The I-switch Tracks Intracellular pH in Salmonella

To determine the impact of acid stress on Salmonella intracellular pH (pH_i), we used the I-switch (I_A488/A647) labeled at its 5′ and 3′ termini with Alexa-488 and Alexa-647, respectively, as a cytoplasmic pH sensor. The I–switch consists of cytosine-rich unpaired regions that form an anti-tetraplex CH⁺.C by alternate Watson and Crick base pairing in the presence of protons. This leads to a “closed” conformation of DNA, enabling FRET to occur between the fluorophore pairs. This process is reversible, and at neutral pH, the I-switch dissociates into an open, extended conformation because of electrostatic repulsion between the duplex arms [14,15]. We first compared the fluorescence intensity of the donor alone (520 nm, "D") and the acceptor alone (666 nm, "A") with the fluorescence of the I-switch containing both donor and acceptor. This provided a determination of the cross-talk between the fluorophores, which was negligible (S1A Fig). I-switch labeled with Alexa 488 and Alexa 647 was suspended in buffers ranging from pH 5.0 to 7.2. The samples were excited at 480 nm and spectra were recorded in a Tecan spectrophotometer (GENios) from 510 to 700 nm. A D/A curve as a function of pH was plotted from the ratio of donor (520 nm) to acceptor (666 nm) intensities to generate the in vitro standard curve (Fig 1A). The I-switch exhibited a sigmoidal increase over this range due to the formation of the I-motif in the closed state.

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Fig 1. Functional integrity of the I-switch in Salmonella.

Wild-type (WT) Salmonella was electroporated with 6 μM I-switch and recovered in SOC for 1 h at 37°C, followed by three washes with PBS. Bacteria were then clamped at various pH values in K⁺-rich buffer containing 40 μM nigericin for 1 h at room temperature (RT) and imaged on an Applied Precision Delta Vision inverted wide-field fluorescence microscope. The fluorescence intensity of both donor and acceptor channels of 50 cells were measured for every pH value indicated in the figure. The ratio of D/A intensities were obtained and the normalized D/A ratios were plotted as a function of pH, which yielded a sigmoidal intracellular calibration curve (A). The intracellular curve is overlaid on the in vitro calibration curve. Error bars represent mean ± standard deviation (SD). (B) Representative Donor (D), FRET (A) and D/A ratio images of WT Salmonella clamped at various pH values. Scale bar, 3 μm, color scale, bottom, right. Pseudocolor images were generated by calculating the D/A ratio per pixel. Using ImageJ software, the pixels were then color-coded using 16 colors with blue (D/A = 0.1) to red (D/A = 1) to indicate the transition from acidic to neutral pH.

https://doi.org/10.1371/journal.pbio.1002116.g001

We next investigated the ability of the I-switch to function inside bacterial cells by incorporating I_A488/A647 into Salmonella by electroporation and verifying its incorporation by fluorescence microscopy (Fig 1B). We confirmed its intracellular location by preparing spheroplasts, which eliminated the outer membrane but still retained the I-switch in the cytoplasm (S1B Fig). We used 6 μM I_A488/A647 for electroporation, since it showed optimum fluorescence at both donor and acceptor channels inside bacterial cells; this concentration was used in all subsequent experiments. The internal pH of Salmonella was clamped to the external pH of K⁺-containing media using 40 μM nigericin [21–23]. An intracellular D/A standard curve (Fig 1A) showed perfect correlation with the in vitro curve, indicating the functionality and integrity of the I-switch in Salmonella. The D/A profiles of cells clamped at acidic pH 5.1 were visibly distinct and showed a 5-fold increase from those clamped at neutral pH 7.1. This study is the first application of the I-switch in bacteria.

Extracellular Acid Stress Acidifies the Salmonella Cytoplasm and Requires EnvZ/OmpR Activation

We next measured the change in cytoplasmic pH (pH_i) in response to varying the pH of the extracellular media (pH_e). Salmonella containing the I-switch were subjected to acid stress and the fluorescence was measured in SPI-2–inducing (pH_e 5.6) and non-inducing (pH_e 7.2) conditions over time (Fig 2A). The pH_i of Salmonella decreased from 6.85 to 6.1 upon incubation at pH_e 5.6, but remained fairly constant when incubated at pH_e 7.2 (Fig 2B), indicating that the Salmonella cytoplasm acidifies in response to extracellular acidic pH. Addition of extracellular media at even lower pH_e, e.g., 4.5, reduced the intracellular pH further, to pH 5.8 (Fig 2C). Acidification was readily reversible upon addition of extracellular media at alkaline pH (Fig 2C), which restored the near neutrality of the cytoplasm. This result further demonstrates the functionality of the I-switch; i.e., it is not degraded, because it can reversibly switch from an open state (low FRET at neutral pH) to a closed state (high FRET at acidic pH) in response to changes in pH_e.

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Fig 2. The Salmonella cytoplasm is acidified upon acid stress and requires OmpR.

(A) Salmonella cultures of I_A488/I_A647-incorporated WT, ompR null mutant, and ompR null complemented with pWSK29-ompR (encoding ompR) were incubated at either acidic pH_e (5.6) or neutral pH_e (7.2) at indicated time points. Representative epifluorescence of the D/A ratio images are shown for WT Salmonella, the ΔompR mutant, and the ΔompR mutant with ompR provided in trans. Scale bar, 3 μm. (B) A plot of the intracellular pH of Salmonella WT at pH_e 5.6 and pH_e 7.2, an ompR null mutant at pH_e 5.6 and pH_e 7.2, and the ompR null mutant complemented with ompR supplied in trans at pH_e 5.6 and pH_e 7.2 over time. The D/A ratios of 50 cells were analyzed at each time point and the pH values were determined from the intracellular standard curve. Error bars represent mean ± standard error of the mean (SEM) (n = 3). (C) To demonstrate the reversibility of the I-switch and mimic vacuolar pH conditions in vitro, WT Salmonella cultures were incubated at pH_e (4.5) for 1 h, and samples were collected at 30 and 60 min for imaging. The cultures were then washed and incubated for an additional 1 h at pH_e 7.2. The D/A ratios of 50 cells were analyzed at the indicated time points. Representative epifluorescence of the D/A ratio images are shown. Scale bar, 3 μm. The average pH_i from 50 cells is listed under the image.

