Journal of General Microbiology (1987), 133, 267-274.
Printed in Great Britain
267
Degradation of Eighteen 1-Monohaloalkanes by Arthrobacter sp.
Strain HA1
By R U D O L F S C H O L T Z , A N N E M A R I E S C H M U C K L E ,
ALASDAIR M . COOK* A N D T H O M A S L E I S I N G E R
Department of Microbiology, Swiss Federal Institute of Technology, ETH-Zentrum,
CH-8092 Zurich, Switzerland
(Received 28 April 1986; revised 12 August 1986)
~~~
~
Four coryneform bacteria were isolated from enrichments with 1-chlorobutane, 1-chloropentane or 1-chlorohexane as sole source of carbon and energy. One organism, strain HAl, was
identified as an Arthrobacter sp. It could utilize at least 18 I-chloro-, 1-bromo-and 1-iodoalkanes,
but not the 1-fluoroalkane tested. Substrate utilization was quantitative and growth yields of
about 5.5 g protein (mol C)-l were observed for hexanol and 1-chlorohexane with specific
growth rates of 0.19 and 0.14 h-l, respectively. The organohalogen atom was released
quantitatively as the halide ion. Cells grown on 1-chlorobutane contained an inducible
enzyme(s) that dehalogenated haloalkanes. The enzyme was soluble, required no cofactors,
membrane components or oxygen for activity, and released halide and the corresponding
n-alcohol from the 1-haloalkane. The halidohydrolase(s) in crude extract had a specific activity
of about 0-7 mkat (kg protein)-' and dehalogenated a much wider spectrum of compounds (at
least 29, including mid-chain and a,o-dichlorosubstituted alkanes) than supported growth.
INTRODUCTION
Chlorinated alkanes were regarded as non-biodegradable until Omori & Alexander (1978a)
and Brunner et al. (1 980) demonstrated aerobic utilization of 1,9-dichlorononane and
dichloromethane by pure cultures, and Bouwer & McCarty (1983a,b) and Vogel & McCarty
(1985) confirmed anaerobic transformations of many haloalkanes. In contrast to work with
chloroalkanoates, in which isolation of organisms seems to be simple (Motosugi & Soda, 1983),
enrichments to degrade haloalkanes have succeeded only with dichloromethane, 1,2-dichloroethane (Stucki et al., 1983; Jannsen et al., 1985), vinyl chloride (Hartmans et al., 1985), 1,9dichlorononane, and, more recently, chloromethane (Hartmans et al., 1986) and 1-chlorobutane
(Yokota et af., 1986); these isolates usually have a very narrow substrate specificity. However,
some of these aerobic dechlorinations have relatively high specific activities in growing cells
[ 2 3 mkat (kg protein)-'], and the application of biodegradation to destroy waste dichloromethane has been demonstrated (Galli & Leisinger, 1985).
The dechlorination of chloroalkanes has been suggested to be NADH-linked (Omori &
Alexander, 1978b), oxidative (Hartmans et al., 1985, 1986; Yokota et al., 1986) or hydrolytic
(Kohler-Staub & Leisinger, 1985; Janssen et al., 1985;Yokota et al., 1986). However, all models
agree that dehalogenation is the first catabolic reaction. In the case of 1,2-dichloroethane,
hydrolytic removal of the first chlorine atom is apparently followed by oxidation of the organic
product to chloroethanoate from which the chlorine atom is removed by a haloalkanoate
halidohydrolase (Janssen et al., 1985). Metabolism of haloalkanes independent of the halogen
atom is also known (Murphy & Perry, 1983, 1984).
Monochloroalkanes have received little attention until now (Hartmans et al., 1986; Yokota et
al., 1986). In biodegradation studies, these sulvents and alkylating agents have the advantage of
chemical and analytical simplicity. Further, some bromo- and iodoanalogues occur naturally
0001-3423 0 1987 SGM
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268
R . SCHOLTZ A N D OTHERS
(Gschwend et al., 1985) and we regard them as models from which biodegradation of more
highly halogenated alkanes can be approached. This paper deals with the enrichment of
biodegradative organisms, the mass balance of the degradation and the mechanism of the
dehalogenation.
