Bioenergetic State of Escherichia coli Controls Aminoglycoside Susceptibility

To investigate whether the type of metabolism used for growth influences the level of AG susceptibility, we measured MIC of the wild type (WT) strain in aerobiosis and anaerobiosis, in Luria-Bertani (LB) medium supplemented with fumarate, NO₃^–, or no electron acceptor, where in the latter instance we assumed mixed-acid fermentation was used (26). Under all anaerobic growth conditions tested, the MIC value of Gm was 4-fold higher than under oxygen, i.e., 8 μg/mL.

As a complementary analysis, we performed time-dependent killing tests using 16 μg/mL of Gm (2× MIC) on strains growing anaerobically in LB supplemented or not with NO₃^– or fumarate. Under these conditions, the susceptibility level of the WT strain varied with the nature of the added electron acceptor (Fig. 1A). Indeed, E. coli survived Gm better in the absence of the added electron acceptor (in LB or LB supplemented with glycerol) than in the presence of fumarate or NO₃^– (Fig. 1A). Interestingly, higher survival was observed for cells grown with fumarate compared with NO₃^– (Fig. 1A). To verify that Gm sensitivity was caused by respiratory activity, we tested survival of strains deficient in respiratory chains of NO₃^– or fumarate. NarGHI is the major nitrate reductase, while fumarate respiration is catalyzed by the multicomponent Frd fumarate reductase (FrdABCD). The MIC value of Gm was equivalent to that of the WT (8 μg/mL) for ΔnarG and ΔfrdA mutants under either NO₃^– or fumarate respiration, respectively. However, the time-dependent killing experiment showed that the ΔnarG and ΔfrdA mutants exhibited an increased level of tolerance to Gm compared to the WT strain (Fig. 1B and C). Overall, these results showed that the respiratory chains have a direct impact on the level of Gm susceptibility and suggest that the lower the energetic yield of the respiratory chain used, the higher the level of Gm survival.

We then focused on the low energy-conserving respiratory chain established under fumarate respiration to pinpoint which components are required for Gm sensitization. Fumarate reduction can be coupled to hydrogen oxidation in an electron transfer chain, including Hyd-2, menaquinol-fumarate oxidoreductase FrdABCD as a terminal reductase, and MK and/or DMK (27) as electron carriers (28). To test the contribution of Hyd-2 and/or MK/DMK, we used the ΔhybC mutant, lacking the large subunit of Hyd-2, which has primarily hydrogen-uptake activity, and the ΔmenA mutant, lacking a 1,4-dihydroxy-2-naphthoate octaprenyltransferase required for MK/DMK biosynthesis (29). Time-dependent killing assays were performed in LB medium supplemented with 10 mM fumarate and 0.2% glycerol and using Gm at 16 μg/mL. The ΔhybC and ΔmenA mutants showed increased tolerance to Gm compared to the WT strain (Fig. 2A and E). These results indicated that under fumarate respiration, sensitivity to Gm requires functional Hyd-2 and MK/DMK. Hyd-2 also oxidizes H₂ produced by the FHL complex. The FHL complex includes formate dehydrogenase (Fdh-H), which produces 2H⁺, 2e⁻, and CO₂ from formate, and hydrogenase 3 (Hyd-3), which generates molecular H₂ from 2H⁺ and 2e⁻. We therefore tested the contribution of the FHL complex to the sensitivity to Gm under fumarate respiratory conditions. Specifically, we tested the influence of mutations in genes that affected synthesis of the large subunits of Hyd-3 (hycE), formate dehydrogenase H (fdhF) and FdhD, an auxiliary maturing factor. Time-dependent killing assays were performed in LB medium supplemented with 10 mM fumarate and 0.2% glycerol and using Gm at 16 μg/mL. The three mutants ΔhycE, ΔfdhF, and ΔfdhD showed increased tolerance of Gm compared with the WT strain (Fig. 2B to D).

E. coli contains another respiratory hydrogenase, Hyd-1, which catalyzes the oxidation of H₂ to protons and electrons, but does not couple H₂ oxidation to fumarate reduction (30, 31). We tested the effect of the Hyd-1 complex on Gm susceptibility by exposing ΔhyaA or ΔhyaB mutants, each lacking a subunit of Hyd-1, to 16 μg/mL Gm. Both mutants exhibited similar antibiotic susceptibility to WT, showing that Hyd-1 does not influence Gm tolerance under such conditions (Fig. 2F).

