of grazing on an adapted population of degrad-ing bacteria will be given below.

5.7Interactions Between Concentration, Growth, Grazing, and Temperature

Many authors found some kind of correlation between the numbers of bacteriaable to degrade 4-Np and the onset and kinetics of 4-Np mineralization [55,83–85]. If the number is small, growth is necessary for the population to thriveand bring about measurable change in the substrate concentration. This growthand the consequential degradation is influenced by several factors, i.e., low tem-perature can severely restrict degradation and high grazing pressure can do thesame, especially as 4-Np is not such a good substrate for growth. Several authorsfound only small numbers of bacteria able to degrade a specific chemical, evenin adapted ecosystems (20/l to 80/ml in unadapted water, up to 780/ml in pre-adapted natural water, compared to a total bacterial number (TBN) of105–107/ml). Therefore it seems logical that so far a correlation between TBNand degradation has never been found.

To give an example, the degradation of 4-Np will be presented under differ-ent conditions typical for ecological stress factors. With this substance it is already known from laboratory experiments that 4-Np is principally bio-degradable. But an adaptation period is usually necessary to provide a largeenough number of bacteria with induced enzymes to attack 4-Np.

A simulation experiment was designed to determine the influence of bioticfactors like growth, the presence of other bacteria, and grazing by protozoa, aswell as abiotic factors like other carbon sources and temperature on degrada-tion. The experimental setup simulated the introduction and dilution of 4-Npand adapted degrading bacteria into seawater as might happen around sewageoutfalls, or when coastal waters are swept out into the open sea by currents.

A coastal seawater sample was incubated with 250 mg/l 4-Np at 20°C till all 4-Np was degraded to get an adapted inoculum. Then 5 vol % of this culture wereadded to differently treated sea water subsamples, so that around 105 degradingbacteria were present per litre. The amount of bacteria introduced was derivedfrom 125 ng 4-Np absolutely or equivalent to 12.5 mg/l 4-Np after dilution. Theycould be expected to multiply roughly up to ten times on the added 8 mg/l ra-dioactive 4-Np, assuming that 1 pg of organic substrate supports the growth ofa single bacterial cell [60]. The uptake, mineralization, and percentage remain-ing in solution were monitored using three replicas and a control.

Three treatments (BW, FF, and CY) were incubated at 20°C to simulate opti-mum summer conditions, two treatments (abbreviated as10°C and 3D) at 10°C,which is the optimum temperature for most psychrophilic bacteria and close tothe annual mean temperature of Baltic Sea surface water [86].

The treatment with sterile aged brackish water – a typical laboratory test set-up – promoted the fastest mineralization (BW, Fig. 6). Most of the 4-Np was de-


graded in the first 4 days before any multiplication of the total population couldbe measured. Therefore the fastest degradation possibly took place with little orno growth. An assumption, which is further proven by the shape of the curve,which looks like a first order reaction and could be fit with a first-order model(MARQFIT curve fitting program by Schmidt and Simkins as described in [49,50]).

In the treatment with natural Baltic Sea water plus cycloheximide, which wasadded to stop protozoan growth, the degradation rate was lower. The final per-centage of CO2 formed was about the same (around 70%). The best curve fit wasobtained with a model for logistic growth which is indicative of growth (CY,Fig. 6). Some growth of the degrading population can be assumed, as this treat-ment showed the highest uptake of 4-Np into the cells (7% uptake after 7 days)through the whole incubation time and a sevenfold increase of the total bacteriacount.

Under conditions coming closest to nature – a small amount of degradersintroduced into a natural water sample with protozoa, other bacteria and nu-trients already present – the mineralization of 4-Np was the slowest (FF, Fig. 6).For the first 4 days the degradation in the fresh seawater treatment (FF) follow-ed the CY treatment, which used the same water. No significant increase of theCO2 formed could then be observed at all for 24 h, and the mean value at day 5was even lower than at day 4. This curve could not be fitted adequately with any


Fig. 6. Mineralization of 8 mg l–1 4-nitrophenol after adaptation. Influence of differently treat-ed water. A mixed inoculum adapted to the degradation of 250 mg l–1 was diluted with newmedia containing only inorganic nutrients (BW), freshly taken seawater without protozoa(CY), or freshly taken seawater with competing bacteria, protozoa, organic, and inorganic nu-trients (FF). The incubation temperature was 20°C. The standard deviation is only shown,when it is higher than 1.5%

model. When the experiment was ended, only 56% was mineralized to CO2(Fig. 6). There were always flagellates present.

To enhance the possible influence of lower temperature and grazing by pro-tozoa on 4-Np degradation, a second set of treatments was incubated at 10°C(10°C and 3D, Fig. 7). Half of the samples were incubated for 3 days without 4-Np and without additional degrading bacteria so that a bloom of heterotrophicnanoflagellates (3.8 ¥ 103/ml) had already developed (3D (3 days), Fig. 7). Thenthe adapted degrading population was added.

After a one-day lag phase, the degradation started in the 10°C treatment un-til 25% CO2 were produced in 7 days (Fig. 7). The mean value of mineralizationdecreased thereafter from day 7 to day 9. Then the mineralization increasedagain, probably leveling of at 43% CO2 after 16 days. The 3D (three-day) treat-ment showed a linear increase of CO2-production to almost 30% in 11 days anda very long stagnation period till day 20 (Fig. 7). After 25 days the CO2-produc-tion stopped again and did not reach more than 48% at the end of the experi-ment (Fig. 7).

The slow growth rate, due to low temperature, grazing, and especially to pre-incubation resulting in higher initial flagellate numbers, led to lag-periods, dur-ing which the CO2-production did not increase. In all mineralization graphsonly the treatments with active flagellates showed high standard deviations(Fig. 6, FF and 7). The described factors combined can result in slow and incomplete mineralization as can be seen in Fig. 7, even when adapted bacteria


Fig. 7. Mineralization of 8 mg l–1 4-nitrophenol after adaptation. Influence of temperature andgrazing. Sample preparation was as described for Fig. 6 (FF), only the incubation temperaturewas 10°C. One set of samples was pre-incubated without 4-Np until a bloom of heterotrophicnanoflagellates could be microscopically observed after 3 days (3D). Then the adapted ino-culum and the 4-Np was added. The standard deviation is only shown when it is higher than1.5%

are utilized. Competition with other bacteria for nutrients or additional carbonsources can change the kinetics as well (Fig. 6).

It is possible that similar effects happen during the first days of 4-Np degra-dation in unadapted samples. The very small population tries to grow, but isslowed down at a much lower level of CO2-production than could be shown in these experiments, thus leading to the often perceived adaptation phenome-non (e.g., Fig. 3).

6What is Persistence, and When is a Substance Biodegradable?

6.1A Practitioner’s View

Halogenated dibenzofurans and dioxins are commonly regarded as typical ex-amples for highly toxic and highly persistent chemicals. The electronegativity ofthe halogen atoms make it very difficult to oxidize the aromatic rings present inthese molecules. Therefore these substances are regarded as not susceptible tobiodegradation. Also the non-halogenated cores of these molecules – dibenzo-furan and dioxin – were regarded as non-biodegradable until special strainswere isolated by Rolf Wittich in 1989 at the University of Hamburg. The NATEClaboratory, however, has found that up to 70–80% of 14C-radiolabeled dibenzo-furan (concentration in soil of 1 mg/kg) is turned over to 14CO2 within 1 year bythe autochthonic microflora in soil – without supplementation of specializedmicrobial strains [87]. Half of this amount was degraded to 14CO2 within1 month. After adding a specialized strain to the soil, the degradation was accel-erated – the residues were degraded within 1 week. At a higher concentration(1 mg/kg), the degradation rates were not as high: the maximum was 50% after1 year (10% after 1 month).

These results show that some degradation processes are only a matter oftime. They also show that the mechanisms resulting in degradability or non-de-gradability can run in several directions: normally higher degradation rateswould be expected at higher start concentrations. If 1 mg/kg can be degraded,normally, 1 mg/kg (if it is not toxic at that concentration) should also be degrad-able – lower concentrations tend to cause more problems since the substancecould be adsorbed into to the soil and would therefore not be available for bio-degradation. The experiments showed that obviously other things happened.The mechanisms are open for speculations: perhaps the bioavailablity of di-benzofuran depended on the water solubility – or maybe the soil microfloragrows a biofilm on surfaces contaminated with dibenzofuran – later on the in-ner bacteria of this biofilm may not be able to graze any more – hindering otherbacteria from reaching the substrate.

Persistence is not a general feature which can be assigned unequivocally toall chemical substances. There is a number of chemical substances which can beregarded as persistent to biodegradation under regularly occurring environ-mental conditions without supplementation of specialized strains (e.g., manyhighly halogenated aromatic compounds). There is also a number of chemical


substances susceptible to ready biodegradation. But there is a large group in be-tween. They may be biodegradable within a certain range of environmentalconditions (temperature, light regime, water activity, available nutrients) andwithin a certain range of substance concentrations in a specific matrix. The bio-degradability may also depend on the history of the contaminated spot, e.g.,have there been contaminations with the same substance or with antagonisticor synergistic substances?

