- Research article
- Open Access
Extracytoplasmic function (ECF) sigma factor σF is involved in Caulobacter crescentus response to heavy metal stress
© Kohler et al.; licensee BioMed Central Ltd. 2012
Received: 4 July 2012
Accepted: 10 September 2012
Published: 18 September 2012
The α-proteobacterium Caulobacter crescentus inhabits low-nutrient environments and can tolerate certain levels of heavy metals in these sites. It has been reported that C. crescentus responds to exposure to various heavy metals by altering the expression of a large number of genes.
In this work, we show that the ECF sigma factor σF is one of the regulatory proteins involved in the control of the transcriptional response to chromium and cadmium. Microarray experiments indicate that σF controls eight genes during chromium stress, most of which were previously described as induced by heavy metals. Surprisingly, σF itself is not strongly auto-regulated under metal stress conditions. Interestingly, σF-dependent genes are not induced in the presence of agents that generate reactive oxygen species. Promoter analyses revealed that a conserved σF-dependent sequence is located upstream of all genes of the σF regulon. In addition, we show that the second gene in the sigF operon acts as a negative regulator of σF function, and the encoded protein has been named NrsF (Negative regulator of sigma F). Substitution of two conserved cysteine residues (C131 and C181) in NrsF affects its ability to maintain the expression of σF-dependent genes at basal levels. Furthermore, we show that σF is released into the cytoplasm during chromium stress and in cells carrying point mutations in both conserved cysteines of the protein NrsF.
A possible mechanism for induction of the σF-dependent genes by chromium and cadmium is the inactivation of the putative anti-sigma factor NrsF, leading to the release of σF to bind RNA polymerase core and drive transcription of its regulon.
Several heavy metals play important roles as trace elements in the metabolism of all kingdoms of life. Whether a trace element is useful or harmful depends on its concentration. Particularly, chromium and cadmium are known to be much more toxic than useful for most microorganisms [1, 2]. Chromium is commonly present in solutions as chromate and dichromate oxyanions (Cr(VI)), the most redox-reactive and soluble forms of the metal . Due to its similar chemical structure to sulfate anions, chromate crosses membranes via sulfate uptake systems . On the other hand, cadmium is a non-redox-reactive metal with high affinity for thiol groups [1, 2]. Once inside cells, chromate, dichromate and cadmium exert their toxic effects by directly damaging cellular components and by inducing oxidative stress [1, 2].
In order to reduce the toxicity of chromate, dichromate and cadmium, some microorganisms eliminate these metals from the cytoplasm by using active transport efflux pumps [1, 2]. Cadmium can also be sequestered within the cells by metal-chelating proteins, while chromate and dichromate are reduced to the less toxic and insoluble trivalent cation Cr(III) by specific NAD(P)H-dependent enzymes under aerobic conditions or in the electron transport chain of bacteria such as Pseudomonas fluorescens LB300 in anaerobic environments [4–9]. In addition, several enzymes work to counteract the deleterious effects of the oxidative stress induced following cell exposure to chromate, dichromate and cadmium.
Caulobacter crescentus, an oligotrophic free-living α-proteobacterium, is able to grow in polluted habitats [10–12]. Not surprisingly, its genome encodes some homologues of genes involved in heavy metal resistance. In a previous report, the set of genes responding to Caulobacter exposure to chromate, dichromate and cadmium was identified . The main actions triggered in response to these metals are protection against oxidative stress (strong induction of superoxide dismutase, glutathione S-transferase, thioredoxin, glutaredoxins and DNA repair enzymes) and reduction of intracellular metal concentration (down-regulation of a sulfate transporter under chromate and dichromate stresses that could reduce nonspecific uptake of these oxyanions, and up-regulation of multiple efflux pumps that could play a key role in removing cadmium from the cytoplasm). However, the signal transduction and control processes involved in the bacterial response to these heavy metals are still poorly characterized.
The C. crescentus genome encodes 13 e xtrac ytoplasmic f unction (ECF) sigma factors . Two of them, the paralogous σT and σU, are involved in the response to various environmental stress conditions, including chromium and cadmium stresses [12, 14]. Additionally, σE mediates a rapid transcriptional response to cadmium, organic hydroperoxide, singlet oxygen and UV-A . In a previous report, σF was found to be required for bacterial survival under hydrogen peroxide stress in the stationary growth phase, but no σF-mediated transcriptional response to hydrogen peroxide could be observed . Thus, the involvement of σF in a transcriptional response to environmental stresses still needs to be characterized. The observation that genes CC2906, CC3255 and CC3257, previously found to be dependent on σF, are induced following C. crescentus exposure to chromate, dichromate and cadmium  suggested to us that σF could be involved in the transcriptional response to these heavy metals.
In the present work, we demonstrate the involvement of σF in chromium and cadmium stress responses. We also identified the set of genes regulated by σF by using global transcriptome analysis and characterized the promoter region of these genes by 5´RACE experiments and β-galactosidase assays. Furthermore, we investigated the role of the protein encoded by the second gene in the sigF operon (CC3252), here named NrsF, and two conserved cysteine residues in this protein on the σF-mediated response to heavy metals.
σF is involved in chromium and cadmium responses in C. crescentus
It is assumed that heavy metal ions cause oxidative stress inside cells [1, 12, 17]. This raises the possibility that induction of σF-dependent genes by chromium and cadmium is a direct consequence of oxidative stress. To test this hypothesis, we stressed the parental and the sigF mutant strains with hydrogen peroxide, t-butyl hydroperoxide, paraquat (source of superoxide anion) or diamide (causes depletion of thiols). According to qRT-PCR experiments, expression levels of CC3255 and CC3252 were not increased more than twofold in the parental strain during these stress conditions (Figure 1). In agreement, transcript levels of CC3255 and CC3252 were also not influenced by any of these stressors in cells lacking sigF. Concentrations of hydrogen peroxide and t-butyl hydroperoxide used in our analyses were previously found to be sufficient to increase expression of other genes in C. crescentus[15, 18]. Taken together, these data suggest that chromium and cadmium are able to induce the σF regulon in an oxidative stress independent manner.