METHODS
Materials. The inoculum for enrichments was sludge J l % dry wt) from the settling tank of an industrial
biological treatment plant which has treated chlorinated solvents for many years. The haloalkanes (99% purity)
were purchased from Fluka, except for chloroethane (99%; Matheson) and 1-fluoropentane (Aldrich); purity
(299%) was confirmed by GLC and the identities of 1-chlorohexane, 1-bromobutane and 1-iodoethane were
confirmed by MS. Ethylene oxide, o-chlorinated alkanoic acids and other chemicals were purchased from Fluka
and were of the highest purity available. Liquid cultures were grown in screw-cap Erlenmeyer flasks closed with
Mininert valves (Hartmans et al., 1986).
Apparatus and analyses. GLC with flame ionization detection (Jutzi et al., 1982), GLC-MS (Cook et al., 1984),
spectrophotometric analyses, ODsj6 measurements and anaerobic experiments were done with apparatus
described previously (Thurnheer et al., 1986; Cook et al., 1984). Haloalkanes and alcohols were assayed by GLC
after separation on a 1.8 m x 2 mm stainless steel column packed with Porapak P (cf. Stucki et al., 1981); for
GLC-MS, a 10 m x 530 pm methylsilicone capillary column was used. Chloride ions were converted to 2chloroethanol (Russel, 1970) which was then assayed by GLC on a 1.8 m x 2 mm Porapak Q column (at 170 "C)
with a nitrogen flow rate of 30 ml min-I. Chloride ions were also assayed with an ion-specific electrode (Cook &
Hiitter, 1984). Halide release was determined by the method of Bergmann & Sanik (1957): the standard curve
always contained blank samples appropriate to the experiment because the colour development was inhibited in
the presence of some ions. Growth was quantified as protein content in a modification of the method of Kennedy
& Fewson (1968). Portions (5 ml) of cultures were brought to 0.5 M with respect to TCA and frozen. Each thawed
sample was centrifuged (27000g for 30 min) and the pellet suspended in 0.75 ml 1.32 M-NaOH before incubation
at 80 "C for 20 min and then at 30 "C for 24 h. The sample was then diluted to 1.5 ml, centrifuged to remove
turbidity (which did not contain protein), and the supernatant fluid was assayed. The Gram-reaction was
deteimined directly by staining, indirectly by the Bactident L-aminopeptidase test (Merck), and was deduced from
the surface characteristics of cells observed in the electron microscope (Hartmans et al., 1986).
Growth medium and the isolation of organisms. The growth medium was a mineral salts solution (Hartmans et al.,
1986), containing trace elements (Cook & Hutter, 1981) and vitamins (Schlegel, 1981), to which a carbon source
was added. The haloalkanes were found to be sterile and so were injected directly into the culture through the
septum. Cultures (30 and 100 ml) were grown at 30 "C on an orbital shaker (180 r.p.m.); the volume of air above
the culture was ten times the culture volume. If a compound was described as light-sensitive, the culture-flask was
wrapped in aluminium foil.
The inoculum for the enrichments was added directly to non-sterile medium (30 ml, without vitamins) in closed
vessels to which 90 pmol of a chlorinated alkane was added. Cultures were evaluated for growth (OD546above the
negative control) and substrate utilization after a week, and cultures were transferred to fresh medium as
appropriate. All subcultures, at some stage after the initial enrichment, needed a vitamin supplement. After three
subcultures, positive enrichments were streaked on nutrient agar plates, and a representative of each colony type
was inoculated into appropriate sterile selective medium. When an isolate that degraded the chlorinated alkane
gave rise to a single colony type on three successive platings, it was considered pure, and was maintained on a slant
of selective medium solidified with 1.5% (w/v) agar. Limited taxonomy was done (Krieg, 1984) to ascertain that
the organisms were unlikely to be pathogens and tentative identification followed Seiler's (1983) key.
We also examined Pseudomonas sp. strain P4, a 1,2-dichloroethane-utilizingorganism kindly supplied by
Professor W. Frommer (Bayer, Wuppertal, FRG). The medium for this strain always contained 0.05% yeast
extract.