Fumarate respiration could also involve Nuo or GlpABC complexes as primary electron donors (19). Accordingly, we tested the effect of mutations in genes encoding these two complexes, e.g., nuoC and glpA, on Gm-mediated killing. Neither the ΔnuoC nor the ΔglpA mutation, alone or in combination, altered the level of Gm sensitivity (see Fig. S1 in the supplemental material).

Taken together, these results demonstrate that in E. coli growing anaerobically in the presence of fumarate (and glycerol), Gm sensitivity is caused by the activity of the FHL-Hyd-2-MK/DMK-FrdA electron transfer chain.

It has been proposed that the RavA-ViaA complex acts as a chaperone by assisting in the assembly of fumarate reductase (22). Therefore, we tested the effect of the RavA-ViaA complex on the AG sensitivity of E. coli grown under anaerobic fumarate respiration conditions. The ΔravA-viaA mutant exhibited the MIC value of Gm during fumarate respiration equivalent to that for the WT (8 μg/mL). However, a time-dependent killing experiment using a Gm concentration equivalent to 2× MIC (16 μg/mL) revealed a significant increase in Gm tolerance of the mutant compared to the WT strain (Fig. 3A). The combination of ΔravA-viaA and ΔfrdA mutations showed no additive effect (Fig. 3A).

A previous analysis showed that RavA-ViaA exerted a slight negative effect on FrdA enzymatic activity (22). We therefore tested whether RavA-ViaA might cause an energetic disadvantage in strains growing using fumarate respiration. WT and ΔravA-viaA strains were mixed in a 1:1 cellular ratio and grown together in M9-glycerol and fumarate under anaerobic conditions for 48 h. The competitive index was determined by counting CFU of the ΔravA-viaA strain, using its chloramphenicol-resistance phenotype, and CFU of the WT at t₀ and t₄₈. The competitive index (CFU_mutant/CFU_wt)t₄₈/(CFU_mutant/CFU_wt)t₀ yielded a median value of 1.3, revealing no growth advantage of one strain over the other (Fig. 3B). These results showed that RavA-ViaA in vivo does not influence growth under fumarate respiration and presumably does not modulate FrdA enzymatic activity to a significant extent, if at all.

Taken together, these results indicate that under fumarate respiration conditions, the RavA-ViaA complex sensitizes E. coli to Gm via a FrdA-dependent mechanism.

Next, we tested whether fumarate was required for the RavA-ViaA complex to exert its sensitizing effect. Thus, E. coli was grown in the absence of an exogenously added electron acceptor, i.e., in LB medium, LB-0.2% glycerol or LB-0.2% glucose. First, MIC values for Gm in all these media were found to be 8 μg/mL. Next, we performed a killing assay using Gm at 16 μg/mL (2× MIC) in LB (Fig. 3C) and in LB glycerol (Fig. 3D), and at 30 μg/mL (~4× MIC) in LB glucose (Fig. 3E). The ΔravA-viaA mutant showed increased tolerance to Gm compared to the WT strain in all media. Notably, complementation of the ΔravA-viaA mutant with a multicopy plasmid (pRV plasmid) containing the tac_p::ravA-viaA operon allele, suppressed its increased tolerance (Fig. 3E). These results indicated that RavA-ViaA exerts a sensitizing effect on E. coli under anaerobic conditions even in the absence of an added exogenous electron acceptor.

Next, we tested whether RavA-ViaA was able to sensitize E. coli to Gm under NO₃^– respiration. First, we determined the MIC value of Gm in LB medium supplemented with 10 mM NaNO₃ and 0.2% glycerol and found it to be 8 μg/mL for both WT and ΔravA-viaA strains. Under the same conditions, a time-dependent killing assay done with Gm at 16 μg/mL showed that the ΔravA-viaA mutant was as susceptible as the WT strain (Fig. 3F). These results indicated that the sensitization of E. coli to Gm under NO₃^– respiration is independent of RavA-ViaA.

