Regulation of catabolic enzymes during long-term exposure of Delftia acidovorans MC1 to chlorophenoxy herbicides

Dirk Benndorf1, Ian Davidson2 and Wolfgang Babel1

1 UFZ – Centre for Environmental Research Leipzig-Halle, Department of Environmental Microbiology, Permoserstr. 15, 04318 Leipzig, Germany
2 University of Aberdeen, Department of Molecular and Cell Biology, Institute of Medical Sciences, Foresterhill, Aberdeen AB25 2ZD, UK

Correspondence
Dirk Benndorf
dirk.benndorf{at}ufz.de


   ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Delftia acidovorans MC1 is able to grow on chlorophenoxy herbicides such as 2,4-dichlorophenoxypropionic acid (2,4-DCPP) and 2,4-dichlorophenoxyacetic acid as sole sources of carbon and energy. High concentrations of the potentially toxic organics inhibit the productive degradation and poison the organism. To discover the target of chlorophenoxy herbicides in D. acidovorans MC1 and to recognize adaptation mechanisms, the response to chlorophenoxy acids at the level of proteins was analysed. The comparison of protein patterns after chemostatic growth on pyruvate and 2,4-DCPP facilitated the discovery of several proteins induced and repressed due to the substrate shifts. Many of the induced enzymes, for example two chlorocatechol 1,2-dioxygenases, are involved in the metabolism of 2,4-DCPP. A stronger induction of some catabolic enzymes (chlorocatechol 1,2-dioxygenase TfdCII, chloromuconate cycloisomerase TfdD) caused by an instant increase in the concentration of 2,4-DCPP resulted in increased rates of productive detoxification and finally in resistance of the cells. Nevertheless, the decrease of the (S)-2,4-DCPP-specific 2-oxoglutarate-dependent dioxygenase in 2D gels reveals a potential bottleneck in 2,4-DCPP degradation. Well-known heat-shock proteins and oxidative-stress proteins play a minor role in adaptation, because apart from DnaK only a weak or no induction of the proteins GroEL, AhpC and SodA was observed. Moreover, the modification of elongation factor Tu (TufA), a strong decrease of asparaginase and the induction of the hypothetical periplasmic protein YceI point to additional resistance mechanisms against chlorophenoxy herbicides.


Abbreviations: 2,4-DCP, 2,4-dichlorophenol; 2,4-DCPP, 2,4-dichlorophenoxypropionic acid

The protein sequence data reported in this article will appear in the SWISS-PROT and TrEMBL knowledgebase under accession numbers Q8KN28, Q9RNZ9, Q93T12, Q9R5K5, P83709, P83707, P83710, P83712, P83708 and P83711.


   INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Micro-organisms are metabolically versatile. Their metabolic activities contribute to the natural cycles of compounds in the environment. In addition, they are able to attack compounds of anthropogenic origin, for example chlorophenoxy acids (Duxbury et al., 1970; Evans et al., 1971; Pemberton & Fisher, 1977; Kilpi, 1980), which were widely used in crop control. So far, several species mineralizing these compounds have been isolated (Pieper et al., 1988; Horvath et al., 1990). The biodegradation of chlorophenoxy acids was elucidated by the purification of their respective enzymes and the sequencing of the corresponding tfd genes (Fukumori & Hausinger, 1993; Kaphammer et al., 1990). Using molecular probes, the distribution of tfd genes has been shown at contaminated sites as well as in pristine soils never treated with chlorophenoxy acids (Kamagata et al., 1997).

However, the presence of genetic information is not sufficient to guarantee biodegradation of chlorophenoxy acids at strongly contaminated sites, since micro-organisms may be poisoned by growth substrate or metabolites, if the concentrations are high and the substances are potentially toxic. First, chlorophenoxy acids are hydrophobic compounds and weak organic acids; therefore, they are potentially toxic agents which may disturb the energy conservation system of bacteria located in the cytoplasmic membrane (Loffhagen et al., 1997). Second, the lipophilicity and the reactivity of compounds may cause damage to several biomolecules as described in Acinetobacter calcoaceticus, which was concluded from the enhanced synthesis of heat-shock proteins and oxidative-stress proteins (Benndorf et al., 1999, 2001). However, the presence of the chlorophenoxy herbicide 2,4-dichlorophenoxypropionic acid (2,4-DCPP) and its metabolites 2,4-dichlorophenol (2,4-DCP) and 3,5-dichlorocatechol during growth of Delftia acidovorans MC1 (Benndorf & Babel, 2002) on pyruvate caused no significant induction of the respective proteins, whereas the induction of catabolic enzymes indicates that productive detoxification is also a component of the response to chemostress in a bacterium which is able to metabolize these compounds (Benndorf & Babel, 2002).