6.2An Ecologist’s View

For an ecologist the environment determines almost everything. The individualorganism and one specific substance are only one combination of interactingfactors in a large array of abiotic and biotic factors, some of which have beendescribed before. One half-life or one rate constant as the all encompassingnumber to characterize the behavior of a chemical in nature (as happens moreand more frequently in hazard assessment) is not an acceptable option for anecologist.

First, it depends largely on the type of test system used what kind of a rateconstant is determined. What bacteria are in it, what concentration of chemicalis used? Are natural conditions simulated (if yes, which?) or are the conditionsoptimized to enhance biodegradation (i.e., 20°C, plus suspended sediment, plusnutrients and so forth)?

Second, what about the rate “constant,” when the rate increases only after acertain time during the degradation process? When ecologists take samples intothe laboratory they are painfully aware that a lot of changes will happen in themicrobial community, making the reaction differ from their “natural one” inunpredictable ways. This so-called bottle effect starts immediately after takinga sample and is the more pronounced the longer a sample is incubated.

For these two reason alone it can be deduced that it cannot be enough to usestandard tests to determine half-lives of the chemical in question.

In addition to these system-specific and microbiological factors, the en-vironmental conditions are at least as important. They are at work at the sametime as the microbe-chemical interactions, which are normally studied isolatedin the laboratory. To name but a few abiotic factors like currents and temper-ature regime, salinity, water quality and sediment content and characteristicshave to be taken into account. On the biotic side, competition, predation, andbiodiversity can play a major role. On top of these basic ecological descriptors,some interactions of the microbe-chemical system happen mainly on an eco-system level, i.e., the pollution history, the dilution factor, photo-oxidation, se-dimentation, and adsorption phenomena. Therefore, a substance can be de-gradable in a (polluted) river estuary, but not in the open sea or in an oligotro-phic lake. This may happen not because the right bacteria are not there, butbecause the environmental conditions are not the right ones for substantialmineralization to happen.

Important is the time scale – for rivers the residence time of a chemical isshort, for estuaries and oceans longer. There should not be a persistence prob-


lem if the major part of the yearly load can be mineralized in several tide cyclesup to one season, a period corresponding to half-lives less than 80 days [88]. Butif the chemical or its primary degradation products are toxic and/or bioaccu-mulate, such a long residence time may cause other problems. Additionally thelonger a substance stays in the system without biodegradation, the higher is thechance in an open system for dilution to levels too low for degradation to start,for evaporation/volatilization and further transport, or for sedimentation andtransfer to a different environment.

The environmental factors correlated with the protection of chemicals frommicrobial attack in waters have not been adequately defined – as the well knownmicrobial ecologist M. Alexander said in 1980 [89]. Therefore the ecologist hasto stress the importance of a clear differentiation between potential for degra-dation estimated in lab tests and proof of mineralization in situ. The examplesmentioned before may have already shown that recent research has led to someanswers and to even more questions.

Keeping all this in mind, the following tentative definitions of terms relevantto biodegradation are given from an ecological point of view:

– Persistance: no mineralization in one growing season or in an appropriatetime span for the ecosystem under study, therefore potential for accumula-tion and toxicity if input is permanent or recurring.

– Biodegradable: mineralization of a concentration encountered in nature un-der natural temperature and other conditions, in appropriate time span forecosystem under study. If a substance or its metabolites significantly adsorbto sediment, the sediment should be integrated into the study.� In rivers, half-life should be days.� In lakes, half-life should be no more than half of the time between the two

mixing periods.� In estuaries, half-life should be no more than half of the residence time.� In oceans, half-life may not be a useful parameter (if first-order model).� Adaptation should be possible in situ with concentrations measured in

situ, and a sufficient number of bacteria able to degrade the chemical inquestion should be present.

� In sediments/soils, mineralization should be possible in undisturbed sedi-ments/soils and under naturally occurring conditions (oxygen, temper-ature, etc.). The time spans should be as described for the different eco-systems before.

– Detoxification: no mineralization but transformation, i.e., transfer intohumic substance, leading to products which are tightly bound and will not be released in their original form anymore. Proof has to be obtained that notoxic intermediates are generated.

7Outlook

To sum up the text of this chapter and its main conclusions, some suggestionsfor further research and some management advice will be given. A leading role


has been taken by some recently updated international conventions for the pro-tection of the northern oceans (North-East Atlantic and the Baltic Sea). An at-tempt has been made to incorporate the precautionary and the polluter paysprinciples. It is stated that:

“hazardous substances are those which are persistent, liable to accumulate or toxic … Thetarget [is] their cessation within one generation (25 years) with the ultimate aim of concen-trations in the environment … close to zero concentrations for man-made synthetic sub-stances” (OSPAR Convention, Convention for the Protection of the Marine Environment ofthe North-East Atlantic).

This statement has recently been updated with the enlightened ecological in-terpretation that other substances may require a similar approach, even if theydo not meet all the criteria mentioned above, but give rise to an equivalent le-vel of concern. Transformation products, which generate concern, even if theoriginal chemical does not, are explicitly mentioned. Persistence is defined inthe draft as:

“if the conversion of a substance or of its degradation products in the marine environmentand in particular in the water column is slow enough to permit long-term occurrence and wi-despread distribution from its point of release” (OSPAR Draft 1997 [90]).

This shows clearly that persistence should be seen in the context of residencetimes and typical environmental factors in the ecosystem under study as elabo-rated in this chapter. Acclimation and adaptation phenomena have to be con-sidered when studies are designed to determine the fate of a specific chemicalin the environment. After lab tests established that a chemical can (only) be de-graded after adaptation the guiding questions should be: are adapted bacteriapresent in the receiving environment and, if not, can the natural microbial com-munity adapt to the chemical under study under close to natural conditions?

Interactions between several factors are the main problem for predictability,i. e., low concentration and temperature, or low temperature and low oxygencontent or the additional necessity of adaptation (enzymatically and/or becausethe population is too small). Natural ecosystems are highly dynamic and notmade to maintain stable or optimum conditions for fast degradation.

Therefore one step of a strategy to ease the chemical burden on the environ-ment should be an optimized understanding and functioning of sewage treat-ment facilities where these factors can be controlled much better. For thosechemicals which have to be used in open systems and which will not reachsewage treatment plants a more detailed array of tests is recommended takingthe described factors and influences into account.

7.1Some Research Suggestions for Better Predictability

It seems that at least for the protection of the oceans the first steps have been ta-ken to prevent further pollution. But the work of prioritizing which chemicalsshould be monitored, controlled, reduced, or eliminated still needs to be done. Thefollowing text tries to give some advice how microbiological research can help:



– Instead of – or additionally to – simulation tests, an estimation of the initialnumber of potentially degrading bacteria in the habitat under study, perhapswith two different concentrations, might help to find problem areas whereregulation of input should be a priority.

– Better kinetic models taking growth or no growth into account are needed.They should not be independent of concentration of the chemical (andperhaps even take second substrates into account).

– Ecosystem modeling should include other members of the food chain (com-petition, grazing), other transformation pathways (photodegradation,(bio)sorption), and all relevant compartments (sediment, surface layer).

– Ecological knowledge should be better integrated into the design of sewagetreatment facilities. A better understanding of processes in sewage treatmentplants may lead to new forms of controlled small scale treatment, tailored toproblem chemicals.

7.2Some Steps Towards Sustainable Development

As has been said before, when the knowledge about a chemical’s behavior in na-ture is still generally unknown or the results, e.g., of degradation tests are con-tradictory, the precautionary principle should be applied and the use of the sub-stance restricted and controlled until the results allow a final decision. Final ismeant in a relative sense as new toxic effects may be found, e.g., the ozone dam-aging substances some years ago or the hormone mimicking substances re-cently. On the other hand, new treatment methods, e.g., with adapted inocula,may allow controlled uses of formerly restricted chemicals.

Encouraging the development of principally degradable chemicals and theconcurrent methods for their disposal after use might lead to a thorough green-ing of the chemical industry and make its products more socially acceptable.Responsible care or other environmental management initiatives can be en-couraged by clear framework regulations and research incentives by the rele-vant government or international authorities.

It should be legitimate to assume that a new substance or substance group,which contain features proven to slow down degradation like branched sidechains, chloro-or nitro-groups, isomeric mixtures, belong to the persistent cate-gory until the opposite has been proven at the cost of the party interested in itsintroduction. If this rule was applied, new problem chemicals like toxaphene ornonylphenol (polyethoxylates) would never have been introduced after the ex-periences with the first generation of detergents and PCB.