σF controls a small set of genes under chromium stress
Expression analysis of σ F -dependent genes upon dichromate stress
Gene number a
Putative identification e
ΔsigF Cr/ WT Cr
WT Cr/ WT no stress
ΔsigF Cr/ΔsigF no stress
ΔsigF Cr/WT Cr
sulfite oxidase subunit YedY
protein of unknown function
protein of unknown function
predicted integral membrane protein
negative regulator of σF
ECF sigma factor σF
predicted integral membrane protein
protein of unknown function
protein of unknown function
protein of DoxX family
The CC2907 gene is predicted to be transcribed divergently from CC2906-CC2905 in the chromosome of CB15 strain. However, the corresponding gene was not included during annotation of the more recent genome sequencing of C. crescentus (NA1000 strain). In the chromosome of NA1000, an open reading frame (CCNA_03001) was proposed to be located between genes CCNA_03000 (corresponding to CC2906) and CCNA_03002 (corresponding to CC2908). Nevertheless, CCNA_03001 appears to be co-transcribed with CCNA_03000 and CCNA_03002. In addition, we could observe co-occurrence of CCNA_03001 with other σF-dependent genes. As the nucleotide sequence between CC2906 and CC2908 in CB15 strain is identical to the region between CCNA_03000 and CCNA_03002 of NA1000 strain, we conclude that CC2907 was incorrectly annotated in the genome of CB15 strain and this gene is the first one of the operon CC2907-CC2906-CC2905 (Figure 2A). As evaluated with probes corresponding to the upstream region of CC2906, the entire coding region of CC2907 is down-regulated in sigF mutant cells relative to the parental strain (Table 1). Therefore, the complete transcriptional unit CC2907-CC2906-CC2905 is controlled by σF.
A thorough re-annotation of genes regulated by σF suggested that CC3257 codes for a putative membrane protein belonging to the DoxX family, whose members are involved in sulfur metabolism. The CC2748 gene, which encodes the putative sulfite oxidase subunit YedY, is another protein with a potential role in sulfur metabolism. All of the remaining σF-dependent genes (CC2905, CC2906, CC2907, CC3254, CC3255 and CC3256) code for proteins with conserved domains of unknown functions. Interestingly, the pairs of genes CC2907 and CC3254, CC2906 and CC3255, as well as CC2905 and CC3256 are probable paralogous genes, with amino acid sequence identities of 36%, 43% and 23%, respectively. Therefore, it is possible that the gene products of both operons exert similar functions. No other gene in the genome of C. crescentus displays significant nucleotide sequence similarity to the above mentioned pairs of paralogous genes or to the functionally annotated genes CC2748 and CC3257.
Proteins encoded by CC2905 and CC3256 present a DUF2063 domain at their N-terminus. This domain was described to be a DNA-binding domain in NGO1945 from Neisseria gonorrhoeae. NGO1945 is involved in the transcriptional regulation of msrAB, which codes for a methionine sulfoxide reductase . However, in our microarray experiments, we could not observe differences in the expression of msrA homologs in C. crescentus (CC0994 and CC1039). Thus, we conclude that the role of NGO1945 in N. gonorrhoeae and CC2905 or CC3256 in C. crescentus is most likely different under these circumstances.
To confirm results obtained in transcriptome analysis, we investigated the expression levels of five genes supposedly dependent on σF (CC2748, CC2905, CC2906, CC3255 and CC3257) by qRT-PCR experiments. These analyses showed that expression of these selected genes under dichromate stress is more than twofold higher in the parental strain relative to the sigF deletion mutant (Table 1). Interestingly, induction of CC2748 expression in the presence of dichromate was only partially dependent on σF (Table 1), suggesting the involvement of an additional regulatory protein in the control of CC2748 expression under this stress condition. Taken together, these results confirm the data obtained in global transcriptional analysis.
Promoter sequence motifs of CC2907 and CC3254 genes are highly similar to those of sigF
To identify putative σF-dependent promoters upstream of CC2907 and CC3254 genes, we performed 5’RACE (rapid amplification of cDNA-ends) experiments using primers that hybridize in the beginning of the coding region of the corresponding genes. For these experiments, RNA samples from cells exposed to dichromate were used, as this stress condition leads to increased expression levels of CC2907 and CC3254. This approach led to the identification of a transcriptional start site (TSS) for CC2907 at position −7 relative to the translational start site +1 proposed here (Figure 2B). A TSS was also determined at position
−61 with respect to the translational start site of CC3254 predicted here (Figure 2B). As expected, no TSS could be observed when an additional 5´RACE experiment was performed using primers that hybridize to the beginning of the coding region of CC3254 proposed by the TIGR annotation. Together, these data confirmed our microarray data with respect to expression of the operons CC2907-CC2906-CC2905 and CC3254-CC3255-CC3256-CC3257.
The putative promoter sequences found for CC2907 and CC3254 were very similar to each other and also quite similar to the promoter sequence previously determined for sigF (Figure 2B). Additionally, analyses of the region upstream of the translational start site +1 of CC2748 also revealed a putative σF-dependent sequence (Figure 2B), suggesting a direct control of this gene by σF. Accordingly, the putative σF-dependent promoters reported here are highly similar to sequences found upstream from sigF homologs in other bacteria .
