ABSTRACT
During batch growth of Ralstonia eutropha (previously named Alcaligenes eutrophus) on phenol in the presence of acetate, acetate was found to be the preferred substrate; this organic acid was rapidly metabolized, and the specific rate of phenol consumption was considerably decreased, although phenol consumption was not abolished. This decrease corresponded to a drop in phenol hydroxylase and catechol-2,3-dioxygenase specific activities, and the synthesis of the latter was repressed at the transcriptional level. Studies with a mutant not able to consume acetate indicated that the organic acid itself triggers the repression. Other organic acids were also found to repress phenol degradation. One of these, benzoate, was found to completely block the catabolism of phenol (diauxic growth). A mutant unable to metabolize benzoate was also unable to develop on benzoate-phenol mixtures, indicating that the organic acid rather than a metabolite involved in benzoate degradation was responsible for the repression observed.
Phenolic compounds are abundant in the biosphere as components of the complex polymer lignin, as humic acids, and as environmental pollutants resulting from various industrial activities. The bacterial degradation of structurally simple, readily degradable aromatic compounds has been studied with the hope that this will facilitate work on more recalcitrant members of the group. However, such studies have generally been performed under optimal laboratory conditions which do not necessarily reflect the complex situation in natural environments, in which growth is subject to a variety of nutritional and physicochemical constraints. One factor likely to modify the degradative potential of microorganisms is the presence of alternative carbon substrates. For substrates such as organic acids, Babel et al. (4) postulated that carbon sources in the environment may often be used simultaneously, although this mixotrophic behavior most likely predominates when only low (often limiting) carbon substrate concentrations are encountered. The repression of the catabolism of less favorable substrates by other carbon sources (often referred to as catabolite repression) has been extensively described for both enteric bacteria (6) and gram-positive bacteria (27). In the Pseudomonasgroup, catabolite repression control (CRC) has also been observed (18). This phenomenon has been shown to be cAMP independent (11, 17, 18, 24), but the mechanism is not yet understood.
So far, succinate has been used as a model compound for studying the catabolite repression of aromatic compounds. In Pseudomonas putida mt-2, succinate has been shown to repress the expression of the upper pathway for catabolism of toluene and xylene at the transcriptional level and, to a lesser extent, the metapathway (7, 8). More recently, Müller et al. (21) demonstrated that succinate also represses the transcription of the chromosomally encoded meta pathway (phl genes) of P. putida H. The regulatory effect of succinate on various catabolic pathway enzymes has now been established (7, 8, 23), but less attention has been paid to the role of other organic acids, such as acetate, on aromatic compounds despite the common occurrence of acetate in natural environments. In addition, little is known about catabolite repression in members of the beta subgroup of the Proteobacteria, such as Ralstonia eutropha (previously named Alcaligenes eutrophus[33]).
R. eutropha is capable of mineralizing phenol through a chromosome-encoded meta pathway (12, 13). The mechanism of molecular control of the expression of the metapathway of R. eutropha is not known; inPseudomonas sp. strain CF600, the dmp genes are under the control of a sigma 54 transcriptional activator (DmpR) which is directly regulated by effectors such as phenol and methylphenols (29, 30). Shingler and Moore (29) have also shown that benzoate does not activate the transcription of the dmpoperon in the presence of DmpR. However, it is not known whether benzoate can prevent the action of a natural effector (e.g., phenol) through, for example, inhibition of the DmpR-phenol interaction, when benzoate and a natural effector are simultaneously present in the cell. Recently, it was shown for P. putida H that CRC of phenol degradation is mediated by a negative controlling factor (21). The authors also suggested that the transcriptional activator, PhlR, is the target of this control. In R. eutropha, as in Pseudomonas spp., the metapathway results in the production of acetate and pyruvate, which are then metabolized by the central pathways. This interface is a logical site for regulatory control. The repression by acetate of a number of enzymes necessary for sugar assimilation by Pseudomonas aeruginosa is well documented (32). Hughes and Bayly (12) have reported that the biodegradation of phenol via themeta pathway is delayed in the presence of acetate inR. eutropha, but no physiological or mechanistic details are available. Only recently, acetate was shown to exert a repression effect on the promoters Ps and Pu in P. putida mt-2 (11), although the mechanistic basis for this remains obscure. On the other hand, the expression of acoE coding for acetyl coenzyme A (acetyl-CoA) synthetase was shown to be repressed by catechol during rapid growth on benzoate-acetate mixtures (1). This enzyme is the only pathway for acetyl-CoA formation from acetate in R. eutropha (31), and hence acetate consumption was blocked by benzoate.