To investigate the spectrum of the sensitizing effect of the RavA-ViaA complex, we measured the survival rate of the ΔravA-viaA strain to another AG (tobramycin), a protein synthesis inhibitor (tetracycline), a fluoroquinolone (nalidixic acid), and a β-lactam (ampicillin). We observed that the ΔravA-viaA mutant was more tolerant to tobramycin than the WT strain (Fig. 4A). A modest tolerance effect was noted upon short time exposure to tetracycline (Fig. 4B). The ΔravA-viaA mutation had no effect on the potency of nalidixic acid and ampicillin (Fig. 4C and D). Overall, these results showed that under anaerobic conditions, RavA-ViaA sensitizes E. coli specifically to AG.

To understand the role of RavA-ViaA in AG sensitization under anaerobic conditions, we performed a Gm uptake assay using ³H-Gm. The accumulation of ³H-Gm in the WT strain gradually increased to 1,200 ng Gm/10⁸ cells after 2.5 h (Fig. 5). In contrast, in the ΔravA-viaA mutant, ³H-Gm accumulation remained below 100 ng of Gm/10⁸ cells after 2.5 h. Our results indicated that RavA-ViaA sensitizes E. coli to Gm by increasing its uptake and, consequently, its intracellular concentration.

We decided to investigate whether the RavA-ViaA complex could have a sensitizing effect under aerobic conditions. The MIC of Gm was found to be 2 μg/mL for both wild type (WT) and ΔravA-viaA strains. In a time-dependent killing experiment using a concentration of Gm equivalent to 2.5× MIC (5 μg/mL), E. coli WT and ΔravA-viaA strains exhibited similar sensitivities to Gm (Fig. 6A). However, because the expression of the ravA-viaA operon is under Fnr control (22), we reasoned that the genes might not be expressed at a high enough level under the conditions used. Therefore, we bypassed Fnr-mediated activation using the pRV plasmid, and tested whether this could sensitize E. coli to Gm. We observed significantly impaired survival of the WT/pRV strain (Fig. 6B), demonstrating the ability of RavA-ViaA to sensitize E. coli to Gm-mediated killing under aerobiosis. Thus, we concluded that the RavA-ViaA complex can also sensitize E. coli to Gm under aerobic respiration, but only when they are produced above a threshold value, a situation that does not appear to be achieved in exponentially growing cells harboring a chromosomal copy of the ravA-viaA operon.

Survival of the WT/pRV strain exposed to Gm was tested in the presence of cyanide-m-chlorophenylhydrazone (CCCP), an ionophore, which dissipates pmf. The data showed that addition of CCCP prevented Gm from killing the WT/pRV strain (Fig. 6C), demonstrating that pRV-mediated killing required pmf. Because pmf results from respiratory metabolism, we tested whether pRV-mediated sensitization was dependent on electron transfer chain (ETC)-forming components. We analyzed the survival of strains defective for the synthesis of ubiquinones, the lipid that acts as an electron transporter in aerobic ETCs. A ΔubiA strain was highly resistant to Gm treatment and the pRV plasmid failed to sensitize this strain to Gm (Fig. 6D). Similarly, a Δnuo Δndh mutation altering NADH dehydrogenase I and II abrogated pRV-mediated sensitization (Fig. 6E). These data supported the idea that respiration is required for pRV-mediated killing of E. coli by Gm in aerobiosis.

The E. coli K-12 strain MG1655 and its derivatives used in this study are listed in Table 1. Deletion mutations (ΔhyaA::Kan^R, ΔhyaB::Kan^R, ΔhybC::Kan^R, ΔhycE::Kan^R, ΔfdhD::Kan^R, ΔfdhF::Kan^R, ΔfrdA::Kan^R, ΔglpA::Kan^R, ΔmenA::Kan^R, ΔnarG::Kan^R, Δndh::Kan^R, ΔnuoC::Kan^R) from the KEIO collection were introduced by P1 transduction. The ΔravA-viaA::Cm^R mutant was constructed using the procedure described by Datsenko K. and Wanner B. (37), using oligonucleotides ravAwanner_up and viaAwanner_do (Table 2), followed by a transduction in MG1655 background. Transductants were verified by PCR, using primer pair hybridizing upstream and downstream the deleted genes. When performed, excision of the antibiotic cassette was done using the pCP20 plasmid (37). Oligonucleotides used in this study are listed in Table 2. E. coli strains were grown at 37°C in Luria-Bertani (LB) rich medium or in minimal M9 medium. Glucose (0.2%), glycerol (0.2%), IPTG (1 mM), CCCP (5 μg/mL), fumarate (10 mM), or NO₃^– (10 mM) were added when indicated. Solid media contained 1.5% agar. For standard molecular biology techniques, antibiotics were used at the following concentrations: chloramphenicol at 25 μg/mL, kanamycin at 50 μg/mL, and ampicillin at 50 or 100 μg/mL.

