RpiR Homologues May Link Staphylococcus aureus RNAIII Synthesis and Pentose Phosphate Pathway Regulation

Yefei Zhu; Renu Nandakumar; Marat R Sadykov; Nandakumar Madayiputhiya; Thanh T Luong; Rosmarie Gaupp; Chia Y Lee; Greg A Somerville

doi:10.1128/JB.05930-11

. 2011 Nov;193(22):6187–6196. doi: 10.1128/JB.05930-11

RpiR Homologues May Link Staphylococcus aureus RNAIII Synthesis and Pentose Phosphate Pathway Regulation ^{^▿}^†

Yefei Zhu ¹, Renu Nandakumar ², Marat R Sadykov ¹, Nandakumar Madayiputhiya ², Thanh T Luong ³, Rosmarie Gaupp ¹, Chia Y Lee ³, Greg A Somerville ^1,^*

PMCID: PMC3209195 PMID: 21926234

Abstract

Staphylococcus aureus is a medically important pathogen that synthesizes a wide range of virulence determinants. The synthesis of many staphylococcal virulence determinants is regulated in part by stress-induced changes in the activity of the tricarboxylic acid (TCA) cycle. One metabolic change associated with TCA cycle stress is an increased concentration of ribose, leading us to hypothesize that a pentose phosphate pathway (PPP)-responsive regulator mediates some of the TCA cycle-dependent regulatory effects. Using bioinformatics, we identified three potential ribose-responsive regulators that belong to the RpiR family of transcriptional regulators. To determine whether these RpiR homologues affect PPP activity and virulence determinant synthesis, the rpiR homologues were inactivated, and the effects on PPP activity and virulence factor synthesis were assessed. Two of the three homologues (RpiRB and RpiRC) positively influence the transcription of the PPP genes rpiA and zwf, while the third homologue (RpiRA) is slightly antagonistic to the other homologues. In addition, inactivation of RpiRC altered the temporal transcription of RNAIII, the effector molecule of the agr quorum-sensing system. These data confirm the close linkage of central metabolism and virulence determinant synthesis, and they establish a metabolic override for quorum-sensing-dependent regulation of RNAIII transcription.

INTRODUCTION

Staphylococcus aureus is an important human and animal pathogen that is capable of infecting nearly all host anatomic sites. The pathogenicity of S. aureus depends on its ability to synthesize virulence factors that facilitate colonization, immune evasion, and nutrient acquisition. Virulence factor synthesis is controlled by a complex network of regulatory proteins, including the agr quorum-sensing system and the SarA family of regulators (6, 29). In addition, tricarboxylic acid (TCA) cycle activity is important for the regulation of staphylococcal virulence factor synthesis (33, 39–41, 48). Since the two most common types of regulation are genetic regulation and metabolic regulation, TCA cycle-dependent regulation most likely occurs via one or both of these mechanisms. Genetic regulation occurs through the repression or induction of enzyme synthesis, while metabolic regulation controls enzyme activity through the availability of substrates and cofactors. An example of staphylococcal metabolic regulation is the synthesis of capsular polysaccharide, which is regulated by TCA cycle activity through the supply of phosphoenolpyruvate for gluconeogenesis (33). Other virulence factors, such as polysaccharide intercellular adhesin (PIA), are genetically regulated by TCA cycle activity through transcriptional repression of the operon encoding the enzymes of PIA biosynthesis (i.e., icaADBC) (34, 44). This TCA cycle-dependent genetic regulation likely depends on response regulators that react to metabolic changes associated with TCA cycle activity fluctuations (35, 41).

In Staphylococcus epidermidis, TCA cycle stress (i.e., any environmental stressor, such as iron limitation, that is capable of altering TCA cycle activity) increases the intracellular ribose concentration, indicating that carbon flow through the pentose phosphate pathway (PPP) is increased during TCA cycle stress (35). This suggests that if there is a regulator that can respond to the concentration of ribose, or another PPP metabolite, then the activity of that regulator will likely be altered. The PPP-responsive regulator prototype, RpiR, was first identified in Escherichia coli as a regulator of ribose-5-phosphate isomerase B (rpiB), which catalyzes the reversible isomerization of ribulose-5-phosphate and ribose-5-phosphate (42). Members of the RpiR family often act as transcriptional regulators of sugar catabolism, and RpiR homologues have been identified as repressors and activators in both Gram-negative and Gram-positive bacteria, including E. coli, Pseudomonas putida, and Bacillus subtilis (8, 42, 46). As sugar-responsive regulators, members of the RpiR family of proteins have N-terminal helix-turn-helix DNA binding motifs and C-terminal sugar isomerase binding (SIS) domains (1).

TCA cycle stress alters the intracellular ribose concentration in S. epidermidis and also alters the temporal expression of virulence factors in S. epidermidis and S. aureus (34, 35, 39, 40). These observations led us to hypothesize that an RpiR homologue may link the PPP to virulence factor regulation in staphylococci. A search of the S. aureus strain Mu50 genome (18) returned three open reading frames (ORFs) with significant amino acid homology to RpiR (21 to 23% amino acid identity and 45 to 46% amino acid similarity): SAV0317, SAV0193, and SAV2315. For simplicity, these homologues were designated RpiRA (SAV0317), RpiRB (SAV0193), and RpiRC (SAV2315). To determine whether these RpiR homologues link the PPP to virulence factor synthesis in S. aureus, three single mutants, three double mutants, and a triple mutant of the rpiR homologues were constructed in S. aureus strain UAMS-1, and the effects on PPP activity, RNAIII transcription, capsular polysaccharide biosynthesis, PIA accumulation, and the ability to form a biofilm were assessed.

MATERIALS AND METHODS

Bacterial strains and growth conditions.

