Department of Molecular and Cell Biology, Institute of Medical Sciences, University of Aberdeen, Foresterhill, Aberdeen AB25 2ZD, UK1
Institut für Mikrobiologie, ETH Zürich, ETH-Zentrum/LFV, CH-8092 Zürich, Switzerland2
Author for correspondence: Stéphane Vuilleumier. Tel: +41 1 632 33 57. Fax: +41 1 632 11 48. e-mail: svuilleu{at}micro.biol.ethz.ch
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ABSTRACT |
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Keywords: glutathione, dichloromethane, intracellular pH, chloride, formaldehyde
Abbreviations: DCM, dichloromethane; FDH, formaldehyde dehydrogenase; GSH, glutathione; GST, glutathione S-transferase; pHi, intracellular pH; , membrane potential
a Present address: Department of Biology, Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
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INTRODUCTION |
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For the majority of bacteria, the physiological functions of GSTs are unknown, although, as with their mammalian counterparts, they are likely to be instrumental in detoxification processes (Zablotowicz et al., 1995 ; Gutheil et al., 1997
; Vuilleumier, 1997
). For others, however, a GST enzyme is actually the central means of carbon acquisition, as in the case of methylotrophic bacteria able to assimilate carbon from dichloromethane (DCM) via a DCM dehalogenase/GST-type enzyme (Leisinger et al., 1994
; Vuilleumier, 1997
). Chlorinated methanes are volatile chemicals, which have been extensively used in industry and consequently are frequent environmental pollutants. Bacteria have evolved the capacity for the metabolism of these compounds (Leisinger et al., 1994
; Leisinger, 1996
). Dehalogenation of chlorinated alkanes by bacteria poses several potential threats to cell viability. In the case of DCM, dehalogenation produces in the cytoplasm 2 mol HCl per mol substrate, plus 1 mol of a toxic product, formaldehyde, via two postulated GSH adducts, S-chloromethylglutathione and S-hydroxymethylglutathione (reaction 1).
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S-Chloromethylglutathione has been shown to alkylate DNA (Dechert, 1995 ) and this is believed to be the basis of the moderately strong mutagenicity in the Salmonella typhimurium Ames tester strain TA1535 expressing mammalian DCM dehalogenase (Thier et al., 1993
; Gisi et al., 1999
). In E. coli and related bacteria, it has been shown that GSH-linked detoxification is intimately associated with potassium efflux systems and the modulation of cytoplasmic pH (Ferguson et al., 1998
). Protection against the toxic compounds methylglyoxal and N-ethylmaleimide, which are detoxified via GSH adducts, is achieved by acidification of the cytoplasm (Ferguson et al., 1993
, 1995
, 1997
). GSH is known to be involved in acid tolerance in some bacteria (Riccillo et al., 2000
) and it has been shown that protection against mutagens can be achieved by incubation with weak organic acids that also lower the cytoplasmic pH (Oktyabrsky et al., 1993
). Dehalogenation reactions in E. coli, such as the conjugation of the model GST substrate 1-chloro-2,4-dinitrobenzene, are inhibitory to growth (Ness et al., 1997
). However, the low expression and specific activity of GSTs may limit the impact of dehalogenation on the metabolism of E. coli cells (Zablotowicz et al., 1995
; Vuilleumier, 1997
). To probe the capacity of E. coli cells to cope with the multiple stresses posed by dehalogenation of chlorinated methanes, we have investigated the physiology of E. coli cells expressing DCM dehalogenase/GST of Methylophilus sp. strain DM11 (Bader & Leisinger, 1994
; Vuilleumier & Leisinger, 1996
; Gisi et al., 1999
). We demonstrate that cells experience no long-term damage from the activity of this enzyme despite transient inhibition of growth during the metabolism of DCM. During dehalogenation cells excrete formaldehyde and chloride and the cytoplasmic pH is lowered. Growth inhibition probably arises from a combination of the stresses of internal acid production, toxicity of a reactive intermediate in the reaction and accumulation of formaldehyde. The data demonstrate that E. coli cells have the ability to cope with intracellularly generated HCl, a previously unremarked ability of this organism.
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METHODS |
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Growth media and growth conditions.