Quantification of growth and substrate utilization. Bacterial growth yields with limiting carbon sources were
determined from 30 ml cultures in shaken 300 ml flasks (Hartmans et al., 1986). Samples for the determination of
halide ion concentration were centrifuged (20000 g ; 20 min; 4 "C) and the supernatant fluid was stored frozen in a
screw-cap vial; data were expressed as a percentage of the theoretical value analogous to the calculation of growth
yield.
Growth kinetics were followed in 100ml cultures in 11 flasks. Samples were taken at intervals for the
determination of OD546and chloride ion concentration. The ODsd6was converted to a protein concentration by
using standard curves for the appropriate cell type: the values varied by about 10% amongst different carbon
sources.
Preparation of cell extracts and enzyme assay. Cultures (1 %, v/v, inoculum) were grown in 100 ml portions, to
which was added 1 mmol 1-chlorobutane. Cells were harvested (20000g; 30 min; 4 "C) in the stationary phase
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Biodegradation of 1-monohaloalkanes
269
after 2.5 d, washed twice in cold 10 m~-Tris/H,S0, buffer, pH 7.5, resuspended to about 250 mg wet wt ml-1 in
cold extraction buffer (10 m~-Tris/H,S0, buffer, pH 7.5, containing 1 mM-EDTA and 2 mM-DTT), and
disrupted by two passages through an ice-cold French pressure cell at about 130 MPa. Cell debris and intact cells
were removed by centrifugation (30000 g; 20 min; 4 "C) and the clear supernatant fluid was used for experiments.
Enzyme activity was measured in 1 ml reaction mixtures (initial volume) at 30 "C in 4 ml screw-cap tubes
capped with Mininert valves. The mixture contained Tris/H,SO,, pH 7.5, and 1-chlorobutane ( 5 pmol), and the
reaction was started by the addition of about 1.5 mg protein through the valve. The reaction rate was measured by
halide release: duplicate 40 p1 samples were withdrawn at intervals with a syringe and the reaction was stopped by
addition to the acidic reagent of the Bergmann & Sanik (1957) test, after which the colour reaction was developed.
was determined. Enzyme activity under
Protein was removed by centrifugation (12000g; 5 min) and the
anaerobic conditions was examined after preparation of the reaction mixture in a glove box: there was too little
oxygen in the gas phase (3 ml, 21 p.p.m. 0 2 i.e.
, 50.1 nmol) to allow significant oxygenation of the substrate
(5 pmol).
RESULTS
Enrichment cultures
Enrichments were prepared with 1-chloroethane, 1-chloropropane, 1-chlorobutane, 1chloropentane and 1-chlorohexane, but only with the latter three substances were growth and
substrate disappearance observed. Four strains, all requiring vitamins, were isolated : strain
HA1 (from 1-chlorohexane), strain HA2 (from 1-chloropentane) and strains HA3 and HA4
(from 1-chlorobutane). All were coryneform bacteria with pleomorphic, Gram-positive rods or
cocci (depending on the state of growth and substrate) and frequent V and Y forms. As strains
HA1, HA2 and HA3 were apparently identical (in their spectrum of substrate utilization and
morphology), only strains HA1 and HA4 were examined further. Strain HA1 gave an eight (of
eight) character fit in Seiler's (1983) group A and was identified as an Arthrobacter sp. Strain
HA4 gave at best a six (of eight) character fit in group C and was not further identified.
Growth physiology
Strains HA1 and HA4 were found to have wide substrate specificity for halogenated
compounds. Strain HA1 could utilize at least one haloalkane more than strain HA4, as well as 3chloropropionate (as opposed to the 4-chlorobutyrate utilized by strain HA4); as it gave a linear
growth response to substrate concentration (as opposed to poor growth of strain HA4 at low
substrate concentrations), we chose Arthrobacter sp. strain HA 1 for detailed work.