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The Assessment of Biodegradation and Persistence

Bernd Beek, Stella Böhling, Christian Franke, Ulrich Jöhncke,Gabriele Studinger, Elisabeth Thumm

Federal Environmental Agency, Seecktstrasse 6-10, D-13581 Berlin, Germany,E-mail: [email protected]

Testing and assessment strategies for biodegradation and persistence of chemicals and theirimpact on microbial activity within the framework of environmental legislations are out-lined. An integrated assessment concept for biodegradation and persistence of chemicals inthe environment is presented, taking into account primary degradation, mineralization andbound residues. Results of simulation tests from soil and water/sediment systems are assign-ed to four classes of these three criteria and aggregated to an overall assessment resulting infour persistence categories allowing for a more comprehensive estimation of fate and behav-ior of a chemical in soils, surface waters, and sediments. Examples are given for an applica-tion of this assessment concept comparing data sets from simulation studies with plant pro-tection agents in soils and water/sediment systems. The proposed assessment scheme mayalso be applied for risk assessment in context with registration and notification proceduresof any kind of chemical substances by environmental authorities. An assessment scheme isalso proposed for the biodegradation and elimination in sewage treatment plants as well asthe toxic impact of substances on microbial activity with regard to impaired biodegradationpotentials in sewage treatment plants, soils, surface waters, and sediments. Deficiencies andfuture needs are addressed for achievement of a more realistic risk assessment of fate and be-havior of chemicals in the environment.

Keywords. Biodegradation, Persistence, Microbial toxicity, Risk Assessment, Deterioration

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 292

2 Some Basic Conditions for the Determination of Biodegradation 294

2.1 Aerobic vs Anaerobic Biodegradation . . . . . . . . . . . . . . . . 2942.2 Primary Degradation vs Mineralization . . . . . . . . . . . . . . . 294

3 Test Systems for Biodegradation and Elimination . . . . . . . . . 296

3.1 Screening Tests for Ready Biodegradation . . . . . . . . . . . . . . 2983.2 Screening Tests for Inherent Biodegradation . . . . . . . . . . . . 3003.3 Simulation Tests and Persistence Categories . . . . . . . . . . . . . 3013.3.1 Simulation Tests for Surface Waters and Soils . . . . . . . . . . . . 3033.3.2 Simulation Tests for Sewage Treatment Plants . . . . . . . . . . . . 307

4 Biodegradation Rate Constants . . . . . . . . . . . . . . . . . . . . 308

4.1 Determination of Biodegradation Rate Constants from Screening Tests . . . . . . . . . . . . . . . . . . . . . . . . . . 308

CHAPTER 5


4.2 Prerequisites for the Derivation of Biodegradation Rate Constants from Simulation Tests . . . . . . . . . . . . . . . . 309

4.3 Field Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310

5 Microbial Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . 311

5.1 Assessment Criteria . . . . . . . . . . . . . . . . . . . . . . . . . . 3115.2 Intrinsic Properties of Chemicals and Consequences for

Choice and Performance of Tests . . . . . . . . . . . . . . . . . . . 3145.3 Risk Assessment and Safety Factors . . . . . . . . . . . . . . . . . 316

6 Deficits and Perspectives . . . . . . . . . . . . . . . . . . . . . . . 317

7 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 318

1Introduction

In addition to natural substances, a large number of anthropogenic chemicalscirculate in the environment today, the disposal of which frequently poses greatproblems for humans and the environment. The most important process mini-mizing this hazard potential in water and soil is biodegradation. In the en-vironmental compartment air degradation is mediated by physico-chemicalprocesses.

The demand for products of high persistence is opposed to the demand fortheir almost complete degradation after use, apart from further utilization(e.g., recycling). Although nearly every substance is liable to transformation,i.e., degradation, continuous discharge can result in the achievement of asteady-state concentration in the environment or, if the decrease is slow, in asteady accumulation (geoaccumulation). Furthermore, chemicals can accumu-late in environmental compartments to persist there for very long periods oftime and also in organisms (bioaccumulation; see [1]) even after long distancetransport as far as into arctic and antarctic regions where conditions are par-ticularly impeded as to biodegradation (Persistent Organic Pollutants, POPs).Spectacular examples encountered today in many environmental samples aredioxines, polychlorinated biphenyls (PCBs), and DDT.

Biodegradation has always been one of the main fields of microbiological re-search, focussing on the identification of degradation pathways and metabolicprocesses. Consequently, the conditions of these tests aimed rather at the opti-mization of test conditions and selection of degradation-potent micro-organ-isms rather than simulation of environmentally relevant parameters.

Yet it was not until the 1950s and 1960s that the issue of biodegradation inthe environment became a matter of public interest when, resulting from thewidespread use of detergents, poorly degradable surfactants reached surfacewaters and foam formation made the problem visible. As a result more degrad-able detergents were produced and test methods were developed in order toquantitatively pursue their biodegradability.

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In the time that followed, all industrial nations developed degradation testswhich, aiming at mutual acceptance of test results, were internationally har-monized by the Organisation for Economic Co-operation and Development(OECD) and later in the European Union (EU).

Since the implementation of the German Chemicals Act [2] in 1982, new no-tified substances are subjected to biodegradation tests according to Annex V [3,4, 12] of the Council Directive 67/548/EEC [5].

For existing substances the Government of the Federal Republic of Germanyhas in its Existing Chemicals Programme chosen the means of cooperation be-tween Industry, Research, and Governmental Institutions, and this was followedby all EU member states.After priority setting, all available information on exis-ting environmentally relevant chemicals, including biodegradation, is collected,assessed, and summarized in dossiers containing proposals for appropriatemeasures.

Since 1986 the German Federal Environmental Agency has been included asan authority of consent into the implementation of the German PlantProtection Act [6], decisive information on biodegradation of many hazardoussubstances being obtained from sophisticated testing.

In contrast to the Chemicals Act, which requires the determination of biode-gradability only in the aquatic milieu, the Plant Protection Act stipulates man-datory degradation testing of pesticides in soil and sediments, thus making dataavailable to provide a deeper insight into the complex proceedings of biodegra-dation processes.

Data on degradation of detergents gathered from the implementation of theGerman Detergents and Cleansing Agents Act [7] increasingly broaden knowl-edge of degradation processes. This also applies to data concerning the biode-gradation of non-agricultural biocides.

From the environmental point of view, however, nearly all tests applied with-in the different fields of environmental legislation have in common that they ex-amine biodegradation in too high test concentrations and under non-represen-tative test conditions. Some standardized tests simulating environmental con-ditions in water and soil and in sewage treatment plants (STP) are available butlaborious and relatively costly.

Numerous chemicals classified according to laboratory tests as readily biodegradable are, however, detected in the environment. For a realistic assess-ment of biodegradation in the environment this uncertainty forces us to re-consider present strategies and also requires in part more sophisticated testmethods.

The objective of this contribution to investigation and assessment of bio-degradation within the scope of the implementation of environmental acts is to impart the present state of knowledge and to show up current activi-ties regarding the progress of further developments in testing and assessmentstrategies.

The Assessment of Biodegradation and Persistence 293

2Some Basic Conditions for the Determination of Biodegradation

2.1Aerobic vs Anaerobic Biodegradation

Biochemical-enzymatic transformations performed in the presence of oxygen es-pecially by micro-organisms such as fungi and bacteria leading to the oxidationof the substance characterize the aerobic biological degradation of organic sub-stances. In the various transformation steps generally more polar, hydrophilicmetabolites are produced, equally subject to further decomposition.As a rule endproducts of aerobic degradation are carbon dioxide, water, and inorganic salts.

In the absence of oxygen, i.e., under anaerobic conditions, ultimate degrada-tion leads to the formation of methane, carbon dioxide, and inorganic salts asend products (cf. Table 5).

Where oxidative degradation processes, e.g., catalyzed by oxygenases orperoxidases, are hindered due to lack of oxygen, reductive degradation proces-ses, e.g., the reduction of nitro-groups, will prevail. Halogenated hydrocarbonscan to a great extent be transformed anaerobically.

Anoxic conditions occur not only in the digesters of sewage treatment plants,but also in many surface water sediments, deeper soil layers, parts of groundwater, and in dumping sites. Especially poorly water-soluble and highly adsorb-ing substances are predominantly transferred to anaerobic sites. The impor-tance of anaerobic biodegradation processes seems to have been underestima-ted until now (see contribution by Reineke, this volume).

2.2Primary Degradation vs Mineralization

For the determination of primary degradation or mineralization, i.e., ultimatebiodegradation of a substance, different analytical parameters are used. Table 1lists the analytical parameters of tests most commonly applied within the prac-tice of chemicals legislation.

Test methods measuring ultimate biological degradation are mostly based onthe determination of summary parameters such as oxygen consumption (bio-chemical oxygen demand, BOD), carbon dioxide evolution (CO2), and dissolvedorganic carbon (DOC) or chemical oxygen demand (COD) removal.

The use of radiolabeled substances allows for both the analysis of primary-and ultimate degradation, especially of substrates in low concentrations. Bymeans of appropriate labeling an unspecific parameter such as CO2 may be-come a substance specific parameter (14CO2).

By measuring the oxygen consumption or the carbon dioxide or methaneproduction, no discrimination is possible between energy metabolism and bio-mass production. DOC and total organic carbon (TOC) both represent a mea-sure for the organic substances present; the degree of oxidation of the substanceor its metabolites has no influence on these parameters, as opposed to the COD-determination.

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If abiotic elimination processes (adsorption, hydrolysis, photolysis, volati-lization) can be excluded, in the case of single substance determinations the extent of the DOC- or TOC-removal is a measure of the degree of biological degradation. However, possible preceding oxidative degradation steps not lead-ing to an elimination of C-atoms from the parent molecule remain uncon-sidered.