All experiments were conducted in Kx minimal medium (where x is equal to the millimolar concentration of potassium; Epstein & Kim, 1971 ) with 0·2% glucose as carbon source. Cells were grown to stationary phase by overnight culture on a shaking incubator at 37 °C in K0·2 containing 0·2% glucose and 1 µg thiamine ml-1. Cells were then diluted 15-fold into 30 ml K0·2 (OD650=0·1) and grown aerobically at 25 °C in the presence of 1 mM IPTG. Growth was monitored until OD650 reached 0·3 and the test compounds (DCM or formaldehyde) were added. DCM was added from a 100 mM stock solution in water to either 0·3 or 0·6 mM final concentration. Formaldehyde was purchased as a 0·19% solution (Fluka). The formaldehyde concentration in the growth medium was assayed as described previously (Vuilleumier & Leisinger, 1996
). Growth was monitored by light scattering at 650 nm (accuracy±0·01). In experiments involving DCM, cells were grown in gas-tight 300 ml Erlenmeyer flasks with mininert caps (Supelco). All experiments were repeated at least three times.
Potassium efflux assays.
These experiments were conducted essentially as described previously (Elmore et al., 1990 ; Ferguson et al., 1997
). Overnight cultures of the appropriate strain were diluted into fresh medium to an OD650 of 0·1 and grown to an OD650 of 0·3 in K0·2 minimal medium. An aliquot of cells (50 ml) was then harvested by filtration (Millipore, 0·45 µm pore size, 4·5 cm) and washed with 5 ml K10 buffer. Cells were then resuspended to give a final OD650 of 0·6 in 25 ml K0 buffer in the presence or absence of 0·2% (w/v) glucose. The resuspended culture was transferred to two 300 ml Erlenmeyer flasks (at 25 °C), containing a magnetic stirring bar, and sealed with mininert caps. A 1 ml sample was kept for OD650 measurement. The culture was aerated by constant stirring. DCM was added from a 100 mM stock solution in water as required. At intervals 1 ml samples were removed, transferred to Eppendorf tubes and centrifuged at 14000 r.p.m. (Jouan A14) for 1 min. The supernatant was then removed by aspiration and the pellets were resuspended in 1 ml distilled water, boiled for 5 min to release the potassium and centrifuged (as above) for 1 min to pellet cell debris. Potassium in the supernatant was measured by flame photometry (Corning 400 and Sherwood 410) as described previously (Elmore et al., 1990
; Ferguson et al., 1997
). All experiments were repeated at least three times (SEM<5%). The data shown are typical data obtained in replicate experiments.
Intracellular pH (pHi) measurements.
The pHi after DCM addition was determined using the distribution of a radiolabelled weak acid according to a previously described method (Kroll & Booth, 1983 ; McLaggan et al., 1994
) by incubating bacteria at an OD650 of 0·6 with [14C]benzoic acid (4·5 µM, 3·7 x 103 Bq ml-1; Sigma) and [3H]inulin (1·0 µM, 3·7 x 104 Bq ml-1; Sigma) as the extracellular marker. The cell pellet was separated from the supernatant by spinning the cells through bromododecane oil (McLaggan et al., 1994
) and pHi was calculated as described previously (Kroll & Booth, 1983
). All experiments were repeated three times. For calculation of mean pHi prior to and after addition of DCM, the pHi values for each condition in each experiment were averaged and the SD of the replicate experiments was calculated to be lower than 0·05 units in all cases. For samples prior to addition of DCM at least two measurements were made and after DCM addition at least four measurements were made.
Preparation and analysis of cell-free extracts and measurements of DCM dehalogenase and formaldehyde dehydrogenase (FDH) activity.
Cell-free extracts were obtained by three passages through a French press apparatus (14000 p.s.i.) followed by centrifugation at 30000 g for 30 min. Protein concentration was determined in the resulting cell-free extracts using commercial Bradford reagent (Bio-Rad) with bovine serum albumin (Sigma) as standard. Specific rates of DCM dehalogenation were determined spectrophotometrically by assaying the rate of formaldehyde production, as described previously (Vuilleumier & Leisinger, 1996 ). One unit is defined as the amount of enzyme catalysing the conversion of 1 µmol DCM min-1. FDH activity was calculated from the rate of NADH formation at 340 nm using
=6220 M-1 cm-1 (Gutheil et al., 1997
). The reaction mixture contained 0·5 mM NAD+ in a final volume of 0·5 ml 100 mM potassium phosphate (pH 8·0). To evaluate the contribution of GSH-independent FDHs, the assay was conducted either with or without 10 mM GSH in the reaction mixture. GSH in cell-free extracts was assayed using a GSH assay kit (Calbiochem) and gave values in the expected range [3060 nmol (mg protein)-1]. The experiments were repeated with independent extracts.