Strain HA1 utilized at least 18 haloalkanes (Table l), each of which was stable in sterile
control experiments. This organism thus has the widest substrate spectrum of all organisms
reported to degrade haloalkanes. These growth substrates were 1-monosubstituted alkanes;
disubstituted (a,o)compounds and those with mid-chain substituents were not utilized. In
contrast to the chloro-, bromo- and iodosubstituents, neither the fluorosubstituted (1fluoropentane tested) nor the unsubstituted hydrocarbon (hexane) was utilized. The fact that 1chloroethane was not utilized by our new isolates (Table 1) or by other strains (Janssen et al.,
1985; Hartmans et al., 1986) does not mean that it is not biodegradable. We found that
Pseudomonas sp. strain P4 could degrade ethanol or 1-chloroethane with a yield of about 4 g
protein (mol C)-l.
Utilization of each substrate by strain HA1 was quantitative as measured by substrate
disappearance and, where measurable, by release of halide (Table 1). In the case of the
chlorohydrocarbons, the identity of the halide released was confirmed by using an ion-specific
electrode and, after derivatization to 2-chloroethanol, by GLC. It was not possible accurately to
quantify all data for halide release because substrate toxicity was frequently too great (Table 1).
In general, the propyl or butyl derivative was the least toxic. The longer the chain and the larger
the halogen atom, the more toxic was the substrate. Table 1 shows the non-toxic limit above
which the growth rate was reduced; at substrate concentrations about 120% of this value, no
growth occurred.
Strain HA1 quantitatively utilized n-alcohols with chain lengths from C2 to C8with a growth
yield of 5.5 g protein (mol C)-l (Table l), but the yield decreased rapidly if the growth rate was
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R . SCHOLTZ AND OTHERS
Table 1. Alkanes substituted in the 1-position as growth substrates f o r Arthrobacter sp.
strain HA1
Growth yields are presented as g protein (mol C)-l. Specific growth rates and specific degradation rates
[mkat (kg protein)-'] are given. Non-toxic limits for non-growth substrates were tested during growth
with 1-chlorohexane. The solubility of C1-C, 1-haloalkanes is higher than the non-toxic limit; larger
compounds are described in handbooks as 'insoluble'.
Number of carbon atoms in substrate molecule
f
1-Hydroxysubstitution
Growth
Growth yield
Specific growth rate (h-l)
Specific degradation rate
1-Chlorosubstitution
Growth
Growth yield
Specific growth rate (h-I)
Specific degradation rate
Chloride released (%)
Non-toxic limit (mM)
1-Bromosubstitution
Growth
Growth yield
Specific growth rate (h-I)
Specific degradation rate
Bromide released (%)
Non-toxic limit (mM)
1-1odosubstitution
Growth
Growth yield
Specific growth rate (h-l)
Specific degradation rate
Iodide released (%)
Non-toxic limit (mM)
c*
c3
+
+
4.2
c4
c5
+
+
5.6
0.23
3
+
2.7
+
104
2.1
-
4
+
5.6
0.11
1.3
100
14
c
7
c,
c9
+
+
+
-
-
5-8
+
+
5.5
5.5
0.14
1-2
100
3.6
1.3
0.4
+
+
+
1-7
0.6
0.4
-
+
5.2
0.14
1.5
+
4.0
0.12
2.8
95
7.3
+
+
96
9.3
97
2.7
+
+
+
+
+
103
1.8
0.7
0.4
+
4.1
4.2
0.09
2.0
116
100
2-3
3.1
1
0-19
1.5
2-9
-
c,
-
depressed by the toxicity of the substrate. This is a typical value for complete utilization of
hydrocarbons (Anthony, 1982; Cook et a/., 1983; Thurnheer et al., 1986). Yields with more
oxidized compounds (e.g., succinate and citrate) were about 3 g protein (mol C)-l.
Strain HA1 also utilized the chlorohydrocarbons with a yield of about 5.5 g protein (mol C)-l,
where this could be tested (Table 1). The chlorinated compounds were thus utilized for growth as
efficiently as were the n-alcohols: negligible amounts of dissolved organic carbon were found in
growth media. We thus have mass balance of carbon from the (ha1o)alkane into cell material and
carbon dioxide, and mass balance of the organochlorine as the chloride ion.
The growth yields from bromo- and iodohydrocarbons were lower (Table 1). These
compounds, however, supported slower growth than the alcohols and chloroalkanes so we
presume the difference in yield to be due to maintenance energy requirements (Tempest &
Neijssel, 1984).