Plasmid pRV was first constructed by PCR amplification of the coding region of ravA-viaA from the E. coli MG1655 chromosomal DNA using the following primer pair: ravA UP BamHI/viaA DO HindIII (Table 2). The PCR product was then digested by BamHI and HindIII and cloned into the BamHI/HindIII linearized pTrc99A vector (38). The sequence of the inserted fragment was checked by DNA sequencing.

Overnight cultures were diluted (1/100) and grown aerobically or anaerobically in specific medium as indicated in the figure legends at 37°C to an OD₆₀₀ of 0.2. At this point (t0) antibiotic at the indicated concentration was added to the cells. At different incubation times, 100 μL of cells were diluted in sterile phosphate-buffered saline solution (PBS buffer), spotted on LB agar, and incubated at 37°C for 24 to 48 h. Cell survival was determined by counting CFU per mL (CFU/mL). The absolute CFU at time point 0 (used as the 100%) was ≈ 5 × 10⁷ CFU/mL. Survival rate in anaerobic conditions was performed in an anaerobic chamber (Coy and Jacomex Chambers). Materials (medium, tubes, plates, etc.) were all equilibrated in the anaerobic chamber for at least 18 h prior to use.

The MICs were determined by the microdilution method in a 96-well plate according to the Clinical and Laboratory Standards Institute (CLSI) guideline. Briefly, serial dilutions of Gm in a 2-fold manner were done in 100 μL cation-adjusted Müller-Hinton or in LB supplemented or not with either glucose (0.2%), fumarate (10 mM), or nitrate (10 mM). E. coli inoculum was prepared by suspending colonies grown overnight on LB agar using 1×PBS to achieve a turbidity of 0.5 McFarland (1 × 10⁸ CFU/mL) and the final concentration of the inoculum in each well was around 5 × 10⁵ CFU/mL. The plates were incubated at 37°C for 18 h under aerobic or anaerobic conditions. MIC was defined as the lowest drug concentration that exhibited complete inhibition of microbial growth. All MICs were determined from at least three independent biological replicates.

[³H]-Gm (20 μCi/mg; Hartmann Analytic Corp.) was added at the indicated final concentration and cultures were incubated at 37°C on a rotary shaker. At given times, 500 μL aliquots were removed and collected on a 0.45 μM-pore-size HAWP membrane filter (Millipore) pretreated with 1 mL of unlabeled Gm (250 μg/mL). Filters were subsequently washed with 10 mL of 3% NaCl, placed into counting vials, and dried for 30 min at 52°C, whereafter 8 mL of scintillation liquid were added and incubated overnight at room temperature. Vials were counted for 5 min. Gm uptake efficiency is expressed as total accumulation of Gm (ng) per 10⁸ cells.

The two strains tested were first grown separately overnight in M9 medium supplemented with casamino acids (0.1%). The cell density of each suspension was measured by OD_600nm reading and by CFU count. Each overnight culture containing approximately 3 × 10⁸ cells/mL was diluted 1/100-fold and mixed in a ratio of 1:1 to inoculate 25 mL of M9 supplemented with casamino acids (0.1%) (time 0 h) and incubated for 24 h at 37°C for a competitive growth. The coculture was diluted 1/100 in 25 mL of fresh M9 medium and grown for another 24 h at 37°C. The initial density of each strain was determined in the initial coculture (0 h) from CFU data by diluting and plating population samples onto LB agar and LB agar supplemented with appropriate antibiotic. Similarly, the final density of each strain was determined.

Cells grown in LB (100 mL) to an OD_600nm of 0.6 were harvested by centrifugation, washed once in 50 mM phosphate buffer pH 7.5, and suspended in 50 mM phosphate buffer pH 7.5 (6 mL), lysed using a French press, aliquoted, and frozen immediately in liquid nitrogen. Nuo activity was assayed at 30°C by adding thawed samples to 50 mM phosphate buffer pH 7.5 containing reduced nicotinamide hypoxanthine dinucleotide (deamino-NADH) (250 mM) as specific substrate, and by following A_340nm. Protein concentration was determined by measuring the absorbance at A_280nm using a NanoDrop 2000 spectrophotometer.