Chloride assay.
The concentration of chloride ions in the culture supernatant was determined as described by Bergmann & Sanik (1957 ). Experiments were designed as described for potassium efflux determination (above) and 0·6 ml samples were withdrawn from the sealed flasks that contained cultures metabolizing DCM. To each 600 µl aliquot of culture supernatant 200 µl 0·25 M ferric ammonium sulfate, dissolved in 9 M nitric acid, and 200 µl saturated mercuric thiocyanate in ethanol were added. After 10 min incubation at room temperature the absorbance at 460 nm was determined. A standard curve of 0·11 mM NaCl was used for calibration. All experiments were repeated twice.
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RESULTS |
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The toxicity of formaldehyde was investigated in strain MJF274(pME1983) (DCM+) (Fig. 2a). Addition of formaldehyde reduced the growth rate, but was only strongly inhibitory above 0·6 mM (Fig. 2a
). Therefore, since growth inhibition by 0·3 mM DCM was immediate (Fig. 1a
), but strongly inhibitory concentrations of the formaldehyde product were not achieved (Fig. 1b
), we conclude that this product of dehalogenation is not the cause of growth inhibition. The growth rate sustained by MJF274(pME1983) after completion of DCM dehalogenation (µ=0·22±0·01 h-1; n=3) was similar to that achieved in the presence of the equivalent concentration of formaldehyde (µ=0·18±0·01 h-1; n=3). Cells washed free of DCM and the products of dehalogenation soon after addition of DCM re-established growth rates identical to those of cultures not treated with DCM (µ=0·28±0·01 h-1; n=3; Fig. 2b
). Similarly, cells incubated in the presence of 0·3 mM formaldehyde recovered normal exponential growth after washing and resuspension in fresh growth medium (Fig. 2b
). These data suggest that cells do not sustain significant damage when dehalogenating DCM under the chosen conditions.
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DISCUSSION |
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At 0·3 mM DCM neither the formaldehyde produced, the lowering of cytoplasmic pH nor the chloride ions generated in the cytoplasm appear to cause significant long-lasting damage to the cells. Transfer of cells incubated with either DCM or formaldehyde to fresh growth medium resulted in immediate exponential growth at the same rate (Fig. 2b). The cells respond to the intracellular generation of formaldehyde by the induction of the E. coli GSH-dependent FDH (Table 1
). GSH-dependent FDH has been shown previously to be a class III alcohol dehydrogenase that also acts on hydroxymethylglutathione (Gutheil et al., 1992
), which is readily formed from formaldehyde and GSH in aqueous solution (Uotila & Koivusalo, 1974
). Induction of GSH-dependent FDH by the addition of formaldehyde to the growth medium has also been demonstrated (Gutheil et al., 1997
) and has been suggested to represent a significant component of the detoxification machinery for this reactive compound (Uotila & Koivusalo, 1989
; Kummerle et al., 1996
). Our data are quite consistent with these observations. In addition, our inability to detect significant levels of GSH-independent FDH (Table 1
) also explains the observed lack of metabolism of this compound by the GSH-deficient strain MJF335.