This study was undertaken to clarify the growth behavior and mechanisms regulating phenol utilization in the presence of acetate or other organic acids.
MATERIALS AND METHODS
Bacterial strains, plasmids, and growth conditions. R. eutropha 335 (= ATCC 17697) was obtained from Laboratorium Microbiologie Gent (Brussels, Belgium), and strain B9, a benzoate mutant derived from strain 335 lacking 1,2-dihydro-1,2-dihydroxybenzoate (DHB) dehydrogenase (26), was kindly provided by George Hegeman. Escherichia coliCM990 harboring pUTluxAB, a Tcr and Apr transposon delivery plasmid for mini-Tn5 luxAB (15), and E. coli CM404 containing the Kmr helper plasmid RK600 (10) were kindly provided by Dirk Springael. E. coli XL1-Blue harboring plasmid pVI 1:00 as described by Bartilson and Shingler (5) was kindly provided by Victoria Shingler. pVI 1:00 is a pBluescript derivative with the dmpB gene coding forPseudomonas sp. strain CF600 catechol-2,3-dioxygenase inserted at the EcoRV site as aSmaI-HpaI fragment.
The mineral salts medium (MSM) used for growth of R. eutropha strains has been described previously (1) and was supplemented with various carbon substrates as described below. Cultures were grown in 2-liter bioreactors (working volume, 1.5 liters) with the pH and partial oxygen pressure maintained at 7 and 70% saturation, respectively, as previously described (2). Precultures were grown on phenol (5 mM). After inoculation, samples were periodically withdrawn from the bioreactors with sterile syringes for analytical procedures.
E. coli XL1-Blue(pVI 1:00) was grown on Luria-Bertani (LB) medium (28) supplemented with tetracycline (12.5 mg/liter) and ampicillin (100 mg/liter). E. coli CM990 and CM404 were grown on LB medium containing tetracycline (20 mg/liter) and kanamycin (100 mg/liter), respectively.
Transposon mutagenesis.Triparental matings were used to mobilize plasmid pUTluxAB into R. eutropha 335 as described by Kristensen et al. (15). To do this, the donor strain (E. coli CM990), the recipient strain (R. eutropha 335), and the helper strain (E. coli CM404) were grown separately overnight in LB medium (supplemented with antibiotics for the E. coli strains), washed with 10 mM MgSO4, mixed at a ratio of 1:5:1, and applied as 100-μl drops to LB agar plates. After 8 h, the cells were scraped off the surface, dispersed in 1 ml of 10 mM MgSO4, diluted as required, and plated onto MSM supplemented with gluconate (20 mM) and tetracycline (20 mg/liter) for counterselection of the donor and helperE. coli strains. Acetate-negative strains were then selected as strains that were not able to grow on MSM supplemented with 20 mM acetate and tetracycline.
Analytical methods.Biomass was measured by determining the cell dry weight.
The concentrations of substrates and products were determined by high-performance liquid chromatography (HPLC) with a model HP 1050 apparatus (Hewlett-Packard, Grenoble, France) equipped with an integrator (model HP 3396A) and an automatic sampler (model SP 8775; Spectra Physics France, Les Ulis, France). Detection was at 210 nm with a Hewlett-Packard series 1050 variable-wavelength detector. Separation was obtained with an AminexR HPX-87H column (300 by 7.8 mm; Chemical Div., Bio-Rad, Richmond, Calif.), and the operating conditions were as follows: temperature, 65°C; mobile phase, H2SO4 (5 mM)–CH3CN (7%, vol/vol); and flow rate, 0.8 ml/min.
Intracellular metabolites were extracted as previously described (2). The intracellular acetyl-CoA concentration was estimated by HPLC. The HPLC equipment described above was used for this. Separation was obtained with a C18 Nucleosil column (Touzart et Matignon, Courtaboeuf, France), and the operating conditions were as follows: temperature, 30°C; mobile phase, gradient consisting of sodium phosphate and CH3CN in sodium phosphate; and flow rate, 0.9 ml/min.