Conserved sequences upstream of CC3254 and sigF are necessary for expression of these genes
As mentioned above, the promoter sequence of the operon CC3254-CC3255-CC3256-CC3257 is highly similar to that located upstream from sigF. To verify if sigF expression was also dependent on these putative promoter elements, we analyzed the upstream region of the sigF gene in β-galactosidase assays using two different plasmid constructs: pCKlac53-1 containing the promoter elements upstream from sigF and construct pCKlac53-2 that lacks the sigF promoter (Figure 3A). β-galactosidase activity measured in parental cells harboring the construct pCK53-2 (Figure 3B) was found to be quite similar to that observed in cells with the empty vector. On the other hand, higher β-galactosidase activity was observed in the parental strain carrying construct pCK53-1, which contains the complete sigF promoter sequence (Figure 3B). Cells from sigF mutant harboring the construct pCKlac53-1 presented β-galactosidase activity slightly lower than that observed in parental cells with the same construct, but still higher than that observed in cells harboring the construct pCK53-2 (Figure 3B). Altogether, these data indicate that the promoter sequence upstream from sigF is necessary for expression of the sigF operon, but in a manner that is not exclusively dependent on σF. This observation suggests that another sigma factor could also be capable of recognizing the region upstream from sigF. Thus, we have investigated the effect of two other ECF sigma factors involved in oxidative and heavy metal stresses, σT and σE, upon sigF promoter activity, but no significant decrease in β-galactosidase activity was observed in mutant strains ΔsigT and ΔrpoE when compared with parental cells, all harboring construct pCKlac53-1 (data not shown). Additionally, qRT-PCR experiments confirmed these results, as no change in transcription levels of the sigF gene was observed in a sigT or rpoE mutant (data not shown). These observations allowed us to rule out the participation of σT and σE in the control of sigF expression.
To further verify if the promoter region upstream of sigF is controlled by σF, we overexpressed sigF in the parental strain from an additional plasmid-encoded copy of the gene under the control of a constitutive promoter (construct pCM30) and measured β-galactosidase activity in these cells harboring either pCKlac53-1 or pCKlac53-2. Overexpression of sigF in cells with the construct containing the complete sigF promoter (pCK53-1) led to an increase in β-galactosidase activity, whereas no difference was observed in cells harboring the promoterless construct pCKlac53-2 (Figure 3B). Similarly, higher β-galactosidase activity was observed in sigF overexpressing cells bearing the construct containing the promoter sequence motifs upstream from CC3254 (pCKlac54-1) when compared to the parental strain carrying the same construct or sigF overexpressing cells harboring the construct containing only the −10 motif of the promoter sequence of CC3254-CC3255-CC3256-CC3257 (pCKlac54-2) (Figure 3B). Therefore, these results confirm that specific and highly similar promoter sequence motifs found upstream from sigF-CC3252 and CC3254-CC3255-CC3256-CC3257 are required for the control of these transcriptional units by σF.
CC3252 negatively regulates σF regulon expression
A further attempt to investigate the role of nrsF as a possible negative regulator of σF function was carried out by trying to construct a null mutant strain in gene nrsF. However, it was not possible to construct a mutant strain by deleting nrsF in the parental strain (data not shown). On the other hand, nrsF could be deleted in the absence of a functional copy of sigF (data not shown), suggesting that high σF activity is apparently responsible for the failure of disrupting nrsF in cells with functional sigF.
σF is released into the cytoplasm during chromium stress and in cells carrying point mutations in conserved cysteines of NrsF
Neither σF nor σF-dependent genes CC2906 and CC3255 are essential for Caulobacter resistance to metal stress
To investigate the requirement of sigF for resistance of C. crescentus cells to dichromate or cadmium, the sensitivity of the parental strain and the sigF deletion mutant to exposure to these metals was monitored. Both strains displayed similar sensitivity profile to dichromate or cadmium (data not shown), suggesting that sigF is not essential for bacterial survival under this stress condition. As the deduced protein sequences of CC2906 and CC3255 are highly similar, we constructed a single deletion mutant strain in each gene (SG19 and SG20) as well as a double mutant (SG21) and tested the resistance of these strains to the metal stresses. Similar to what was found for the sigF deletion mutant, no increased sensitivity was observed for these mutant strains following exposure to either dichromate or cadmium, when compared to parental cells (data not shown). Together, these data suggest that σF-mediated transcriptional response to chromium or cadmium is not essential for survival of C. crescentus to exposure to these metal ions.
In this report, we clearly show that C. crescentus σF is involved in the transcriptional response to chromium and cadmium in an oxidative stress independent manner. Transcriptome analysis of cells under dichromate stress revealed that σF controls a small regulon comprised of eight genes, which are distributed in three transcriptional units. Although a conserved domain was predicted for the deduced protein sequence of all σF-dependent genes, only two of these sequences could be assigned to a possible function. The protein encoded by CC2748 belongs to the group of sulfite oxidases, which catalyze the oxidation of the toxic and very reactive sulfite to the inert sulfate anion . The product of CC3257 is a member of the DoxX family. Although nothing is actually known about the physiological role of bacterial proteins belonging to this family, the archaeal counterparts are involved in the elemental sulfur oxidation pathway [23, 24]. Therefore, both σF-dependent genes with a putative assigned function appear to play a role in sulfate acquisition by cells. Interestingly, Hu et al. (2005) found a strong down-regulation of a Caulobacter sulfate ABC transport system under chromate and dichromate exposure. While this detoxification strategy apparently contributes to decrease the concentration of chromate and dichromate in the cells , sulfate uptake from the extracellular environment might be significantly affected. Alternative sources such as degradation of sulfur-containing amino-acids  and organosulfonate metabolism  can be used to counteract this sulfur uptake limitation [1, 27–29]. It is therefore conceivable that induction of CC2748 and CC3257 could supply cells with sulfate. This is consistent with the observation that in Arthrobacter sp. strain FB24 and Pseudomonas putida, chromate exposure also results in increased levels of proteins potentially involved in reversing the effects of cellular sulfur limitation, such as transporters of alternative sulfur sources [27, 28].