In the case of adsorbing, poorly water-soluble, or volatile substances, a cleardistinction between biological degradation processes and elimination (e.g., viaadsorption, precipitation, stripping effects) is not possible. These latter proces-ses can only be measured and quantified by means of a sterile adsorption con-trol. For substances with the properties mentioned above, it must generally beassumed that, if such controls are missing, measurements based on DOC-ana-lysis cannot be regarded as degradation but only as elimination tests, and theresults assessed accordingly.

Using summary parameters for the degradation of a mixture of substances,all organic compounds present in the test are measured jointly; a decrease ofthe amount of the different compounds cannot be differentiated. Furthermore,if metabolites are formed their amounts cannot be quantified. Thus it cannot bedifferentiated whether the observed partial degradation results from the com-plete degradation of one constituent as opposed to others being not degradedat all or all substances undergoing only partial degradation.

Tests based on summary parameters are therefore only applicable to singlesubstances.


Table 1. Analytical parameters of biodegradation tests

Criterion ParameterPrimary degradation 1. Specific analysis for groups of substances, e.g.,

Anionic tensides: decline of MBASa

Non ionic tensides: decline of BiASb

Cationic tensides: decline of DSBASc

Hydrocarbons: decline of IR d absorption2. Analysis of single substances, e.g., active ingredients

of plant protection productsUltimate degradation 1. Summary parameter(mineralization) Consumption of O2 (BOD e in % ThOD f)

Evolution of CO2 (CO2 in % ThCO2g)

Removal of DOC h

2. Analysis of single substances (in case of radioactive labeling),e.g., active ingredients of plant protection products

a MBAS = Methylene Blue Active Substance.b BiAS = Bismuth Active Substance.c DSBAS = Disulfin Blue Active Substance.d IR = Infra Red.e BOD = Biochemical Oxygen Demand.f ThOD = Theoretical Oxygen Demand.g ThCO2 = Theoretical Carbon Dioxide evolution.h DOC = Dissolved Organic Carbon.

3Test Systems for Biodegradation and Elimination

In the beginning of the 1960s the first biodegradability tests for detergents weredeveloped, aiming at the prediction of the fate of the substances in the environ-ment from the results. It is therefore not surprising that the oldest legislative re-gulations to examine biological degradation originate from the field of deter-gents and cleansing agents.

Of these tests, some are still, although in a modified version, used today.Generally, substance or group specific methods were applied to analyze primarydegradation. In the meantime test methods were expanded to include all or-ganic compounds, at the same time both broadening the test program and in-creasing the stringency of some tests with respect to biodegradation potential.In general, testing for ultimate biological degradation, complete mineralizationis prescribed; however exceptions to this rule still exist for several substancegroups, e.g., tensides.

Today in chemicals legislation, a great number of tests exist, predominantlyto be conducted according to standardized test procedures and thus possessingmutual international acceptance:

– For the testing of chemicals including the determination of their biodegrad-ability the Organisation for Economic Co-operation and Development(OECD) recommends a three-tier testing hierarchy naming appropriate testmethods for each of these tiers [8].

– The EU has adopted this tiered program in its guidelines for New Substances[9]; depending on the amount placed on the market or imported quantities,the stipulated test methods provided are to a large extent identical with thoseof the OECD.

– The German Chemicals Act [2] based on the EU-Directives and in Article 2,paragraph 4 of the Ordinance on Test Certifications and other Registrationand Notification Documents under the Chemicals Act (ChemGPrüfV) [10]explicitly refers to the test methods cited there.

Furthermore, a great number of so far internationally not standardized testmethods to examine biological degradation exists, especially for the testing ofbiodegradation under environmentally more realistic conditions. In the light ofa testing strategy tailored to fit every single substance, greater access to thesetests should gain in importance.

Screening tests such as the tests for ready and inherent biodegradability de-scribed below cannot consider the various circumstances of natural conditions.The investigation of the degradation behavior of a substance in an appropriatesimulation model is extremely difficult, taking adequately into account the reallocal environment into which the substance is discharged or released in theevent of an accident. Nevertheless, to assess ultimately a substance with respectto its biodegradability or persistence it is necessary to investigate biodegrad-ability in a suitable simulation model.

Principally two different types of degradation tests can be distinguished:screening tests and simulation tests.

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First, in screening tests with relatively simple test conceptions, substances aretested in high concentrations compared to those normally found in the envi-ronment (but not inhibiting bacteria) in an aqueous solution or suspension intest vessels together with a small amount of a polyvalent inoculum. This meansthat no specialized or adapted micro-organisms are added. Instead the inocu-lum is generally taken from municipal sewage treatment plants (STP), river wa-ter, and/or soil suspension in order to represent a realistic spectrum of degrad-ing organisms present in the environment. The test substance as sole source ofcarbon is incubated in the dark for 28 days under conditions favoring bio-degradation with respect to pH-value, O2-content, and temperature. Biodegra-dation is generally followed by means of summary parameters.

Depending on the test substance loading, two test types are distinguished:

1. Tests with low inoculum concentration and high initial substrate concentra-tion (tests for ready biodegradation). Low micro-organism concentrationsare encountered, e.g., in surface waters.

2. Tests with high inoculum concentration and, compared with the inoculumamount, low initial substrate concentration (tests for inherent biodegrada-tion/elimination). Such conditions prevail in municipal STP.

Screening tests should allow for a general statement concerning the biode-gradation potential of a substance and do not simulate any specific environ-mental situation. Hence the degradation rates obtained cannot be transferred toenvironmental conditions.

Tests for ready biodegradation were designed to be stringent. It can be assum-ed that substances reaching the degradation pass levels in tests for ready bio-degradation within a defined period of time (10 days-window) will be ultima-tely degraded in the environment within surveyable periods of time (see Fig. 1).


Fig. 1. The concept of the 10 days-window. Idealized curve of a DOC-Die-Away-Test for readybiodegradability

Second, simulation tests should simulate the degradation behavior in a spe-cific environmental compartment as far as possible.

The results from simulation tests enable the classification of a substance intopersistence categories (see below). Furthermore, field-studies may be per-formed, the evaluation of which permit an assessment of the biodegradabilityof the substance under concern on an ecosystematic level.

3.1Screening Tests for Ready Biodegradation

If the amount of a new chemical placed on the market exceeds 100 kg/year or500 kg cumulative (base-set level), it is first subjected to testing for ready bio-degradation in a screening test.

Table 2 gives an overview of the internationally standardized tests of thebase-set level [11, 12].

A further development of the CO2 Evolution-Test is the CO2 Headspace-Test(ISO DIS 14593) [13]. Apart from the CO2 in the gaseous phase, this modifica-tion of the CO2 Evolution-Test also measures the solubilized CO2 in the aqueousphase. This test is suited for water-soluble, poorly water-soluble, and volatilesubstances.

Decisive for the evaluation of the tests for ready biodegradation are both thedegradation percentage after 28 days and the fulfillment of the 10 days-windowcriterion.

The biodegradation assessments are as follows.If:– ≥ 60% of ThOD 1 OR– ≥ 60% of ThCO2

2 OR– ≥ 70% DOC removal AND

the criterion of the 10 days-window 3 is fulfilledthe assessment will be readily biodegradable4

Reaching the pass-levels mentioned above but failing the 10 days-window re-sults in the assessment readily biodegradable, but failing 10 days-window.

In both cases further testing for inherent biodegradability on tier 2 is not ne-cessary.

If the pass levels are not met, the substance is classified as not readily biode-gradable. In this case further testing for inherent biodegradability on tier 2 isnecessary.

Classifications other than the above stated from the tests for ready biodegrad-ability within the scope of chemicals legislation for New Substances are notfeasible.

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1 ThOD = Theoretical Oxygen Demand2 ThCO2 = Theoretical Carbon Dioxide evolution3 exceptions: (1). The 10 days-window criterion does not apply to the Mod. MITI I-Test [11,

12] (2). According to the instructions of the Closed Bottle-Test [11, 12] weekly measure-ments are permitted. Consequently only a 14 days-window applies.

4 According to Technical Guidance Document (TGD) of the EU [14].

Given only a BOD5/COD ratio (BOD5=BOD after 5 days of incubation), a ra-tio > 0.5 is classified as indication of ready biodegradability 5.

Due to less stringent test conditions this quotient is from our point of viewnot comparable with results from screening tests for biodegradation and cantherefore only be assessed as an “indication.” Consequently, a ratio BOD5/COD< 0.5 is classified as no indication of ready biodegradability.