The growth rate of strain HA1 was examined with several substrates as sole source of carbon
and energy for growth: data for 1-chlorohexane are shown in Fig. 1. Growth was concomitant
with substrate utilization (release of halide ion), and the rate was about 75% of that with the
homologous alcohol (Table 1). Specific degradation rates were calculated to lie between 1.2 and
2.9 mkat (kg protein)-' for the different substrates (Table 1).
Dechiorination
Resting cells of strain HA1 degraded 1-chlorobutane with a specific rate of 800 pkat (kg
protein)-', whether cells were harvested during exponential growth or in the stationary phase.
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27 1
Biodegradation of'1-monohaloalkanes
100
1
r-'
1
I
1 " " " "
0
8 16
24 32 40 48 56
Time (h)
Fig. 1. Growth of Arthrobacter sp. strain HA1 with 1.7 mhl-I-chlorohexane ( 0 )or 3.2 mM-n-hexanol
(a)as the source of carbon and energy. Inset are the utilization of 1-chlorohexane (as release of Cl-) and
the residual concentration of n-hexanol during growth.
Table 2. Substrate spec$city of the haloalkane halidohydrolase(s) in cell-free extracts of strain
HA1 grown with 1-chlorobutane
Reaction rates are presented relative to the reaction with 1-chlorobutane [loo%; 700 pkat (kg
protein)-']. All tests were done under the standard conditions with 5 pmol substrate per 1 ml reaction
mixture.
Reaction rate (%) with the following substrates:
Substituent
1-Chloro1-Bromo1-1odo-
h
f
C,
0
NA
53
Cz
C3
C,
C5
C,
10
23
163
186
100
78
89
90
40
58
67
56
28
159
148
NA,
C7
C8
C9
34
98
24
116
19
125
NA
NA
22
J
Cl0
15
NA
NA
Not assayed.
Whole cells could degrade at least one substance, 1,6-dichlorohexane, which did not support
growth, and both chlorine atoms were released.
Crude extracts from strain HA1 dechlorinated 1-chlorobutane (Table 2). In the reaction,
substrate was quantitatively converted to product: 5-0 2 0.2 mM-Cl- was released from 5 mM-1chlorobutane and, in parallel assays by GLC, the substrate disappeared with concomitant
release of putative n-butanol(5 mM). The organic product from 1-chlorohexane, 1-bromobutane
or 1-iodoethane was identified as the corresponding n-alcohol by co-chromatography with
authentic material and by GLC-MS. Both the organic product and the halide were released
stoichiometrically [ 1 mol (mol substrate)-']. The substrate was stable in the absence of enzyme
and the enzyme preparation released no halide in the absence of substrate. The rate was
proportional to the amount of extract present to a maximum of 1 mg crude extract in a 2 ml
assay, and to the incubation time for at least 2 h.
The specific rate of enzymic dechlorination was 700 pkat (kg protein)-' ;this is very similar to
the rate observed in growing cultures, so presumably the activity accounted for the reaction in
intact cells. The reaction was catalysed by a soluble enzyme(s) which did not require cofactors,
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272
R. SCHOLTZ A N D OTHERS
because removal of membrane proteins by ultracentrifugation (1 10000g; 1-5h) and removal of
small molecules by chromatography on Sephadex G-25 did not significantly alter the
dechlorination activity. The reaction was presumably hydrolytic, since it occurred as rapidly
anaerobically as aerobically.
The substrate spectrum for dehalogenation was wider than the spectrum of growth substrates
(Table 2). Thus, substrates with chain lengths longer and shorter than those of the growth
substrates were dehalogenated. Further, 2-chlorobutane [ 120 pkat (kg protein)-'] and 3chloroheptane [250 pkat (kg protein)-'] were substrates and a,o-dichlorohydrocarbons (at least
C4-C6) were dechlorinated rapidly to give 2 mol C1- although they did not support growth.