Curiously, none of the most representative functional categories up-regulated under chromate, dichromate or cadmium exposure (protection against oxidative stress and reduction of intracellular metal concentration) were found to be controlled by σF, indicating that additional molecular systems are engaged in C. crescentus response to these metals. In fact, we previously reported the involvement of the paralogous sigma factors σT and σU in the control of response to chromium and cadmium [14, 15, 30] and σE in response to cadmium [14, 15, 30]. The observation that σF, σE and σT/σU regulate distinct sets of genes indicates that each of these sigma factors make a different contribution to the C. crescentus response to metal stress. Together, σF, σE, σT and σU are responsible for the induction of 20% of the genes previously found to be up-regulated under cadmium stress and σF, σT and σU control the expression of about 12% of genes induced following Caulobacter exposure to chromate or dichromate (Additional file 1: Table S1). Therefore, transcriptional regulators other than σF, σE, σT and σU appear to be involved in the response to chromate, dichromate and cadmium. The existence of several molecular systems contributing to the transcriptional response to metal stresses could explain why the absence of sigF, CC2906 or CC3255 does not decrease the viability of Caulobacter cells under dichromate or cadmium stresses. In agreement, we previously reported that σE elicits a rapid response to cadmium, but cells lacking rpoE are not impaired in survival to this stress condition [14, 15, 30].
Interestingly, sigF is not highly induced under either chromium or cadmium stress, different from what was observed for other ECF sigma factor genes such as rpoE and sigT in C. crescentus[14, 15, 30] and rpoJ, rpoK, rpoI, cnrH and rpoQ in C. metallidurans. This indicates that sigF is obviously not strongly auto-regulated under heavy metal stress conditions. Although the experimentally determined promoter sequences of sigF and CC3254 are highly similar to each other, promoter activity analyses supported our observation that CC3254 is solely regulated by σF, while the sigF operon is transcribed under the control of σF and a still unknown transcriptional regulator. Interestingly, both σF and the additional regulator depend on sequences located from −37 to +37 relative to the transcriptional start site (+1) of sigF. An apparent competition between these proteins might be the reason why sigF promoter activity is less responsive to high levels of σF when compared to the CC3254 promoter, which is solely controlled by σF. The existence of a second regulator of the sigF operon would be important to maintain a certain basal level of σF and consequently to allow a rapid response when cells experience environments contaminated with heavy metals. In the literature, one can find various examples of ECF sigma factor genes dependent on a second ECF sigma factor [32, 33]. In the present study, we could exclude Caulobacter rpoE and sigT as possible regulators of σF, since no difference in sigF expression was observed in the absence of either one of these ECF sigma factor genes.
In most cases, the activity of ECF sigma factors is modulated by a cognate anti-sigma factor [34–36]. Here, we showed that the second gene (CC3252) in the sigF operon acts as a negative regulator of σF function, as overexpression of the putative membrane protein encoded by CC3252 abolishes the transcriptional activation of sigF and its regulon under dichromate stress. Thus, CC3252 was here denominated nrsF. An interesting question about the nature of σF inhibition came from the observation that most of the protein encoded by nrsF is predicted to lie in the inner membrane of the bacterium: six transmembrane helices separated by five linkers ranging from 6 to 19 amino acid residues and an N-terminal segment of 25 residues. Usually, anti-sigma factors bind their cognate sigma factor through an extensive surface interaction, in which a domain of the first protein is sandwiched between domains σ2 and σ4 of the sigma factor . It is possible that several of the linkers of NrsF contact σF, resulting in a more stable interaction surface. However, we cannot discard the presence of a third component in this system able to directly bind both σF and NrsF and transduce the signal leading to activation of the sigma factor, to compensate this apparent lack of sufficient cytoplasmic segments in NrsF to contact σF. Attempts to obtain soluble recombinant full-length NrsF failed, probably because the protein cannot correctly fold in the absence of the hydrophobic environment found in the membrane compartment of bacterial cells. Therefore, it was not possible to test whether the recombinant protein encoded by nrsF directly binds σF.
As previously observed for other ECF sigma factors of C. crescentus[14, 15, 30], we were not able to delete nrsF, probably due to the toxic effect of high levels of σF under no stress conditions. However, we could isolate strains in which one or both of the conserved cysteine residues of NrsF were replaced for serine. As suggested by Western blot analysis, isolation of these point mutation strains was possible probably because most of σF molecules are still directly or indirectly sequestered in an inactive state to the inner membrane by NrsF. Substitution of the conserved cysteines might have caused structural changes in NrsF and hence resulting in a lower capacity to bind σF. In fact, σF was found to accumulate in the soluble fraction of cells expressing NrsF mutated in both cysteine residues even when cells were cultured under unstressed conditions. The presence of σF in the soluble fraction was also detected following treatment of parental cells with dichromate. Therefore, we could observe accumulation of σF in the soluble fraction in situations in which lower affinity of NrsF for σF is expected. Interestingly, two conserved cysteine residues in ChrR, the anti-sigma factor of Caulobacter σE, were also shown to be important for the response to cadmium mediated by that sigma factor [14, 15, 30]. Furthermore, the sensor histidine kinase PhyK, involved in the control of the anti-anti-sigma factor PhyR of Caulobacter σT, which as mentioned above responds to dichromate and cadmium, also presents a conserved cysteine that is important to PhyK activity [14, 15, 30]. Thus, cysteines in the probable sensor proteins (NrsF, ChrR and PhyK) of ECF sigma factor mediated systems seem to play a key role in triggering the response to heavy metal stress in C. crescentus.
Based on the fact that dichromate and cadmium are able to directly bind thiol groups [2, 38], it is conceivable that these metals could disrupt contacts mediated by the conserved cysteines of NrsF, leading to changes in its conformation similar to those expected in the mutant proteins with one or both of the cysteine residues substituted. However, activation of σF might also be caused by direct interaction of chromate, dichromate and cadmium with other amino acid residues in NrsF or even with another yet unknown sensory component of the system. The finding that single substitutions of the conserved cysteine residues still allows for induction of σF-dependent genes ruled out the formation of an intramolecular bond between Cys131 and Cys181 residues under stress conditions. Nevertheless, we could not discard the possibility that NrsF functions as a dimer/multimer using intermolecular bonds for sensing the metals in the extracytoplasmic environment.
