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. Author manuscript; available in PMC: 2014 Sep 8.
Published in final edited form as: Mol Microbiol. 2009 Jan 23;71(5):1228–1238. doi: 10.1111/j.1365-2958.2009.06597.x

Acid stress activation of the σE stress response in Salmonella enterica serovar Typhimurium

Cécile Muller 1,§,, Iel-Soo Bang 1,†,, Jyoti Velayudhan 1,‡, Joyce Karlinsey 2, Kai Papenfort 3, Jörg Vogel 3, Ferric C Fang 1,2,*
PMCID: PMC4157319  NIHMSID: NIHMS621020  PMID: 19170886

Summary

The alternative sigma factor σE is activated by unfolded outer membrane proteins (OMPs) and plays an essential role in Salmonella pathogenesis. The canonical pathway of σE activation in response to envelope stress involves sequential proteolysis of the anti-sigma factor RseA by the PDZ proteases DegS and RseP. Here we show that σE in Salmonella enterica sv. Typhimurium can also be activated by acid stress. A σE-deficient mutant exhibits increased susceptibility to acid pH and reduced survival in an acidified phagosomal vacuole. Acid activation of σE-dependent gene expression is independent of the unfolded OMP signal or the DegS protease but requires processing of RseA by RseP. The RseP PDZ domain is indispensable for acid induction, suggesting that acid stress may disrupt an inhibitory interaction between RseA and the RseP PDZ domain to allow RseA proteolysis in the absence of antecedent action of DegS. These observations demonstrate a novel environmental stimulus and activation pathway for the σE regulon that appear to be critically important during Salmonella–host cell interactions.

Introduction

Bacteria adapt to their environment by sensing conditions and responding through the co-ordinated expression of gene networks. The alternative sigma factor σE senses perturbations in the outer membrane and periplasm and activates a subset of genes in response to heat stress (Rouviere et al., 1995; Missiakas and Raina, 1998). σE is required for Escherichia coli viability at all temperatures (De Las Penas et al., 1997), implying that the σE regulon plays important roles in addition to thermal adaptation.

Under non-stress conditions, σE is sequestered by the inner membrane-spanning anti-sigma factor RseA (Missiakas et al., 1997). The cytoplasmic domain of RseA interacts with the RNA Polymerase (RNAP) binding domain of σE, thereby inhibiting its assembly with core RNAP. Heat stress leads to the periplasmic accumulation of unfolded outer membrane proteins (OMPs) such as OmpC, which trigger the sequential proteolytic cleavage of RseA by the proteases DegS and RseP (YaeL) (Alba and Gross, 2004). DegS is an inner-membrane-anchored serine protease of the HtrA family (Clausen et al., 2002) with proteolytic and PDZ domains positioned in the periplasm. RseP is a zinc metalloprotease possessing four transmembrane segments, a periplasmic PDZ domain and an inner membrane/cytoplasmic border proteolytic domain (Kanehara et al., 2001; 2002; Koide et al., 2007). The PDZ domains have been shown to bind in a sequence-specific manner to the C-termini of target proteins and inhibit protease activity (Kanehara et al., 2003; Koide et al., 2007).

Specifically, recognition of the C-termini of unfolded OMPs by the DegS PDZ domain allows DegS to cleave the periplasmic C-terminus (site-1) of RseA (Walsh et al., 2003; Hasselblatt et al., 2007). This converts RseA into a substrate for RseP, which cleaves the transmembrane segment (site-2) of RseA (Alba et al., 2002; Kanehara et al., 2002; Akiyama et al., 2004). The glutamine (Gln)-rich region of RseA has been shown to interact with the RseP PDZ domain to inhibit RseP proteolytic activity. DegS-mediated processing removes the Gln-rich regions to relieve inhibition of RseP protease activity (Kanehara et al., 2003). In the absence of a functional RseP PDZ domain, RseA proteolysis by RseP becomes DegS-independent (Ades et al., 2003; Kanehara et al., 2003; Bohn et al., 2004). Following cleavage by RseP, the RseA/σE complex is released from the membrane, and the cytoplasmic portion of RseA is rapidly degraded by proteases such as ClpXP (Alba et al., 2002; Kanehara et al., 2002; Chaba et al., 2007).

Once released from the cytoplasmic membrane, free σE binds core RNAP and directs the resulting holoenzyme to a subset of promoters resulting in the expression of target genes, including rpoE itself, rpoH, fkpA, surA and rybB (Missiakas et al., 1996; Dartigalongue et al., 2001; Thompson et al., 2007). The rybB gene encodes a small RNA that accelerates the global decay of OMP mRNAs (Papenfort et al., 2006). RybB also appears to downregu-late expression of rpoE (Papenfort et al., 2006; Thompson et al., 2007), probably by limiting OMP-dependent upstream signals that lead to σE activation. Additional regulation is provided by the periplasmic protein RseB, which increases RseA affinity for σE (Collinet et al., 2000) and retards its cleavage by DegS (Cezairliyan and Sauer, 2007), and by the inner membrane protein RseC, which may promote σE activation (Missiakas et al., 1997; Alba and Gross, 2004). It has been suggested that RseB may limit access of RseP to RseA until RseA is first cleaved by DegS (Grigorova et al., 2004; Kim et al., 2007).