The strains and plasmids used in this study are listed in Table 1. E. coli strains were grown in 2× YT broth (36) or on 2× YT agar, and S. aureus strains were grown in tryptic soy broth (TSB) (BD Biosciences) or on TSB containing 1.5% agar. TSB is a complex medium that contains glucose (0.25%, wt/vol) and stachyose. Stachyose is a plant carbohydrate that S. aureus cannot catabolize. Unless otherwise stated, all bacterial cultures were inoculated at 1:200 from an overnight culture (normalized for growth) into TSB, incubated at 37°C, and aerated at 225 rpm with a flask-to-medium ratio of 10:1. Antibiotics were purchased from Fisher Scientific or Sigma Chemical and, when used, were used at the following concentrations: for E. coli, ampicillin at 100 μg/ml; for S. aureus, erythromycin at 8 or 10 μg/ml, chloramphenicol at 10 to 15 μg/ml, and tetracycline at 10 μg/ml.

Table 1.

Strains and plasmids used in this study

Plasmid or strain	Relevant genotype and/or characteristic(s)^a	Source or reference
Plasmids
pBluescript II KS(+)	E. coli phagemid cloning vector	Stratagene
pTS1	S. aureus-E. coli temperature-sensitive shuttle vector; Amp^r Cam^r	14
pTS1-d	Derivative of pTS1 with deletion of plasmid-encoded 3′ region of ermC	34
pEC4	pBluescript II KS(+) with ermB inserted into ClaI site	3
pJF12	Plasmid pCR2.1 containing tetM; Amp^r Min^r	J. Finan and G. Archer
pYF-7	pBluescript II KS(+) containing a portion of SAV0317	This study
pYF-8	pYF-7 containing an ermB cassette inserted into the NdeI site of SAV0317	This study
pYF-9	SAV0317::ermB product from pYF-8 inserted into BamHI/SacI-digested pTS1	This study
pYM-4	Derivative of pTS1 with SAV0193::cat fragment	This study
pYM-5	Derivative of pTS1 with SAV2315::tetM fragment	This study
pCL15	Expression vector; derivative of pSI-1; Cam^r	Chia Lee
pCL15-ermB	Replacement of cat with ermB in expression plasmid pCL15; Erm^r	This study
pYF-10	pCL15 with the SAV0317 gene under the control of the P_spac promoter; Cam^r	This study
pYF-11	pCL15-ermB with the SAV0193 gene under the control of the P_spac promoter; Erm^r	This study
pYF-12	pCL15 with the SAV2315 gene under the control of the P_spac promoter; Cam^r	This study
Strains
RN4220	Restriction-negative S. aureus	30
DH5α	E. coli cloning host	Invitrogen
UAMS-1	S. aureus clinical isolate	13
UAMS-1-rpiRA	SAV0317 insertion mutant of UAMS-1; Erm^r	This study
UAMS-1-rpiRB	SAV0193 deletion mutant of UAMS-1; Cam^r	This study
UAMS-1-rpiRC	SAV2315 deletion mutant of UAMS-1; Min^r	This study
UAMS-1-rpiRAB	SAV0317 SAV0193 double mutant of UAMS-1; Erm^r Cam^r	This study
UAMS-1-rpiRAC	SAV0317 SAV2315 double mutant of UAMS-1; Erm^r Min^r	This study
UAMS-1-rpiRBC	SAV0193 SAV2315 double mutant of UAMS-1; Cam^r Min^r	This study
UAMS-1-rpiRABC	SAV0317 SAV0193 SAV2315 triple mutant of UAMS-1; Erm^r Cam^r Min^r	This study

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Amp^r, ampicillin resistant; Cam^r, chloramphenicol resistant; Erm^r, erythromycin resistant; Min^r, minocycline resistant.

Construction of S. aureus rpiR mutants.

To inactivate rpiRA (ORF SAV0317), a 2.559-kb fragment was PCR amplified using primers SAV0316-BamHI and SAV0318-SacI (Table 2), and the product was cloned into the SmaI site of pBluescript II KS(+) (Stratagene) to generate plasmid pYF-7. The ermB cassette of pEC4 was amplified using primers pEC4ErmBNdeIF and pEC4ErmBNdeIR (Table 2) and was ligated into the NdeI site within rpiRA of pYF-7 to yield plasmid pYF-8. The rpiRA::ermB fragment of pYF-8 was cloned into the BamHI and SacI sites of pTS1 to create plasmid pYF-9. The temperature-sensitive plasmid pYF-9 was isolated from S. aureus strain RN4220 and was introduced into strain UAMS-1 by electroporation. Transformed bacteria were used to construct the rpiRA mutant using the temperature shift method of Foster (11).

Table 2.