https://doi.org/10.1371/journal.pbio.1002116.g002

Because OmpR was implicated in the Acid Tolerance Response in Salmonella ([24]; see also S2 Fig), we measured the response of an ompR null strain of Salmonella under similar acid stress conditions. In the ompR null strain, the intracellular pH did not decrease in response to extracellular pH stress (Fig 2A and 2B). OmpR-dependent intracellular acidification was completely restored upon supplying ompR in trans on a plasmid (Fig 2A and 2B). This result provides the first evidence for a role of OmpR as a regulator of acid stress in Salmonella by regulating genes that alter intracellular pH. In addition, the envZ null strain did not respond to acid stress, but acidification was restored upon supplying the cytoplasmic domain of envZ (envZc) in trans (Fig 3A and 3B). The requirement of the cytoplasmic domain of the EnvZ kinase only (EnvZc) further indicates that the I-switch is in the cytoplasm and the cytoplasmic pH becomes acidic under these conditions. Acidification was not dependent on growth phase, but was similar at both exponential and stationary phase (S3 Fig).

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Fig 3. The cytoplasmic domain of EnvZ is required for intracellular acidification upon acid stress.

(A) Salmonella cultures of I_A488/I_A647-incorporated WT, envZ null mutant, and envZ null complemented with pMPM-envZc (encoding cytoplasmic domain of envZ) were incubated at acidic pH_e (5.6) at indicated time points. Representative epifluorescence of the D/A ratio images are shown. Scale bar, 3 μm. (B) A plot of the intracellular pH at pH_e 5.6 of Salmonella WT, an envZ null mutant and the envZ null mutant complemented with envZc supplied in trans over time. The D/A ratios of 50 cells were analyzed at each time point and the pH values were determined from the intracellular standard curve. Symbols represent the mean ± SEM (n = 3).

https://doi.org/10.1371/journal.pbio.1002116.g003

2',7'-Bis-(2-Carboxyethyl)-5-(and-6)-Carboxyfluorescein, Acetoxymethyl Ester (BCECF-AM) Reports Similar Acidification As the I-switch in Response to Extracellular Acid Stress

To further ensure that the I-switch was a high-fidelity reporter of intracellular pH, we compared it to the fluorescent probe BCECF-AM, which has been widely used to measure intracellular pH in mammalian cells [25,26], plant cells [27], yeast [28,29], and bacteria [23,30,31]. It allows ratiometric detection (S4A and S4B Fig) and is permeable to bacterial membranes. Once it gains access to the intracellular compartment, it is cleaved by intracellular esterases to create the active fluorescent form. Although the range of BCECF-AM (pH 6–7) is more limited than the I-switch (pH 5.1–7.1), a similar acidification of the Salmonella cytoplasm was observed in response to extracellular acid stress (S4C Fig), corroborating the use of the I-switch in reporting the pH of the Salmonella cytoplasm. Our observation that BCECF (which is intracellular) and the I-switch reported similar intracellular pH values corroborates the intracellular location of the I-switch, and the I-switch was also clearly visible in spheroplasts (S1B Fig).

Cytoplasmic Acidification Occurs Immediately After Entry into the Macrophage SCV

Because Salmonella acidified its cytoplasm in response to external acid stress (Figs 2 and 3), we reasoned that uptake of Salmonella into the acidified SCV should also lead to a decrease in intracellular pH. This internal acidification would in turn provide a signal to EnvZ/OmpR to induce the SPI-2 type III secretion system by activating SsrA/B [16,18], which is critical for the survival of Salmonella within the phagocyte. To measure the intracellular pH of Salmonella during infection in RAW264.7 macrophages, we used confocal microscopy to directly visualize I-switch containing Salmonella inside macrophages. At each time point, the D/A ratio of individual cells was determined as a measure of intracellular pH (Fig 4). Salmonella exhibited a remarkable, rapid intracellular pH drop (6.85 to 5.75) immediately post-infection and reached a plateau value of 5.65 after 3 h. Interestingly, the Salmonella ompR null mutant did not experience a decrease in intracellular pH upon entry into the SCV (Fig 4). Again, the dependence of ompR on cytoplasmic acidification could be fully complemented by supplying ompR in trans (Fig 4). This result identifies OmpR as a key regulator of Salmonella intracellular pH, not only for in vitro acid stress but also during macrophage infection. The I-switch containing bacteria in the vacuole were viable because they were capable of SPI-2 secretion (see below).

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Fig 4. The Salmonella cytoplasm is rapidly acidified upon macrophage infection and requires OmpR.

RAW264.7 macrophages infected with WT Salmonella and the ΔompR null mutant containing I_A488/I_A647 were imaged over time on a Nikon A1R confocal microscope. The D/A ratio in the ΔompR strain showed minimal FRET, indicating ompR was required for cytoplasmic acidification. RAW264.7 macrophages were pre-treated with 25 nM of the V-ATPase inhibitor bafilomycin A1 (BAF) for 30 min prior to infection, and macrophages were infected with I_A488/I_A647-electroporated Salmonella in the presence/absence of BAF for 3 h and imaged as before. BAF neutralized the SCV and prevented Salmonella acidification. (A) Representative images of D/A ratios are shown at 30 and 180 min post-infection and compared to WT Salmonella prior to infection. Infection was carried out as described in Materials and Methods. Scale bar, 3 μm. (B) Prior to infection, the D/A ratios of Salmonella were determined and the intracellular pH was approximately 6.8. The intracellular pH of Salmonella remained fairly constant in macrophages upon BAF treatment (approximately 6.8) or in the ΔompR null strain in macrophages, unlike untreated WT Salmonella. It showed a remarkable intracellular pH drop to 5.65, similar to the ompR null mutant complemented with ompR supplied in trans. Dataset is represented as the mean ± SEM (n = 3).