The σE pathways of Salmonella enterica sv. Typhimurium (henceforth referred to as Salmonella) and E. coli are highly conserved with regard to organization and mechanism of activation (Rhodius et al., 2006). However, σE is not essential in Salmonella (Humphreys et al., 1999; Testerman et al., 2002), allowing the construction of an rpoE null mutation and analysis of the contribution of σE to bacterial virulence. S. enterica is a globally important cause of food- and water-borne illness with clinical manifestations ranging from gastroenteritis to bacteremia and systemic infection (Ohl and Miller, 2001). Although it is not surprising that σE plays an essential role in Salmonella pathogenesis (Humphreys et al., 1999; Testerman et al., 2002; Onufryk et al., 2005), an rpoE mutant strain is unexpectedly less virulent than a degS mutant (Rowley et al., 2005), suggesting that σE might be activated by an alternative mechanism during the interaction of Salmonella with its host.

During passage through the gastrointestinal tract and replication in host phagocytic cells, Salmonella must withstand a number of stresses including nutrient deprivation, membrane damage, reactive oxygen and nitrogen species, and acid stress (Foster and Spector, 1995). Salmonella is exposed to acid stress during passage through the stomach and within the Salmonella-containing vacuole (SCV) of activated macrophages, which acidifies to pH 4–5 (Rathman et al., 1996). Acid pH also represents a signal that triggers the expression of Salmonella virulence genes required for intracellular survival (Alpuche Aranda et al., 1992; Rathman et al., 1996) and the trans-location of virulence effectors into the cytosol of host macrophages (Beuzon et al., 1999). The involvement of σE in the response of Salmonella to acid stress and the mechanism of σE activation was therefore investigated. We report here that acid pH activates σE in Salmonella via a non-canonical mechanism independent of DegS-mediated proteolysis of RseA.

Results

σE promotes Salmonella survival within the acidified vacuoles of RAW264.7 macrophages

To investigate if vacuolar acidification stimulates a σE-dependent Salmonella adaptive response in macroph-ages, bacterial survival was monitored following uptake by RAW264.7 murine macrophages with or without the addition of NH4Cl to inhibit vacuolar acidification. Wild-type bacteria (WT) exhibited approximately 10-fold reduced survival in non-acidified vacuoles (Fig. 1), consistent with previous observations, demonstrating that acidification enhances the survival of intracellular Salmonella (Rathman et al., 1996). A previous study showed that σE is required for the intracellular survival of Salmonella in macrophages (Humphreys et al., 1999), so we examined whether σE participates in acidification-dependent Salmonella survival in the SCV. An isogenic rpoE mutant strain (Testerman et al., 2002) exhibited reduced survival in the absence of NH4Cl (Fig. 1). σE enhanced Salmonella survival only within acidified vacuoles, but interestingly, not in NH4Cl-treated cells. These results implicate the σE regulon in the acid pH-triggered adaptation of Salmonella to the intracellular environment.

Fig. 1.

Fig. 1

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σE-dependent survival of Salmonella in acidified macrophage vacuoles. RAW264.7 macrophages, treated (filled symbols) or untreated (open symbols) with NH4Cl, were infected with complement-opsonized WT (diamonds) or rpoE mutant (squares) Salmonella.

σE is necessary for Salmonella tolerance of acid pH

Reduced survival of rpoE mutant Salmonella in the acidified SCV might reflect increased susceptibility of σE-deficient bacteria to acid stress per se. We therefore measured the replication of rpoE mutant Salmonella in acidified medium (pH 5.0) and found substantially delayed growth compared with WT (Fig. 2A). In addition, the survival of an rpoE mutant was dramatically reduced under more extreme acid stress conditions (pH 3.0), and survival was restored to WT levels when rpoE was provided on a complementing plasmid (Testerman et al., 2002) (Fig. 2B). During the acid tolerance response (ATR) (Foster and Spector, 1995), Salmonella exposure to relatively mild levels of acid stress (pH > 4) induces the expression of genes that enable the bacteria to tolerate more extreme acidic conditions (Foster, 1999). We therefore examined whether σE participates in the ATR. An rpoE mutation dramatically reduced Salmonella survival at pH 3.0 and eliminated adaptive tolerance (Fig. 2C), demonstrating that the ATR requires σE. Collectively, these results indicate that the σE regulon makes an important contribution to the ability of Salmonella to adapt to and withstand acidic conditions.