This report deals with the role and regulation of C. crescentus σF under stress conditions and provides new interesting information about this conserved but still poorly characterized ECF sigma factor: i) σF-dependent genes are induced in the presence of heavy metals in a manner independent of oxidative and disulfide stress; ii) σF directly controls a small regulon including genes involved in sulfur metabolism and iii) σF is negatively regulated by a putative membrane bound protein, here named NrsF, which contains two conserved cysteine residues that are important for its function and are located in the periplasmic portion of the protein.
Bacterial strains and growth conditions
The strains and plasmids used in this study are described in Additional file 1: Table S2. C. crescentus strains were cultured at 30°C in M2 minimal salts medium plus glucose . When appropriate, the growth medium was supplemented with chloramphenicol (1 μg ml-1), kanamycin (10 μg ml-1) or tetracycline (2 μg ml-1). Plasmids were propagated in Escherichia coli strain DH5α (Invitrogen) and mobilized into C. crescentus by bacterial conjugation using E. coli strain S17-1 . E. coli strains were grown at 37°C in LB broth .
Deletion of genes CC2906 and CC3255 in C. crescentus
Single mutant strains for CC2906 (SG20) and CC3255 (SG19) were obtained by an in-frame deletion in the coding region of these genes. For that, two fragments flanking the regions to be deleted were amplified by PCR (a complete list of primers used in this study is in Additional file 1: Table S3) and subcloned into pNPTS138 . Constructs into pNPTS138 were transferred to C. crescentus strain NA1000  by conjugation with E. coli S17-1 and the deletion of the wild-type copy of the gene in the NA1000 background was achieved by two homologous recombination events. Mutant strains were isolated by screening colonies by PCR and DNA sequencing. For the construction of a double mutant strain (SG21), the single mutant strain SG20 was used for the two homologous recombination events of the CC3255 deletion.
Construction of point mutations in CC3252 and overexpression of CC3252 in C. crescentus
Codons for the conserved cysteine residues of the protein encoded by CC3252 (C131 and C181) were replaced for a codon corresponding to serine by overlapping PCR with a pair of complementary primers (Additional file 1: Table S3) designed for each substitution. Each part of CC3252 was amplified separately by PCR using one of each complementary primer set and a primer hybridizing upstream or downstream from CC3252. The partially complementary PCR products were used together as templates in a second amplification reaction with the primers hybridizing upstream and downstream from CC3252. The amplicons obtained were cloned into pGEM-T (Promega) and sequenced. The inserts were excised from vectors and subcloned into pNPTS138. Constructs into pNPTS138 were transferred to C. crescentus strain NA1000  by conjugation with E. coli S17-1 and replacement of the wild-type copy of the gene for the corresponding mutated copy in the NA1000 background was achieved by two homologous recombination events. Mutant strains were isolated by screening colonies by PCR and DNA sequencing. To overexpress CC3252 in C. crescentus cells, a fragment corresponding to the coding region of the gene was first amplified by PCR. This fragment was excised from the vector and ligated into pJS14. The construct was introduced into C. crescentus NA1000 by conjugation with E. coli S17-1.
For quantitative real time-PCR (qRT-PCR) analysis, cultures of different C. crescentus strains were grown to exponential phase (OD600 0.5), submitted for 30 minutes to stress (55 μM dichromate, 55 μM cadmium, 100–500 μM hydrogen peroxide, 50–200 μM t-butyl hydroperoxide, 100–500 μM paraquat or 50–200 μM diamide) or kept under no stress conditions and cells (four aliquots of 2 ml from each treatment) were collected by centrifugation in a microcentrifuge for 1 min. For microarray experiments, total RNA was extracted from the parental NA1000 and the sigF mutant strain SG16 at the exponential growth phase exposed to 55 μM dichromate for 30 min.
The cell pellets were suspended in 1 ml of Trizol Reagent (Invitrogen), and after the extraction procedure according to manufacturer’s instructions, the integrity of the RNA was checked by agarose gel electrophoresis and tested for the absence of DNA contamination by PCR.
Quantitative real-time PCR
Reverse transcription for qRT-PCR was performed using 5 μg of total RNA, 200 U of Superscript III reverse transcriptase (Invitrogen) and 500 ng of random primer, following manufacturer’s instructions. Quantitative PCR amplification of the resulting cDNA was performed with Platinum SYBR Green (Applied Biosystems) and gene-specific primers (see Additional file 1: Table S3). These primers were designed using the Primer Express software (Applied Biosystems). Results were normalized using CC0088 gene as the endogenous control, which was previously used [15, 30] and shown to be constant in the samples analyzed. Relative expression levels were calculated using the 2-ΔΔCT method .
RNA 5’ ends of genes of interest were determined using the 3′/5′RACE kit (Roche). For that, the RNA was reverse transcribed using a gene-specific primer (Additional file 1: Table S3), purified and poly(dA) tailed at their 3′ends. The resulting cDNA was amplified by PCR using the forward poly(dT)-anchor primer provided by the kit to anneal at the poly(dA) tail and a second gene-specific primer. The PCR products were used in a second PCR reaction using a primer complementary to the poly(dT)-anchor primer and a distinct gene-specific nested primer. PCR products were ligated into the pGEM-T vector (Promega) and several distinct clones were sequenced.