The goal of the present research is to elucidate and understand the long-term adaptation of D. acidovorans strain MC1 during continuous growth on low and high concentrations of chlorophenoxy herbicides simulating both low bioavailability and excess concentration of substrates at contaminated sites. Proteome analysis has been proven to deliver an overall picture of gene expression that represents the metabolic state of an organism (Peng & Shimizu, 2003), as well as indicating the presence of adaptive responses (Vasseur et al., 1999). It was used in this study as this technique also allows recognition of post-translational modifications which may also play a role in response(s) to chemostress.


   METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Bacterial strain and growth conditions.
D. acidovorans MC1 (Müller et al., 1999) was grown in minimal medium (Müller & Babel, 1986) in shake flasks for batch experiments and in a fermenter (Biostat Q, Braun, working volume 250 ml) for continuous cultivation. For continuous growth on 2,4-DCPP, the fermenter was inoculated with cells pre-cultivated on a mixture of 50 ml PYE medium (3 g peptone l–1, 3 g yeast extract l–1 and 10 mM fructose) and 100 ml minimal medium pH 8·5 containing 13·7 mM 2-oxoglutarate, overnight. The growth temperature was 30 °C, the dissolved oxygen concentration ranged from 50 to 95 % of saturation and the pH was maintained at 8·5 by automatic titration with 0·2 M KOH or 0·2 M H2SO4, as appropriate. Afterwards, the culture was grown chemostatically on 8·5 mM 2,4-DCPP (s0) or on 18·2 mM sodium pyruvate with a dilution rate (D) of 0·05 h–1.

For growth on excess concentrations of 2,4-DCPP, the pH-auxostat principle (Krayl et al., 2003) was used. Briefly, the reservoir medium containing the substrate was used to adjust the pH of the culture. Several concentrations between 4·2 and 29·5 mM 2,4-DCPP and different volumes of 2 M KOH were added to the medium, whereas the buffer capacity of the medium had to be lower than the amount of acid which would be released if all 2,4-DCPP were consumed. For batch experiments, D. acidovorans MC1 was pre-cultivated on minimal medium pH 7 containing 3 g sodium pyruvate l–1 in batches, overnight. Then, 5 ml inoculum was added to 20 ml medium (pH ranging between 6 and 8·5) containing 3 g sodium pyruvate l–1 and several concentrations of 2,4-DCPP.

Growth was measured spectrophotometrically by monitoring optical density at 700 nm.

Determination of 2,4-DCPP, its metabolites and measurement of enzyme activities.
Concentrations of 2,4-DCPP and 2,4-DCP were determined by HPLC analysis (Oh & Tuovinen, 1990). The activities of chlorocatechol 1,2-dioxygenase (Müller et al., 2001) and dichlorprop/2-oxoglutarate dioxygenases were measured as described previously (Westendorf et al., 2003).

Sample preparation, 2D electrophoresis and electroblotting.
The bacteria were harvested and the cells were lysed as described previously (Benndorf & Babel, 2002). The protein content was determined as described previously (Holtzhauer & Hahn, 1988). For 2D electrophoresis, 50 µg protein for analytical gels and 2000 µg for micropreparative gels were precipitated with ice-cold acetone, resolubilized, loaded on 18 cm long Immobiline DryStrip pH 3–10 NL and 2D electrophoresis was carried out as described previously (Benndorf & Babel, 2002). Analytical gels were silver-stained as described by Blum et al. (1987) and dried in a stream of unheated air from a GelAir Dryer (Bio-Rad). With respect to reproducibility, we carried out two independent experiments with at least two gels per sample. The best gels of each replicate were selected for image analysis. For mass spectrometry (MS) and internal protein sequencing, micropreparative gels were stained with Coomassie blue, whereas for amino terminal sequencing, the gels were first electroblotted overnight on PVDF (Bio-Rad) membranes using the CAPS buffer system (Jin & Cerletti, 1992) and stained with Coomassie blue afterwards (Benndorf & Babel, 2002).