The dehalogenation was inducible. Cells grown with butanol had no dechlorinating activity;
this was observed only in cells grown with haloalkanes. Extracts of cells grown with 3chloropropionate as sole added source of carbon and energy could dechlorinate 3chloropropionate [data were corrected for spontaneous hydrolysis of the substrate (4 % d-*)I, but
not 1-chlorobutane, and extracts of cells grown with 1-chlorobutane could not dechlorinate 3chloropropionate. Degradation of the haloalkanoates was thus independent of the metabolism
of haloalkanes (cf. Janssen et al., 1985).
DISCUSSION
We have obtained from a single inoculum at least two different coryneform bacteria (strains
HA1 and HA4) able to degrade a very wide spectrum of haloalkanes. Similar successes were
reported by Janssen et al. (1985) and Hartmans et al. (1985, 1986), in contrast to the work of
Omori & Alexander (1978a), who obtained only three successful enrichments from over 500 soil
types. This comparison emphasizes the advantage of inocula chosen for their likelihood to
contain organisms which have evolved to utilize specific xenobiotics (Cook et al., 1983). 1Bromopentane occurs naturally (Gschwend et al., 1985), so its degradation might be expected
(Dagley, 1975), and evolution to utilize the chloro-analogue is not difficult to envisage. We used
a simple mineral salts medium in which successful enrichments are easily scored (OD, GLC,
halide release) and, fortunately, non-toxic substrate concentrations. The nature of the observed
toxicity (or toxicities) is unclear but longer chains were more toxic, whereas with the alkanes the
reverse is true (Britton, 1984). The toxicity of 1-iodoethane to bacteria may be due to its
efficiency as an alkylating agent. However, the toxicity to growing cells is independent of the
reaction rate of the halidohydrolase(s) with the different substrates (cf. Tables 1 and 2). The
enrichment of Arthrobacter and related strains is not surprising, because they classically degrade
aliphatic and aromatic hydrocarbons (e.g., Schlegel, 198l), and they are known to degrade
chlorinated derivatives (Janssen et al., 1985; Stanlake & Finn, 1982). However, our strains do
not utilize the unsubstituted alkanes.
The present work extends that of Janssen et al. (1985), Omori & Alexander (1978a,b) and
Yokota et al. (1986), and, with data from Hartmans et al. (1986), shows that the
monochloroalkanes from C to c8 are all biodegradable, though different organisms utilize
substrates of different chain lengths. Longer chain compounds are probably degradable, but the
only data available (Murphy & Perry, 1983, 1984) do not include mass balances. Ours is the first
report of the degradation of all the c2-c8 monobromo- and C2-C7 monoiodohydrocarbons. The
relatively high specific activities and the quantitative dehalogenation could allow the organisms
to be used to destroy these chlorinated solvents in waste streams (cf. Galli & Leisinger, 1985).
The mechanism of dechlorination seems to be hydrolytic, corresponding to the well defined
halidohydrolase of Janssen et al. (1985) (see also Kohler-Staub & Leisinger, 1985)and the system
of Yokota et al. (1986), although it is not possible to make quantitative comparisons with the
latter results. There is no evidence for the oxidative systems known or suspected for
chloromethane, vinyl chloride (Hartmans et al., 1985, 1986) and 1,9-dichlorononane (Yokota et
al., 1986) or the NADH-coupled system implied by Omori & Alexander (1978b). Much of the
dehalogenation activity in crude extracts can be explained by a single halidohydrolase which is
currently being purified (R. Scholtz, A. M. Cook & T. Leisinger, unpublished).
The enzymic dehalogenation is inducible. Some substrates for the enzyme (e.g. 1Downloaded from www.microbiologyresearch.org by
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Biodegradation of I -monohaloalkanes
273
chloropropane) are apparently not inducers though the reaction product is a growth substrate
(Table l), so regulatory and catalytic proteins recognize different molecules. In addition, many
substrates (e.g. with mid-chain substituents) of the halidohydrolase do not support growth, so, if
degradative pathways for the dehalogenated products were present, yet more haloalkanes would
be biodegradable.
We are grateful to Dr K. Ramsteiner, Ciba-Geigy AG, Basel, for obtaining the mass spectra, to Dr D. Studer for
taking the electron micrographs, and to Dr T. Egli for measuring dissolved organic carbon. The investigation was
supported in part by a grant from the Swiss Federal Institute of Technology, Zurich.
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