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Génétique Microbienne, Institut National de la Recherche Agronomique, 78352 Jouy-en-Josas cedex, France,1 Groupe de Recherches Métaboliques, Christian de Duve Institute of Cellular Pathology, 75 av. Hippocrate, B-1200 Brussels, Belgium,2 Department of Molecular Genetics, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Kerklaan 30, 9751 NN Haren, The Netherlands3
Received 5 November 2004/ Accepted 2 March 2005
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ABSTRACT |
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INTRODUCTION |
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Although non-PTS uptake systems for fructose utilization have been described (5), sugar-specific PTS appear to be the most frequent system of fructose utilization. Several fructose (fru) operons encoding EIIFru enzyme and 1-phosphofructokinase have been described in different bacterial groups, such as Spiroplasma citri (a mollicute), Streptococcus mutans, and Streptococcus gordonii (firmicutes). In the first bacterium, the fru operon has been shown to be involved in phytopathogenicity of S. citri, the causal agent of the citrus "stubborn" disease (11). In the two latter bacteria, high-affinity sugar utilization systems such as the PTSFru enhance survival of oral streptococci during periods between meals, while acid production from sugar contributes directly to human tooth decay. Moreover, it was shown that the fru operon is involved in biofilm formation by S. gordonii (19), a process allowing bacterial accumulation, proliferation, and persistence on oral surfaces. Interestingly, in these distantly related bacteria, the two fructose utilization genes are preceded by a regulator of the DeoR repressor family. The three genes were shown to be cotranscribed from an upstream promoter. In silico analysis shows that the genetic organization of most fructose utilization operons is the same in different genera, such as Bacillus, Staphylococcus, Lactococcus, Enterococcus, Lactobacillus, Streptococcus, Streptomyces, Corynebacterium, Clostridium, and Fusobacterium (our personal analysis, but most information can be retrieved at http://theseed.uchicago.edu or http://string.embl.de).
The regulation of the fru operon and the role of the FruR regulator have been investigated in S. citri and S. gordonii (12, 19). The transcription of these operons is enhanced by the presence of fructose in the culture medium. Surprisingly, FruR was found to be an activator in S. citri, whereas it was shown to be a repressor in S. gordonii (12, 19). This difference could not be explained from the sequence analysis of the two proteins, which share more than 35% identity (55% similarity) over their entire length. Moreover, putative motifs of regulation were suggested in both cases, but there are no experimental data available to confirm their involvement in fru operon transcription. Finally, the signal sensed by FruR has not been identified.
In this paper, we present a study of the fru operon from Lactococcus lactis. The genes were initially annotated lacR, lacC, and fruA (2), as the products encoded by the first two genes share high identity to the repressor-of-lactose-utilization operon and tagatose-6-phosphate kinase from L. lactis, respectively (28). We show that fruR (lacR), fruC (lacC), and fruA are involved in the main fructose utilization pathway in L. lactis. Furthermore, we present evidence that FruR is the repressor of the fructose operon and that its activity is modulated by fructose-1-phosphate. In this regulatory process, FruA, encoding a potential EIIABCFru PTS unit, is necessary to produce fructose-1-phosphate, while FruC, encoding a putative 1-phosphofructokinase, plays an indirect role in fructose operon regulation even in the absence of fructose in the culture medium. The specificity of FruR regulation in L. lactis was shown by DNA microarrays, and the DNA-binding site for FruR was identified by genetic experiments. Finally, comparative genomics analysis indicates that the L. lactis fructose regulation model may exist in many low-GC gram-positive bacteria.
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MATERIALS AND METHODS |
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DNA manipulation procedures. Procedures for DNA manipulations, transformation of E. coli cells, and cloning were done essentially as described elsewhere (21). Electrotransformation of L. lactis was carried out as described previously (15). Southern hybridization and detection were performed according to the Amersham ECL protocol (Amersham, Freiburg, Germany). DNA was sequenced on both strands using the 370A DNA sequencer according to the manufacturer's instructions.