Three distinct biological RNA samples isolated from each strain analyzed were reverse transcribed and labeled using the FairPlay III Microarray Labeling system (Agilent). Briefly, the cDNA was synthesized from total RNA (20 μg) in the presence of amino allyl modified dUTP and random primer. After purification, the resulting amino-modified cDNA was chemically labeled by incorporation of the dyes Alexa Fluor 555 (Cy3) or Alexa Fluor 647 (Cy5). Labeled cDNAs were combined, mixed with Agilent hybridization buffer, and competitively hybridized to custom-designed Agilent microarrays according to the manufacturer’s instructions (Agilent). Data extraction and normalization was performed using Agilent Feature Extraction Software 22.214.171.124 (Agilent). The custom-designed arrays contain 9–11 probes covering a region around the translational start site (−300 to +200 relative to the translational start site +1) of each gene. Only those probes downstream of the translational start site were considered for estimating the fold change in gene expression. Ratios obtained for probes corresponding to the same gene were averaged and genes showing a ratio log2 (mutant/parental) < −1 or log2 (mutant/parental) > 1 in all three biological replicates were considered as differentially expressed between the strains analyzed. Complete microarray dataset was deposited in GEO (GSE 32406).
Cell fractionation and Western blot analysis
Protein extracts were obtained from cultures of parental strain NA1000 and a CC3252 mutant with both C131 and C181 replaced for serine before and after treatment with 55 μM dichromate for 30 min. Cells were cultured until OD600 0.5, harvest by centrifugation and washed once with 0.2 M Tris–HCl pH 8.0. Cells were then resuspended in 1 ml 60 mM Tris–HCl pH 8.0, 0.2 M sucrose, 0.2 mM EDTA, 200 μg ml-1 of lysozyme and incubated for 10 min at room temperature. After brief sonication (three 10 s pulses), cell debris were removed and the supernatant was centrifuged at 150,000 x g for 1 h. The pellet was washed once with 60 mM Tris–HCl pH 8.0 and resuspended in 1 ml 60 mM Tris–HCl pH 8.0, 0.2 M sucrose, 0.2 mM EDTA. Equal amounts of total protein (20 μg) were resolved through SDS-PAGE and transferred to nitrocelulose membrane, as previously described . Membranes were incubated overnight at 4°C with anti-σF (1:500)  or anti-FtsH (1:2000) (kindly provided by T. Ogura, Kumamoto University, Japan) antibody in 10 mM Tris–HCl pH 8.0 containing 150 mM NaCl, 0.02% Tween 20, and 0.03% Triton X-100. The blots were developed using fluorescent CF680 Goat Anti-Rabbit IgG (1:10000- Uniscience) and imaged using Odyssey Imager- LI-COR (Biosciences).
Promoter activity assay
β-galactosidase assays were carried out with cells carrying a CC3255-lacZ transcription fusion (pCKlac54-1 or pCKlac54-2) or a sigF-lacZ transcription fusion (pCKlac53-1 or pCKlac53-2). For that, cells were cultured to exponential phase, harvested and used for the enzymatic assay. The empty plasmid placZ290  was used as the control in the experiments. β-galactosidase activity was measured as previously described . All experiments were performed in duplicates and repeated on three different occasions.
Stress sensitivity tests
Exponentially growing cells were exposed to 55 μM dichromate or kept under unstressed conditions. To measure cell viability, aliquots were removed, serially diluted and plated on M2 minimal medium for counting colony-forming units.
Sequence alignment and structure prediction
This work was supported by a grant to S.L.G. from Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP). R.F.L. and C.K. were postdoctoral fellows from FAPESP, G.M.A. is a pre-doctoral fellow of FAPESP, and S.L.G. was partially supported by Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq-Brazil). We thank Michael T. Laub for assistance with the microarray analysis, Cristina E. Alvarez-Martinez for important discussions and construct pCM30, Chuck S. Farah for careful reading of the manuscript, and Anne Kohler, Luci D. Navarro and Sandra M. Fernandes for expert technical assistance.
- Ramirez-Diaz MI, Diaz-Perez C, Vargas E, Riveros-Rosas H, Campos-Garcia J, Cervantes C: Mechanisms of bacterial resistance to chromium compounds. Biometals. 2008, 21 (3): 321-332. 10.1007/s10534-007-9121-8.PubMedView ArticleGoogle Scholar
- Nies DH: Microbial heavy-metal resistance. Appl Microbiol Biotechnol. 1999, 51 (6): 730-750. 10.1007/s002530051457.PubMedView ArticleGoogle Scholar
- Barceloux DG: Chromium. J Toxicol Clin Toxicol. 1999, 37 (2): 173-194. 10.1081/CLT-100102418.PubMedView ArticleGoogle Scholar
- Cervantes C: Reduction and efflux of chromate in bacteria. In NiesDH and Silver S (eds) Molecular Biology of Heavy Metals. 2007, Berlin: Springer-VerlagGoogle Scholar
- Ohtake H, Komori K, Cervantes C, Toda K: Chromate-resistance in a chromate-reducing strain of Enterobacter cloacae. FEMS Microbiol Lett. 1990, 55 (1–2): 85-88.PubMedView ArticleGoogle Scholar
- Gonzalez CF, Ackerley DF, Lynch SV, Matin A: ChrR, a soluble quinone reductase of Pseudomonas putida that defends against H2O2. J Biol Chem. 2005, 280 (24): 22590-22595. 10.1074/jbc.M501654200.PubMedView ArticleGoogle Scholar