Determination of enzyme activities.Phenol hydroxylase (EC1.14.13.7; phenol, NADH:oxygen oxidoreductase [2-hydroxylating]) activity was estimated with whole cells that were obtained directly from the bioreactor, washed with 100 mM Tris-HCl (pH 7.5), and resuspended in the same buffer. The cells were placed in a biological oxygen monitor (model YSI 5300; Yellow Springs Instrument Co., Yellow Springs, Ohio) along with 3 ml of the same buffer containing 200 μM phenol. Blanks without phenol were prepared for each assay. Phenol hydroxylase activity was expressed as millimoles of O2consumed per gram of dry cells per hour. This method was adapted from that described by Farr and Cain (9).
For all other enzymes, cell extracts were prepared as follows. Approximately 50 to 100 mg (wet weight) of freshly harvested cells was washed twice in 100 mM Tris-HCl (pH 7.5) at 4°C and resuspended in 10 ml of Tris-carballylate buffer (pH 7.8) (9 mM tricarballylic acid, 35 mM Tris-HCl, 5 mM MgCl2, 20% [vol/vol] glycerol). The cells were disrupted by sonication, and the resulting crude extracts were centrifuged (15,000 × g, 20 min, 4°C) to obtain soluble extracts, which were used to assay enzyme activities. Acetyl-CoA synthetase (EC 6.2.1.1; acetate:CoA ligase [AMP forming]) activity was determined by the enzyme assay procedure of Oberlies et al. (22) in which the formation of AMP from ATP is monitored by coupling the reaction to the oxidation of NADH via adenylate kinase, pyruvate kinase, and lactate dehydrogenase. Blanks without CoA and ATP were prepared for each extract. Isocitrate lyase (EC 4.1.3.1) and malate synthase (EC 4.1.3.2) activities were determined at pH 7.5 by the procedures described by Maloy et al. (19). Catechol-2,3-dioxygenase (EC 1.13.11.2; catechol:oxygen 2,3-oxidoreductase) activity was assayed by the procedure described by Kataeva and Golovleva (14), except that the buffer was replaced by 100 mM Tris-HCl (pH 7.5). Blanks without substrate were prepared for each extract. Protein was determined as described by Lowry et al. (16). Activities are expressed below as milli-international units per milligram of protein (i.e., nanomoles per minute per milligram of protein).
dmpB mRNA probe preparation.Plasmid pVI 1:00 was extracted by using the rapid extraction procedure (28). After digestion with PstI, a 1,474-bp fragment containing the dmpB gene was eluted from the agarose gel by using a GeneClean kit (Stratagene, La Jolla, Calif.) and used as a probe. Priming was performed by the Mega-prime random priming method (28). A 20- to 50-ng portion of DNA was denatured for 5 min at 100°C and incubated for 2 h with [α32-P]dCTP.
Preparation of RNAs.RNAs were extracted by the sodium dodecyl sulfate (SDS)-EDTA procedure as previously described (1).
Dot blot analysis.RNAs were dotted onto a Hybond-N membrane (Amersham, Somerville, N.J.). Prehybridization was done for 2 h at 42°C in a solution containing 50% (wt/vol) formamide, 5× SSPE, 1× Denhardt’s solution, 1% (wt/vol) SDS, and 300 mg of tRNA per liter (1× SSPE is 0.15 M NaCl, 0.01 M NaH2PO4, and 0.37% [wt/vol] EDTA [pH 7.4]; 1× Denhardt’s solution is 0.2% Ficoll, 0.2% polyvinylpyrrolidone, and 0.2% [wt/vol] bovine serum albumin). The same solution was used for hybridization after 20 to 50 ng of the dmpB probe was added. After hybridization at 42°C for 15 h, the membrane was washed twice in 2× SSPE–0.1% (wt/vol) SDS at 37°C for 20 min and then twice in 0.2× SSPE–0.1% (wt/vol) SDS at 55°C for 25 min and subjected to autoradiography (28).
Chemicals.All chemicals were analytical grade. All substrates, enzymes, coenzymes, and radioistopes were obtained from Sigma Chimie (St. Quentin Fallavier, France) or C. F. Boehringer & Soehne (Mannheim, Germany).
RESULTS
Growth of R. eutropha on single substrates. R. eutropha was grown in bioreactors with acetate (5 or 60 mM) or phenol (5 mM) as a single carbon source. Growth on acetate was exponential; the specific growth rate (μ) was 0.38 ± 0.02 h−1, the specific rate of acetate consumption (q acetate) was 12.7 ± 1.0 mmol · g−1 · h−1, and a growth yield of 0.5 ± 0.06 g of biomass · g of acetate−1was observed throughout the experiment. Growth on phenol was not exponential due to the inhibitory effect of the aromatic substrate. As the residual phenol concentration decreased, both the μ and the specific rate of phenol consumption (q phenol) increased until maximal values of μ (0.36 ± 0.02 h−1) and q phenol (5.7 ± 0.2 mmol · g−1 · h−1) were attained. The growth yield was constant at 0.67 ± 0.1 g of biomass · g of phenol−1 throughout the experiment.