https://doi.org/10.1371/journal.pbio.1002116.g004

An alternative method employed by many Salmonella researchers for examining intracellular bacteria involves Triton solubilization at various time points post-infection to recover bacteria from macrophages. Although this method is less rigorous than direct visualization by confocal microscopy, the results obtained using this approach were in agreement with the microscopy (S5 Fig). The pHi of cells recovered after solubilization was determined from the reference intracellular standard curve. The pHi of Salmonella rapidly dropped by nearly 1 unit (from 6.81 to 5.87) immediately within 30 min post-infection and reached a final value of 5.6 at 3 h post-infection. Bacteria recovered from the vacuole were capable of growth on agar plates, further indicating that they were still viable. Taken together, both approaches firmly establish that uptake of Salmonella into the SCV leads to an OmpR-driven immediate drop in intracellular pH. This result has significant ramifications in interpreting the pH control of the gating of the SPI-2 type III secretion apparatus [20] (see Discussion).

Inhibition of Vacuolar Acidification by Bafilomycin Prevents Salmonella Acidification

Phagosomal pH is maintained between 4 to 5 by the vacuolar H⁺-ATPase (V-ATPase) and remains acidified for at least 5–6 h post-infection [9,32]. Treatment with the specific inhibitor Bafilomycin A1 (BAF) prevents SCV acidification [1]. To examine the contribution of vacuolar SCV acidification to the low intracellular pH of Salmonella during macrophage infection, we used BAF, which is known to raise the pH of endosomes to neutral in both HeLa [33] and RAW264.7 cells [19]. In the presence of BAF, the Salmonella cytoplasm remained approximately pH 6.7 throughout the infection and failed to acidify (Fig 4). The pH_i in the presence of BAF was similar to the ompR null mutant. Our results with BAF clearly indicate that the acidic pH of the SCV leads to a decrease in intracellular pH of Salmonella in macrophages (Fig 4). Pre-treatment with and maintenance of 25 nM BAF throughout infection completely abrogated the intracellular pH decrease in Salmonella-infected macrophages. This effect was rapidly reversible [34], as removal of BAF after 1 h restored the ability of Salmonella to reduce intracellular pH, suggesting a complete correlation between vacuolar acidification and an intracellular pH drop (S6 Fig). Together, these data provide strong evidence that the cytoplasm of Salmonella rapidly follows the pH of the macrophage vacuole and requires OmpR (Fig 4).

Well-Characterized OmpR-Regulated Genes Were Not Responsible for Intracellular Acidification

Because OmpR regulates the SsrA/B two-component system that, in turn, activates the type III secretion system encoded by SPI-2 [16–18,35,36], we wanted to determine whether or not SPI-2 was the target that was essential for cytoplasmic acidification. We performed a similar analysis on a mutant Salmonella strain that does not make a type III secretory apparatus. An ssaC null strain lacks the gene encoding the outer membrane ring protein SsaC of the SPI-2 type III secretion system, and thus is not capable of type III secretion. The ssaC null strain was fully capable of cytoplasmic acidification, as shown in (S7A and S7B Fig). This was not surprising, because we observed a similar OmpR-dependent response to external acid in E. coli, which lacks SPI-2 (unpublished results). OmpR is best characterized for its regulation of the outer membrane proteins OmpF and OmpC [12]. An ompC mutant and the double ompC/ompF mutant lacking the genes encoding the outer membrane proteins OmpC and OmpF were similarly capable of acidification (S7A and S7B Fig). A previous study identified MgtC as an inhibitor of the FoF1 ATPase, leading to acidification [37], but an mgtC null strain was as acidified as the WT (S7A and 7B Fig). Thus, OmpR regulates an unidentified target(s) that is responsible for cytoplasmic acidification.

OmpR Directly Represses cadC/BA to Acidify the Salmonella Cytoplasm

In order to identify genes that were regulated by OmpR, we performed a microarray analysis and compared the transcriptome of the ompR null and the WT strain under both acid and osmotic stress. We identified genes belonging to the cadC/BA operon to be highly up-regulated in the ompR mutant. We validated the microarray results by investigating the expression of the cadC/BA operon under acid stress by quantitative real-time PCR. Transcripts for genes cadC (2.3-fold), cadB (3.2-fold) and cadA (4.9-fold) were all up-regulated in the ompR mutant, suggesting that OmpR functions as a repressor at this locus (Fig 5A). Known targets of OmpR activation, including ssrA and ompR (ompB), were highly down-regulated in the absence of ompR (Fig 5A). The ssrA transcript was down-regulated in the envZ null strain; it was restored to the WT levels upon supplying envZc in trans. This result further demonstrates that OmpR activation is via EnvZc (Fig 5B). The envZ null strain showed higher expression of cadA (1.2-fold), cadB (1.4-fold), and cadC (5.9-fold) and failed to acidify its cytoplasm upon acid stress (Fig 3).

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Fig 5. OmpR directly represses cadC/BA to prevent recovery from acidification in the macrophage vacuole.