Identification of proteins by amino terminal and internal sequencing and MS.
For peptide-mass mapping, proteins of interest were excised from micropreparative 2D gels digested by trypsin in-gel and analysed using a Voyager-DE STR MALDI-TOF mass spectrometer at the Aberdeen Proteome Facility (Cash et al., 1999). Furthermore, proteins were identified by amino terminal and internal amino acid sequencing using a model 491cLC protein sequencer (Applied Biosystems). Before internal sequencing, the protein spots were excised from micropreparative 2D gels and digested in-gel as described above except that Lysyl Endopeptidase (LysC) from Achromobacter lyticus (Wako Chemicals; 2 µg in 200 µl 25 mM Tris/HCl pH 8·0) was used instead of trypsin. The peptides were separated by HPLC using a self-packed column (column length 150 mm, i.d. 0·5 mm, Self Pack POROS 10 R2 Reversed Phase Packing, PerSeptive Biosystems). A gradient with a flow rate of 50 µl min–1 and with increasing acetonitrile concentrations was used (buffer A – 0·1 % TFA in water; buffer B – 0·085 % TFA in 70 % acetonitrile, increase from 0 % B to 60 % B within 90 min). The peptides were detected by UV absorption at 214 nm and fractions containing peptides were collected manually. The fractions were dried down to 10 µl, mixed with 100 µl of 0·1 % TFA in water and applied to Prosorb cartridges (Applied Biosystems) according to the manufacturer's instructions.

Comparison of 2D protein patterns.
Silver-stained dried gels were scanned using a UMAX Power Look 2000 Scanner with an eight bit dynamic range and 200 d.p.i. resolution. Gel images were analysed by PHORETIX 2D 5.01 software (NonLinear Dynamics). For comparing protein patterns, we used only spots that were present in both gels of replicated experiments. Protein spots with a twofold, or greater, normalized volume (density) than the corresponding spots in control gels were considered to be amplified, and spots with half, or less, normalized volume than the corresponding spots in control gels were considered to be diminished. Spots which were observed following imposition of stress conditions, but not in the control protein pattern, were considered to be newly synthesized proteins. Molecular mass and pI were calibrated using internal standards defined by calibration with the 2D SDS-PAGE Standards kit (Bio-Rad).


   RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Sensitivity against 2,4-DCPP
Xenobiotic compounds such as 2,4-DCPP are sources of carbon and energy; on the other hand, they become toxic at higher concentrations. They damage bacteria at several sites. The extent of damage can be evaluated by measuring physiological parameters, for example viability, growth rate, respiration rate and ATP synthesis (Krayl et al., 2003). The exposure of bacteria during growth on non-toxic substrates to 2,4-DCPP (Benndorf & Babel, 2002) and other chlorophenoxy acids (Loffhagen et al., 2003) causes diminishing of growth rates. In the presence of 2,4-DCPP, the diminishing of growth rate of D. acidovorans MC1 depends on pH (Table 1). It indicates that this compound may act as an uncoupler of electron transport phosphorylation. This has been proved in Comamonas testosteroni by measuring respiration rates and rates of ATP synthesis in the presence of several chlorophenoxy acids (Loffhagen et al., 2003). To withstand such challenges and to increase their resistance, cells often stabilize their cytoplasmic membranes by changing their fatty acid composition (Heipieper et al., 1992; Loffhagen et al., 1995) or induce stress proteins. The measurement of growth as one physiological parameter to quantify the sensitivity of D. acidovorans MC1 against the impact of 2,4-DCPP showed that cells growing on 2,4-DCPP are more stable than cells growing on pyruvate, since a threefold higher concentration of the 2,4-DCPP was necessary to inhibit growth on 2,4-DCPP to the same extent as growth on pyruvate (see table elements with grey background in Table 1). This phenomenon was promising for detailed studies of resistance at the molecular level.


View this table:
[in this window]
[in a new window]
 
Table 1. Growth rates of D. acidovorans MC1 in the presence of several concentrations of 2,4-DCPP

Grey boxes, conditions causing a similar decrease of growth rates in cells growing on pyruvate or 2,4-DCPP at pH 8·5.