Growth of R. eutropha 335 (wild type) on a phenol-acetate mixture. (i) Kinetic analysis.Cells of R. eutropha 335 pregrown on phenol were transferred to a medium containing both acetate (60 mM) and phenol (5 mM). After a period of rapid acceleration, a high growth rate (μ > 0.3 h−1) was measured, although no true exponential phase was observed (Fig.1). During this rapid growth, theq acetate was high (>12.5 mmol · g−1 · h−1) and similar to that observed with acetate alone. On the other hand, theq phenol, although not equal to zero (it was ≈0.8 mmol · g−1 · h−1), was much lower than the value obtained when phenol was the only substrate. Only when acetate was almost totally degraded (at an acetate concentration of <5 mM) did the q phenolincrease.
Kinetics of growth and substrate consumption forR. eutropha 335 (wild type) grown in batch cultures on a phenol-acetate mixture. (A) ▪, biomass; ⧫, phenol; •, acetate. (B) ——, q phenol; –––,q acetate; ····, μ. The lines for substrate and biomass concentrations were based on interpolation (degree 4) of the raw data. The lines for μ,q acetate, and q phenolwere directly derived from the lines for substrates and biomass.
(ii) Enzymatic analysis.The activities of phenol hydroxylase and catechol-2,3-dioxygenase, the enzymes that catalyze the first two steps of the meta pathway for phenol degradation, were monitored throughout the culture described above (Fig.2). In addition, the activities of the enzymes specific to acetate catabolism (acetyl-CoA synthetase, malate synthase, and isocitrate lyase) were measured. From the onset of growth, the phenol hydroxylase activity decreased to a value close to 50% of the maximal activity measured with phenol alone (12 mmol of O2 · g−1 · h−1). A specific activity of 5 to 6 mmol of O2 · g−1 · h−1 was maintained until acetate was almost totally consumed. Finally, the phenol hydroxylase specific activity increased to its initial value. The catechol-2,3-dioxygenase specific activity followed the same pattern, but the residual activity measured while acetate was rapidly degraded was less than 10% of the maximal activity observed on phenol alone. This activity also increased toward the end of the growth of the culture. A different profile was observed for the enzymes involved in acetate catabolism. Acetyl-CoA synthetase, an enzyme also necessary for the catabolism of phenol, was present throughout the culture. The initial specific activity was 60 ± 5.8 mIU · mg of protein−1 and corresponded to the specific activity observed on phenol alone. It then increased to 100 to 120 mIU · mg of protein−1, values similar to the value observed during growth on acetate alone (1). On the other hand, the enzymes of the glyoxylate shunt (malate synthase and isocitrate lyase) were present at only low levels in the phenol-grown cells used to inoculate the culture, but were progressively induced so that they reached specific activities typical of those associated with acetate metabolism during the period of rapid acetate consumption.
Key enzymes for the catabolism of phenol and acetate during growth of the culture described in the legend to Fig. 1. Symbols: ○, phenol hydroxyase; □, catechol-2,3-dioxygenase; •, acetyl-CoA synthetase; ⧫, malate synthase; ◊, isocitrate lyase.
(iii) Transcription of the catechol-2,3-dioxygenase gene.Total mRNA was isolated from samples taken throughout the culture described above, as well as during rapid growth on phenol or acetate alone. Dot blots containing 20 ng of each RNA sample were hybridized with a Pseudomonas sp. strain CF600 catechol-2,3-dioxygenase gene (dmpB) heterologous probe, as Bartilston and Shingler (5) found very high levels of identity among the enzymes responsible for the meta cleavage of unsubstituted catechol in various pseudomonads; these results were confirmed by Moon et al. (20), who found levels of homology of 84 to 92% between the catechol-2,3-dioxygenase gene of Alcaligenes sp. strain KF711 and the xylE (TOL plasmid), nahH (NAH7 plasmid), and dmpB genes. Kinetic results showed that the rate of transcription of the R. eutrophacatechol-2,3-dioxygenase gene fell to a low level 15 min after the beginning of the culture, whereas high levels of transcription were detected when phenol was used as the single carbon source (Fig.3). No transcription of catechol-2,3-dioxygenase was detected when acetate was the only carbon source.