Primers used in this study

Primer target	Primer designation	Nucleotide sequence (5′–3′)
SAV0317	SAV0316-BamHI	GCTGGATCCCGACTGAACAATGAACGCCTAAGTC
SAV0317	SAV0318-SacI	CCTGAGCTCATCAACGCCGGACAACAAAAGTG
ermB	pEC4ErmBNdeI-f	GCGCATATGCGTTAGATTAATTCCTACCAGTGAC
ermB	pEC4ErmBNdeI-r	GCGCATATGCTCATAGAATTATTTCCTCCCG
SAV0193	BamHI-SAV0192-f	CCAGGATCCAGAACGAATTATTGCTGCAGTAGG
SAV0193	cat-SAV0193-r	CCACTTTATCCAATTTTCGTTTGTTGTTCACCGTCATATCAATGATTTTATGTGG
cat	SAV0193-cat-f	CCACATAAAATCATTGATATGACGGTGAACAACAAACGAAAATTGGATAAAGTGGG
cat	SAV0193-cat-r	GCAAGATGCTTCCGGTAATTATCAAGCGACTGTAAAAAGTACAGTCGGC
SAV0193	cat-SAV0193-f	GCCGACTGTACTTTTTACAGTCGCTTGATAATTACCGGAAGCATCTTGC
SAV0193	SacI-SAV0195-r	GCAGAGCTCGTTGAATAAGTGCTTCTACCGCATAC
SAV2315	BamHI-SAV2312-f	CCAGGATCCGATTCCTAAACTATGGAGTCGATGGG
SAV2315	tetM-SAV2315-r	CGTAAGAGCATATTTGTAAAGGAATCTCCGAGACCTCATTTTAATCACCTTTTGAGG
tetM	SAV2315-tetM-f	CCTCAAAAGGTGATTAAAATGAGGTCTCGGAGATTCCTTTACAAATATGCTCTTACG
tetM	SAV2315-tetM-r	GAAGTTGTTGCTCCCATATGCATCCGATCTCCTCCTTTCCACTTTAATTC
SAV2315	tetM-SAV2315-f	GAATTAAAGTGGAAAGGAGGAGATCGGATGCATATGGGAGCAACAACTTC
SAV2315	KpnI-SAV2316-r	GAAGGTACCAATGGATTGTAGTTGGTATGAGTGAG
RNAIII	SARNAIII-f	GAAGGAGTGATTTCAATGGCACAAG
RNAIII	SARNAIII-r	GGCTCACGACCATACTTATTATTAAGGG
rpiA	rpiaf	GTGACATGACGCTGGGAATTGG
rpiA	rpiar	GTATCCTGTCTCAAACACACCTGTCAG
SAV1505	SAV1505f	GCACCACAATTCTTTGGCGTTATTTC
SAV1505	SAV1505r	AGTACGAATATAGAATGGTACACCAGCC
SAV0317	BamHI-SD-rpiR-f	CAAGGATCCATTAAGATGAAGGGGTGACACAATG
SAV0317	SacI-rpiR-r	CAAGAGCTCAATCACGATGATTGTCTACAGTTGC
SAV0193	BamHI-SAV0193-f	CTAGGATCCATGACAAATATTTTATATCGCATTGATAAACAGTTGAG
SAV0193	SAV0193-r	CAAACAACTGAATCACATCAAAAACTTCAATTG
SAV2315	BamHI-SAV2315-f	CTAGGATCCATGTCAAACGTACTAACAGAAATAGATAGTCAATATCC
SAV2315	SAV2315-r	GCGTATGTTATACAAGATAAAAGACATGTAAGCTTTG

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To inactivate rpiRB (ORF SAV0193), the technique of gene splicing by overlap extension (15) was used to replace a 741-bp internal region of rpiRB with the cat gene from plasmid pTS1. For PCR, genomic DNA from S. aureus strain UAMS-1 was used as a template for the amplification of regions flanking rpiRB. PCR primers BamHI-SAV0192-f and cat-SAV0193-r (Table 2) were used for the amplification of a 1.5-kb region upstream of rpiRB, and a 1.5-kb region of the rpiRB downstream region was amplified using primers cat-SAV0193-f and SacI-SAV0195-r (Table 2). The cat gene was amplified from pTS1 using primers SAV0193-cat-f and SAV0193-cat-r (Table 2). The resulting 3.9-kb PCR product consisted of an internal 816-bp cat gene with DNA flanking the rpiRB gene. The 3.9-kb PCR product contained BamHI and SacI sites that were used for ligation into pTS1-d digested with SacI and BamHI to generate pYM-4. Plasmid pYM-4 was used to construct an rpiRB mutant (UAMS-1-rpiRB::cat) by using the temperature shift method of Foster (11).

Gene splicing by overlap extension was used to replace a 614-bp internal region of rpiRC (ORF SAV2315) with the tetM gene from plasmid pJF-12 (Table 1). A 1.5-kb region upstream of rpiRC was amplified using primers BamHI-SAV2312-f and tetM-SAV2315-r (Table 2), and primers tetM-SAV2315-f and KpnI-SAV2316-r (Table 2) were used for amplification of a 1.6-kb downstream region. tetM was amplified from pJF-12 using primers SAV2315-tetM-f and SAV2315-tetM-r (Table 2). A 5.4-kb PCR product consisting of the 2.3-kb tetM gene and DNA flanking the rpiRC gene with BamHI and KpnI sites was inserted into pTS1-d digested with BamHI and KpnI to generate pYM-5. Plasmid pYM-5 was used to construct a strain UAMS-1 rpiRC mutant (UAMS-1-rpiRC::tetM) by using temperature shifts. To minimize the possibility that any phenotype(s) was the result of random mutations occurring during temperature shifts, all resulting mutations were back-crossed into wild-type strain UAMS-1 using transducing phage φ85 (11). All mutants were verified by PCR and Southern blot analysis. The rpiR double mutants and triple mutant were constructed using transducing phage φ85.

Construction of rpiR complementing plasmids.

Plasmids pCL15 and pCL15-ermB (Table 1), containing a P_spac promoter, were used to construct the rpiRA, rpiRB, and rpiRC complementation plasmids pYF-10, pYF-11, and pYF-12. The promoterless genes from S. aureus strain UAMS-1 were PCR amplified using the primers listed in Table 2 and were ligated into plasmid pCL15 or pCL15-ermB. Plasmids were isolated from S. aureus strain RN4220 and were electroporated into the UAMS-1 rpiRA, rpiRB, and rpiRC mutants.

Northern blot analysis.

To determine whether rpiR inactivation affected the transcription of PPP genes, Northern blot analysis was performed on ribose 5-phosphate isomerase A (rpiA) and glucose-6-dehydrogenase (G6PD) (zwf; ORF SAV1505). RNAIII transcript levels were also evaluated in order to determine the effect of rpiR inactivation on the agr system. Northern blotting was performed as described previously (36), except that total RNA was isolated using the FastRNA Pro Blue kit (Qbiogene) and was purified using an RNeasy kit (Qiagen). Probes for Northern blotting were generated by PCR amplification of unique internal regions of RNAIII, rpiA, and zwf (Table 2) and were labeled using the North2South random prime labeling kit (Pierce). Detection was performed using the chemiluminescent nucleic acid detection module (Pierce).

Glucose-6-dehydrogenase activity assay.

To determine whether rpiR inactivation affected PPP activity, G6PD activity was measured as described previously (7). Protein concentrations were determined using a modified Lowry assay (Pierce Chemical).

Western blot analysis.

To determine whether rpiR inactivation affected protein A biosynthesis, protein A was collected as described previously (45), and Western blot analysis was performed (43).

Hemolytic assay.