- Kwak YH, Lee DS, Kim HB: Vibrio harveyi nitroreductase is also a chromate reductase. Appl Environ Microbiol. 2003, 69 (8): 4390-4395. 10.1128/AEM.69.8.4390-4395.2003.PubMedPubMed CentralView ArticleGoogle Scholar
- Mazoch J, Tesarik R, Sedlacek V, Kucera I, Turanek J: Isolation and biochemical characterization of two soluble iron(III) reductases from Paracoccus denitrificans. Eur J Biochem. 2004, 271 (3): 553-562. 10.1046/j.1432-1033.2003.03957.x.PubMedView ArticleGoogle Scholar
- Ackerley DF, Gonzalez CF, Park CH, Blake R, Keyhan M, Matin A: Chromate-reducing properties of soluble flavoproteins from Pseudomonas putida and Escherichia coli. Appl Environ Microbiol. 2004, 70 (2): 873-882. 10.1128/AEM.70.2.873-882.2004.PubMedPubMed CentralView ArticleGoogle Scholar
- Lapteva NA: Ecological features of distribution of bacteria of the genus Caulobacter in freshwater bodies. Mikrobiologiya. 1987, 56: 537-543.Google Scholar
- Poindexter JS: The caulobacters: ubiquitous unusual bacteria. Microbiol Rev. 1981, 45 (1): 123-179.PubMedPubMed CentralGoogle Scholar
- Hu P, Brodie EL, Suzuki Y, McAdams HH, Andersen GL: Whole-genome transcriptional analysis of heavy metal stresses in Caulobacter crescentus. J Bacteriol. 2005, 187 (24): 8437-8449. 10.1128/JB.187.24.8437-8449.2005.PubMedPubMed CentralView ArticleGoogle Scholar
- Nierman WC, Feldblyum TV, Laub MT, Paulsen IT, Nelson KE, Eisen JA, Heidelberg JF, Alley MR, Ohta N, Maddock JR: Complete genome sequence of Caulobacter crescentus. Proc Natl Acad Sci U S A. 2001, 98 (7): 4136-4141. 10.1073/pnas.061029298.PubMedPubMed CentralView ArticleGoogle Scholar
- Lourenco RF, Kohler C, Gomes SL: A two-component system, an anti-sigma factor and two paralogous ECF sigma factors are involved in the control of general stress response in Caulobacter crescentus. Mol Microbiol. 2011, 80 (6): 1598-1612. 10.1111/j.1365-2958.2011.07668.x.PubMedView ArticleGoogle Scholar
- Lourenco RF, Gomes SL: The transcriptional response to cadmium, organic hydroperoxide, singlet oxygen and UV-A mediated by the sigmaE-ChrR system in Caulobacter crescentus. Mol Microbiol. 2009, 72 (5): 1159-1170. 10.1111/j.1365-2958.2009.06714.x.PubMedView ArticleGoogle Scholar
- Alvarez-Martinez CE, Baldini RL, Gomes SL: A caulobacter crescentus extracytoplasmic function sigma factor mediating the response to oxidative stress in stationary phase. J Bacteriol. 2006, 188 (5): 1835-1846. 10.1128/JB.188.5.1835-1846.2006.PubMedPubMed CentralView ArticleGoogle Scholar
- Molecular mechanisms in metal carcinogenesis: role of oxidative stress, In O. I. Aruoma and B. Halliwell (ed.), Molecular biology of free radicals in human diseases. Edited by: Klein C, Snow E, Frenkel K. 1998, London, England: OICA InternationalGoogle Scholar
- Italiani VC, da Silva Neto JF, Braz VS, Marques MV: Regulation of catalase-peroxidase KatG is OxyR dependent and Fur independent in Caulobacter crescentus. J Bacteriol. 2011, 193 (7): 1734-1744. 10.1128/JB.01339-10.PubMedPubMed CentralView ArticleGoogle Scholar
- Das D, Grishin NV, Kumar A, Carlton D, Bakolitsa C, Miller MD, Abdubek P, Astakhova T, Axelrod HL, Burra P: The structure of the first representative of Pfam family PF09836 reveals a two-domain organization and suggests involvement in transcriptional regulation. Acta Crystallogr Sect F Struct Biol Cryst Commun. 2010, 66 (Pt 10): 1174-1181.PubMedPubMed CentralView ArticleGoogle Scholar
- Gunesekere IC, Kahler CM, Ryan CS, Snyder LA, Saunders NJ, Rood JI, Davies JK: Ecf, an alternative sigma factor from Neisseria gonorrhoeae, controls expression of msrAB, which encodes methionine sulfoxide reductase. J Bacteriol. 2006, 188 (10): 3463-3469. 10.1128/JB.188.10.3463-3469.2006.PubMedPubMed CentralView ArticleGoogle Scholar
- Staron A, Sofia HJ, Dietrich S, Ulrich LE, Liesegang H, Mascher T: The third pillar of bacterial signal transduction: classification of the extracytoplasmic function (ECF) sigma factor protein family. Mol Microbiol. 2009, 74 (3): 557-581. 10.1111/j.1365-2958.2009.06870.x.PubMedView ArticleGoogle Scholar
- Kappler U: Bacterial sulfite-oxidizing enzymes. Biochim Biophys Acta. 2011, 1807 (1): 1-10. 10.1016/j.bbabio.2010.09.004.PubMedView ArticleGoogle Scholar
- Muller FH, Bandeiras TM, Urich T, Teixeira M, Gomes CM, Kletzin A: Coupling of the pathway of sulphur oxidation to dioxygen reduction: characterization of a novel membrane-bound thiosulphate:quinone oxidoreductase. Mol Microbiol. 2004, 53 (4): 1147-1160. 10.1111/j.1365-2958.2004.04193.x.PubMedView ArticleGoogle Scholar
- Purschke WG, Schmidt CL, Petersen A, Schafer G: The terminal quinol oxidase of the hyperthermophilic archaeon Acidianus ambivalens exhibits a novel subunit structure and gene organization. J Bacteriol. 1997, 179 (4): 1344-1353.PubMedPubMed CentralGoogle Scholar
- Griffith OW: Mammalian sulfur amino acid metabolism: an overview. Methods Enzymol. 1987, 143: 366-376.PubMedView ArticleGoogle Scholar