(A) mRNA levels of cadA, cadB, cadC, ssrA, and ompR genes were determined by quantitative real-time polymerase chain reaction (qRT-PCR) from WT and ompR null strains as described in Materials and Methods. The error bars represent the mean ± SD (n = 6). The ompR null strain showed increased expression of cadC/BA genes, indicating OmpR repression. (B) mRNA levels of cadA, cadB, cadC, and ssrA genes were determined by qRT-PCR from WT, envZ null mutant and the envZ null mutant complemented with envZc as described in Materials and Methods. The error bars represent the mean ± SD (n = 6). The envZ null strain showed increased expression of cadC/BA genes, indicating a role for EnvZ in OmpR-mediated repression. The cytoplasmic domain of envZ alone could restore ssrA expression levels when provided in trans, indicating that it was responding to cytoplasmic signals. (C–F) Electrophoretic mobility shift assays (EMSA) experiments were conducted to examine the interaction between OmpR and cadC/BA. OmpR was incubated with 10 fmol of biotin end-labeled cadA (C), cadB (D), cadC (E), along with 60 bp biotin end-labeled EBNA DNA (F) as a negative control. Addition of 50-fold excess unlabeled DNA is indicated by an asterisk. (G) Representative images of D/A ratios are shown at 30 and 180 min post-infection of WT, the cadBA double null mutant, an ompR/cadBA triple null strain and a cadBA over-expressed strain of Salmonella. Infection is described in Materials and Methods. Scale bar, 3 μm. (H) Plot of the D/A ratios of various mutants of Salmonella. The D/A ratio of approximately 30 cells were calculated at the indicated time points post-infection. Symbols represent the mean ± SEM (n = 3).

https://doi.org/10.1371/journal.pbio.1002116.g005

To examine whether OmpR repression was a result of a direct interaction, we performed electrophoretic mobility shift assays (Fig 5C–5F). DNA sequences upstream of cadC (-354 to -3), upstream of cadB (-362 to -1) and 264-bp upstream to 34-bp downstream of cadA relative to the ATG in each case were amplified using primers that were biotinylated on the 5′-ends. The resulting biotinylated DNA fragment was used in the assay with purified OmpR or OmpR~P prepared by phosphorylation from acetyl phosphate (Fig 5C) [38]. The formation of an OmpR~P-DNA complex is clearly visible in the presence of 0.3 μM OmpR~P at both cadB and cadC promoters (Fig 5D and 5E). The addition of 50-fold excess of unlabeled cadB/C DNA resulted in the release of the labeled probes (Fig 5D and 5E, OmpR~P*), confirming that the complexes formed were due to a specific interaction of OmpR~P with DNA. Neither OmpR nor OmpR~P bound to a 60 bp biotin end-labeled Epstein-Barr Nuclear Antigen (EBNA) DNA, a control for non-specific binding (Fig 5F). In the absence of phosphorylation, OmpR bound to DNA (Fig 5D and 5E), but with considerably lower affinity [38]. Interestingly, no binding was evident at cadA (Fig 5C), suggesting cadB and cadA may be co-transcribed. Thus, OmpR represses cadC/BA by directly binding to upstream DNA at cadC and cadB.

We next measured the intracellular pH of the cadBA null strain in the macrophage vacuole and compared it to the WT and ompR null strains (Fig 5G and 5H). If OmpR functions as a repressor of cadC/BA (Fig 5A), we would expect the cadBA null strain to be similar to the WT strain in terms of cytoplasmic acidification, i.e., knocking out cadBA would be the same as OmpR repressing cadBA. Indeed, the pH_i of the cadBA null strain was similar to WT Salmonella and unlike the bafilomycin-treated strain or the ompR null strain (see Figs 4, 5G and 5H). This result identifies the cadC/BA operon as the major system that eliminates protons upon acid stress and maintains pH homeostasis when Salmonella is in the macrophage vacuole. OmpR represses the cadC/BA system to prevent recovery from acidification. When the cadBA genes are not present, the presence of OmpR is not required for acidification (Fig 5G and 5H), indicating a key role for OmpR in repressing the cad system in the vacuole. This acidification is required for activation of SPI-2 effector expression and secretion (see below).

Over-expression of cadBA was able to completely over-ride the OmpR-dependent acidification, and the Salmonella cytoplasm remained neutral both during macrophage infection and in response to in vitro acid stress (Fig 5G and 5H; S8 Fig). The pH was similar to an ompR null strain (Fig 2B) or to BAF-treated cells (Fig 4), clearly confirming that cadBA enables the ompR mutant to restore pH homeostasis in the macrophage vacuole. The cadBA null strain acidified more rapidly than the WT strain, presumably because it takes some time for OmpR to repress the cad system (S8 Fig). However, the intracellular pH was nearly identical to the WT strain by 120 min.

OmpR Up-Regulates atpB, Increasing Intracellular ATP Levels

The F₁F_o ATP synthase employs the proton gradient to synthesize ATP [39]. Because the Salmonella cytoplasm was acidified when grown at pH_e 5.6, we anticipated cytosolic ATP levels would be higher than at neutral pH. Indeed, switching Salmonella from pH 7.2 to pH 5.6 increased intracellular ATP levels 1.8-fold compared to the ompR null strain (S9A Fig). ATP was restored to WT levels when the ompR null strain was complemented with ompR supplied in trans. Control experiments with an atpB null strain or in the presence of the uncoupler carbonyl cyanide m-chlorophenyl hydrazone (CCCP) drastically reduced intracellular ATP. In our microarray, ATP synthase genes were down-regulated in the absence of ompR by 2- to 3-fold. We then compared the mRNA levels of atpB, a gene encoding the a F_o subunit, by qRT-PCR in the WT and ompR null strains. The transcript for atpB was up-regulated (2.1-fold) in the WT strain, suggesting a role for OmpR in activation of atpB (S9B Fig). Thus, OmpR appears to play a dual role in acid stress, by activating proton translocation via increased F₁F_o ATP synthase coupled with elimination of proton consumption via repression of the cad system. Taken together, these combined effects enable WT Salmonella to reduce intracellular pH upon acid stress.

Acidification Leads to SseB Translocon Movement Away from the Salmonella Cell Surface

We next investigated the kinetics of translocon secretion by immuno-labeling SseB in macrophages at various times post-infection. Salmonella were visualized using anti-LPS antibody up to 6 h. SseB was detectible immediately 30 min post-infection at one locus of the cell, and the number of SseB-labeled cells increased over time. After 3.5 h, SseB was no longer associated with a majority of Salmonella cells, but was removed from the cell surface (Figs 6A and 7). SseB was also detected in Salmonella-infected macrophages treated with BAF, but SseB remained on the bacterial cell surface (Figs 6B and 7A).