 
Induction of catabolic genes by substrate shift
To identify the enzymes which were involved in the biodegradation of chlorophenoxy acids, D. acidovorans MC1 was chemostatically grown with a dilution rate of D=0·05 h–1 on pyruvate or for induction of catabolic genes on 2,4-DCPP. At steady-state, neither 2,4-DCPP nor its early metabolite 2,4-DCP was detectable (Table 2). The 2D protein patterns after chemostatic growth on pyruvate and on 2,4-DCPP, each containing about 600 spots, were compared (Fig. 1); 17 amplified, 11 newly synthesized and 13 diminished proteins were detected during growth on 2,4-DCPP. The similar genetic background of chlorophenol degradation in D. acidovorans MC1 (Müller et al., 2001) and in Ralstonia eutropha (Fukumori & Hausinger, 1993; Kaphammer et al., 1990) facilitated the identification of all enzymes of the catabolic pathway (Table 3) except for dienelactone hydrolase TfdE, although its presence in gels could be expected from their calculated isoelectric points and molecular masses. Differing from R. eutropha, the initial step of degradation of 2,4-DCPP in D. acidovorans MC1 requires two enantioselective 2-oxoglutarate-dependent dioxygenases, RdpA and SdpA (Westendorf et al., 2002, 2003). By gel comparison, it became obvious that both enzymes are already expressed in the absence of 2,4-DCPP, indicating that they are not co-regulated with the Tfd proteins. The possibility to observe these changes in metabolism by measuring the concentrations of catabolic enzymes in 2D gels is a good precondition to evaluate also the responses of D. acidovorans MC1 to stress, particularly to high concentrations of 2,4-DCPP.


View this table:
[in this window]
[in a new window]
 
Table 2. Conditions of continuous growth of D. acidovorans on 2,4-DCPP and 2,4-DCP

Growth in the pH-auxostat was initiated by the addition of 15 mM 2,4-DCPP and 0·5 mM 2,4-DCP to cells pre-cultivated in a chemostat D=0·05 h–1 on 2,4-DCPP, which also represents the control of the experiment.

 


View larger version (35K):
[in this window]
[in a new window]
 
Fig. 1. 2D protein pattern of D. acidovorans MC1 after chemostatic growth on pyruvate (A) and 2,4-DCPP (B). Boxed spots, more than twofold amplified proteins; circled spots, new proteins; diamonds, proteins diminished by at least 50 %.

 

View this table:
[in this window]
[in a new window]
 
Table 3. Identification of proteins of D. acidovorans MC1 by amino terminal and internal protein sequencing

 
Regulation of catabolic pathway in response to excess concentrations of 2,4-DCPP
For long-term exposure of D. acidovorans MC1 to excess concentrations of 2,4-DCPP, the pH-auxostat regime was used, since it enabled us to adjust high and stable concentrations of 2,4-DCPP, which was used as the carbon source on the one hand and acted as a toxic compound on the other. After 48 h continuous growth on excess concentration of 2,4-DCPP (s0=25·5 mM), a dilution rate of 0·05 h–1 was measured and the residual concentrations of 2,4-DCPP and its metabolites rose to 14·12 mM 2,4-DCPP and 0·56 mM 2,4-DCP. For a detailed study, the pH-auxostat experiment was repeated and 15 mM 2,4-DCPP and 0·5 mM 2,4-DCP, corresponding to the concentrations accumulated in the experiment before, were added initially. During the first 3 h the dilution rate increased from 0·05 to 0·144 h–1 and remained at this level (Table 2). Sixteen hours after the addition of both compounds the dilution rate started to decrease and reached the final level of 0·054 h–1 after about 30 h. Apparently, the poisoning of cells needs some time.

Afterwards, the protein pattern after chemostatic growth on 2,4-DCPP representing low residual concentration was taken as a control pattern for comparison with growth at high residual concentration of 2,4-DCPP (pH-auxostat regime). Seventy-four changed protein spots (34 amplified, 22 newly synthesized, 18 diminished, see Fig. 2) indicate that D. acidovorans MC1 is able to adapt at the protein level to excess 2,4-DCPP. To measure the dynamics of protein pattern in response to 2,4-DCPP, protein samples were also taken from the time-course experiment with instant addition of 15 mM 2,4-DCPP and 0·5 mM 2,4-DCP. The corresponding 2D gels (whole gels not shown) showed that induction profiles of the identified proteins were very different (Figs 3 and 4). The further induction of catabolic enzymes (chlorocatechol 1,2-dioxygenase TfdCII, chloromuconate cycloisomerase TfdD) can probably increase the rate of biodegradation and reduce the concentrations of metabolites. Furthermore, the activities or specificities of some enzymes, particularly of TfdD, are additionally regulated by post-translational modifications because the occurrence of isoforms 3 and 4 was favoured in the presence of excess 2,4-DCPP (Figs 4 and 5). However, an inactivation of enzymes resulting from chemical reactions with reactive metabolites, for example 3,5-dichlorocatechol or others, cannot be excluded, since the culture was sometimes brownish coloured. However, 3,5-dichlorocatechol was never detected by HPLC. Moreover, the SdpA synthesis was repressed in the presence of high concentration of 2,4-DCPP. On the one hand, its decrease after one day reveals a potential bottleneck because this enzyme catalyses the first step of biodegradation of the S-enantiomer of 2,4-DCPP. On the other hand, the decrease of this enzyme, which was confirmed by the measurement of specific activity in cell-free extracts (Table 4), may indicate that the enzymic release of the more toxic 2,4-DCP from 2,4-DCPP was reduced. A detailed analysis of the induction pattern and of the transcription is necessary to clarify if the decrease happens due to less stability of the enzyme or due to decreased transcription of the sdpA gene.