Fig. 2.

Fig. 2

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σE-dependent acid resistance of Salmonella.

A. Growth of WT (triangles) and rpoE mutant (squares) Salmonella was monitored at pH 7.0 (open symbols) or pH 5.0 (filled symbols).

B. Survival of stationary-phase WT (diamonds), rpoE mutant (squares) or rpoE mutant complemented with pBR3-rpoE (triangles) was determined following a 2 h acid challenge (pH 3.0).

C. The ATR was compared by adapting WT (grey bars) or rpoE mutant (white bars) at pH 5.0 for 2 h before challenge at pH 3.0.

Acid shock increases σE expression

The ATR requires de novo protein synthesis (Foster and Hall, 1991), so we hypothesized that σE itself might be acid-inducible. The expression of the rpoE gene was monitored in log- or stationary-phase cells submitted to acid stress. Quantitative real-time RT-PCR (qRT-PCR) revealed that rpoE mRNA levels begin to rise within 10 min at pH 4.5 and eventually increase 50- and 58-fold over basal levels after 60 min acid stress in log- and stationary-phase cells respectively (Fig. 3A).

Fig. 3.

Fig. 3

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Acid induction of σE expression.

A. Levels of rpoE mRNA were measured by qRT-PCR in log-phase (OD600 ~0.4) and stationary-phase (overnight) cells of the WT (grey bars) and rpoE (white bars) strains, after 10, 30 and 60 min of an acid stress (pH 4.5) compared with pH 7.0 and normalized to reference gmk mRNA.

B. β-Galactosidase expression was measured in stationary-phase cells harbouring P3rpoE–lacZ (grey bars) or P1rpoE–lacZ (white bars) fusions after re-suspension in pH 7.0 or pH 4.5 medium and incubation at the indicated time intervals.

Transcription of rpoE is initiated from three promoters. Two of these, P1 and P2, are σ70-dependent, while the P3 promoter is σE-dependent (Miticka et al., 2003). P1rpoE–lacZand P3rpoE–lacZreporter fusions were used to determine the promoter involved in acid-induction (Testerman et al., 2002). We observed that the P3rpoE–lacZ fusion was induced by acid stress whereas the activity of P1rpoE–lacZ was unchanged (Fig. 3B). These results indicate that increased rpoE transcription during acid stress results from activation of σE itself. Abrogation of acid-induced rpoE transcription in an rpoE mutant background supports this interpretation (Fig. 3A).

It should be noted that the fold-induction using lacZ transcriptional fusions was substantially lower than what was observed by qRT-PCR. This can be explained by high background expression from the plasmid-borne reporter.

Acid induction of σE is OMP- and DegS-independent in contrast to heat stress induction

The rate-limiting step in σE activation by heat stress is the sensing of unfolded OMPs by the DegS protease (Chaba et al., 2007). To analyse the mechanism of σE activation by acid stress, relative levels of σE target gene (rpoH, fkpA, surA, rpoE) expression were monitored by qRT-PCR during acid stress and compared with pH 7.0. The rpoH gene encodes σ32, which is involved in the cytoplasmic heat stress response. The FkpA and SurA proteins are peptidyl-prolyl isomerases, which function as protein folding catalysts in the periplasm. All of these σE-dependent loci were induced 20- to 80-fold at pH 4.5 in WT strain compared with the reference gene gmk (data not shown).

Papenfort et al. (2006) demonstrated that overexpression of the σE-regulated sRNA RybB promotes the decay of major OMP mRNAs by direct interaction, resulting in the rapid cessation of OMP synthesis. A plasmid (pBAD-RybB) expressing rybB under control of the arabinose-inducible ParaBAD promoter and pBAD vector control (Guzman et al., 1995) producing a short nonsense RNA (Papenfort et al., 2006) were used to compare the effects of rybB overexpression on heat and acid stress induction of σE (Fig. 4). Thus, the effect of OMP presence (no induction of rybB expression) or absence (overproduction of rybB) on the σE activation pathway was studied. The σE-regulated genes rpoH, fkpA, surA and rpoE were induced by heat stress, and RybB overproduction substantially reduced this induction, as anticipated (Fig. 4A, WT pBAD versus WT pBAD-RybB). Inactivation of rpoE or deletion of the DegS PDZ domain eliminated heat stress induction of σE (Fig. 4A, WT pBAD versus rpoE pBAD versus degSΔPDZ pBAD). These data confirm the involvement of OMPs, DegS and σE itself in the canonical pathway of σE activation during heat stress.

Fig. 4.

Fig. 4

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Role of OMPs and DegS in σE induction. Expression of the σE-dependent rpoE, rpoH, fkpA and surA genes was measured by qRT-PCR. Fold induction represents relative gene expression during stress in comparison to control conditions after normalization to reference gene (gmk) mRNA levels. Cells were obtained at OD600 ~0.4, re-suspended in stress media with 0.2% arabinose for induction of RybB or OmpC expression, and incubated for 1 h. WT and mutants Salmonella were subjected to heat stress (50°C) (A), acid stress (pH 4.5) (B, D) or control conditions (37°C, pH 7.0) (C).