Strain UAMS-1 is lysogenized with an hlb-converting phage and has a nonsense mutation in hla; hence, it does not produce the major hemolysins alpha-toxin and beta-toxin (4). The mRNA for delta-toxin is contained within RNAIII (16). To determine whether inactivation of any rpiR homologue altered delta-toxin accumulation, a semiquantitative microtiter plate assay was carried out as described previously (10). Briefly, horse red blood cells (RBCs; Colorado Serum Company) were washed three times in phosphate-buffered saline (PBS) (pH 7.2) and were suspended at 2% (vol/vol) in PBS. Bacteria were grown in TSB for 15 h and were then centrifuged at 16,100 × g for 5 min; supernatants were collected, and 2-fold serial dilutions were made in PBS. Hemolytic assays were started by mixing 100 μl of freshly prepared 2% horse RBCs with 100 μl of serial 2-fold dilutions of the appropriate culture supernatant. The microtiter plates were incubated at 37°C for 30 min, followed by 12 h at 4°C. After incubation, the supernatant fluids were collected, and hemoglobin release was measured at 595 nm. Each experiment was repeated three times, and the mean and standard error of the mean (SEM) were calculated.

Polystyrene primary attachment assay.

The primary attachment assay was performed as described by Lim et al. (19). Briefly, bacterial cultures (2 h postinoculation) were diluted into TSB to yield approximately 300 CFU. Bacteria were poured onto polystyrene petri dishes (Fisher Scientific) and were incubated at 37°C for 30 min. Following incubation, the petri dishes were rinsed three times with sterile PBS (pH 7.5) and were covered with 15 ml of TSB containing 0.8% agar maintained at 48°C. The percentage of bacteria attached to the polystyrene was defined as the number of CFU remaining in the petri dishes after washing compared to the number of CFU in unwashed TSB plates. The experiment was repeated three times, and the mean and SEM were calculated.

Capsule immunoblot assay.

To determine whether rpiR inactivation altered capsule biosynthesis, capsule accumulation was quantified by immunoblotting as described previously (22), except that immunoblots were developed using a chemiluminescent horseradish peroxidase (HRP) substrate (Millipore). For the capsule blots, bacteria (1.25 optical density at 660 nm [OD₆₆₀] units) were harvested after overnight growth in tryptic soy broth at 37°C, with a flask-to-medium ratio of 20:1, and were aerated at 225 rpm.

PIA immunoblot assay.

PIA accumulation was determined after 2, 4, and 6 h of growth as described previously (47).

Biofilm formation in flow cell chambers.

S. aureus strains were grown in flow cell chambers (Stovall Life Science) as described previously (47). To assess bacterial growth, at 12 h postinoculation and every 4 h thereafter, effluent samples were collected, the pH was measured, and the chamber was photographed.

Proteomic analyses.

Bacterial cells (2 h and 6 h postinoculation) were harvested by centrifugation and were suspended in 1.0 ml of lysis buffer containing 50 mM ammonium bicarbonate, 8 M urea, and 1.5 mM phenylmethylsulfonyl fluoride (PMSF). The samples were homogenized for 40 s at 6.0 m/s in a FastPrep instrument (MP Biomedical), and the lysate was centrifuged for 5 min at 20,800 × g and 4°C. Bacterial proteins were subjected to in-solution trypsin digestion as described previously (28). Briefly, the proteins were reduced with 10 mM dithiothreitol and were alkylated with 40 mM iodoacetamide, followed by trypsin (Roche) digestion (trypsin/protein ratio, 1:50) overnight at 37°C. The tryptic peptides were desalted and concentrated using PepClean C₁₈ spin columns according to the manufacturer's instructions (Thermo Scientific).

Fully automated 2-dimensional (2D) chromatographic experiments were performed with an UltiMate 3000 Proteomics multidimensional liquid chromatography (MDLC) system (Dionex Corporation) integrated with a nanospray source and an LCQ (liquid chromatography quadrupole) Fleet ion trap mass spectrometer (Thermo Scientific). The first-dimension LC separation (strong cation-exchange [SCX] chromatography) with fraction collection was followed by the second-dimension LC separation (reverse-phase chromatography) and detection by tandem mass spectrometry (MS-MS). The first-dimension separation was performed on an SCX column (polysulfoethyl; inside diameter [i.d.], 1 mm; length, 15 cm; particle size, 5 μm; pore size, 300 Å; Dionex). Twenty microliters of the sample was loaded onto the first-dimension SCX column and was eluted using a salt gradient (0 to 600 mM) for 45 min. Based on the UV absorbance of the eluted peptides, selected fractions were subjected to second-dimension analysis. The second-dimension separation included on-line sample preconcentration and desalting using a monolithic C₁₈ trap column (PepMap; i.d., 300 μm; length, 5 mm; particle size, 5 μm; pore size, 100 Å; Dionex). The sample was loaded onto the monolithic trap column at a flow rate of 300 nl/min. The desalted peptides were then eluted and separated on a C₁₈ PepMap column (i.d., 75 μm; length, 15 cm; particle size, 3 μm; pore size, 100 Å) by applying an acetonitrile (ACN) gradient (ACN plus 0.1% formic acid; 90-min gradient at a flow rate of 300 nl/min) and were introduced into the mass spectrometer using the nanospray source. The LCQ Fleet mass spectrometer was operated with the following parameters: nanospray voltage, 2.0 kV; heated capillary temperature, 200°C; full-scan m/z range, 400 to 2,000. The mass spectrometer was operated in the data-dependent mode with 4 MS-MS spectra for every full scan, 5 microscans averaged for full scans and MS-MS scans, a 3 m/z isolation width for MS-MS isolations, and 35% collision energy for collision-induced dissociation.

The MS-MS spectra were searched against the S. aureus MRSA252 database using MASCOT (version 2.2; Matrix Science). The database search criteria were as follows: enzyme, trypsin; missed cleavages, 2; mass, monoisotropic; fixed modification, carbamidomethyl (C); peptide tolerance, 1.5 Da; MS-MS fragment ion tolerance, 1 Da. Probability assessment of peptide assignments and protein identifications were accomplished by Scaffold (version 3.0; Proteome Software Inc.). Only peptides with ≥90% probability were considered. Criteria for protein identification included the detection of at least 2 unique identified peptides and a protein probability score of ≥90%. Relative quantitation of proteins was done by use of the label-free method of spectral counting (20) using the normalized spectral counts for each protein. For ease of reference, the NCBI GenInfo Identifier (gi) numbers have been included in this report and in Tables S1 and S2 in the supplemental material.