Transcription of the catechol-2,3-dioxygenase gene during growth of the culture described in the legend to Fig. 1. RNA isolated from cells grown on phenol alone was used as the control (lane C).
Intracellular acetyl-CoA concentrations were measured for cells grown on either phenol or the phenol-acetate mixture. In both cases, the level of acetyl-CoA was high, and the concentrations measured were not significantly different (166 ± 40 μmol · g−1 on phenol versus 218 ± 61 μmol · g−1 on phenol-acetate).
Study of R. eutropha mutants unable to grow on acetate. (i) Isolation of R. eutropha mutant strains.Organic acids are frequently claimed to be responsible for catabolite repression phenomena in pseudomonads, although it is not clear whether the organic acid itself or an intermediate in the central metabolism is responsible for the repression effect. In order to further characterize this phenomenon, we constructed and studied mutants of R. eutropha that are unable to grow on acetate as a single carbon source. Mutants were obtained by random transposon mutagenesis with a Tn5 derivative. Transconjugant clones were identified on minimal medium containing gluconate (20 mM) and tetracycline (20 mg/liter). The frequency of insertion of the mini-Tn5 luxABtransposon was 7 × 10−7. Seven clones were found to be unable to grow rapidly on acetate. Mutant T31 did not grow on 60 mM acetate. With 10 mM acetate, no growth was observed in the first 24 h. Then slow apparent growth (μ < 0.04 h−1) was observed, but microscopic observations revealed the presence of large polyhydroxybutyrate (PHB) granules in the cells, suggesting that the acetate degraded was transformed into polyhydroxybutyrate and not into true biomass. On the other hand, mutant T31 grew normally on gluconate or benzoate. T31 cells grown in the presence of acetate showed no detectable malate synthase activity, but isocitrate lyase and acetyl-CoA synthetase were expressed at levels similar to those found for strain 335 (wild type).
(ii) Growth of acetate mutants on phenol and a phenol-acetate mixture.When grown on phenol (5 mM) as a single carbon source, strain T31 behaved exactly like wild-type strain 335 (data not shown), showing that the glyoxylate shunt is not necessary for growth on phenol.
When cells of R. eutropha T31 pregrown on phenol were transferred to a medium containing both acetate (60 mM) and phenol (5 mM), growth was very slow and linear (Fig.4). The specific rate of phenol consumption rapidly decreased, whereas acetate was not significantly degraded. A 50% decrease in phenol hydroxylase specific activity was also observed, as well as a drop in the catechol-2,3-dioxygenase specific activity. No malate synthase was detected, and only low levels of isocitrate lyase (<5 mIU · mg of protein−1) were measured.
Kinetics of growth and substrate consumption forR. eutropha T31 (malate synthase mutant) grown in batch cultures on a phenol-acetate mixture. (A) ▪, biomass; ⧫, phenol; •, acetate. (B) ——, q phenol; ····, μ; ··–··, theoretical washing curve. The lines for substrate and biomass concentrations were based on interpolation (degree 4) of the raw data. The lines for μ andq phenol were directly derived from the lines for substrates and biomass.
Repression of phenol catabolism by other organic acids.The abilities of other organic acids to repress the catabolism of phenol were also tested. Two of these compounds, pyruvate and gluconate, did not repress phenol degradation. On the other hand, benzoate, fumarate,m-hydroxybenzoate, lactate, malate, and succinate strongly diminished the ability of strain 335 to catabolize phenol. Interestingly, benzoate (5 mM) was found to completely inhibit phenol degradation. Cells pregrown on phenol and transferred to a medium containing both phenol (5 mM) and benzoate (5 mM) showed a diauxic pattern of growth. During the first phase of growth, phenol degradation was halted, while benzoate was rapidly consumed; phenol was metabolized in the second phase only after benzoate had been totally degraded (Fig.5).
Kinetics of growth and substrate consumption forR. eutropha 335 (wild type) grown in batch cultures in the presence of phenol-benzoate (▪), phenol (⧫), or benzoate (•).