In contrast, rybB overexpression had only a modest 1.3-to 2-fold effect on the expression of σE-regulated genes in response to acid stress (Fig. 4B, WT pBAD versus WT pBAD-RybB). Deletion of the DegS PDZ domain similarly had little or no effect on σE-regulated gene expression (Fig. 4B, WT pBAD versus degSΔPDZ pBAD).A degS null mutation produced effects on both heat and acid stress induction identical to those observed in the degSΔPDZ mutant strain (Fig. S1), with the exception that the degS-ΔPDZ mutant exhibited a global increase in the expression of σE-regulated genes that may reflect loss of the inhibitory effect of the PDZ domain on DegS protease activity (Walsh et al., 2003; Sohn et al., 2007). qRT-PCR was used to confirm a reduction of ompC mRNA following rybB overexpression (data not shown). These observations indicate that acid stress induction of the σE regulon, unlike heat stress induction, does not require OMPs or the DegS protease.

Overproduction of OMPs leads to increased transcription at σE-dependent promoters (Mecsas et al., 1993) by activating the DegS protease and triggering the degradation of RseA. We reproduced this condition by placing the ompC gene under control of the ParaBAD promoter (Bang et al., 2005). OmpC overexpression at neutral pH induced the expression of rpoE, rpoH, surA and fkpA by 84-, 25-, 19- and 10-fold respectively (Fig. 4C, WT pBAD versus WT pBAD-ompC). OmpC-induction of the σE regulon was completely dependent on σE (Fig. 4C, rpoE pBAD versus rpoE pBAD-ompC) and the DegS PDZ domain (Fig. 4C). However, OmpC overproduction during acid stress only modestly augmented the response of sE (Fig. 4D, WT pBAD versus WT pBAD-ompC), and did not enhance σE target gene expression in rpoE and degS mutants (Fig. 4D), suggesting that σE activation by unfolded OMPs and acid stress occurs by distinct mechanisms.

It should be noted that some acid induction of σE target genes was detected in the rpoE mutant strain (Fig. 4B and D, rpoE pBAD and rpoE pBAD-RybB/ompC), suggesting that some rpoE-dependent genes can be partially induced by acid stress in a σE-independent fashion.

In summary, the induction of σE during acid stress cannot be attributed to DegS-sensing of unfolded OMPs as occurs during heat stress or OmpC overproduction, because acid induction is largely indifferent to changes in OMP expression or the presence of the protease DegS.

Acid induction of σE requires RseP-dependent proteolysis of the anti-σE factor RseA

Degradation of RseA is the key control point in σE activation. In the canonical pathway, initial cleavage by DegS is required before subsequent proteolysis by RseP can occur (Grigorova et al., 2004). However, our discovery that DegS is dispensable for σE activation during acid stress suggested that RseP might be able to cleave RseA in the absence of DegS under acidic conditions. To determine the contribution of regulatory proteins of the sE pathway, we measured σE target gene expression during acid stress by qRT-PCR. These experiments demonstrated that acid induction is abolished in an rseA or rseP mutant (Fig. 5). Elimination of RseA results in derepression and constitutive activation of σE, whereas elimination of RseP abrogates acid induction without affecting baseline activation (Fig. S2). These results eliminate the possibility that additional regulators can activate σE in response to acid stress and show that acid induction requires RseA and RseP. Possible involvement of RseB or RseC was also investigated. Interaction of RseB with the C-terminal domain of RseA modulates the affinity of the anti-sigma for σE (Collinet et al., 2000). RseB and DegS independently repress RseP protease activity, and the elimination of RseB and DegS in E. coli promotes the cleavage of RseA by RseP and increases σE activity (Grigorova et al., 2004), suggesting that acid stress might activate σE by disrupting RseB–RseA interactions. The elimination of RseB had a modest effect on baseline σE activity (Fig. S2). However, we observed that acid induction of σE-dependent gene expression is preserved in Salmonella mutant strains lacking RseB or RseC, as well as in an rseB degS double mutant (Fig. 5A).

Fig. 5.

Fig. 5

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Effects of rseA, rseB, rseC and rseP mutations on acid induction of σE. Expression of the σE-dependent rpoE, rpoH, fkpA and surA genes was measured by qRT-PCR in WT, rseA, rseB, rseB degS rseC (A) or rseP and rsePΔPDZ (B) mutant Salmonella subjected to 1 h acid stress (pH 4.5) during early stationary phase and compared with expression at pH 7.0. Steady-state mRNA levels were normalized to levels of reference mRNA (gmk) under the same conditions.