- Cook AM, Denger K: Metabolism of taurine in microorganisms: a primer in molecular biodiversity?. Adv Exp Med Biol. 2006, 583: 3-13. 10.1007/978-0-387-33504-9_1.PubMedView ArticleGoogle Scholar
- Henne KL, Turse JE, Nicora CD, Lipton MS, Tollaksen SL, Lindberg C, Babnigg G, Giometti CS, Nakatsu CH, Thompson DK: Global proteomic analysis of the chromate response in Arthrobacter sp. strain FB24. J Proteome Res. 2009, 8 (4): 1704-1716. 10.1021/pr800705f.PubMedView ArticleGoogle Scholar
- Thompson DK, Chourey K, Wickham GS, Thieman SB, VerBerkmoes NC, Zhang B, McCarthy AT, Rudisill MA, Shah M, Hettich RL: Proteomics reveals a core molecular response of Pseudomonas putida F1 to acute chromate challenge. BMC Genomics. 2010, 11: 311-10.1186/1471-2164-11-311.PubMedPubMed CentralView ArticleGoogle Scholar
- Brown SD, Thompson MR, Verberkmoes NC, Chourey K, Shah M, Zhou J, Hettich RL, Thompson DK: Molecular dynamics of the Shewanella oneidensis response to chromate stress. Mol Cell Proteomics. 2006, 5 (6): 1054-1071. 10.1074/mcp.M500394-MCP200.PubMedView ArticleGoogle Scholar
- Alvarez-Martinez CE, Lourenco RF, Baldini RL, Laub MT, Gomes SL: The ECF sigma factor sigma(T) is involved in osmotic and oxidative stress responses in Caulobacter crescentus. Mol Microbiol. 2007, 66 (5): 1240-1255. 10.1111/j.1365-2958.2007.06005.x.PubMedView ArticleGoogle Scholar
- Grosse C, Friedrich S, Nies DH: Contribution of extracytoplasmic function sigma factors to transition metal homeostasis in Cupriavidus metallidurans strain CH34. J Mol Microbiol Biotechnol. 2007, 12 (3–4): 227-240.PubMedGoogle Scholar
- Dona V, Rodrigue S, Dainese E, Palu G, Gaudreau L, Manganelli R, Provvedi R: Evidence of complex transcriptional, translational, and posttranslational regulation of the extracytoplasmic function sigma factor sigmaE in Mycobacterium tuberculosis. J Bacteriol. 2008, 190 (17): 5963-5971. 10.1128/JB.00622-08.PubMedPubMed CentralView ArticleGoogle Scholar
- Raman S, Song T, Puyang X, Bardarov S, Jacobs WR, Husson RN: The alternative sigma factor SigH regulates major components of oxidative and heat stress responses in Mycobacterium tuberculosis. J Bacteriol. 2001, 183 (20): 6119-6125. 10.1128/JB.183.20.6119-6125.2001.PubMedPubMed CentralView ArticleGoogle Scholar
- Osterberg S, Del Peso-Santos T, Shingler V: Regulation of Alternative Sigma Factor Use. Annu Rev Microbiol. 2010Google Scholar
- Missiakas D, Raina S: The extracytoplasmic function sigma factors: role and regulation. Mol Microbiol. 1998, 28 (6): 1059-1066. 10.1046/j.1365-2958.1998.00865.x.PubMedView ArticleGoogle Scholar
- Helmann JD: The extracytoplasmic function (ECF) sigma factors. Adv Microb Physiol. 2002, 46: 47-110.PubMedView ArticleGoogle Scholar
- Campbell EA, Tupy JL, Gruber TM, Wang S, Sharp MM, Gross CA, Darst SA: Crystal structure of Escherichia coli sigmaE with the cytoplasmic domain of its anti-sigma RseA. Mol Cell. 2003, 11 (4): 1067-1078. 10.1016/S1097-2765(03)00148-5.PubMedView ArticleGoogle Scholar
- Brauer SL, Hneihen AS, McBride JS, Wetterhahn KE: Chromium(VI) Forms Thiolate Complexes with gamma-Glutamylcysteine, N-Acetylcysteine, Cysteine, and the Methyl Ester of N-Acetylcysteine. Inorg Chem. 1996, 35 (2): 373-381. 10.1021/ic941452d.PubMedView ArticleGoogle Scholar
- Ely B: Genetics of Caulobacter crescentus. Methods Enzymol. 1991, 204: 372-384.PubMedView ArticleGoogle Scholar
- Simon R, Priefer U, Pühler A: A Broad Host Range Mobilization System for In Vivo Genetic Engineering: Transposon Mutagenesis in Gram Negative Bacteria. Nat Biotechnol. 1983, 1: 784-790. 10.1038/nbt1183-784.View ArticleGoogle Scholar
- Miller JH: Experiments in Molecular Genetics. Cold Spring Habor. 1972, New York: Cold Spring Habor Laboratory Press, ed.Google Scholar
- Tsai JW, Alley MR: Proteolysis of the McpA chemoreceptor does not require the Caulobacter major chemotaxis operon. J Bacteriol. 2000, 182 (2): 504-507. 10.1128/JB.182.2.504-507.2000.PubMedPubMed CentralView ArticleGoogle Scholar
- Evinger M, Agabian N: Envelope-associated nucleoid from Caulobacter crescentus stalked and swarmer cells. J Bacteriol. 1977, 132 (1): 294-301.PubMedPubMed CentralGoogle Scholar
- Livak KJ, Schmittgen TD: Analysis of relative gene expression data using real-time quantitative PCR and the 2(−Delta Delta C(T)) Method. Methods. 2001, 25 (4): 402-408. 10.1006/meth.2001.1262.PubMedView ArticleGoogle Scholar
- Towbin H, Staehelin T, Gordon J: Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. 1979. Biotechnology. 1992, 24: 145-149.PubMedGoogle Scholar
- Gober JW, Shapiro L: A developmentally regulated Caulobacter flagellar promoter is activated by 3' enhancer and IHF binding elements. Mol Biol Cell. 1992, 3 (8): 913-926.PubMedPubMed CentralView ArticleGoogle Scholar
- Corpet F: Multiple sequence alignment with hierarchical clustering. Nucleic Acids Res. 1988, 16 (22): 10881-10890. 10.1093/nar/16.22.10881.PubMedPubMed CentralView ArticleGoogle Scholar
- Letunic I, Doerks T, Bork P: SMART 7: recent updates to the protein domain annotation resource. Nucleic Acids Res. 2012, 40 (Database issue): D302-D305.PubMedPubMed CentralView ArticleGoogle Scholar
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