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Fig 6. The translocon protein SseB is released from the Salmonella surface over time, and acidification is required.

(A) WT Salmonella were electroporated with unlabeled I-switch, recovered, and used for infection in RAW264.7 macrophages for various indicated times. 25 nM BAF (B), or an equal volume of DMSO, was used to pre-treat the macrophages for 30 min prior to infection and was maintained throughout the infection time. The cells were fixed, permealized, and stained for Salmonella LPS (green) and translocon protein SseB (red). Scale bar, 3 μm.

https://doi.org/10.1371/journal.pbio.1002116.g006

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Fig 7. The percentage of detached SseB increased over time upon acidification.

(A) The percentage of detached SseB was plotted over time post-infection as described in the text. (B) The distance (in μm) of 50 SseB protein spots from the nearest bacterial cell surface was plotted as a function of time post-infection. Symbols are represented as the mean ± SEM.

https://doi.org/10.1371/journal.pbio.1002116.g007

Measurement of SseB from the Salmonella Cell Surface

To measure the extent to which SseB protein was separated from the bacterial cell surface, the Distance Transformation function of Imaris Software (version 7.7.1) was employed. The distance of the red SseB translocon protein to the nearest green Salmonella cell was measured by applying the distance transformation on the green channel surface to generate a new channel whose intensity indicated the shortest distance to the surface object border. Hence, for each red SseB protein spot, the minimum distance to its nearest green cell was obtained. For each time point post-infection, the distance of 50 SseB protein spots from the nearest cells were measured. A distance value of 0 indicated that SseB was attached to the cell surface, whereas a distance value larger than 0 indicated that SseB was separated from the surface. The percentage of distant SseB was plotted over time post-infection (Fig 7A). Using this approach, it was evident that the vast majority of SseB (>60%) were no longer surface-associated by 3.5 h, and by 6 h post-infection this value increased to 88% (Fig 7A). When acidification was prevented by treatment with BAF, the number of cells in which SseB was detached was only 5%–10%. Measurements of the distance (in μm) of SseB from the bacterial cell surface as a function of time post-infection are shown in (Fig 7B). At 6 h post-infection, the average distance from the Salmonella cell surface was approximately 3 μm, indicating that SseB was no longer cell-surface associated.

SseB Co-localizes with the SCV Membrane after it Separates from the Bacterial Surface

Salmonella translocated effectors (SifA, SseF, SseG, PipB, and SpiC) localize to the SCV and to its tubular membranous structures termed Salmonella-induced filaments (Sifs) [40–42]. The Sifs extend from and connect SCVs during Salmonella replication inside host cells [43]. Therefore it was possible that once SseB was separated from the cell surface, it might be in association with endosomal tubules of vacuolar membrane origin. To address this question, we used immunofluorescence to compare the localization of SseB with the endosomal membrane marker, LAMP-1. RAW macrophages were infected with Salmonella WT harboring pFPV25.1 for constitutive expression of GFP and immunolabeled for LAMP-1 (blue) and SseB (red). Confocal microscopic images indicated that when SseB was separated from the Salmonella surface, it was co-localized with LAMP-1 positive membranes, resulting in magenta staining of SseB (Fig 8A). Nearly 90% of the translocon protein SseB was co-localized with LAMP-1 by 3.5 h and this level did not change up to 6 h post-infection (Fig 8B).

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Fig 8. As SseB moves away from the Salmonella surface, it is associated with the vacuolar membrane.

(A) RAW264.7 macrophages were infected with WT Salmonella harboring pFPV25.1 for the constitutive expression of GFP (green). Cells were fixed, permeabilized, and immunostained for mouse monoclonal anti-LAMP-1 (blue) and rabbit polyclonal anti-SseB (red) followed by Alexa568- or Alexa647-labeled secondary antibodies, respectively, for various times as indicated. The merged image indicates that once SseB was detached from the bacterial cell surface, it was co-localized with the host endosomal membrane (i.e., LAMP-1–positive membranes). Images were obtained using the Nikon A1R confocal microscope. Localization resulted in magenta staining of Salmonella SseB as analyzed by Image J software. Scale bar = 3μm. (B) Results of three independent experiments in which macrophages were infected with WT GFP-expressing Salmonella and immunostained with LAMP-1 and SseB at 3.5 h, 4 h, 5 h, and 6 h post-infection. The values represent the mean ± SEM.

https://doi.org/10.1371/journal.pbio.1002116.g008

In the macrophage vacuole, a single focus of SseB was evident on the bacterial surface (Fig 6). This was similar to the localization of SseB that we observed in vitro (S10A Fig, bottom panel). Other components of the T3SS exhibited similar patterns of localization, including the outer membrane ring protein SsaC (top panel) and the inner membrane ring protein SsaJ (middle panel). Thus, the SPI-2 needle complex is localized primarily near to one end of the cell pole, unlike SPI-1 needles, which are distributed all over the bacterial surface [6,44]. An ssaC null strain was deficient for secretion and SseB was retained in the cytoplasm, substantiating that the focal assembly of the SPI-2 needle complex was competent for SPI-2 effector secretion (S10B Fig). Our observation that only approximately 13% of the population expressed SPI-2 needles on their surface in vitro highlights that, in contrast to SPI-1, SPI-2 is not abundant.