To further investigate the role of RseP in the acid induction of σE, RseA proteolysis was monitored by immunoblot following heat or acid induction in WT and degS or rseP mutant backgrounds. An RseA derivative containing a N-terminal haemagglutinin epitope (HA-RseA) was constructed as described (Kanehara et al., 2003) and expressed from a low-copy plasmid. A C-terminal truncated form of HA-RseA [HA-RseA(ΔP)] resulting from site-1 cleavage by DegS has been shown to accumulate in an E. coli rseP mutant strain (Kanehara et al., 2002), with lower levels of HA-RseA(ΔP) detectable in an rseP-ΔPDZ mutant (Kanehara et al., 2003). Under non-stress conditions in Salmonella, intact RseA accumulated in WT, degS and degSΔPDZ strains (Fig. 6A, left panel). Both HA-RseA and HA-RseA(ΔP) species were detected in an rseP mutant, reflecting partial degradation of RseA by DegS from basal activity of this protease. In an rsePΔPDZ strain, nearly all RseA was degraded due to disinhibition of RseP protease activity (Alba et al., 2002; Kanehara et al., 2003).

Fig. 6.

Fig. 6

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Effects of heat and acid stress on RseA degradation. The pACYC177-HA-RseA plasmid was introduced into WT, degS, degSΔPDZ (A), rseP, rsePDPDZ (A), or degS rseP (B) strains. Cells at OD600 ~1.5 were subjected to heat (centre) or acid stress (right) and compared with baseline (left). HA-tagged RseA accumulation was measured by anti-HA immunobloting.

During heat stress, RseA was rapidly degraded in WT cells corresponding with activation of the σE regulon (Fig. 6A, centre panel). This degradation requires DegS, and RseA therefore persisted in heat-stressed degS and degSΔPDZ mutant cells. Loss of the inhibitory PDZ domain enhances RseA processing by DegS (Walsh et al., 2003; Cezairliyan and Sauer, 2007; Hasselblatt et al., 2007) and might have been predicted to reduce RseA levels. However, a degSΔPDZ allele only increased basal σE activity by 1.5- to 2-fold (Walsh et al., 2003), suggesting the presence of other constraints on DegSΔ-PDZ in intact cells. In an rseP mutant, RseA was degraded by OMP-activated DegS but could not be further processed so that the HA-RseA(ΔP) product accumulated, whereas complete degradation of RseA was observed in an rsePΔPDZ mutant strain. These results confirm the σE activation model in which DegS is activated by unfolded OMPs accumulating during heat shock and cleaves RseA to allow a second cleavage by RseP.

During acid stress, HA-RseA levels were reduced in WT, degS and degSΔPDZ strains (Fig. 6A, right panel), indicating that RseA degradation under these conditions is primarily DegS-independent. Basal levels of DegS-mediated cleavage were revealed by the presence of HA-RseA(ΔP) in an rseP mutant strain, as observed under non-stress conditions. Again, an rsePΔPDZ mutant showed unregulated protease activity with no detectable HA-RseA. These results corroborate the qRT-PCR data, demonstrating that acid stress induction of σE differs from heat stress induction in its lack of a requirement for the DegS protease.

As RseP is important for σE acid induction, and the PDZ domain regulates its protease activity, we hypothesized that this domain might play a role in the acid pH sensing. Measurement of σE-dependent gene expression by qRT-PCR revealed that the RseP PDZ domain is indispensable for the σE acid induction (Fig. 5B). The absence of acid induction is attributable to constitutive proteolysis of RseA by RsePΔPDZ protein; σE-dependent gene transcription is elevated in the rsePΔPDZ mutant at both neutral and acid pH (Fig. S2). The putative inhibitory interaction between the RseA periplasmic domain and the RseP PDZ domain may be destabilized at acid pH, to allow the activation of RseP with subsequent RseA cleavage and release of σE.

A Salmonella mutant strain with inactivation of both degS and rseP was constructed to exclude the presence of a novel site-1 protease that might be activated during acid pH stress. Western blot analysis of HA-RseA was repeated under conditions identical to those used in Fig. 6A, and demonstrated an absence of RseA cleavage in the double mutant (Fig. 6B). Thus, DegS-independent RseP-dependent proteolysis of RseA is induced by acid stress and does not appear to involve a novel site-1 protease.

Involvement of DegS and RseP in the survival within the acidified vacuoles of RAW264.7 macrophages

To validate our results concerning the role of the proteases DegS and RseP in σE regulon acid-induction and corroborate these in vitro observations with the acidification-dependent Salmonella survival in SCVs, we used the Salmonella degS and rseP mutants to infect macrophages treated or untreated with NH4Cl. As shown in Fig. 7, in the absence of NH4Cl and 6 h after infection, survival of degS and rseP mutants was decreased 10-and 100-fold compared with WT cells respectively. Thus, both proteases appear important for Salmonella survival in SCV. However, the degS mutant was more sensitive to the presence of NH4Cl, whereas the rseP strain displayed the same behaviour as an rpoE mutant. Thus, these results confirm that σE, RseP and DegS are important for Salmonella pathogenesis, but only σE and RseP contribute to the acid stress response.