Construction of lux transcriptional fusions and negative mutants in L. lactis. The integrative plasmids used to obtain strains carrying lux transcriptional fusions and the inactivated gene(s) are described in Table 1. Briefly, PCR products containing the PfruR promoter or an internal fragment of the gene to be inactivated were cloned in the pGEM-T vector in E. coli and their sequences were verified. The specific primers used in this work are listed in Table 2. The integrative plasmids were obtained by fusing the derived pGEM-T vectors with the L. lactis integrative vector pJIM2374 at the SalI restriction sites. Out of the two possible plasmids obtained by this method, we chose those where the luciferase gene of pJIM2374 and the fru gene insert have the same orientation. In some constructions, the pGEM-T vector was removed to allow transcription of the downstream genes from the erythromycin gene promoter. These plasmids were integrated in the chromosome of L. lactis by a single crossover event with pJIM1276 as a helper plasmid (14). The resulting strains were verified by PCR amplification and Southern blotting. The strains JIM8233 and JIM8253 contained the luxAB genes downstream of the cloned promoter region followed by a copy of the intact gene (Fig. 1). The strains JIM8234 to -8240 and JIM8245, JIM8246, and JIM8254 contained the duplicated target gene deleted either at its 3' end or at its 5' end (Fig. 1). The strains JIM8641 to -8644 contained luxAB downstream of the intact promoter region followed by the fru operon transcribed from a modified promoter (see Fig. 4, below).
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Fructose-1-phosphate assays on perchloric acid extracts. Cells were grown in CDM containing appropriate sugars at a final concentration of 0.5%. For each condition, two independent cultures were carried out. When cultures reached an OD600 of 0.3 to 0.4, they were rapidly chilled to 4°C in an ethanol-dry ice bath and subsequently harvested by centrifugation. Pellets were resuspended in 2 volumes of ice-cold 20% perchloric acid solution. The perchloric acid extracts were neutralized by the addition of 2.5 volumes of tri-n-octylamine-CH3Cl (1:3.6 [vol/vol]) mixture according to the methods of Khym (16). After centrifugation, supernatants were extracted with the same volume of CH3Cl and centrifuged. This neutralization step was repeated twice. The final supernatants were kept at –80°C until further analysis. Assays of fructose-1-phosphate were carried out using recombinant purified E. coli 1-phosphofructokinase according to the methods described by Veiga-da-Cunha et al. (30).
RNA isolation. Cells were grown in CDM containing appropriate sugars until the OD600 reached 0.3 to 0.4. They were quickly centrifuged for 2 min at 6,000 x g, frozen in liquid nitrogen, and broken using 500 mg of glass beads, 500 µl of phenol-chloroform, 30 µl of 3 M sodium acetate, and 15 µl of 20% sodium dodecyl sulfate. RNAs were isolated using the High Pure RNA isolation kit (Roche, Mannheim, Germany) according to the manufacturer's instructions.
RT-PCR and 5'-RACE. Reverse transcriptase PCR (RT-PCR) was carried out on 500 ng of total RNA with the OneStep RT-PCR kit (QIAGEN, Courtaboeuf, France) as recommended by the manufacturer, using primers fruA2 and lacR2. Reaction conditions were a reverse transcription of 30 min at 45°C; an initial PCR activation step of 15 min at 95°C; 25 cycles of 10 s at 94°C, 50 s at 50°C, and 3 min at 68°C; and a final extension step of 10 min at 68°C. The 5'/3' RACE kit (Roche) was used according to the supplier's instructions. A 1.5-µg aliquot of total RNA was used to obtain the cDNA by primer extension with primer lacC2. Following the 3' tailing reaction with a dATP string, the cDNA was amplified by PCR using the reverse primer lacR3 and the forward primer [oligo(dT)-anchor primer] supplied with the kit. The 5' end of the transcript was then determined by sequencing the PCR product.