Acidification Leads to SseJ Effector Secretion After SseB Translocon Surface Release

To gain insight into the fate of effector secretion, we also compared the kinetics of SseJ secretion in Salmonella-infected macrophages by immunofluorescence microscopy. A strain expressing SseJ-HA was completely comparable to WT in terms of its intracellular pH response and replication in macrophages. SseJ-HA secretion was first detected 3.5 h post-infection in macrophages, similar to a time course of SseF-HA expression in HeLa cells reported previously [20]. The percentage of cells secreting SseJ increased between 3.5 and 6 h post-infection (Fig 9, visible as red puncta in the macrophage cytoplasm). In contrast to SseB secretion (Fig 6A), SseJ was not translocated in the presence of BAF, indicating that cytoplasmic acidification was essential for effector secretion (Fig 9B, lower panel; i.e., no red puncta are visible in the BAF-treated macrophages). SseJ secretion was subsequent to SseB release from the bacterial surface (Fig 6A and 6B). These results are in agreement with previous results [19] but significantly differ from other studies that reported that a neutralization step from the host was required for SPI-2 effector secretion [20]. Our results indicate that formation of the translocon pore is essential and precedes effector secretion. To verify that the translocon complex was required for effector secretion, we monitored the kinetics of SseJ secretion in an sseB null strain expressing SseJ-HA. No translocation of SseJ was evident, even after 6 h post-infection in the macrophages in the absence of SseB (S11 Fig).

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Fig 9. Acidification is required for secretion of effector SseJ.

(A) RAW264.7 macrophages were infected with I-switch-electroporated Salmonella WT harboring psseJ-HA for expression of SseJ-HA. Cells were immunostained for Salmonella LPS (green) and HA epitope (red), followed by Alexa488- or cy5-labeled secondary antibodies, respectively, for various time points as indicated. (B) 25 nM BAF or an equal volume of DMSO (A) was used as a control. Samples were imaged by confocal microscopy and analyzed using ImageJ software. Scale bar, 3 μm. SseJ secretion was observed (red puncta) after SseB release from the surface after 3.5 h.

https://doi.org/10.1371/journal.pbio.1002116.g009

Cell Culture, Bacterial Strains, and Antibodies

RAW 264.7 cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA, United States) and grown in a humidified 37°C, 5% CO₂ tissue culture incubator. Cell culture reagents were from Life Technologies (Carlsbad, CA, US) unless otherwise stated. RAW 264.7 were maintained in Dulbecco’s Minimal Essential Media (DMEM) containing 2 mM L-glutamine, 1 mM sodium pyruvate and 17.5 mM D-glucose with 10% heat-inactivated fetal bovine serum (FBS). RAW 264.7 cells were always cultured in antibiotic-free media. Low passage number cells (<15 after receipt from ATCC) were seeded in 24-well dishes (Nunclon) and grown O/N before infection. The cells were washed gently with PBS and removed from the flask by scraping. The cells were suspended at a density of 1 × 10⁵ cells per well in 24-well plates. WT S. enterica serovar Typhimurium 14028s and the ompR null derivative (ΔompR) were used for all infection experiments unless otherwise indicated. DMSO was obtained from Sigma-Aldrich (St Louis, MO). To determine the acid stress response, bacterial strains were grown as in [60] in a modified N-minimal medium (MgM) buffered with 20 mM Tris (pH 7.2) or 20 mM MES (pH 5.6) containing 7.5 mM (NH₄)₂SO₄, 5 mM KCl, 0.5 mM K₂SO₄, 1 mM KH₂PO₄, 10 mM MgCl₂, 2 mM glucose, and 0.1% Casamino acids. The following antibodies were used for Immuno-labeling experiments: rabbit polyclonal anti-SseB, rabbit monoclonal anti-HA (Sigma), mouse monoclonal anti-Salmonella LPS (Abcam), rabbit anti-LAMP-1 (Abcam), mouse Alexa-488 IgG (Invitrogen), rabbit Alexa-488 IgG (Invitrogen), and rabbit cy5 IgG (Invitrogen).

Transformation of the I-switch in Salmonella

I-switch sample preparation was performed as previously described [14] with the following modifications: 50 μM O1, O2, and O3 were mixed in equimolar ratios in 20 mM potassium phosphate buffer at the desired pH containing 100 mM KCl. Primers were annealed using touch-down PCR with a decrease in temperature of 1°C per 3 min from 95°C to 25°C and equilibrated overnight at 4°C before use. A concentration of 6 μM of I-switch DNA was used to transform electro-competent Salmonella cells. After electroporation, cells recovered for 1 h at 37°C in Super Optimal Broth (SOB) containing 20 mM glucose and 10 mM MgCl₂ (SOC) unless otherwise stated. Cells were washed three times with PBS (Life technologies) and used for analysis.

pH Clamping of Salmonella

pH clamping protocols were modified based on [14]. Double-labelled I-switch (I_A488/A647) was diluted to 60 nM in clamping buffers (120 mM KCl, 5 mM NaCl, 1 mM MgCl, 1 mM CaCl, 20 mM HEPES buffered at various pH). The samples were excited at 488 nm and emission collected between 500–750 nm in a Tecan Spectrophotometer (GENios). The fluorescence intensity at 520 nm (D) was divided by the fluorescence intensity at 666 nm (A) and the D/A ratio was plotted over the pH range. For in vivo pH clamping experiments, bacterial cells electroporated with the I-switch were incubated in a K⁺-rich clamping buffer at various pH at RT for 1 h in the presence of 40 μM nigericin. Cells were placed on microscope slides (Marienfeld, Germany) coated with 1% agarose and then imaged by wide field fluorescence microscopy. Three independent measurements were recorded for each pH value and the D/A ratios were plotted as the mean ± SEM.

Salmonella Infection in RAW264.7 Macrophages

Salmonella were first electroporated with the I-switch as described. Cells were seeded onto overnight-grown monolayers of RAW264.7 cells at a multiplicity of infection of 100:1 in 24-well plates. The plates were centrifuged at 500 rpm for 10 min to synchronize the infection. The infection was conducted for 30 min at 37°C in 5% CO₂. Cells were washed three times with PBS and incubated with DMEM containing 100 μg/ml of gentamicin for 1 h at 37°C. Cells were again washed three times with PBS and incubated in DMEM containing 10 μg/ml gentamicin for the remainder of the experiment. For BAF-treated cultures, RAW264.7 cells were pre-treated with 25 nM BAF for 30 min prior to infection and 25 nM BAF was maintained in every step of the infection. At the stated time points, the cells were fixed using 3% PFA for 15 min at RT. The coverslips were mounted on a drop of gold anti-fade reagent (Invitrogen) and sealed with entellan (Merck). Samples were imaged using confocal microscopy (Nikon A1R) and analyzed by Image J imaging software. To inhibit vacuolar acidification, 25 nM BAF or an equal volume of DMSO were added to the macrophage cells 30 min prior to infection and maintained throughout the infection process.