Fig. 7.

Fig. 7

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Survival of WT and rpoE, degS or rseP mutant Salmonella in the SCV. RAW264.7 macrophages treated (dotted lines) or untreated (continuous lines) with NH4Cl were infected with complement-opsonized WT (diamonds), rpoE (squares), degS (crosses) or rseP (triangles) mutant Salmonella.

Discussion

Acid pH is an important environmental stress encountered by Salmonella in its interaction with the host. The SCV of infected macrophages acidify to pH 4–5 within an hour of bacterial uptake (Rathman et al., 1996).Although previous studies have yielded conflicting results regarding the effect of acidification on intracellular Salmonella replication (Rathman et al., 1996; Steele-Mortimer et al., 2000), our observations (Fig. 1) support the report of Rathman et al. (1996) that vacuolar acidification promotes survival and replication in RAW264.7 macrophages. This is consistent with the stimulation of Salmonella virulence gene expression (Alpuche Aranda et al., 1992; Rathman et al., 1996) and effector protein translocation (Beuzon et al., 1999) by acid pH. Study of the survival of degS or rseP mutants in SCV revealed that both proteases are important for the basal survival of Salmonella in macrophages. However, neutralization of macrophage vacuolar pH with NH4Cl revealed that survival of Salmonella within the acidified phagosome requires RseP but not DegS (Fig. 7); this requirement for RseP suggests that σE is involved in the acid pH-triggered adaptation of intracellular Salmonella.

The σE sigma factor is required for Salmonella virulence in mice (Humphreys et al., 1999; Testerman et al., 2002) and for resistance to two stress conditions imposed by host innate immunity, oxidative stress (Testerman et al., 2002) and anti-microbial peptides (Onufryk et al., 2005). We now further show that σE promotes Salmonella replication within the acidified SCV of macrophages (Figs 1 and 7), facilitates growth at pH 5.0 in vitro, and is essential for the ATR (Foster, 1999) (Fig. 2). These observations demonstrate for the first time that the σE regulon is involved in the maintenance of pH homeostasis.

In conjunction with the Cpx system, σE plays a critical role in activating the expression of genes encoding membrane proteins, enzymes involved in phospholipid, lipopolysaccharide and outer membrane biosynthesis, and primary metabolic and signal transduction pathways (Dartigalongue et al., 2001) that help to maintain cellular integrity under stress conditions. Studies in E. coli have demonstrated that the σE regulon can be activated by the presence of unfolded OMPs in the periplasm, which are recognized by the DegS PDZ domain and lead to sequential regulated proteolysis of the RseA anti-σ (Alba et al., 2002; Kanehara et al., 2002; Walsh et al., 2003; Chaba et al., 2007). The present study demonstrates that σE can also be activated by acid stress (Fig. 3). However, the well-characterized mechanism of DegS-dependent σE activation in response to heat shock and unfolded OMPs does not appear to be responsible for activation at acid pH. In contrast to the heat stress induction of rpoE and the σE-dependent genes rpoH, fkpA and surA, acid stress induction of the σE regulon is not abolished by rybB over-expression (i.e. is not OMP-dependent) and is not dependent on the PDZ domain of DegS (Fig. 4). The cleavage of RseA by DegS is normally rate-limiting for σE activation (Chaba et al., 2007), so that RseA degradation and the release of free σE during heat stress are determined by the OMP signal. Activation by acid stress is independent of DegS (or RseB and RseC) but remains dependent on RseP and RseA (Fig. 5), and RseP but not DegS is necessary for RseA degradation under this condition (Fig. 6). Moreover, RseP appears sufficient to trigger the σE-mediated response because qRT-PCR assays demonstrate acid pH-induction of σE target genes even in a degS mutant (Figs 4 and 5, Fig. S1). Thus, σE activation by acid stress does not even require basal levels of RseA processing by DegS (Fig. 6) that may result from low steady-state levels of unfolded OMPs in the periplasm (Sohn et al., 2007). No RseA proteolysis was observed in a degS rseP double mutant, thus excluding the possibility that an acid-inducible protease is responsible for site-1 processing in the absence of DegS.