QRT-PCR. Differential expression of genes was checked by real-time quantitative RT-PCR (QRT-PCR). De novo cDNAs were prepared as described previously. One microliter of 40-fold-diluted cDNAs was used for each 25-µl PCR mixture containing 1x Absolute QPCR SYBR Green ROX (ABgene, Epsom, Surrey, United Kingdom) and a 200 nM concentration of each primer. All reactions were carried out in duplicate using an ABI Prism 7700 sequence detector (Applied Biosystems, Foster City, Calif.) with the following cycle parameters: one cycle of 95°C for 10 min, followed by 40 cycles of 95°C for 15 s and annealing and extension at 60°C for 60 s. Prior to comparative analysis, each primer pair was tested to determine its efficiency using a genomic DNA scale. The efficiency of the primer pairs was in the range of 80% to 100%. Results were calculated from at least two independent RNA extractions for which measures by QRT-PCR were carried out in duplicate. The tuf gene was used as an internal control with the tuf1 and tuf2 primers.
Microarray experiments and analysis. DNA microarrays contained 2,126 L. lactis IL1403 gene PCR products spotted in duplicate to glass slides as previously described (18). Single-strand reverse transcription and labeling of 20 µg of total RNA were done using the Superscript 3 reverse transcriptase (Invitrogen, Cergy Pontoise, France) and the Amersham CyScribe labeling kit according to the manufacturers' instructions. Slides were prehybridized for 1 h in 4x SSC (1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate), 0.1% sodium dodecyl sulfate, and 0.1 mg ml–1 bovine serum albumin. After removal of the prehybridization buffer, slides were hybridized for 16 h at 42°C in SlideHyb buffer I (Ambion, Huntingdon, Cambridgeshire, United Kingdom) containing Cy3/Cy5-labeled cDNA mix. All comparisons were performed at least twice (once each with Cy3 and Cy5) to check for possible differences in labeling efficiency between fluorophores. Slides were scanned using a confocal laser scanner (Virtek Chipreader, Virtek, Waterloo, Ontario, Canada). Fluorescent signal intensity data were quantified using Imagene 5.5 software (Biodiscovery, Los Angeles, Calif.). Each expression ratio was represented by at least four separate measurements (duplicate spots on each of two slides). The data sets were normalized, and a statistical analysis (z-test) was done using the PreP software (10). Genes having a threefold ratio and a P value of <0.001 were considered to be differentially expressed. DNA microarray data are available on the website http://genome.jouy.inra.fr/efp/base/www.
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RESULTS |
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Role of FruC and FruA in fructose catabolism. In order to determine the role of fruC and fruA, the two genes were inactivated by integration into the chromosome of plasmids derived from pJIM2374 (Fig. 1C). The growth patterns of the resulting strains were followed in CDM containing different sugars: glucose, mannose, trehalose, maltose, or fructose. The strains JIM8240, JIM8236, and JIM8235 inactivated for fruC, fruA, and fruCA, respectively, did not grow in CDM containing 0.5% fructose, whereas their growth on other sugars was unaffected (data not shown). Moreover, API 50 CH profiles, allowing the determination of the fermentation for 22 sugars, showed that the mutants were only affected in fructose metabolism (data not shown). We conclude that FruC and FruA are specifically involved in fructose catabolism.
To test if another lower-affinity pathway for fructose catabolism is present in L. lactis, the fruC or fruA mutants were grown in a medium containing a higher fructose concentration (2%). The high fructose amount allowed weak growth of both mutants (doubling time sixfold higher than the wild-type strain), suggesting that an alternative route for fructose catabolism exits.
Regulation of the expression of the fructose utilization operon. (i) Regulation by FruR. In order to determine the role of the potential regulator FruR in the expression of the fru operon, fruR was inactivated by integration of plasmids derived from pJIM2374 carrying the luxAB genes (strains JIM8234 and JIM8239) into the bacterial chromosome (Fig. 1). In the wild-type strain (JIM8233) and the mutant strains, the expression of the fru operon can be quantified by measuring luciferase activity encoded by the luxAB genes. In CDM containing trehalose or glucose, the luciferase activity was, respectively, 160-fold and 100-fold higher in JIM8239 than in the wild-type strain, JIM8233 (Table 3). Similar results were obtained with strain JIM8234, although it did not grow in medium containing fructose as the sole carbon source, which is likely to be due to a polar effect of pGEMt (Table 3). The strong expression of the promoter fusion in the two fruR mutants compared to the wild type suggests that FruR acts as a repressor.