Wide-Field Fluorescence and Confocal Microscopy

All the wide-field images were collected using an Olympus IX71 Inverted Microscope (Applied Precision DeltaVision Deconvolution microscope system) equipped with 100X, 1.4 numerical aperture (NA) objective lens with a mercury arc bulb (OLYMPUS U-RFL-T: Mercury 150W) as the illumination source. Image capture was performed with the CoolSnap HQ, a fast, high resolution, high quantum efficiency, cooled CCD camera (Photometrics CoolSNAP HQ2 (CCD); 1,392 x 1,040 pixels; 11 fps). Two sets of images were taken corresponding to (i) donor emission wavelength (525/36 nm) upon donor excitation (490 ± 20 nm) (donor image: D), and (ii) acceptor emission wavelength (666 nm long pass) (acceptor FRET: A) upon donor excitation (490/20 nm). Acceptor images (I) acceptor emission wavelength (666 nm long pass) upon acceptor excitation (645 ± 20 nm) were obtained for cells clamped at pH 5.2 and 7.1. Confocal imaging was carried out on an inverted Nikon A1Rsi-Polarizing Module equipped with CFI Plan ApochromatVC 100X, 1.4 numerical aperture (NA) objective lens by using a Coherent CUBE laser at 488 excitation with set appropriate dichroics. Excitation at 470/40 nm wavelength was performed through a 510 nm dichroic mirror, and emission was recorded through a 525/50 nm band pass filter on a Andor DU897 EMCCD camera to yield the donor image (D). Similarly, emission was recorded through a 700/75 nm band pass filter upon similar excitation to generate the FRET intensity (A). Multiple Z sections were taken (0.5 μm apart) and each image is represented as sum slices. Images were analyzed using ImageJ ver.1.42. All of the images were background-subtracted by taking the mean intensity over an adjacent cell-free area. Autofluorescence of unlabeled Salmonella in RAW 264.7 cells was measured and used to correct the determined pH values. However, there was no difference in the intracellular pH values when we corrected for auto-fluorescence compared with the uncorrected values. Because the measurement is ratiometric, auto-fluorescence did not influence the determined pH values. Images showing the D/A values were obtained by using the ratio plus plugin of ImageJ software. The high and low values were color-coded and calibrated to their respective pH values as described in the main text. For all studies in macrophages, the mean D/A intensity of at least thirty bacterial cells was measured and is represented as the mean ± SEM of three independent batches of experiments.

Measurement of BCECF in Salmonella

The free acid form of BCECF was suspended in buffers ranging from pH 5.0 to 7.2. Fluorescence intensities were measured at excitation wavelengths of 488 and 440 nm and the emission wavelength was recorded at 525 nm in a Tecan spectrophotometer (GENios). This generated the in vitro standard curve. Overnight cultures of WT and the ompR null mutant of Salmonella were clamped in 100 mM potassium phosphate buffer at various pH containing 40 μM nigericin and incubated in the presence of 20 μM BCECF-AM. Paired images were obtained by using excitation wavelengths of either 440 nm (pH-insensitive wavelength) or 488 nm (pH-sensitive wavelength). Fluorescence emission was recorded through a 525/50-nm band pass filter on a Andor DU897 EMCCD camera on an inverted Nikon A1Rsi-Polarizing Module by using Coherent CUBE lasers at 488 and 440 excitation with set appropriate dichroics. The ratios were plotted as a function of pH, and are comparable to the in vitro calibration curve. WT and the ompR null mutant of Salmonella were incubated at either acidic pHe (5.6) or neutral pHe (7.2) for 2 h. 20 μM BCECF-AM was added to the cultures for 30 min before imaging. To determine whether acidification was growth phase dependent, cells were grown until O.D. = 0.56 (log phase) or O.D. ≈ 2.3 (stationary phase) before the addition of the probe. The ratios of the fluorescence intensities of emission channel (525 nm) upon 488 nm excitation and 440 nm excitation were obtained as before. Cells were placed on microscope slides (Marienfeld Germany) coated with 1% agarose and then imaged. Images were first corrected for background fluorescence by subtracting an image of a cell-free area and analyzed by Image J imaging software.

Preparation of Salmonella Spheroplasts

I-switch single labeled with Alexa 647 (I_A647) was transformed in Salmonella as described above. Spheroplasts were prepared as described in [61] with the following modifications; Cells were resuspended after recovery in cold hypertonic solution A (0.75M sucrose 10 mM TrisHCl pH 7.8). 5 mg/ml of lysozyme solution was gently added to the cell suspension and the mixture was kept on ice for 5 min. EDTA solution (1.5 mM EDTA pH 7.5) was then added slowly over 10 min. Cells were centrifuged at 4,000 rpm for 20 min and re-suspended in solution D (0.25M sucrose 3 mM Tris-HCl pH 7.8, 1 mM EDTA pH 7.5). Cells were imaged on Applied Precision DeltaVision Deconvolution microscope system.

Construction of Mutants and HA/GFP/mCherry Tagged Strains

The sseJ-HA fusion or ompR DNA sequences were amplified using primers XhoI-PsseJ #F and sseJ-HA-BamHI #R or KpnI-PompR #F and envZ-HindIII #R from Salmonella 14028s genomic DNA. The PCR products and low copy promoter-less pWSK29 vector were double-digested with restriction enzymes XhoI and BamHI or KpnI and HindIII (Thermo Fisher Scientific), and the digested samples were ligated with the Rapid DNA Ligation Kit (Thermo Fisher Scientific). The cadA, cadB, cadC, ompC, ompF, mgtC, atpB, and sseB genes were deleted from the Salmonella chromosome using λ-Red recombination techniques as described previously [62,63]. A cadBA over-expressed strain was generated by cloning cadBA into plasmid pBR322, replacing the bla gene with cadBA. SsaC-GFP and SsaJ-mCherry were constructed as follows: plasmid pWSK29 containing ssaC with the spiC promoter fused before GFP and ssaJ with the sseA promoter fused before mCherry were inserted in ssaC and ssaJ null strains, respectively. Primers are listed in S1 Table. A list of strains and plasmids are provided in S2 Table.