Table 2. Screening tests for ready biodegradability – comparison of present and former na-mes of tests

Present name Guideline Former name Guideline Commentof test of test

OECD EU OECD EU[11] [12] [8] [3]

DOC Die 301 A C.4-A Modified 301 A C.4 a

Away-Test AFNOR-TestCO2 Evolution- 301 B C.4-C Modified Sturm- 301 B C.5 a

Test TestModified MITI 301 C C.4-F Modified MITI 301 C C.7 b

I-Test I-Test Closed Bottle- 301 D C.4-E Closed Bottle- 301 D C.6 c

Test Test Modified OECD 301 E C.4-B Modified OECD 301 E C.3 d

Screening-Test Screening-Test(MOST)Manometric 301 F C.4-D a,e

Respirometry-Test

a By standardization of inoculum concentration resulting in a maximum of 30 mg substance(dry weight) per liter test medium or 100 ml effluent of sewage treatment plant per liter testmedium.

b By specification of inoculum concentration, i.e., measurement of cell density per liter testmedium; restriction of valid range of temperature and approximation of this range with therange in other tests.

c By enhancement of inoculum concentration to a maximum of 5 ml effluent of sewage tre-atment plant per liter test medium.

d By enhancement of inoculum concentration to a maximum of 0.5 ml effluent of sewage tre-atment plant per liter test medium.

e Newly included; coincides with the European version of the Japanese Modified MITI I-Testwith only one inoculum and decreased range of temperature.

5 This classification deviates from the TGD. Some EU member countries accept the BOD5,others do not. In our opinion the BOD5/COD-ratio cannot replace a complete and valid testfor ready biodegradability.

3.2Screening Tests for Inherent Biodegradation

On level 1, i.e., at 100 tonnes/year or already at 10 tonnes/year triggered by cer-tain hazard criteria, substances classified as not readily biodegradable are sub-jected to further investigation of their biodegradation potential in screeningtests for inherent biodegradation.

In these tests the amount of inoculum is increased as compared to the base-set tests, thus leading to better conditions for biodegradation.

The EU-guidelines [12] or OECD-guidelines [11] comprise 2 or 3 test me-thods: Mod. S.C.A.S.-Test [4, 8], Zahn-Wellens-Test [4, 8, 11], and Mod. MITI II-Test [8].

The German Federal Environmental Agency accepts the performance of theMod. MITI II-Test with only one inoculum from municipal STP as opposed tothe requirement of using ten different inocula in the original Japanese version.In both cases a direct parameter for mineralization is measured (O2-consump-tion). Of equal scientific value and therefore also accepted are CO2 Evolution-[11, 12] or Respirometer-Tests [11, 12], in which, however, the substrate/inocu-lum relationship is in inverse proportion to the respective test for ready biode-gradability (C.4-C or C.4-D). This is in accordance with the differences betweenthe Mod. MITI I-Test [4, 8] and Mod. MITI II-Test [8].

In general, both the Zahn-Wellens-Test and the S.C.A.S.-Test do not distin-guish between biological degradation and other elimination mechanisms (mea-sured parameter DOC, open system). In the assessment this is taken into ac-count by the supplement “biodegradable/eliminable.” This differentiation is notfound in the Technical Guidance Document (TGD) of the EU [14]. In both teststhe only assessment provided for is in terms of “degradability.” Also no assess-ment of results from the tests on inherent biodegradability failing the pass le-vel, i.e., 20–70%, is adequately considered in the TGD. Since in this range theoccurrence of stable metabolites cannot be excluded, the term “partial degra-dation” is used here. To derive degradation rate constants for biodegradation(see below) the TGD clearly states that results from the Zahn-Wellens-Test orfrom the Mod. MITI II-Test are only to be considered if biological degradationis clearly identified. To enable such a conclusion, various requirements whichare to be met in the tests are listed below.

The S.C.A.S.-Test is not comparable with other tests for inherent biodegra-dability due to its test design (i.e., discontinuous operation modus, high inocu-lum concentration, nutrients addition, long adaptation phase). A positive resultin the S.C.A.S.-Test can therefore only be regarded as an “indication” for inhe-rent biodegradability.

Biodegradation < 20% Non-biodegradable/eliminableBiodegradation 20–70% Indication of partial biodegradability/ eliminationBiodegradation > 70% Indication of inherent biodegradability/elimination

Even if positive results are achieved, the TGD does not put the S.C.A.S.-Test onan equal level with the other tests for inherent biodegradability and assigns adegradation rate constant of 0.

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For the Zahn-Wellens-Test the following terminology of assessment is used:

Biodegradation < 20% Non-biodegradable/eliminableBiodegradation 20–70% Partially biodegradable/eliminable (with indication

of formation of stable metabolites)Biodegradation > 70% Inherently biodegradable/eliminable

According to TGD the assessment “inherent biodegradable” can only be appliedif the following criteria are fulfilled: the biodegradation pass-level of 70% mustbe reached within seven days, log-phase may be no longer than three days, andelimination (e.g., by adsorption) prior to begin of biodegradation must be lessthan 15%.

If by appropriate test controls abiotic elimination processes, e.g., adsorption,volatilization, precipitation can be excluded, the term “eliminable” may be omit-ted.

For the Mod. MITI II-Test the following terminology of assessment is used:

Biodegradation < 20% Non-biodegradableBiodegradation 20–70% Partially biodegradable (with indication of forma-

tion of stable metabolites)Biodegradation > 70% Inherently biodegradable (with indication of mine-

ralization)

According to TGD, the assessment “inherent biodegradable” can only be appliedif the following criteria are fulfilled: the biodegradation pass level of 70% mustbe reached within 14 days and the log-phase may be no longer than 3 days.

A tabular compilation of the classification of biodegradation potentials of asubstance on the basis of screening tests for ready and inherent biodegradabi-lity is given in Table 3.

3.3Simulation Tests and Persistence Categories

Substances are subjected to simulation tests to verify their degradation poten-tials and to investigate their degradation behavior in specified, exposure rele-vant compartments by means of test designs as close to environment as possi-ble. From the results of such investigations a classification into persistence clas-ses may be derived.

Depending on which environmental compartment degradation and dissipa-tion processes are to be simulated, various test systems are currently available,essentially belonging to three groups:

– Simulation tests for surface waters– Simulation tests for soils– Simulation tests for sewage treatment plants (STP)

The simulation of sewage treatment plants represents a special case, since notan environmental milieu but a technical plant is to be simulated. From this dif-ference a separate assessment concept results for sewage treatment plant simu-lation tests.


Considering physico-chemical characteristics of the substance, exposurescenarios, and the results from screening tests, the appropriate tests are selectedin dialogue with the notifier.

Currently the following standardized simulation tests are available inGermany:

– Degradation and fate of plant protection agents in a water/sediment system.BBA-guideline, part IV, 5–1 [17]

– Fate of plant protection agents in soil – degradation, transformation, andmetabolism – BBA-guideline, part IV, 4–1 [18]

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Table 3. Classification of biodegradation potential

Readily Not readily biodegradablebiodegradable

Indication of Mineralizable Inherently bio- Partially bio- Non bio-persistence degradable and degradable and degradablecategory I indication of indication of(see Table 4) mineralizationa formation of

stable metabolites

Persistence of a substance

Screening tests for ready Screening tests for inherent biodegradabilitybiodegradability

Biodegradation: Biodegradation:≥ 60 % of ThOD ≥ 70 % > 20 % to < 70% ≥ 20 %≥ 60 % of ThCO2≥ 70 % DOC removaland fulfilling and not ful-10 days-window filling 10 days-criterion window criterion

Closed Bottle-Test Modified MITI II-Test [8]Modified MITI I-TestManometric Respirometry-Test BODIS-Test [15]CO2 Evolution-TestModified OECD Screening-Test Zahn-Wellens-Test [4, 8]DOC Die-Away-Test [11, 12] Modified S.C.A.S-Test [4, 8](relation of BOD5/COD > 0.5)

ISO 11734 (Test on anaerobic biodegradability, ECETOC-Test) [16]

Stringency of tests

a Equivalent to inherently biodegradable fulfilling specific criteria according to TechnicalGuidance Document.

For deriving biodegradation rate constants for low concentrations of chemicalsin surface waters without sediment, an ISO guideline is currently in the processof adoption: ISO/CD 14592 Water Quality – Evaluation of the aerobic biodegrad-ability of organic compounds at low concentrations, Part 1: shake flask batchtest with surface water or surface water/sediment suspensions [19].

3.3.1Simulation Tests for Surface Waters and Soils

Simulation tests for surface waters (water/sediment systems) and soils are as-sessed according to identical criteria. In addition to the quantitative base para-meters primary degradation (dt50, disappearance time of 50% of the substance),mineralization, and bound residues, further qualitative parameters, especiallythe degradation kinetics and the metabolism scheme, are included in theassessment and formation of persistence classes.

A proposal for a comprehensive assessment concept is presented in Table 4.From the use of different sediments, soils, or temperature in different tests,

different ranges of results may arise. This has to be kept in mind. By inclusionof these parameters the persistence class may change.

In accordance with the assessment practice of the TGD the results of degra-dation tests are taken for the calculation of the predicted environmental con-centration (PEC).

An example of the application of this assessment concept is presented in thefollowing.

In context with the registration of plant protection products, fate studies onthe degradation and distribution of active substances in soil and water/sedi-ment simulation test systems were evaluated and assessed based on the para-meters primary degradation, mineralization, and bound residues.

The studies were conducted based on the above-mentioned guidelines [17,18].