Hydrogen peroxide susceptibility assay.

To determine if rpiR inactivation affects hydrogen peroxide susceptibility, S. aureus strain UAMS-1 and all of the rpiR mutant strains were grown in TSB for 15 h and were then diluted to an OD₆₀₀ of 0.05 into sterile medium containing increasing concentrations of hydrogen peroxide (Fisher Scientific). Cultures were grown at 37°C with shaking (225 rpm) for 4 h. Bacterial densities were determined by measuring the OD₆₀₀.

RESULTS

Characterization of rpiR mutants.

Analysis of the Mu50 genome (18) revealed the presence of three RpiR homologues: RpiRA (SAV0317), RpiRB (SAV0193), and RpiRC (SAV2315). Each rpiR homologue was deleted either individually or in tandem with one or both of the other rpiR homologues in strain UAMS-1 (Table 1). To assess the effects of inactivation of the rpiR homologue genes on growth, the optical densities and pH of the culture medium (TSB) were measured over time (Fig. 1). Inactivation of any single rpiR homologue in UAMS-1 did not alter the growth rate, growth yield, or pH profile of the culture medium (Fig. 1A). Similarly, the double and triple mutants had growth rates and growth yields equivalent to those of the wild-type strain UAMS-1 (Fig. 1B). Of note, the pH profile of the culture medium for the triple mutant showed an increased rate of alkalization relative to that for the wild-type strain, suggesting that this strain had an increased rate of acetic acid utilization or an increase in ammonia generation due to amino acid catabolism (Fig. 1B). These results demonstrate that the growth of the rpiR mutants is equivalent to that of the isogenic wild-type strain.

Fig. 1. — Deletion of any *rpiR* homologue in strain UAMS-1 does not alter the growth profile in TSB medium. (A) Growth curves and culture medium pH profiles are shown for strain UAMS-1 and for the UAMS-1-*rpiRA*, UAMS-1-*rpiRB*, and UAMS-1-*rpiRC* mutants (A) and the UAMS-1-*rpiRAB*, UAMS-1-*rpiRAC*, UAMS-1-*rpiRBC*, and UAMS-1-*rpiRABC* mutants (B).

RpiR homologues regulate PPP activity.

As stated above, RpiR was first identified in E. coli as a repressor of the PPP gene rpiB (42). Similarly, the Pseudomonas putida RpiR homologue HexR regulates zwf, which codes for glucose 6-phosphate dehydrogenase (G6PD), the rate-controlling enzyme of the PPP (8). These data led us to hypothesize that one or more of the RpiR homologues in S. aureus would regulate the transcription of PPP genes. To test this hypothesis, transcription of the PPP genes rpiA (ribose-5-phosphate isomerase A) and zwf (sav1505; coding for G6PD) in the rpiR mutant strains was assessed by Northern blot analysis of total RNA isolated during the exponential phase of growth (2 h) (Fig. 2 A and data not shown). Deletion of rpiRB or rpiRC decreased the transcription of both rpiA and zwf relative to that in the parental strain UAMS-1; however, rpiRA inactivation had only a minor effect on rpiA and zwf mRNA levels (Fig. 2A and data not shown). Complementation of the UAMS-1 rpiRB and rpiRC mutants increased the levels of rpiA mRNA (Fig. 2B), confirming that the transcriptional changes are due to the inactivation of the mutated rpiR genes. Interestingly, deletion of rpiRA in either an rpiRB or an rpiRC mutant strain restored the level of rpiA mRNA to that found in strain UAMS-1 (Fig. 2A), suggesting an antagonistic effect between RpiRA and both RpiRB and RpiRC. Because zwf mRNA migrates on an agarose gel near rRNA, and in order to confirm the Northern blot data, the activity of G6PD was assessed in the wild-type and rpiR mutant strains. In agreement with the Northern blot data, mutation of rpiRB or rpiRC led to decreased G6PD enzymatic activity in the exponential-growth phase (2 h) (see Fig. S1 in the supplemental material) relative to that for the wild-type strain UAMS-1. Also consistent with the Northern blot data are the antagonistic effects of RpiRA on G6PD activity in both the rpiRB and rpiRC mutant backgrounds. In contrast to the findings for the exponential-growth phase, only rpiRB inactivation significantly decreased G6PD enzymatic activity during the post-exponential-growth phase (8 h) relative to that for the wild-type strain (see Fig. S1). Overall, these data demonstrated that RpiRB and RpiRC have a positive regulatory function in PPP regulation and that RpiRA is antagonistic to this function.

Fig. 2. — Inactivation of *rpiR* homologues alters *rpiA* mRNA accumulation. (A) Northern blot analysis demonstrating that inactivation of *rpiRB* or *rpiRC* decreases the transcription of ribose phosphate isomerase A (*rpiA*). (B) Northern blot analysis demonstrating that complementation of *rpiRB* or *rpiRC* restores *rpiA* transcription. Ethidium bromide-stained agarose gels showing 23S and 16S rRNA are included in each panel to demonstrate the equivalent loading of total RNA. The results are representative of at least two independent experiments.

Inactivation of rpiRC delays biofilm development and decreases the synthesis of cell wall-associated virulence determinants.