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Interestingly, the disruption of fruC, encoding the 1-phosphofructokinase (strain JIM8240), resulted in the induction of the expression of the operon independently of the sugar present in the growth medium (Table 3). Since this pattern of expression is similar to that of a fruR mutant, we have verified that the inactivation of fruC does not affect the expression of fruR in cis. For this purpose the plasmid pJIM5517, expressing fruR, was introduced in different genetic backgrounds. This plasmid was able to restore a wild-type repressing activity when introduced in the fruR background of JIM8245 (Table 3). Clearly, the introduction of pJIM5517 in the fruC background of JIM8246 did not suppress the constitutive high expression from the fru operon promoter. This result shows that the induction observed in JIM8240 is not due to fruR mis-expression but to the lack of activity of its product, FruR.
Furthermore, the inactivation of fruC could lead to an accumulation of FruR effector in the cell and thus to the inactivation of its repressor activity. To test this hypothesis, the intracellular concentration of fructose-1-phosphate was measured. The fructose-1-phosphate content of cells grown in glucose was 156 and 35 nmol of fructose-1-phoshate/g (dry weight) in JIM8240 (fruC) and in JIM8233 (wild type), respectively. A similar result was obtained in trehalose-grown cells, where the level of fructose-1-phospate was of 156 nmol/g in the fruC mutant, against 73 nmol/g in the wild-type strain. Thus, it appears that a basal expression of FruC is required to degrade fructose-1-phosphate, which is likely generated by spontaneous dephosphorylation of the fructose-1,6-bisphosphate coming from the glycolysis pathway or a side activity of a dephosphorylating enzyme with a phosphatase-like activity that remains to be identified. The induction of the fru operon in the fruC background leading to an increase of fructose-1-phosphate in the cell is a supplementary argument supporting that fructose-1-phosphate is the FruR effector.
(iii) Catabolite repression by CcpA. Expression of sugar catabolic genes is usually repressed by glucose in the medium. Part of this repression is mediated by the CcpA repressor in low-GC gram-positive bacteria. In the case of the fru promoter, the presence of glucose in CDM supplemented with fructose resulted in a 13-fold decrease of luciferase activity (Table 3), suggesting that expression of the fru operon was down-regulated by catabolite repression. To test whether the glucose repression is CcpA dependent, the transcriptional fusion between the promoter of the fru operon and luxAB genes was compared in a ccpA background and in a wild-type background. In the presence of an intact fruR gene, the luciferase activity in strains JIM8233 (wild type) and JIM8253 (ccpA) remained low and almost identical for all the sugars tested except fructose (Table 3). However, in a fruR background, the luciferase activity increased sixfold when ccpA was inactivated (JIM8239 versus JIM8254) in glucose-supplemented CDM. These results indicate that CcpA effectively represses the expression of the fru operon in the presence of glucose.
Specificity of FruR regulation at the genome scale. To analyze whether genes other than those of the fru operon are regulated by FruR, we determined genome-wide expression profiles, using total RNAs extracted from the fruR and fruC mutants (JIM8239 and JIM8240, respectively). Since the presence of glucose in the medium could interfere with the expression of sugar utilization genes, this experiment was performed in CDM with trehalose. The only genes that appeared to be differentially expressed relative to the wild-type IL1403 strain were those from the fru operon (Fig. 2). A similar result was obtained by comparing the fructose-1-phosphate-accumulating fruC (JIM8240) mutant to the wild-type strain (Fig. 2). Therefore, it appears that the FruR regulatory target is unique in L. lactis and that accumulation of fructose-1-phosphate has no effect other than modulating the regulatory activity of FruR.