The evaluation is based on 294 comparable data sets for soil systems and 253data sets for water/sediment systems, respectively, and have been classified ac-cording to the assessment scheme outlined above (Table 4).

As can be seen in Fig. 2, primary degradation rates in soils and water/sedi-ment systems reveal that the disappearance from the water phase is much fasterthan in soils (class I, rapid primary degradation, i.e., dt50< 10 days).

Whereas in water/sediment systems the rapid disappearance from the waterphase is mostly due to transfer and adsorption of the parent compound to se-diment, the elimination of the parent compound in soil systems is caused byprimary degradation, i.e., the transformation of the molecule. Primary degra-dation is a slower process than physico-chemical reactions, leading to the ob-served rapid disappearance from the water phase.

As shown in Fig. 3, mineralization in water/sediment systems is less effectivethan in soil systems.

A comparison of especially class I in Figs. 2 and 3 shows that the mineraliza-tion in water/sediment systems is remarkably lower than the primary degrada-tion (dissipation) from the water phase.


As can be seen in Fig. 4, higher amounts of bound residues have been foundin soil systems (classes III and IV) as compared to sediment systems (classes Iand II).

In spite of the differences found in primary degradation, mineralization, andthe amount of bound residues between soil and water/sediment systems, theoverall assessments reveal comparable persistence categories, i.e., comparable

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Table 4. Persistence classes and persistence categories

1st criterion: primary degradationdt50 Class Assessment

< 10 days I Rapid primary degradation10–30 days II Delayed primary degradation30–100 days III Slow primary degradation> 100 days IV Negligible primary degradation

2nd criterion: mineralization (after 100 d)CO2 Class Assessment

> 50% I Extensive mineralization25–50% II Moderate mineralization10–25% III Limited mineralization< 10% IV Negligible mineralization

3rd criterion: bound residues (after 100 d)Amount Class Assessment

< 10% I Low plateau10–25% II Moderate plateau25–50% III High plateau> 50% IV Very high plateau

Calculation of Persistence CategoryThe three criteria mentioned above, respectively the resulting classes are equally taken forcalculation of the overall persistence category (average by rounding): sum of single classes:number of parameters = persistence category

I Low persistenceII Moderate persistenceIII High persistenceIV Not biodegradable

On a case-by-case basis the degradation curve as well as the metabolism scheme is consid-ered for obtaining the overall persistence category

Example:A substance shows the following properties:dt50 3 days I Rapid primary degradationCO2 12% III Limited mineralizationBound residues 60% IV Very high plateau

8:3 = 2.7 (rounded: 3)Consequently the substance has to be considered as highly persistent (persistence categoryIII). This example clearly demonstrates that a classification on the basis of primary degrada-tion alone (class I) would have resulted in a wrong assessment of the real persistence


Fig. 2. Primary degradation (class I–IV; see Table 4) of plant protection substances in water/sediment (water phase only) and soil systems

Fig. 3. Mineralization (class I–IV; see Table 4) of plant protection substances in water/sedi-ment and soil systems

biodegradation/elimination behavior in both systems as presented in Figs. 5and 6.

Conclusions and consequences from the results of the test systems used are:– Primary degradation and mineralization must be assessed separately.– Primary degradation/disappearance in water in the presence of sediment is

faster than in soil.

– Mineralization in soil is more effective than in sediment.– Amount of bound residues in soils is higher than in sediments; this is rele-

vant because bound residues may be bioavailable for soil/sediment orga-nisms.

– The overall degradation/disappearance rates in soils and water/sediment sy-stems (persistence categories) are almost equal.

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Fig. 4. Bound Residues (class I–IV; see Table 4) of plant protection substances in water/sedi-ment and soil systems

Fig. 5. Overall assessment of persistence categories (I–IV; see Table 4) of plant protectionsubstances in water/sediment systems

3.3.2Simulation Tests for Sewage Treatment Plants

Simulation tests for sewage treatment plants must permit an assessment of thedegradation and dissipation behavior of a substance in an STP. An approachorientated solely at measuring the elimination capacity is insufficient as, e.g.,the transfer of a problematic compound from the aqueous compartment to thesoil compartment via sludge or into air by stripping (volatilization) is not con-sidered.

For an assessment information on the following should be available:

– Primary degradation and formation of metabolites– Mineralization– Adsorption onto sewage sludge– Volatilization

An inclusion of these requirements into the internationally harmonized testprotocols is still lacking.

Representing a simulation test on a laboratory scale, in chemicals legislationonly the Coupled Units-Test [8] is currently available. Due to its test design(open system) and its analytics (DOC) it is applicable only to sufficiently water-soluble, non-adsorbing, and non-volatile substances. The test does not enable adifferentiation between biodegradation and abiotic elimination mechanisms,like adsorption and volatilization, and hence cannot be considered to be a truesimulation test. Since alternative test methods are lacking at present, the resultsfrom a Coupled Units-Test are taken into consideration on a case-by-case basisfor a quantitative estimation of the elimination capacity of a mechanical-bio-logical sewage treatment plant.

The OECD 303A is currently under revision and will comprise two parts, in-cluding far-reaching improvements.


Fig. 6. Overall assessment of persistence categories (I–IV; see Table 4) of plant protectionsubstances in soil systems

Aspects such as the percentage connection to STP, variable loading rates,operational disturbances due to intoxication or fluctuating substance concen-trations (e.g., campaign operation, other temporal fluctuations), as well as dis-charges resulting from rainwater overflows must be considered in a final ex-posure analysis. The qualitative assessment of the removal efficiency inGermany is based on the legal requirements of the Federal Water Act (WHG)[20], Appendix 22, in compliance with generally accepted rules of the state ofthe art.

The elimination capacities measured in the Coupled Units-Test are not suffi-cient as a basis for a quantitative exposure estimation. Kinetic rate constantscould be derived if the appropriate parameters like sludge age and hydraulic re-tention time are taken into consideration as provided by the updated draft ofthe OECD 303A.

For a tentative estimation of the elimination capacity of STP, the eliminationclassification from results of STP simulation tests given in Table 5 may be used.

4Biodegradation Rate Constants

Biodegradation rate constants, i.e., the biodegradation within defined time in-tervals, are essential for the estimation of the fate of a chemical in the differentenvironmental compartments. For this estimation first order biodegradationkinetics are assumed. Due to mostly low concentrations of chemicals and rela-tively low microbial density, biodegradation processes in the environment fre-quently follow such kinetics.

4.1Determination of Biodegradation Rate Constants from Screening Tests

The determination of biodegradation rate constants from screening tests isgenerally not possible.

For exposure estimation, based on results from biodegradation tests onready and inherent biodegradability, kinetic rate constants for the biodegrada-tion of a substance in various environmental compartments (STP, surface wa-ters, sediments, soils) have nevertheless been derived as default values, present-ed in Tables 6 and 7.

These parameters were agreed upon internationally within the context of ad-opting the TGD and are used for exposure estimation as long as no substancespecific data are available from higher quality degradation tests (simulation

308 B. Beek et al.

Table 5. Classification of elimination performance of sewage treatment plants

Degree of elimination Class Assessment

> 95% I Substantial elimination75–95% II Medium elimination< 75% III Poor elimination

tests), which may substitute the tentative rate constants derived from screeningdegradation tests.

4.2Prerequisites for the Derivation of Biodegradation Rate Constants from Simulation Tests

To derive biodegradation rate constants from results of simulation tests, at leastthe following prerequisites should be fulfilled:

– The elimination portion resulting from adsorption in an aquatic test systemhas to be determined. If no direct experimental data are available this por-tion is estimated by an additional adsorption control in a test for inherentbiodegradability, e.g., Zahn-Wellens-Test [4, 8, 11], by an experimentally de-termined Koc [21], or by a calculated Koc (from the Kow or from water solub-ility).

– The abiotic part of elimination either by adsorption, precipitation, or volatil-ization is subtracted from the total elimination. This gives the amount of de-gradation, from which the biodegradation rate constant will be calculated.

As a general rule, the test conditions should simulate real conditions as close aspossible.


Table 6. First order degradation rate constants in sewage treatment plants derived from re-sults of screening tests for ready or inherent biodegradability

Tests for Ready Biodegradability (28 days)Tests 92/69/EEC C.4 A – F respectively OECD 301 A – F or tests which

are considered scientifically equal (“expert judgement”)Pass-level 60/70% with 60/70% without < 60/70%

10 d-window 10 d-windowAssessment Readily bio- Readily biodegrad- Not readily

degradable able, but failing biodegradable10 d-window

Degradation rate in ka1st order = 1 h–1 k1st order = 0.3 h–1 k1st order = 0 h–1

models for sewage treatment plants

Tests for Inherent Biodegradability (28 days)Tests 88/302/EEC respectively OECD 302 B – C or tests which are con-

sidered scientifically equal (“expert judgement”)Pass-level 70% ≥ 20 to < 70% < 20%Assessment Inherently bio- Partially biodegrad- Not bio-

degradable, fulfilling able/eliminable degradablespecific criteria

Degradation rate in k1st order = 0.1 h–1 k1st order = 0 h–1 k1st order = 0 h–1

models for sewage treatment plants

a Equivalent to dt50 = 0.7 h.