Nuclear magnetic resonance (NMR) metabolomic analysis indicated that the intracellular ribose concentration in S. epidermidis changes in response to stressors that induce biofilm formation and PIA accumulation (35). Because the RpiR homologues have been reported to respond to PPP intermediates in other bacteria (8), we assessed the effects of rpiR inactivation on biofilm formation and PIA accumulation (Fig. 3 A and data not shown). The deletion of any rpiR homologue, singly or in tandem, did not significantly alter the accumulation of PIA (data not shown). In S. aureus, biofilms can form in the absence of PIA biosynthesis (2); therefore, the lack of any significant effect of rpiR inactivation on PIA accumulation did not preclude the possibility that one or more of the RpiR homologues would affect biofilm formation. Consistent with this premise, deletion of rpiRC delayed biofilm maturation (Fig. 3A). While biofilm maturation was delayed, the gross morphologies of the biofilms formed by the wild-type and rpiRC mutant strains were similar after 24 h of growth. The delay in biofilm maturation and the absence of any attenuation of PIA accumulation were consistent with a defect in bacterial attachment or adhesion. To determine whether inactivation of rpiRC decreased adhesin synthesis, polystyrene attachment assays were used to assess the abilities of wild-type and rpiR mutant strains to adhere to surfaces (Fig. 3B). In agreement with the delay in biofilm formation, strains containing a mutation in rpiRC had a significantly decreased ability to attach to polystyrene relative to that of the parental strain (Fig. 3B). Taken together, these data suggest that the synthesis of cell wall-associated adhesins was decreased by rpiRC inactivation.

Fig. 3. — Deletion of *rpiRC* delays biofilm maturation by inhibiting adhesion and the synthesis of cell-associated virulence determinants. (A) Growth of *S. aureus* strains UAMS-1, UAMS-1-*rpiRA*, UAMS-1-*rpiRB*, UAMS-1-*rpiRC*, and UAMS-1-*rpiRABC* in three-chamber flow cells. Bacterial strains were grown at 37°C with a continuous flow (0.5 ml min⁻¹ per chamber) of TSB containing 0.5% glucose and 3% NaCl. The results are representative of at least two independent experiments. (B) Adhesion of *S. aureus* strains to polystyrene. The data are presented as means and SEMs for three independent experiments. Significant differences, as determined using Student's t test, are indicated by asterisks (**, P < 0.01; *, P < 0.05). (C) Western blot analysis of protein A. The blot is representative of three independent experiments.

The association of protein A with biofilm formation (26) and the decreased ability of rpiR mutant strains to adhere to polystyrene suggested that cell-associated adhesin synthesis was impaired by inactivation of one or more RpiR homologues. Protein A is synthesized primarily during the exponential-growth phase and is considered representative of cell wall-associated protein synthesis. To determine whether rpiR inactivation altered the exponential-growth-phase expression of protein A and potentially of other cell wall-associated proteins, the exponential-growth-phase accumulation of protein A was assessed by Western blotting (Fig. 3C). Mutations in either rpiRA or rpiRB did not affect the accumulation of protein A relative to that for strain UAMS-1; however, inactivation of rpiRC completely inhibited the exponential-growth-phase accumulation of protein A (Fig. 3C). Interestingly, rpiRA inactivation did not antagonize the expression of protein A in the rpiRC mutant background, suggesting that the antagonistic effects of RpiRA are confined to regulation of the PPP. In total, these data suggest that RpiRC acts as a regulatory bridge between the PPP and virulence factor synthesis in S. aureus.

RpiRC represses RNAIII transcription or message stability.

RNAIII is the effector RNA of the agr quorum-sensing system and a negative regulator of protein A (spa) (29). Mutation of rpiRC eliminated the exponential-growth-phase accumulation of protein A (Fig. 3C), raising the possibility that RNAIII transcription or stability was increased. To determine whether rpiR inactivation affected RNAIII levels, Northern blot analysis of RNAIII was performed on all rpiR mutant strains throughout a 12-h growth cycle (Fig. 4 A and B). As expected, rpiRC inactivation increased the RNAIII transcript level relative to that for the parental strain during the exponential-growth phase (2 h) (Fig. 4A). Complementation of the rpiRC mutation with pYF-12 decreased the level of RNAIII relative to that for the rpiRC mutant strain, confirming that the increased RNAIII level was due to rpiRC inactivation (Fig. 4B). In agreement with the results of the Western blot analysis of protein A and the attachment assays (Fig. 3B and C), we did not observe an antagonistic effect of rpiRA inactivation on the exponential-growth-phase (2 h) transcription or stability of RNAIII in either an rpiRB or an rpiRC mutant background (Fig. 4A). Although RNAIII levels were largely independent of RpiRA or RpiRB in the exponential-growth phase, rpiRB inactivation increased the post-exponential-growth-phase RNAIII transcript levels (Fig. 4B). RNAIII is both a riboregulator and the coding sequence for delta-toxin (16); therefore, if RNAIII levels are increased, it is likely that delta-toxin synthesis is increased. (Strain UAMS-1 is lysogenized with an hlb-converting phage and has a nonsense mutation in hla; hence, it does not produce the major hemolysins alpha-toxin and beta-toxin [4].) By use of a hemolytic titer assay, the increased RNAIII levels correlated with an increase in hemolysis activity (Fig. 5 A). In total, these data indicate that RpiRC represses RNAIII transcription during the exponential-growth phase, while RpiRB represses RNAIII transcription during the post-exponential-growth phase.

Fig. 4. — Deletion of *rpiRC* increases the transcription and/or stability of RNAIII. (A) Temporal Northern blot analysis of RNAIII. (B) Complementation of *rpiR* homologues moderately restores RNAIII levels after 2 h of growth. Ethidium bromide-stained agarose gels showing 23S and 16S rRNA are included in each panel to demonstrate the equivalent loading of total RNA. All Northern blotting was performed at least twice using independently isolated total RNA.

Fig. 5. — Inactivation of *rpiR* homologues alters virulence factor synthesis. (A) Hemolytic activities of culture supernatants from strain UAMS-1 and the *rpiR* mutant strains against washed rabbit erythrocytes. The data are presented as the means and SEMs for three independent experiments. (B) Immunoblotting for capsule polysaccharide. The blot is representative of at least two independent experiments.