The precise molecular mechanism by which acid stress is sensed in the σE activation cascade awaits further investigation. An rsePΔPDZ mutation allows RseP-mediated proteolysis of RseA to take place in a DegS-independent fashion (Ades et al., 2003; Kanehara et al., 2003; Bohn et al., 2004). We have observed that an rseP-ΔPDZ mutation abolishes acid induction of σE-dependent gene expression (Fig. 5B), which suggests that the putative interaction between RseA and the RseP PDZ domain might be sensitive to changes in pH. Reversible protonation of His (pKa ≈ 6) serves as a pH sensor in some regulatory proteins. However, we replaced the single His residue (His188) in the RseA periplasmic domain with Ala and observed no effect on acid induction of σE-dependent target gene expression (data not shown). Alternatively, interaction between the two Gln-rich regions of RseA with the RseP PDZ domain might be sensitive to changes in pH and responsible of the RseP protease activation. Indeed, removal of the Gln-rich regions by DegS is responsible for relieving inhibition of RseP by its PDZ domain in response to unfolded OMPs (Kanehara et al., 2003; Akiyama et al., 2004). Further studies will be required to establish the mechanism of RseP activation in response to acid stress.

In conclusion, the expression of σE-dependent genes can be triggered by heat stress sensed by the PDZ domain of the DegS protease, or by acid stress, possibly sensed by the PDZ domain of the RseP protease (Fig. 8). Both heat stress and acid stress are important stress conditions encountered by bacteria in the environment or during interactions with mammalian hosts. The complex regulation of σE activation by multiple proteases permits the integration of diverse environmental signals to trigger a common stress response, an emergent theme in gene regulation by alternative sigma factors (Fang, 2005).

Fig. 8.

Fig. 8

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Pathways of σE activation by heat and acid stress. σE can be activated by the accumulation of unfolded OMPs during heat stress, which leads to activation of DegS and the sequential proteolysis of the RseA anti-σ by DegS and RseP. Alternatively, acid stress can trigger RseA proteolysis by RseP in an OMP- and DegS-independent fashion.

Experimental procedures

Construction of Salmonella mutants

In this study, Salmonella 14028s (ATCC) was used as WT. Mutations were constructed by the λ-Red recombinase method (Datsenko and Wanner, 2000). DNA primers used for rseB, rseC, degS or rseP mutant constructions are listed in Table S1, and strains are listed in Table 1. After construction, all mutations were transduced into WT using bacteriophage P22HT105/int and non-lysogenic colonies selected by sensi- tivity to lytic P22 H5 (Maloy, 1990). The degSΔPDZ and rsePΔPDZ mutants were constructed as described (Karlinsey, 2007): PCR amplification of the tetRA element was performed using primer pairs CM1/CM2 for degS and CM7/CM8 for rseP (Table S1), and allelic replacement of tetRA was achieved with the double-stranded oligomers CM3 and CM43 respectively. For construction of N-terminal HA-tagged RseA, an HA duplex was created with primers HA1/HA2 and ligated into NcoI–SmaI-digested pBAD24 (Guzman et al., 1995) to produce plasmid pBAD24-HA. The rseA gene was amplified with primers HA3/HA4 and cloned into pBAD24-HA. For constitutive expression, an HA-rseA DNA fragment was amplified from pBAD24-HA-rseA with the primers HA5/HA6 and cloned into pACYC177 (Chang and Cohen, 1978).

Table 1.

Strains and plasmids used in this study.

Strains/plasmids Relevant characteristics Source
Strains
    14028s Wild-type Salmonella enterica sv. Typhimurium ATCC
    IB2 14028s ΔrpoE∷cat Bang et al. (2005)
    IB355 14028s ΔrseC∷cat This study
    IB408 14028s ΔdegS∷cat This study
    IB412 14028s ΔrseP∷cat This study
    IB480 14028s ΔrseB∷cat This study
    JK628 14028s ΔdegS∷aph This study
    JK629 14028s ΔdegS∷aph rseP∷cat This study
    CM10 14028s with deletion of the degS PDZ domain coding sequence This study
    CM35 14028s with deletion of the rseP PDZ domain coding sequence This study
Plasmid
    pACYC177 p15A ori bla aph Chang and Cohen (1978)
    pBAD24 ColE1 ori ParaBAD bla Guzman et al. (1995)
    pBAD30 p15A ori ParaBAD bla Guzman et al. (1995)
    pFB175 pBAD30∷ompC, ompC under the control of ParaBAD Bang et al. (2005)
    pFB188 pACYC177 carrying the HA-RseA DNA sequence This study
    pKD46 oriR101 repA101ts ParaBAD-λRed bla Datsenko and Wanner (2000)
    pKP17-1 pBAD-RybB, rybB under the control of Para Papenfort et al. (2006)
    pKP8-35 pBAD control plasmid, produces short nonsense RNA Papenfort et al. (2006)
    pRB983 pBR3-rpoE, rpoE complementation Testerman et al. (2002)
    pTFP1 pRS1274∷P1rpoE–lacZ Testerman et al. (2002)
    pTFP2 pRS1274∷P3rpoE–lacZ Testerman et al. (2002)
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Culture conditions and physiological assays