View larger version (35K):
[in this window]
[in a new window]
 
Fig. 2. 2D protein pattern of D. acidovorans MC1 after chemostatic growth on 2,4-DCPP (A, control) and growth on high residual concentrations of 2,4-DCPP (B). Boxed spots, more than twofold amplified proteins; circled spots, new proteins; diamonds, proteins diminished by at least 50 %; spots surrounded by dotted lines, proteins already induced during the substrate shift from pyruvate to 2,4-DCPP (see Fig. 1B).

 


View larger version (22K):
[in this window]
[in a new window]
 
Fig. 3. Intensities of selected proteins of D. acidovorans MC1 after 2,4-DCPP shock. Abscissa, time after 2,4-DCPP shock (h); ordinate, intensities are expressed as a percentage of total spots in a gel (%); error bars, SEM of duplicated experiments.

 


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 4. Intensities of isoforms of TfdD of D. acidovorans MC1 after 2,4-DCPP shock. Abscissa, time after 2,4-DCPP shock (h); ordinate, intensities are expressed as a percentage of total spots in a gel (%); error bars, SEM of duplicated experiments.

 


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 5. Enlarged regions of gels showing the isoforms of TfdD and TufA of D. acidovorans MC1 before (control) and after 2,4-DCPP shock.

 

View this table:
[in this window]
[in a new window]
 
Table 4. Activities of (R)/(S)-2,4-DCPP 2-oxoglutarate dioxygenases and chlorocatechol 1,2-dioxygenase during continuous growth on low and high concentrations of 2,4-DCPP at steady-state

Growth in the pH-auxostat was initiated by the addition of 15 mM 2,4-DCPP and 0·5 mM 2,4-DCP to cells pre-cultivated in a chemostat D=0·05 h–1 on 2,4-DCPP, which also represents the control of the experiment.

 
Impact of 2,4-DCPP on the synthesis of other proteins
Apart from DnaK, only a weak or no induction of the well-known stress proteins, for example GroEL, AhpC and SodA, in response to excess concentrations of 2,4-DCPP was observed. Although the data are contrary to the majority of reported results (van Dyk et al., 1994; Cho et al., 2000), they correspond with short-term experiments in D. acidovorans MC1 (Benndorf & Babel, 2002) and affirms our assumption that classical stress proteins play a minor role in adaptation to chemostress. In any case, there are many other proteins affected. Although some of them were identified as component E2 of 2-oxoglutarate dehydrogenase, asparaginase AspG (Davidson et al., 1977), elongation factor Tu (TufA) and hypothetical conserved excreted (periplasmic) protein (Table 3), their contribution for adaptation cannot be completely elucidated. For the identification of TufA and YceI, internal amino acid sequences had to be used because TufA was always amino terminal blocked and the amino terminal sequence of YceI alone gave no significant database hit.