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To examine whether other genes might be regulated by FruR, we searched for a motif, TGDNWRDWWDTKRWWDNWWDTGVNDRDNWWTGVNNGWNWD, obtained by the alignment of the potential FruR motifs of 12 low-GC gram-positive bacteria (Fig. 3), in all potential promoter regions of the corresponding genomes. No motifs other than those upstream of the fructose operon were found, suggesting a high specificity of FruR regulation in these bacteria.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The involvement of FruC and FruA in fructose catabolism was confirmed by the growth defect of the corresponding mutants in a fructose-containing medium. However, at a high fructose concentration, a weak growth of the mutants was observed, possibly due to the lack of specificity of other sugar transport systems present in L. lactis. A similar observation was reported for E. coli (17), where a second route for fructose utilization involves the PTS and membrane-spanning proteins that recognize a variety of sugars possessing the 3,4,5-D-arabino-hexose configuration. Fructose metabolism via this route is only observed when fructose is supplied in large amounts and, in this case, fructose-6-phosphate was formed instead of fructose-1-phosphate. In a number of streptococci, the mannose PTS was reported to catalyze the transport and the phosphorylation of glucose, mannose, 2-deoxyglucose and, to a lesser extent, fructose (7, 27). In L. lactis, a mannose PTS is present in the genome (2) and could be an alternative pathway for the transport of fructose in L. lactis.
The expression of the fru operon is repressed in a medium containing glucose. Such regulation, generally found for most sugar utilization operons, is designated catabolite repression and leads to the preferred use of highly metabolizable sugars. In low-GC gram-positive bacteria, it is exerted at the transcriptional level by the regulator CcpA and at the cellular level by inducer expulsion-exclusion (4). Our expression assays indicate that the fru operon of L. lactis is regulated directly at the transcriptional level by CcpA. This result is supported by the presence of a CRE box overlapping the site +1 of transcription in the promoter region of the operon. However, the level of expression in CDM-glucose is increased only 4-fold by a ccpA mutation, versus 100-fold for a fruR mutation, confirming the predominant role of FruR over CcpA in the regulation of the fru operon. Moreover, a very strong level of control is still exerted by glucose in the ccpA mutant, leading to a 30-fold decrease of luciferase activity when glucose is added to CDM-fructose. The most efficient level of catabolite control is thus mediated at the metabolic level, possibly by inducer exclusion, since the only way to degrade the fructose-1-phosphate appears to be its metabolism toward glycolysis (see the discussion below).
In L. lactis, FruR is the main repressor of the fructose operon. This level of regulation is directly dependent on the presence of fructose in the medium, as shown by the total lack of modulation of luciferase activity in the fruR mutant. In S. gordonii, FruR was also shown to act as a repressor of fructose metabolism (19). It is thus somewhat surprising that FruR from the mollicute S. citri might be an activator (12), as the two proteins have similar structure as shown by multiple alignments with all FruR proteins completed by secondary structure prediction searches (ClustalW and Predator). FruR of S. citri was proposed to bind to two repeats overlapping the –35 box of the fructose operon promoter, a location that usually leads to steric hindrance, making the promoter inaccessible to RNA polymerase. This apparent contradiction could be resolved if the mutant form of the S. citri FruR were trans-dominant over the wild type, as has been described for certain AgaR repressor mutants (24).
Interestingly, not only the fruR mutant but also fruA and fruC mutants are affected in the transcriptional regulation of the fructose operon promoter. Indeed, a nonpolar fruA mutation impairs growth on fructose and also leads to a constitutive low expression of the fru operon, even in the presence of fructose in the medium. This result suggests that uptake and phosphorylation of fructose to fructose-1-phosphate by FruA is necessary to relieve repression by FruR. The analysis of the fruC mutant, impaired in 1-phosphofructokinase activity, confirms this hypothesis. This mutant, which is also unable to grow on fructose as the sole carbon and energy source, displays a constitutive high expression of the fru operon, independent of the fructose content of the medium. Fructose-1-phosphate accumulates in the fruC mutant, even in a medium depleted of fructose. This constitutive induction also occurs in the double fruCA mutant (polar fruC mutation), a result that rules out the possibility that the induction is due to contamination of the medium by fructose. The lack of induction in the fruA mutant and its constitutive induction in the presence of fructose-1-phosphate indicate that fructose-1-phosphate modulates the DNA-binding activity of FruR. This agrees well with the observation that the DNA-binding activity of regulators of the DeoR family proteins is usually regulated by sugar-phosphates produced by the regulated pathway. For example, LacR from the lac operon in L. lactis is induced by tagatose-6-phosphate (29). In the case of fructose metabolism, fructose-1-phosphate is the only specific intermediate that could be used by the cell for this purpose, since this pathway branches to glycolysis in the next step. Lastly, the analysis of the fruC mutant shows that fruC plays a crucial role in the regulation of the fructose operon of L. lactis. Indeed, no other enzyme appears to be able to dephosphorylate fructose-1-phosphate, either formed by FruA or present as an offshoot of glycolysis. It follows that a basal level of FruC is necessary in the cell to degrade the inducer and mediate catabolic control of the fructose operon by inducer exclusion and transcriptional control by CcpA.