Bacterial cells were grown in Luria–Bertani (LB) or E minimal medium (Vogel and Bonner, 1956) containing 0.4% glucose at 30°C, 37°C or 42°C with agitation, or on solid LB medium containing 15 mg ml−1 agar. As necessary, media were supplemented with penicillin (250 µg ml−1) or chloramphenicol (40 µg ml−1). Acidic LB medium was supplemented with 100 mM MES [2-(N-morpholino)ethanesulphonic acid] buffer for pH < 5. The pH of E medium was adjusted with HCl. Bacterial growth rates were measured by determining the optical density at 600 nm (OD600) at 37°C with agitation on a BioScreen C Microbiology Microplate reader (Growth Curves USA). For acid pH survival assays, cells were grown in E medium 0.4% glucose at 37°C to OD600 ~1.5, and incubated at 37°C in fresh medium adjusted to pH 7.0 or pH 3.0. To measure the ATR, bacteria were pre-adapted in mildly acidic medium (pH 5.0) for 2 h before challenge at pH 3.0. For each time point colonies were enumerated on LB agar plates. Per cent survival represents the number of viable cells after exposure to acid stress as a proportion of the number of cells prior to challenge. To measure β-galactosidase activity of P1rpoE–lacZ and P3rpoE–lacZ(Testerman et al., 2002), bacterial cells collected from overnight cultures were divided and re-suspended in neutral (pH 7.0) or acidic (pH 4.5) E medium. At timed intervals, samples were collected for β-galactosidase assays (Maloy, 1990).

Macrophage killing assay

RAW264.7 macrophages were infected with mouse serum-opsonized Salmonella at a multiplicity of infection of 10:1 and allowed to internalize bacteria for 15 min. After phagocytosis, cells were washed with PBS and incubated in RPMI1640 medium containing 6 mg ml−1 gentamicin. To neutralize vacuolar pH, 20 mM ammonium chloride (NH4Cl) was added. At timed intervals, infected macrophages were lysed with 1% Triton X-100 and surviving bacteria enumerated on LB agar plates. Results were expressed as per cent survival.

Real-time reverse transcription PCR (qRT-PCR)

Total RNA was isolated using an RNeasy Mini kit (Qiagen) according to manufacturer’s instructions. For acid stress, cells were grown in E medium until OD600 ~0.4 and incubated at 37°C for 1 h in E medium at pH 7.0 or pH 4.5. Heat stress was administered in E medium at 50°C. Each RNA extraction was conducted on three independent biological samples and qRT-PCR assays performed on cDNA synthesized from those RNA. The housekeeping gene gmk (guanylate mono-phosphate kinase) was used as reference after checking that gmk expression does not change under conditions of heat or acid stress. Expression of the σE-dependent rpoE, rpoH, fkpA and surA genes was quantitatively assessed by qRT-PCR in a Rotor-Gene 3000 (Corbett Research), with gmk from the same sample as reference. Forward (F) and reverse (R) primers are listed in Table S1. For the rpoE insertion mutant, primers used to quantify the rpoE transcript were designed upstream the mutation (Bang et al., 2005). For each reaction, 8 µl of RNA sample (50 ng per reaction) were mixed with 1.8 µl of primer sets (0.9 µM final) and 10 µl of SYBR Green mix (Qiagen) containing 0.2 µl of reverse transcriptase. Each sample was assayed in triplicate for each run. Control RNA from WT cells was used to construct a standard curve. Reaction conditions were: 20 min at 50°C, 15 min at 95°C and 40 cycles at 94°C for 15 s, 52°C for 25 s, 72°C for 20 s.

Immunoblotting

Culture samples (1 ml) were harvested and re-suspended in 200 ml sample buffer (62.5 mM Tris-HCl pH 6.8, 2% SDS, 25% glycerol, 0.01% Bromophenol Blue, 350 mM DTT). After electrophoresis and protein transfer onto a PVDF membrane (Millipore), blots were blocked in 5% non-fat dry milk (NFT) PBST (Phosphate-buffered saline, 0.05% Tween 20). Primary and secondary incubations used anti-HA mouse antibody (Sigma) and goat peroxidase-conjugated anti-mouse HRP (Bio-Rad), respectively, in 5% NFT/PBST. Blots were developed with the ECL Plus Western Blotting Detection System (Amersham).

Supplementary Material

Supp FigureS1-S2 & TableS1

Acknowledgement

This work was supported by a grant from the National Institutes of Health (AI44486) and a grant from the Korea Research Foundation (KRF-2008-314-C00328).

Footnotes

Supporting information

Additional supporting information may be found in the online version of this article.

Please note: Wiley-Blackwell are not responsible for the content or functionality of any supporting materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.

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Supplementary Materials

Supp FigureS1-S2 & TableS1
NIHMS621020-supplement-Supp_FigureS1-S2___TableS1.pdf (240.6KB, pdf)

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