As early as 1 h after the addition of high concentrations of 2,4-DCPP TufA was modified by post-translational modification. Its low molecular mass isoform began to disappear whereas its high molecular mass isoform appeared (Figs 3 and 5). TufA is an essential part of the protein synthesis apparatus, promotes the GTP-dependent binding of aminoacyl-tRNA to the A-site of ribosomes and seems to be implicated in protein folding and protection from stress, because it also shows chaperone activity in vitro (Caldas et al., 1998). More than one isoform of TufA was found in 2D databases of Escherichia coli (SWISS-2DPAGE, http://us.expasy.org/ch2d/) and Mycobacterium tuberculosis (European Bacteria Proteome Project, http://www.mpiib-berlin.mpg.de/2D-PAGE/EBP-PAGE/index.html). A proposed mechanisms for modification is proteolytic cleavage of TufA in E. coli as response to phage T4 infection (Georgiou et al., 1998) and in Salmonella typhimurium as response to starvation of phosphate (Adams et al., 1999). Furthermore, regulation of TufA in response to nutrient deprivation by methylation (Young & Bernlohr, 1991) or phosphorylation of TufA (Lippmann et al., 1993) was described in E. coli. Probably, TufA is proteolytically cleaved at its carboxy terminal end, because both isoforms are blocked at the amino terminal end as also described in E. coli (Jones et al., 1980) and a limited proteolysis seems to be more probable in bacteria than a glycosylation with an estimated mass of more than 2000 Da. However, our data are insufficient to identify the site and kind of modification of TufA in D. acidovorans MC1 as well as its effect on metabolism. As reported by other authors, proteome analysis often delivers much data (Blom et al., 1992) which are difficult to interpret, for example the bulk of unidentified spots (more than 60). In addition, the identification of such stress protein sometimes brings up more questions because proteins such as TufA, AspG, OdhB and YceI are involved in complex metabolic and regulatory networks.

Conclusion
Finally, proteome analysis allowed us to evaluate the response of D. acidovorans MC1 to 2,4-DCPP on a global level. One strategy of adaptation during growth on high residual concentrations of 2,4-DCPP could be the repression (SdpA) as well as the induction (TfdCII, TfdD) of catabolic enzymes probably providing resistance by lowering the concentrations of toxic intermediates. Classical stress proteins play a minor role. Furthermore, the modification of essential proteins such as TufA and the induction of the hypothetical periplasmic protein YceI indicate that further important mechanisms of resistance may exist. Their investigation is indispensable for a comprehensive understanding of the response of bacteria to chlorophenoxy herbicides.


   ACKNOWLEDGEMENTS
 
This work has been supported by a grant from the European Commission within its Fifth Framework Program (Project designation, HERBICBIOREM; Contract No., QLK3-CT-1999-00041). In addition, D. B. was supported by the OECD (Co-operative Research Programme, Biological Resource Management for Sustainable Agricultural Systems; Contract No. AGR/PROG/JA00012773).


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Adams, P., Fowler, R., Howell, G., Kinsella, N., Skipp, P., Coote, P. & O'Conner, C. D. (1999). Defining protease specificity with proteomics: a protease with a dibasic amino acid recognition motif is regulated by a two-component signal transduction system in Salmonella. Electrophoresis 20, 2241–2247.[CrossRef][Medline]

Benndorf, D. & Babel, W. (2002). Assimilatory detoxification of herbicides by Delftia acidovorans MC1: induction of two chlorocatechol 1,2-dioxygenases as a response to chemostress. Microbiology 148, 2883–2888.[Abstract/Free Full Text]

Benndorf, D., Loffhagen, N. & Babel, W. (1999). Induction of heat shock proteins in response to primary alcohols in Acinetobacter calcoaceticus. Electrophoresis 20, 781–789.[CrossRef][Medline]

Benndorf, D., Loffhagen, N. & Babel, W. (2001). Protein synthesis patterns in Acinetobacter calcoaceticus induced by phenol and catechol show specificities of responses to chemostress. FEMS Microbiol Lett 200, 247–252.[CrossRef][Medline]

Blom, A., Harder, W. & Matin, A. (1992). Unique and overlapping pollutant stress proteins of Escherichia coli. Appl Environ Microbiol 58, 331–334.[Abstract]

Blum, H., Beier, H. & Gross, H. J. (1987). Improved silver staining of plant proteins, RNA and DNA in polyacrylamide gels. Electrophoresis 8, 93–99.