The repression by FruR appears to depend on the presence of four repeats of 10 bp. This structure, well conserved in various low-GC gram-positive bacteria, has a consensus motif (TGAWWGWTTT)4. The deletion of two repeats completely prevents repression. Mutations in the first two bases of the consensus (TG
AA) also abolish the repression by FruR. These two bases are almost perfectly conserved, since they are present in 48/48 and 47/48 of the analyzed sequences, respectively (Fig. 3A). Another member of the DeoR family of regulators, LacR from L. lactis, binds to two operators localized in lacR and lacABCDFEGX promoters (framed in Fig. 3B) (29). Two repeats of a 10-bp consensus (TGTTTNWTTT) are present in the lacR and lacA promoters of a set of various gram-positive bacteria (Fig. 3B). This consensus shares similarities with the FruR consensus, suggesting that the two members of the DeoR family recognize similar DNA sequences. This hypothesis is enforced by the similarities of the second helix of the FruR and LacR helix-turn-helix, which is known to be directly involved in the recognition and contact of the DNA target region in the DeoR family regulators (data not shown) (3).
In S. gordonii, fructose metabolism through the PTSFru pathway appears to be essential for biofilm formation, as inactivation of either 1-phosphofructokinase or fructose-specific enzyme II leads to a biofilm-defective phenotype (19). It was suggested that the fructose operon could be involved in a sensory mechanism enabling the switch from a sessile to a planktonic phenotype or that 1-phosphofructokinase plays a role in cross-regulation of a two-component system by phosphorylation of a response regulator. A wider set of genes is thus suspected to be regulated by the fructose system in this bacterium. Our analysis has failed to identify other genes that could be regulated by FruR in completely sequenced genomes of low-GC gram-positive bacteria.
It is interesting that in enteric bacteria, transcription of the fru operon is regulated by a pleitropic regulator, Cra (formerly named FruR), and to a lesser extent by the cyclic AMP-CRP complex (9, 25). Cra represses the expression of genes encoding glycolytic enzymes (i.e., key enzymes in the Embden-Meyerhof and Entner-Doudoroff pathways) and activates expression of genes encoding biosynthetic and oxidative enzymes (i.e., key enzymes in the Krebs cycle, the glyoxalate shunt, the gluconeogenic pathway, and electron transfer) (23, 25). Furthermore, the effect of Cra on transcription is counteracted by a micromolar concentration of fructose-1-phosphate (22). In order to detect possible side effects due to fructose metabolism or its regulation in L. lactis, DNA microarray experiments were carried out. The inactivation of fruC leads to an increase in expression of the fru operon comparable to that due to fruR inactivation, which is most probably a consequence of an elevated content of fructose-1-phosphate in the cells. However, the effect of fruC or fruR inactivation appeared to be restricted to the fru operon under the conditions tested. Moreover, addition of fructose to CDM containing trehalose specifically induces the fru operon and marginally represses the expression of trehalose and some other sugar utilization genes in the wild-type IL1403 strain (data not shown). Thus, metabolism of fructose does not appear to trigger other signaling pathways acting on other genes in L. lactis. The involvement of fructose metabolism as a sensory mechanism in biofilm formation may thus be either specific to S. gordonii or, more likely, the result of a metabolic effect rather than a sensory role.
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ACKNOWLEDGMENTS |
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This work was supported by grant QLK3-2001-010473 under the EU sub-program area "Quality of Life and Management of Living Resources—The Cell Factory."
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FOOTNOTES |
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REFERENCES |
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