Immunofluorescence Detection in Macrophages

For every time point post-infection, cells were fixed with 3% paraformaldehyde for 10 min at RT followed by three PBS washes. Cells were probed with various primary and secondary antibodies in PBS containing 2% BSA and 0.1% saponin. Antibody staining was done sequentially for 1 h, and after each incubation, washes were performed five times with PBS. The HA epitope tag was detected with the monoclonal HA antibody at 1:500 (Covance); LAMP-1 was detected with fluorescein isothiocyanate (FITC)-conjugated rabbit monoclonal antibody at 1:500 (Research Diagnostics); Salmonella was detected with mouse polyclonal anti-lipopolysaccharide (anti-LPS) antibodies at 1:500 (Difco). Secondary antibody detection was performed with various anti-rabbit or anti-mouse antibodies conjugated to Alexa-488 and cy5.

Determination of Intracellular Salmonella Replication in Macrophages

To determine intracellular replication within macrophages, a similar protocol was followed as reported in [64] with slight modification. Salmonella was electroporated with non-labeled I-switch, and used for infection (see text for details). At 2 and 16 h post-infection, cells were washed repeatedly in PBS and lysed in 0.1% Triton-X100 at RT for 15min. Samples were then serially diluted and plated on LB to enumerate the intracellular bacteria. Replication was described as the CFU/ml counted and represented as an average of three plates for all time points.

Electrophoretic Mobility Shift Assay

Electrophoretic mobility shift assays (EMSAs) were performed using the Lightshift chemi-luminescence EMSA kit (Pierce) according to the manufacturer’s instructions described in [65]. The upstream regions of cadC (354 bp), cadB (361 bp), and cadA (264 bp upstream to 34 bp downstream) were amplified using biotinylated oligonucleotides. Ten fmol of biotinylated DNA was used in a 15 μl reaction containing binding buffer (10 mM Tris, pH 7.5, 50 mM KCl) along with 2.5% glycerol, 5 mM MgCl₂, 0.05% Nonidet P-40, and 1μg poly(dI-dC)). OmpR or OmpR~P protein was added at the concentrations indicated, and samples were separated by electrophoresis on 5% non-denaturing acrylamide gels run in 0.5X Tris-acetate buffer with EDTA. Following electrophoresis, DNA was electro-transferred to a nylon membrane and detected using the biotin detection system (Pierce).

RNA Isolation and Quantitative Real-Time RT-PCR

Salmonella strains were grown in SPI-2 inducing Minimal Magnesium Medium (pH 5.6) as described previously [60] to O.D ~0.6. The total RNA was isolated using an RNeasy mini kit (Qiagen). After DNase treatment of the isolated RNA, cDNA was synthesized using the iScript Reverse Transcription Supermix (Biorad). Quantification of cDNA was carried out using SsoFast ^TM Eva Green Supermix (Bio-Rad), and real-time amplification of the PCR products was performed using the iCycler iQ real-time detection system (Bio-Rad). The mRNA expression level of the target gene was normalized relative to the 16S rRNA expression level.

Acid Tolerance Response Assay

Cultures were grown in LPM media (pH 7.0) to O.D ~1.5 as described in [66] with slight modifications. Unadapted cultures were immediately shifted to pH 3.0 acidified by HCl. Adapted cultures were shifted to pH 4.5 for 2 h prior to the challenge at pH 3.0. Viable cells were calculated by plating aliquots of serially diluted cultures by standard plate count.

Measurement of Intracellular ATP in Salmonella

Intracellular ATP levels using Tecan spectrophotometer as described with modifications [37]: Overnight cultures of Salmonella in LB were sub-cultured in MgM (7.2) for 24 h. The cultures were washed in MgM (5.6) and inoculated in 5 ml of the same media for 5 h. Cells were normalized by the OD₆₀₀ and re-suspended in 500 μl of (PBS). Nucleic acids were extracted by adding ice cold 1% Trichloroacetic acid (TCA) and 2 mM EDTA for 15 min. The extracts were neutralized with 100 μl of neutralization buffer and centrifuged for 15 min. The supernatant was diluted 2-fold with L buffer and the luciferase reaction was initiated using an ATP determination kit (Invitrogen) according to the manufacturer's instructions. Intracellular ATP levels (μM) were converted using reference to standards of known concentration.

Immunoblotting

Overnight cultures of Salmonella WT and ssaC null mutant strains in LB were sub-cultured in MgM (7.2) for 24 h. The cultures were washed either in MgM (5.6) or MgM 7.2 and inoculated in 50 ml of the same media for 5 h. For isolation of the secreted protein, cells were removed from the culture by centrifugation (5,500 x g, 20 min, 4°C), and the supernatant was filtered through a 0.22 μm pore size binding filter (Millipore, Billerica, MA). The secreted fraction was isolated by 10% trichloroacetic acid precipitation, and the protein pellet was washed three times with −20°C acetone and then air dried. Equivalent amounts of cellular protein, adjusted according to the optical densities, were used for the whole cell fraction. The protein samples were separated by 12% SDS-PAGE and transferred to PVDF membrane (Millipore). The membrane was incubated with anti-SseB (1:10,000) or anti-GroEL (1:5,000) antibodies in PBS buffer (with 0.05% Tween 20 and 1% Skim Milk) followed by anti-rabbit secondary antibody (1:5,000, Santa Cruz Biotechnology).