Caldas, T. D., Yaagoubi, A. E. & Richarme, G. (1998). Chaperone properties of bacterial elongation factor EF-Tu. J Biol Chem 273, 11478–11482.[Abstract/Free Full Text]

Cash, P., Argo, E., Ford, L., Lawrie, L. & McKenzie, H. (1999). A proteomic analysis of erythromycin resistance in Streptococcus pneumoniae. Electrophoresis 20, 2259–2268.[CrossRef][Medline]

Cho, Y.-S., Park, S.-H., Kim, C.-K. & Oh, K.-H. (2000). Induction of stress shock proteins DnaK and GroEl by phenoxyherbicide 2,4-D in Burkholderia sp. YK-2 isolated from rice field. Curr Microbiol 41, 33–38.[CrossRef][Medline]

Davidson, L., Brear, D. R., Wingard, P., Hawkins, J. & Kitto, G. B. (1977). Purification and properties of L-glutaminase-L-asparaginase from Pseudomonas acidovorans. J Bacteriol 129, 1379–1386.[Medline]

Duxbury, J. M., Tiedje, J. M., Alexander, M. & Dawson, J. E. (1970). 2,4-D metabolism: enzymatic conversion of chloromaleylacetic acid to succinic acid. J Agric Food Chem 18, 199–201.[Medline]

Evans, W. C., Smith, B. S., Moss, P. & Fernley, H. N. (1971). Bacterial metabolism of 4-chlorophenoxyacetate. Biochem J 122, 509–517.[Medline]

Fukumori, F. & Hausinger, R. P. (1993). Purification and characterization of 2,4-dichlorophenoxyacetate/{alpha}-ketoglutarate dioxygenase. J Biol Chem 268, 24311–24317.[Abstract/Free Full Text]

Georgiou, T., Yu, Y.-T. N., Ekunwe, S., Buttner, M. J., Zuurmond, A.-M., Kraal, B., Kleanthous, C. & Snyder, L. (1998). Specific peptide-activated proteolytic cleavage of Escherichia coli elongation factor Tu. Proc Natl Acad Sci U S A 95, 2891–2895.[Abstract/Free Full Text]

Heipieper, H. J., Diefenbach, R. & Keweloh, H. (1992). Conversion of cis unsaturated fatty acids to trans, a possible mechanism for the protection of phenol-degrading Pseudomonas putida P8 from substrate toxicity. Appl Environ Microbiol 58, 1847–1852.[Abstract]

Holtzhauer, M. & Hahn, V. (1988). Biochemische Labormethoden: Arbeitsvorschriften und Tabellen, pp. 2–3. Berlin: Springer.

Horvath, M., Ditzelmüller, G., Loidl, M. & Streichsbier, F. (1990). Isolation and characterization of a 2-(2,4-dichlorophenoxy)propionic acid-degrading soil bacterium. Appl Microbiol Biotechnol 33, 213–216.[Medline]

Jin, Y. & Cerletti, N. (1992). Western blotting of transforming growth factor {beta}2. Optimization of the electrophoretic transfer. Appl Theor Electrophor 3, 85–90.[Medline]

Jones, M. D., Petersen, T. E., Nielsen, K. M., Magnusson, S., Sottrup-Jensen, L., Gausing, K. & Clark, B. F. (1980). The complete amino-acid sequence of elongation factor Tu from Escherichia coli. Eur J Biochem 108, 507–526.[Abstract]

Kamagata, Y., Fulthorpe, R. R., Tamura, K., Takami, H., Forney, L. J. & Tiedje, J. M. (1997). Pristine environments harbor a new group of oligotrophic 2,4-dichlorophenoxyacetic acid-degrading bacteria. Appl Environ Microbiol 63, 2266–2272.[Abstract]

Kaphammer, B., Kukor, J. J. & Olsen, R. H. (1990). Regulation of tfdCDEF by tfdR of the 2,4-dichlorophenoxyacetic acid degradation plasmid pJP4. J Bacteriol 172, 2280–2286.[Medline]

Kilpi, S. (1980). Degradation of some phenoxy acid herbicides by mixed cultures of bacteria isolated from soil treated with 2-(2-methyl-4-chloro)phenoxypropionic acid. Microb Ecol 6, 261–270.

Krayl, M., Benndorf, D., Loffhagen, N. & Babel, W. (2003). Use of proteomics and physiological characteristics to elucidate ecotoxic effects of methyl tert-butyl ether in Pseudomonas putida KT2440. Proteomics 3, 1544–1552.[CrossRef][Medline]

Lippmann, C., Lindschau, C., Vijgenboom, E., Schröder, W., Bosch, L. & Erdmann, V. A. (1993). Prokaryotic elongation factor Tu is phosphorylated in vivo. J Biol Chem 268, 601–607.[Abstract/Free Full Text]

Loffhagen, N., Hartig, C. & Babel, W. (1995). The glucose dehydrogenase-mediated energization of Acinetobacter calcoaceticus as a tool for evaluating its susceptibility to, and defence against, hazardous chemicals. Appl Microbiol Biotechnol 42, 738–743.[CrossRef][Medline]