In water/sediment studies and/or simulation tests for biodegradation in soil,a fractionation into extractable and non-extractable residues (bound residues)has to be conducted in order to differentiate between easily bioavailable andless available fractions. Hence the extraction method applied has to be consid-ered. For the characterization of bound residues soft extraction methods shouldbe used in order to simulate environmental conditions in sediments and soils.

4.3Field Studies

Field studies should give information on the biological degradation and dissi-pation processes on an ecosystematic level thus allowing for a complex assess-ment of the environmental behavior of a substance; since in field studies someprocesses like volatilization, leaching, and metabolism can hardly be investigat-ed in an appropriate manner, a combination of both laboratory and field studiesmay lead to a comprehensive assessment of fate and behavior of a chemical inthe environment. Such studies are to be designed and performed in close co-operation with the assessing authority. Due to the high complexity of a fieldecosystem and the resulting methodological problems in evaluating and inter-

310 B. Beek et al.

Table 7. First order degradation rate constants for different environmental compartments de-rived from results of screening tests for ready or inherent biodegradability (according toTGD)

Compartment Biodegradation 1st Order Potential Degradation Rate

Constant

Fresh water Water Readily bio- k1st order = 0.047 day–1

(river, lake) Phase degradableReadily biode- k1st order = 0.014 day–1

gradable, but failing 10 d-windowInherently bio- k1st order = 0.0047 day–1

degradable, fulfilling specific criteriaInherently bio- k1st order = 0.00047 day–1

degradable, not ful-filling specific criteria

Sediment Water All biodegradation Same values as forPhase Phase potentials water phase

Solid Compared to water k1st order = 0 day–1

Phase phase biodegradation (default)potential is lower

Sea and brackish Compared to water k1st order = 0 day–1

water phase biodegradation (default)potential is lower

preting results from field studies, currently no standardized test system exists,on the one hand fulfilling the requirements of legislation and on the other handthose of the above-mentioned scientific minimum requirements.

5Microbial Inhibition

Testing for toxic or inhibiting effects of a substance to micro-organisms is con-ducted together with biodegradation testing under the following aspects:

– For interpreting test results from biodegradation tests it is essential to haveknowledge of the impact of potentially toxic or inhibiting effects of the sub-stance of concern to the degrading micro-organisms.

– By estimating the risk potential for sewage treatment plants the risk of atechnical malfunction/breakdown due to intoxication can be identified.

– Prospective disturbances of biogenic cycles in the primarily exposed envi-ronmental compartments are to be detected.

A microbial inhibition below 10% or 20% with respect to the control depend-ing on the test system used is considered as not significant due to methodolo-gical variability. For the calculation of a PNEC (predicted no effect concentra-tion) such results (EC10/EC20) are treated as NOEC (no observed effect concen-tration) values.

For a risk assessment concerning microbial toxicity it is necessary to com-pare exposure concentrations (PEC, predicted environmental concentration)with effect concentrations (PNEC) as is usually applied in ecotoxicology.

The PNECmicro-organisms is generally derived from a NOEC supplemented by an appropriate safety/uncertainty factor depending on the test system used(endpoint tested, sensitivity) and environmental compartment under consider-ation (water, soil, STP). Safety factors are applied to minimize the risk of dam-age to more sensitive micro-organisms as used in the respective test systems.

In Tables 8 and 9 standardized test methods on inhibition of microbial ac-tivity are compiled, including the most important parameters of test perfor-mance.

Additionally, a test guideline has recently become available from ISO [22]based upon the inhibition of growth of sewage sludge bacteria.

5.1Assessment Criteria

When assessing the results from microbial inhibition tests in the aquatic milieua qualitative and quantitative assessment of results is performed. In additionthe test results EC50 (inhibition concentration of 50%) or NOEC are given,stating the test method applied, the measured endpoint, and the test duration.

In soils the inhibition of microbial activity of soil micro-organisms can bedetermined according to the guideline IV 1–1 “Auswirkung auf die Aktivität derBodenmikroflora/Side-effects on soil microflora” [32] of the German FederalResearch Center for Agriculture and Forestry.


The endpoints respiration rates, dehydrogenase-activity, and nitrogen turn-over are measured within the registration procedure in fulfillment of theGerman Plant Protection Act.

Accordingly, the test duration is generally 28 days and can – provided effects>15% are encountered – be prolonged up to 56 days or 100 days. The respectiveeffects, i.e., both increase and decrease of activities, are expressed as percent de-viation from an untreated control and assessed according to a model describedby Malkomes [33] as shown in Fig. 7. The results from these tests, carried outwith fivefold and tenfold maximum rate of application in order to simulate theaccumulation of the substance in deeper soil layers, represent a realistic worst-

312 B. Beek et al.

Table 8. Inhibition of microbial activity – comparison of test methods – part 1

Method Inhibition of Inhibition con- Activatet sludge Inhibition ofoxygen control of the respiration dehydrogenase-sumption with closed bottle inhibition test activityPseudomonas test a

putida

References DIN 38 412, OECD 301 D EU 88/302/EEC [26]part 27 [23] [8, 11] L 133 [4]

EU C.4 E [12] OECD 209 [24]ISO 8192 [25]

InoculumOriginBacilliculture P. putidaMixed culture Effluent of sewage Activated sludge Activated sludge

treatment plant or surface water

Density £ 5 ml/l 0.8–1.6 g dw/l 0.4–2.4 g/lApprox. cells/l No information 104–106 109–1010 No information

Temperature 21±1 °C 22±2 °C 20±2 °C 21 °CpH-Value 7.5±0.5 7.4±0.2 7.25–8.0 No information

(optimum)Test duration 30 min 14 d 30 min and/ 15 or 30 min

or 3 hMeasured Inhibition of Inhibition of O2- Inhibition of O2- Inhibition ofparameters O2-consumption consumption of consumption reduction of redox

a readily degrad- dye-stuffs by able reference microbial dehy-substance drogenase

Evaluation Determination Graphical com- Graphical dose- Graphical dose-of dilution grade parison of BOD: effect relation, effect relationship;causing an inhi- reference sub- ship; EC20 , EC50 EC50bition of < 20% stance vs.

reference sub-stance + test substance

a This applies to all tests for ready biodegradation. In all these tests the performance of atoxicity control is optional.

case. Such an accumulation can be observed with strongly-adsorbing sub-stances as well as under extremely dry weather conditions.

In the context of implementing the Council Directive 91/414/EEC of 15th of July 1991 [34] concerning the placing of plant protection products on the mar-ket, the following classification based on the same parameters is currently used:


Table 9. Inhibition of microbial activity – comparison of test methods – part 2

Method Inhibition of Growth inhibi- Inhibition of Side-effect on the nitrification tion test with light emission activity of soil

Pseudomonas microfloraputida

a) Dehydrogenase activityb) Short time respirationc) Metabolic active biomassd) Nitrogen turn-over

References ISO 9509 [27] DIN EN ISO DIN EN ISO BBA Guideline,10712 [28] 11348 Part 1 to 3 Part VI, 1–1, [32]

[29, 30, 31]InoculumOriginBacilliculture P. putida Vibrio fisheriMixed culture Activated Two agricultural

sludge soils with micro-flora of differing activity

Density 1.5 g/lApprox. cells/l No information 109 2 ¥ 109 No informationTemperature 20–25°C 21 ± 1°C 15 ± 2°C 20 ± 2°CpH-Value 7.6 7.4 7.0 ± 0.2 soil 1 :5.5–7.0

soil 2 :6.0–7.5

Test duration 4 h 16 ± 1 h 15 and 30 min Different; depend-ing on parameter to be tested

Measured Inhibition of Inhibition of Decline of a) Reduction ofparameters nitrification cell multipli- luminescence redox dye-stuffs

(oxidation of cation intensity by microbial ammonium) (turbidity) dehydrogenase

b) O2-consumption c) CO2-evolutiond) Determination ofNH4

+, NO3–, (NO2

–),N-total

Evaluation Graphical dose- Graphical dose- Graphical dose- Graphical dose-effect relation- effect relation- effect relation- effect relationship;ship; EC50 ship; EC10; EC50 EC20; EC50 EC50

Category Toxicity Deviation of activity After x daysfrom control (in %)

I very weak < 2.5 28II weak < 25 28III moderate < 25 56IV high < 25 100V very high > 25 100

The first category with a deviation of < 2.5 % from control, however, is in viewof methodological aspects, not applicable because effects will only be signifi-cant above 10–15%. Therefore we propose to omit this category completely.

5.2Intrinsic Properties of Chemicals and Consequences for Choice and Performance of Tests

The issue of which test design is suited to clarify the microbial toxicity of a testsubstance depends on the selection of an appropriate endpoint, the respective

314 B. Beek et al.

Fig. 7. The decrease of microbial toxicity with time according to Malkomes [33]: 1 negligible,2 tolerable, 3 critical, 4 non-tolerable toxicity

environmental compartment, and the intrinsic properties of the substance un-der concern.

As shown in Tables 8 and 9 (see above), the summarized tests exhibit differ-ent toxicological endpoints, sensitivities, and test durations. Generally, short-term measurements in terms of hours (e.g. 10 h) are preferred, in accordancewith the retention time in an STP.