Chloride ion transport is poorly characterized in bacterial cells. In our experiments the rate of conversion of DCM was approximately 3540 nmol DCM min-1 (mg cell dry wt)-1, which corresponds to 7080 nmol chloride ion expelled min-1. This rate is essentially set by the metabolism of DCM and thus may not represent the fastest rate of chloride efflux that E. coli can sustain. Inverted membrane vesicles from E. coli are known to be permeable to chloride ions, but the mechanism underlying this observation is not known (Reenstra et al., 1980 ). A chloride channel has been detected in E. coli (Maduke et al., 1999
), but whether such a channel can participate in the efflux of this anion is unknown. Such a system would be subject to influence by the external chloride concentration. Based on data from Figs 1
and 5
and an intracellular volume of 1·6 µl (mg dry wt)-1, the intracellular concentration of chloride ions would be approximately 40 mM after 1 min of dehalogenation. Allowing for a delay of 23 min (Fig. 5
) before the onset of chloride excretion, the intracellular chloride concentration would be approximately 120160 mM. We have previously shown that a rise in anion concentration of this magnitude can be accommodated in E. coli by the release of glutamate and other physiological anions (Roe et al., 1998
). The external concentration of chloride in the experiments of Roe et al. (1998
) was approximately 0·25 mM. Thus, the considerable transmembrane gradient generated under these conditions would allow a channel to operate to release chloride. Given a membrane potential (
) of greater than -120 mV however, the chloride concentration required to block channel activity can be estimated to be at least 10 M (-120 mV is equivalent to a 100-fold gradient and the estimated internal concentration is 100 mM), which is beyond the upper limit of that tolerable for growth by E. coli. In this work, the presence of 0·1 M NaCl during dehalogenation did not significantly alter growth inhibition. Alternatively, the presence in E. coli of a uniporter energized only by the
would provide another mechanism for chloride ion egress. This would have the potential benefit of chloride efflux becoming independent of any channel gating mechanism and allow the cell to sustain gradients of 100-fold or more (
-120 mV; Booth, 1985
).
Lowering of the cytoplasmic pH is expected to cause growth inhibition. We have previously shown that when the cytoplasmic pH falls below pH 7·4, which is a similar value to that observed in the presence of DCM (Fig. 3), the growth rate falls by 50% (Roe et al., 1998
). During DCM dehalogenation growth was completely inhibited and this suggests that the change in the cytoplasmic pH is not the sole cause of inhibition. The levels of formaldehyde produced by DCM dehalogenation are insufficient to cause the growth inhibition, since the addition of 0·3 mM formaldehyde, which is the highest concentration of formaldehyde reached in the presence of 0·3 mM DCM, does not significantly inhibit growth (Fig. 2a
). The cause of growth inhibition can be rapidly removed, since washing cells restored normal exponential growth almost immediately (Fig. 2b
). One possible explanation for the greater growth inhibition observed during DCM dehalogenation is toxicity of GSH adducts generated during DCM metabolism. The observed mutagenicity of GST-mediated DCM conversion (Thier et al., 1993
; Gisi et al., 1999
) is possibly due to the modification of DNA bases by S-chloromethylglutathione, which was demonstrated in vitro (Dechert, 1995
). Insertion mutants of the DCM-degrading bacterium Methylobacterium dichloromethanicum DM4 generated by mini-Tn5 mutagenesis were isolated that are unable to grow with DCM, but which still possess GSH and active DCM dehalogenase. These mutants grow well with methanol, which is oxidized during growth to formaldehyde, a central metabolic intermediate in methylotrophic bacteria. In one of these mutants, the transposon insertion is located in the gene encoding DNA polymerase I, a key enzyme in DNA repair (Kayser et al., 2000
). Thus, it seems possible that the unexpected severity of the growth inhibition observed during exposure to DCM of E. coli cells expressing DCM dehalogenase is due in part to genotoxic GSH adducts. In this context the fall in the cytoplasmic pH, caused by the production of acid during dehalogenation, may even protect the cells from further damage (Ferguson et al., 1995
, 1997
, 2000
).
This study demonstrates that E. coli experiences growth inhibition during dehalogenation of DCM. In contrast, some DCM-degrading methylotrophic bacteria grow whilst actively dehalogenating DCM (Leisinger et al., 1994 ). It is possible that they are more tolerant of the perturbations of pHi and chloride ion concentration arising from dehalogenation and conceivably they have developed tolerance of the DCM adducts formed with GSH. Growth of these organisms with DCM is associated with rapid acidification of the medium, which is consistent with high rates of DCM metabolism. Nevertheless, the slow maximal growth rates of these organisms compared to that of E. coli (approx. 0·08 h-1) may reflect the burden faced by methylotrophic bacteria growing with DCM due to constant acidification of the cytoplasm, rather than slow rates of carbon and energy acquisition. Whatever the case may be, our study suggests that significant physiological adaptations may have been required by methylotrophic bacteria to utilize chlorinated methanes as a means of generating carbon and energy.
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ACKNOWLEDGEMENTS |
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Received 10 April 2000;
revised 7 July 2000;
accepted 10 July 2000.
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