Loffhagen, N., Hartig, C. & Babel, W. (1997). The toxicity of substituted phenolic compounds to a detoxifying and an acetic acid bacterium. Ecotoxicol Environ Saf 36, 269–274.[CrossRef][Medline]

Loffhagen, N., Härtig, C. & Babel, W. (2003). Energization of Comamonas testosteroni ATCC 17454 for indicating toxic effects of chlorophenoxy herbicides. Arch Environ Contam Toxicol 45, 317–323.[Medline]

Müller, R. H. & Babel, W. (1986). Glucose as an energy donor in acetate growing Acinetobacter calcoaceticus. Arch Microbiol 144, 62–66.

Müller, R. H., Jorks, S., Kleinsteuber, S. & Babel, W. (1999). Comamonas acidovorans strain MC1: a new isolate capable of degrading the chiral herbicides dichlorprop and mecoprop and the herbicides 2,4-D and MCPA. Microbiol Res 154, 241–246.[Medline]

Müller, R. H., Kleinsteuber, S. & Babel, W. (2001). Physiological and genetic characteristics of two bacterial strains utilizing phenoxypropionate and phenoxyacetate herbicides. Microbiol Res 156, 121–131.[Medline]

Oh, K. H. & Tuovinen, O. H. (1990). Degradation of 2,4-dichlorophenoxy acid by mixed cultures of bacteria. J Ind Microbiol 6, 275–278.

Pemberton, J. M. & Fisher, P. R. (1977). 2,4-D plasmids and persistence. Nature 268, 732–733.[Medline]

Peng, L. & Shimizu, K. (2003). Global metabolic regulation analysis for Escherichia coli K12 based on protein expression by 2-dimensional electrophoresis and enzyme activity measurement. Appl Microbiol Biotechnol 61, 163–178.[CrossRef][Medline]

Pieper, D. H., Reineke, W., Engesser, K.-H. & Knackmuss, H.-J. (1988). Metabolism of 2,4-dichlorophenoxyacetic acid, 4-chloro-2-methylphenoxyacetic acid, and 2-methylphenoxyacetic acid by Alcaligenes eutrophus JMP 134. Arch Microbiol 150, 95–102.

van Dyk, T. K., Majarian, W. R., Konstantinov, K. B., Young, R. M., Dhurjati, P. S. & LaRossa, R. A. (1994). Rapid and sensitive pollutant detection by induction of heat shock gene-bioluminescence gene fusions. Appl Environ Microbiol 60, 1414–1420.[Abstract]

Vasseur, C., Labadie, J. & Hébraud, M. (1999). Differential protein expression by Pseudomonas fragi submitted to various stresses. Electrophoresis 20, 2204–2213.[CrossRef][Medline]

Westendorf, A., Benndorf, D., Müller, R. H. & Babel, W. (2002). The two enantiospecific dichlorprop/{alpha}-ketoglutarate-dioxygenases from Delftia acidovorans MC1 – protein and sequence data of RdpA and SdpA. Microbiol Res 157, 317–322.[Medline]

Westendorf, A., Müller, R. H. & Babel, W. (2003). Purification and characterisation of the enantiospecific dioxygenases from Delftia acidovorans MC1 initiating the degradation of phenoxypropionate and phenoxyacetate herbicides. Acta Biotechnol 23, 3–17.[CrossRef]

Young, C. C. & Bernlohr, R. W. (1991). Elongation factor Tu is methylated in response to nutrient deprivation in Escherichia coli. J Bacteriol 173, 3096–3100.[Medline]

Received 18 September 2003; revised 24 November 2003; accepted 28 November 2003.



This Article
Abstract
Full Text (PDF)
Alert me when this article is cited
Alert me if a correction is posted
Citation Map
Services
Email this article to a friend
Similar articles in this journal
Similar articles in PubMed
Alert me to new issues of the journal
Download to citation manager
Google Scholar
Articles by Benndorf, D.
Articles by Babel, W.
Articles citing this Article
PubMed
PubMed Citation
Articles by Benndorf, D.
Articles by Babel, W.
Agricola
Articles by Benndorf, D.
Articles by Babel, W.


HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
INT J SYST EVOL MICROBIOL MICROBIOLOGY J GEN VIROL
J MED MICROBIOL ALL SGM JOURNALS
Copyright © 2004 Society for General Microbiology.