The behavior of a benzoate mutant of strain 335 on benzoate-phenol mixtures was then investigated. In R. eutropha, benzoate is degraded through the ortho pathway (13). The first steps of this pathway are benzoate → DHB (benzoate-1,2-dioxygenase) and DHB → catechol (DHB dehydrogenase). Strain B9 lacks DHB dehydrogenase and is unable to grow on benzoate (26), but its growth is identical to the growth of wild-type strain 335 on phenol. When cells of B9 pregrown on phenol were transferred to media containing phenol-benzoate and phenol-DHB mixtures (5 mM each), no growth was observed in the phenol-benzoate mixture after 48 h, although the growth with phenol-DHB was similar to that obtained with phenol alone (data not shown).
DISCUSSION
Previous work has shown that when R. eutropha is grown in the presence of both benzoate and acetate, benzoate is the preferred substrate and the catabolism of acetate is repressed at the transcriptional level (1). This goes against the commonly accepted idea that organic acids repress the degradation of aromatic compounds. Additional studies provided evidence that this mechanism is not the only mechanism that governs carbon source utilization inR. eutropha; there seem to be several mechanisms which control the hierarchy of substrate utilization (3). One substrate, acetate, was shown to inhibit the catabolism of phenol (3, 12), but until now this inhibition had not been described in detail, and no information on the mechanism responsible for this phenomenon was available. In this work, we showed that acetate blocks the synthesis of phenol hydroxylase, as well as the synthesis of catechol-2,3-dioxygenase. Acetyl-CoA is not responsible for this repression, as high intracellular concentrations were found in both phenol-grown and phenol-acetate-grown cells. Furthermore, mRNA analysis demonstrated that, at least in the case of catechol-2,3-dioxygenase, the repression mechanism operates at the transcriptional stage. This repression can be extended to all of the enzymes of the metapathway as they appear to be organized in a single operon in R. eutropha (12), as they are in other pseudomonads (25). Our results also indicate that although acetate effectively represses phenol degradation, this repression is not total; in the presence of acetate, phenol continues to be degraded slowly. A 50 to 60% repression of phenol hydroxylase specific activity was observed, whereas catechol-2,3-dioxygenase was repressed by more than 90%. These quantitative results are in contrast to those reported previously by Hughes and Bayly (12), who claimed that they observed total repression of phenol hydroxylase and a 50% decrease in the activity of the other enzymes of the meta pathway inR. eutropha, without giving any precise explanation of the kinetics of this repression. The phenol hydroxylase assay that we used in this study included both the transport and the oxidation of phenol (see above). The activity measured with phenol-acetate-grown cells was around 5 to 6 mmol · g−1 · h−1(50 to 60% of the maximal activity measured with phenol alone); since this value was higher than the experimentalq phenol measured, this step cannot be a limiting step in phenol catabolism. In addition, we measured phenol hydroxylase activities with the protocol described above in the presence of various organic acids; this enzyme was not inhibited by any of the organic acids tested (data not shown). In light of this, we concluded that phenol catabolism is not blocked by acetate through transport inhibition.
Experiments performed with a malate synthase mutant unable to grow on acetate showed that rapid growth is not necessary to provoke repression of the meta pathway. The results of such experiments also suggest that the organic acid itself is the signal (repressor) triggering the transcriptional repression. The hypothesis that there is a direct action by the organic acid molecule is further supported by the observation that other organic acids also repressed phenol degradation. However, not all of the acids tested had the same repressive effect (e.g., benzoate completely blocked phenol catabolism, whereas in the presence of acetate, phenol was still slowly degraded). In addition, some organic acids (e.g., pyruvate) had no effect on phenol degradation; these results were in agreement with those published by Müller et al. (21), who found that succinate repressed the expression of the meta pathway ofP. putida, whereas pyruvate did not. So far, no logical explanation has been found in terms of acid strength or pKa to explain this variation in response. In the case of benzoate-phenol mixtures, the total repression observed (diauxic growth) can be attributed to the presence of benzoate.
The results show that there are several mechanisms which lead to catabolite repression phenomena in pseudomonads when an organic acid and an aromatic compound are simultaneously present. To some extent, this effect is correlated with the pathways involved and, as would be expected, with the growth rates obtained with each substrate. So far, all experiments performed with R. eutropha or P. putida have shown that CRC in pseudomonads is exerted at the transcriptional stage and suggest that (i) there are common regulatory mechanisms in these species and (ii) a hierarchy of substrate utilization rules carbon source preferences.
ACKNOWLEDGMENTS
We thank the Institut National de la Recherche Agronomique, the Centre National de la Recherche Scientifique, and the French Midi-Pyrénées region for financial support.
FOOTNOTES
- Received 11 August 1997.
- Accepted 4 October 1997.
- Copyright © 1998 American Society for Microbiology