In S. aureus, strain-dependent differences in the regulation of virulence determinant biosynthesis have been reported (4, 49). To determine whether the effect of RpiR inactivation on RNAIII transcription was common to S. aureus strains from divergent genetic backgrounds, the rpiRB and rpiRC mutations were transduced into S. aureus strain SA564 (38), and Northern blot analysis of RNAIII was performed (see Fig. S2 in the supplemental material). As with strain UAMS-1, inactivation of rpiRC in strain SA564 derepressed RNAIII transcription during the exponential-growth phase and had no effect during the post-exponential-growth phase (see Fig. S2). Similarly, inactivation of rpiRB in strain SA564 had a minimal effect on RNAIII transcript levels during the exponential-growth phase. In contrast to the finding for strain UAMS-1, inactivation of rpiRB in strain SA564 had no apparent effect on RNAIII transcription during the post-exponential-growth phase. These data demonstrate that RpiRC represses the exponential-growth-phase level of RNAIII in divergent genetic backgrounds.

Inactivation of rpiRC dramatically increases capsule accumulation.

RNAIII is a positive regulator of capsule gene (cap) transcription (9, 21, 32); thus, an increase in RNAIII levels should correlate with an increase in capsule biosynthesis. To determine whether rpiR inactivation affects capsule biosynthesis, capsule accumulation was assessed by capsule immunoblotting. In agreement with the increased RNAIII levels, inactivation of all three rpiR genes increased capsule accumulation (Fig. 5B); however, the increased accumulation of capsule was most apparent in strains with a mutation in rpiRC. These data strongly suggest that the RpiR-dependent derepression of RNAIII facilitates virulence determinant expression and that the RpiR proteins act as a bridge between the PPP and virulence factor synthesis.

Inactivation of rpiRC alters the proteome.

To identify changes in cytosolic protein content in strain UAMS-1 and the rpiRC mutant strains, cell-free lysates were prepared from strains UAMS-1, UAMS-1-rpiRC, and UAMS-1- rpiRABC grown to the exponential and post-exponential phases of growth and were analyzed by 2D LC–MS-MS (see Tables S1 and S2 in the supplemental material). Although rpiRC inactivation resulted in numerous proteomic changes, we were specifically interested in changes to PPP enzymes and proteins that might clarify the increased RNAIII transcript levels. Proteomic analysis showed that the PPP enzymes transaldolase (gi∣49242155) and ribose-phosphate pyrophosphokinase (gi∣49240856) were present at lower concentrations in strains UAMS-1-rpiRC and UAMS-1- rpiRABC than in strain UAMS-1, consistent with regulation of the PPP by RpiRC. Interestingly, deletion of rpiRC increased the accumulation of ribosomal proteins (see Tables S1 and S2 in the supplemental material); however, the reason for this remains unknown. Proteomic analysis also suggested that there was an increase in the levels of proteins associated with σ^B; specifically, inactivation of rpiRC increased the concentrations of the alkaline shock protein A (Asp23; gi∣49242531) and RsbU (gi∣49242422) (see Tables S1 and S2). Because asp23 transcription is controlled exclusively by σ^B, Asp23 is used as an indicator of σ^B activity (17, 27). RsbU is a phosphatase that dephosphorylates (activates) the anti-anti-sigma factor RsbV, which then binds the anti-sigma factor RsbW in a competitive manner to increase the concentration of free σ^B (12). In addition to regulating the transcription of asp23, σ^B regulates the transcription of sarA from the sar P3 promoter (27). SarA is a positive effector of agrACDB and RNAIII transcription (5). Inactivation of rpiRC increased RNAIII levels relative to those in the wild-type strain (Fig. 4), suggesting that rpiRC inactivation might increase the availability of SarA. Consistent with this suggestion, rpiRC inactivation increased the cytosolic concentration of SarA (gi∣49240975) during both the exponential- and post-exponential-growth phases (see Tables S1 and S2). These data suggest that the increased RNAIII levels in the rpiRC mutants are due to increased availability of σ^B, which increases sarA transcription and translation, resulting in increased RNAIII transcription.

Inactivation of rpiRC decreases peroxide susceptibility.

In some strains of S. aureus, σ^B has been implicated in susceptibility to oxidative stress (12, 17). This observation and the fact that strain UAMS-1-rpiRC had higher ferritin and catalase levels than strain UAMS-1 (see Tables S1 and S2 in the supplemental material) led us to assess the susceptibilities of strain UAMS-1 and the rpiR mutants to peroxide stress (Fig. 6). As expected, inactivation of rpiRC significantly decreased the susceptibilities of strains UAMS-1-rpiRC, UAMS-1- rpiRAC, UAMS-1-rpiRBC, and UAMS-1-rpiRABC to hydrogen peroxide relative to that of strain UAMS-1 (Fig. 6). Taken together, these data demonstrate that the S. aureus RpiR family of proteins functions in cell survival under conditions of oxidative stress.

Fig. 6. — Deletion of *rpiRC* decreases the susceptibility of *S. aureus* strains to hydrogen peroxide. Data are presented as the means and SEMs for three independent experiments.

DISCUSSION

Three central metabolic pathways (i.e., glycolysis, the PPP, and the TCA cycle) provide the 13 biosynthetic intermediates needed to synthesize all macromolecules produced in bacteria. By default, virulence determinants are synthesized from these 13 biosynthetic intermediates of central metabolism; hence, virulence determinant synthesis is dependent on the endogenous or exogenous availability of these intermediates or by-products of these intermediates. Because of the importance of these intermediates, bacteria have evolved metabolite-responsive regulators (e.g., CcpA, CodY) that “sense” the availability of these intermediates or compounds derived from them (41). Not only do these metabolite-responsive regulators function to maintain metabolic homeostasis; many also regulate virulence determinant synthesis (41). Although metabolite-responsive regulators that respond to changes in the levels of glycolytic and TCA cycle intermediates or derivatives have been identified in S. aureus, none that respond to changes in the levels of PPP intermediates have been identified. To that end, three RpiR family members, RpiRA, RpiRB, and RpiRC, were identified and inactivated in S. aureus strain UAMS-1, and the phenotypic and regulatory changes associated with each RpiR homologue were characterized.

PPP regulation.