For soils only one test system is available in Germany (BBA, 1990) [17]. Twoguidelines for microbial toxicity in soils are currently under development by theOECD (based upon the inhibition of transformation of carbon and nitrogen insoils). The other test systems refer to aquatic compartments and STP.

Concerning the intrinsic properties of substances, poor water solubility andadsorbance are especially problematic.

For such “difficult substances” guidance documents are currently under de-velopment by ISO, OECD, and EU. From a pragmatic point of view, substancesmay be defined as being poorly water soluble if their solubility is below100 mg/l. The use of solubilizers for increasing the solubility of poorly watersoluble substances should be avoided because the results may be misleading.Instead, stirring a nominal of, e.g., 100 mg/l for 24 h ensuring the maximum sol-ubility and subsequent testing is recommended as a limit test.

In such a test performance the real substance concentrations are often belowdetection limits, i.e., unknown. For this the term Water AccommodatedFraction (WAF) may be used as has originally been proposed for mixtures ofpoorly water soluble substances [35].

If testing microbial toxicity in serial dilutions of a stock solution of a poorlywater soluble substance, as is usually done with well soluble substances, mis-leading results will be obtained as exemplified in the following.

Given a poorly water soluble substance with a solubility of 1 mg/l which,however, was not analytically determined, a stock solution with a nominal con-centration of 100 mg/l is prepared exerting a slight but significant effect. Hencethe LOEC (lowest observed effect concentration) would be noted as 100 mg/lnominal. Given further that a 1 :1 dilution would stop the effect, then the NOEC(no observed effect concentration) would be noted as 50 mg/l.

However, preparation of a stock solution of a nominal 50 mg/l would alsocontain 1 mg/l dissolved substance exerting the same effect as 25 mg/l and soon, until 1 mg/l is dissolved and further diluted 1:1, leading now to the trueNOEC of 0.5 mg/l instead of 50 mg/l as above.

Testing should consequently be performed with the supernatant of WAFswithout filtration or centrifugation procedures.

For the OECD 209 “Activated sludge respiration inhibition test” [24] the testduration of 3 h is recommended for poorly water soluble substances to en-counter possible delayed effects. This is true also for the other test systems lis-ted in Tables 8 and 9. The rationale behind this and an actual example is givenin the following.

A poorly water soluble substance (identity confidential) was tested in anOECD 209 test within a notification procedure.After 30 min an EC50> 1000 mg/lwas measured, whereas after 3 h an EC50 of 400 mg/l and a NOEC of 100 mg/lwas obtained.


If after 3 h testing no effects are found, no further testing is required and thesubstance is classified as being not inhibiting the microbial activity up to, e.g.,100 mg/l nominal.

For STP and surface waters, no risk is assumed from microbial toxicity, andno PNECs are derived.

If significant effects occur, WAFs of single concentrations within a concen-tration range should be prepared to establish a dose-response relationship.

With water soluble substances, i.e., water solubility >100 mg/l, testing ofmicrobial toxicity may start with a limit test as well, preferably with 1000 mg/l,and then in case of significant effects (> 10% or 20% depending on the test sys-tem) regular testing with serial dilutions has to be performed as describedabove to establish a dose-response relationship to derive ECx-values and theNOEC for the calculation of the PNECmicro-organisms .

If the exposure concentration exceeds the test concentration of, e.g.,100 mg/l, a test with higher concentration either as a limit test or with serial di-lutions has to be performed.

A further problematic property is the adsorption tendency of a substance,since in this case an adsorption onto suspended solids, soils, sediments, vesselwalls, and onto the inoculum must be expected. This may lead to a reduction ofthe true test concentration in a test system. We assume an essential adsorptionif the substance has a log KOW> 3 (octanol/water partition coefficient) or if anadsorption potential can be proven in an adsorption/desorption screening test(OECD 106, ISO draft).

Moreover, delayed toxicity may occur due to depot effects. Quarternary am-monia compounds for example are known to cause such effects.

Apart from regular tests on microbial toxicity mentioned above, inhibi-tion controls from tests on ready biodegradability may be used for a preli-minary screening for toxic effects. In such assays the influence of a test sub-stance on the biodegradation of a well-degradable reference substance is deter-mined. However, results are only obtained for one single concentration (seeTable 8).

5.3Risk Assessment and Safety Factors

For risk assessment of a sewage treatment plant a PECSTP/PNECmicro-organisms ratiois calculated. As already mentioned, for the calculation of a PNECmicro-organismsthe NOEC is taken as a basic value supplemented with an appropriate safety fac-tor according to the TGD [14] listed in Table 10.

The rationale behind the different safety factors is given by differences insensitivities, end points, and test durations of the respective test system. Thus,test systems with higher safety factors are the less sensitive ones.

For the calculation of the predicted environmental concentration for sewagetreatment plants (PECSTP) the influent concentration (cinf), i.e., in the sewershould be taken rather than, as suggested by the TGD, the effluent concentration(ceff ), because toxicity already occurs in the sewer, possibly giving rise to inhi-bition effects of microbial activity in the sewage sludge.

316 B. Beek et al.

Given a PECSTP/PNECmicro-organisms of <1, no indication of microbial toxicity inSTP is assumed and no further testing is required.

In case of a PECSTP/PNECmicro-organisms of > 1, an indication of microbial tox-icity for STP is given and further testing of microbial toxicity is required usingdifferent toxicological endpoints, preferably with regard to the intrinsic pro-perty of the test substance.

The same procedure is applied for the compartments soil and surface water.Protozoa (ciliates) are an integral and important part of a functioning bio-

cenosis of an STP, mainly due to the elimination of germs and improvement ofcarbon and nitrogen metabolism in an STP (see contribution by Pauli et al., thisvolume). Besides microbial toxicity the role of protozoa in STP has recently beconsidered in a Technical Recommendation of the European Chemicals Bureau[36] as follows:

All valid ciliate growth impairment data should be taken into account for thederivation of a PNECSTP. Protozoa have to be regarded as additional species, notas an additional trophic layer. The PNECSTP should be derived on the basis ofthe most sensitive species regardless of whether this is from a test with activat-ed sludge, relevant bacteria, or ciliated protozoa.

6Deficits and Perspectives

– The modified MITI II-Test (guideline OECD 302 C) [8] should be included inthe test repertoire of the EEC as a test on inherent biodegradability, since thetrue biodegradation is measured separate from adsorption and/or volatiliza-tion. As to the practical performance, the use of one inoculum from a muni-cipal STP instead of ten as in the original Japanese prescription may be ac-cepted for reasons of simplification.

– The Two Phase Closed Bottle-Test (BODIS-Test, ISO 10708) [15] for poorlywater-soluble substances should be included in the inventory of tests for ulti-mate biodegradability. However, the test conditions regarding the agitationprocedure should be standardized.


Table 10. Safety factors in accordance with the TGD

Test Safety factor applied Safety factor appliedto NOEC or EC10/EC20 to EC50

Growth Inhibition Test with 1 10Pseudomonas putida

Inhibition of nitrification 1 10Inhibition of luminescent bacteria 1 10Other tests with single species 1 10Activated Sludge Respiration Inhibition 10 100

Test (OECD 209)Other tests with mixed inoculum 1 or 10 (depending 10 or 100

on sensitivity of test)

– There is a need for standardized test guidelines and assessment schemes forscreening and simulation tests on the anaerobic biodegradation of sub-stances in hydrosphere, soil, and sludge for risk assessment.

– A test system has been developed sponsored by the German FederalEnvironmental Agency for testing fate and behavior of chemicals in surfacewaters (rivers, lakes) including sediment. A draft has recently been submit-ted to ISO.

– There is a need for tests simulating biodegradation and microbial toxicity inestuaries, coastal areas, and the open sea.

– Besides the existing tests on nitrification in water and soil there is a need forfurther development of denitrification test systems.

– At present, standardized tests covering the inhibition of microbial activityunder anaerobic conditions are missing (e.g., inhibition of methane produc-tion). This is also relevant for inhibition controls in anaerobic biodegrada-tion tests mentioned above.

– For testing difficult substances (volatile, poorly water soluble, adsorbing)there is still no appropriate guidance available. However, ISO, OECD, and EUhave respective guidance documents currently in progress.

– Likewise, still no guidance exists for situations where conflicting results onbiodegradability have been obtained from tests of the same level, e.g., baseset screening tests on ready biodegradability as experienced within the noti-fication of New Chemicals according to the German Chemicals Act. Somemember states of the EU have suggested the best test results be taken if va-lidity of the performance had been ensured, while others rely on expertjudgement (“weight of evidence”). In cases where a sufficient number ofindependent and valid test results exists, the calculation of a median value(geometric mean value) is proposed for discussion.

Biodegradation and persistence have become of increasing importance for en-vironmental risk assessment of chemicals during the last decade. This is also re-flected by ongoing international activities of the United Nations EnvironmentalProgram (UNEP) concerning Persistent Organic Pollutants (POPs).

Thus, the above-mentioned deficiencies urgently need to be removed.

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