RpiRB and RpiRC positively regulate the exponential-growth-phase transcription of the PPP genes rpiA and zwf (Fig. 2A). In addition, RpiRC positively affects the expression of transaldolase and ribose-phosphate pyrophosphokinase (see Tables S1 and S2 in the supplemental material). Although RpiRB and RpiRC are paralogues, there appears to be minimal overlap in function between the two regulatory proteins, since inactivation of either rpiRB or rpiRC decreases the transcription of rpiA and zwf to the same extent (Fig. 2A). In other words, RpiRB does not compensate for the loss of RpiRC, and RpiRC does not compensate for the loss of RpiRB. Interestingly, RpiRA has only a slight effect on rpiA and zwf transcription; however, it does antagonize the regulatory effects of both RpiRB and RpiRC (Fig. 2A). In double mutants, inactivation of rpiRA restores the transcription of rpiA and, to a lesser extent, zwf to near-wild-type levels. Interestingly, this antagonism involves only RpiRB- and RpiRC-dependent regulation of rpiA and zwf, not RpiRC-dependent regulation of RNAIII (Fig. 4A and B). Taken together, these data confirm that the S. aureus RpiR homologues positively affect PPP transcription and activity.

RNAIII regulation.

Synthesis of RNAIII is under the control of the agr cell density-sensing system (31); hence, RNAIII transcription usually begins late in the exponential phase of growth (∼4 h) (Fig. 4A). The growth rates and growth yields of the UAMS-1 rpiRA, rpiRB, and rpiRC mutant strains are equivalent to those of the parental strain (Fig. 1A); thus, it was surprising to find that the transcription of RNAIII was derepressed during the early-exponential-growth phase (2 h) in strain UAMS-1-rpiRC compared to that in strain UAMS-1 (Fig. 4A). This RpiRC-dependent derepression persists into the post-exponential-growth phase (4 to 6 h) but declines thereafter (Fig. 4A). The more likely explanations for the RpiRC-dependent derepression of RNAIII transcription are either an increase in the level of expression of the agr cell density-sensing system or an agr-independent increase in the level of RNAIII transcription. Proteomic analysis of the cytosolic fractions of strains UAMS-1, UAMS-1-rpiRC, and UAMS-1-rpiRABC (see Tables S1 and S2 in the supplemental material) demonstrated that rpiRC inactivation increased the intracellular SarA concentration during the exponential (2 h)- and post-exponential (6 h)-growth phases relative to that in the parental strain UAMS-1. The increased level of SarA is likely mediated by an increase in the level of free σ^B due to enhanced RsbU phosphatase activity (gi∣49242422) (see Tables S1 and S2 in the supplemental material). We speculate that the increase in the level of free σ^B is a response to increased oxidative stress. This increase in oxidative stress would occur as carbon flow through glycolysis is increased due to the diversion of carbon away from the PPP. This leads to an increase in the reducing potential, which requires the oxidation of dinucleotides via the electron transport chain to maintain redox homeostasis. An increase in electron transport chain activity would result in an increase in the release of reactive oxygen species. This speculation is supported by proteomic analysis, which revealed increases in the levels of enzymes of glycolysis and the electron transport chain in the rpiRC and rpiRABC mutants relative to those in the wild-type strain. Consistent with an increase in free σ^B levels, proteomic analysis also revealed that rpiRC inactivation resulted in a greater accumulation of the σ^B-regulated alkaline shock protein A (gi∣49242531) in strain UAMS-1-rpiRC. These data suggest that the increased level of RNAIII in strains lacking RpiRC is due to an increase in the SarA-mediated transcription of RNAIII. While these data form the basis for one explanation of how RpiRC can regulate virulence determinant synthesis, it is an incomplete explanation, because data regarding known regulators, such as Rot (25), were not present in the proteomic analysis. That being said, these data confirm a direct linkage between central metabolism (i.e., the PPP) and three major virulence regulators (SarA, σ^B, and RNAIII) in S. aureus. Finally, these data demonstrate that putative metabolite-responsive regulators can override the normal quorum-sensing-dependent temporal pattern of virulence determinant synthesis.

Conclusions.

Richard Novick postulated in a “black-box” model (29) that an energy signal derived from intermediary metabolism would, in an unknown (i.e., black-box) fashion, regulate the transcription of the agr cell density-sensing system. Since the introduction of this black-box model, several regulators (e.g., CcpA and CodY) that link metabolism to the regulation of virulence determinants have been identified (reviewed in reference 41). In the present study, it was observed that RNAIII synthesis is coregulated with central metabolism, specifically the PPP, through the direct or indirect action of three RpiR family regulators. Although the black-box model is largely accurate, based on data presented here and in other studies (23, 24, 37), the energy signal responsible for regulating the transcription of agr is more than likely a carbon signal.

In Pseudomonas putida, the DNA binding activity of the RpiR homologue HexR is modulated by the Entner-Doudoroff pathway intermediate 2-keto-3-deoxy-6-phosphogluconate (8). The three S. aureus RpiR homologues all have sugar isomerase binding domains, suggesting that their regulatory activity may be controlled by intermediates of the PPP. Collaborative studies are under way to identify the metabolites to which the RpiR homologues bind; hopefully, this information will fill in one of the black boxes in S. aureus virulence factor regulation.

Supplementary Material

Supplemental material

supp_193_22_6187__index.html^{(1.3KB, html)}

ACKNOWLEDGMENTS

This article is a contribution of the University of Nebraska Agricultural Research Division, supported in part by funds provided through the Hatch Act and the National Institutes of Health (AI087668) to G.A.S. C.Y.L. was supported by funds provided by the National Institutes of Health (AI-37027 and AI-67857).

We thank Markus Bischoff for critical review of the manuscript and helpful discussions. Additionally, we thank the reviewers for helpful suggestions.

Footnotes

^†

Supplemental material for this article may be found at http://jb.asm.org/.

^▿

Published ahead of print on 16 September 2011.

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PERMALINK

RpiR Homologues May Link Staphylococcus aureus RNAIII Synthesis and Pentose Phosphate Pathway Regulation ▿ †

Yefei Zhu

Renu Nandakumar

Marat R Sadykov

Nandakumar Madayiputhiya

Thanh T Luong

Rosmarie Gaupp

Chia Y Lee

Greg A Somerville