Various Gram-negative (E. coli, P. aeruginosa, Acinetobacter baumannii, Klebsiella Pneumoniae, Shigella flexneri, Salmonella typhimurium, and Aeromonas hydrophila) and Gram-positive (S. aureus, Bacillus subtilis, Bacillus thuringiensis, and Staphylococcus epidermidis) bacterial strains were used in this study and their characteristics are described in Supplementary Table S1. For normal cell culturing, three mediums were used: M9 medium plus 5 g/L glucose, Luria-Bertani (LB) medium, or Mueller-Hinton broth. M9 medium with and without 2 g/L fumarate were used for nutrient shift- and starvation-induced E. coli persister formation, respectively. Yeast nitrogen broth medium was used for starvation-induced persister formation in S. aureus. Antibiotics used in this study include tobramycin, streptomycin, gentamicin, kanamycin, ampicillin, ofloxacin, with their manufacturers and final concentrations for different treatments being described in Supplementary Table S2. Carbonyl cyanide m-chlorophenyl hydrazone (CCCP) and its analog FCCP (carbonyl cyanide-p-trifluoromethoxyphenylhydrazone) were purchased from Sigma-Aldrich. All other chemical reagents are of analytical purity.

Nutrient shift-induced persisters were prepared as previously reported (Radzikowski et al., 2016). In brief, over-night cultures of each bacterial strain were diluted at 1:100 into M9 medium plus 5 g/L glucose or LB medium (37 °C, 220 rpm) and cultured to mid-exponential phase at a cell density of OD₆₀₀ = 0.5–0.6. Cells were centrifuged and washed with M9 medium twice before transferred to M9 medium plus 2 g/L fumarate and agitated for 4 h before antibiotic tolerance test. Starvation-induced persisters were prepared as previously reported (Eng et al., 1991). Briefly, E. coli and S. aureus cells were diluted at 1:500 in Mueller-Hinton broth medium and cultured for 24 h (35 °C, 220 rpm) to a cell density of around 10⁹ CFU/mL. Cells were centrifuged, re-suspended in M9 medium (for E. coli) and in yeast nitrogen broth medium without amino acids (for S. aureus) by dilution to a cell density of around 10⁸ CFU/mL and agitated for 5 h. Antibiotic tolerance test was performed by adding each antibiotic at concentrations as described in Supplementary Table S2 and further agitated the cells for 2 or 3 h. Antibiotic-treated cells were washed twice using phosphate-buffered saline (PBS; 0.27 g/L KH₂PO₄, 1.42 g/L Na₂HPO₄, 8 g/L NaCl, 0.2 g/L KCl, pH 7.4) by centrifugation (13000 g, 30 s), and then 5 μL of tenfold serially diluted cell suspension were spot plated onto LB agar dishes for survival assay. The antibiotic sensitivity of each bacterium was evaluated by incubating the exponential-phase cell culture with ampicillin (100 μg/mL), tobramycin (50 μg/mL) or ofloxacin (5 μg/mL) for 2 h before bacterial survival assay.

Nutrient shift- or starvation-induced persister cells were prepared as described above and hypoionic shock was performed as we previously reported (Jiafeng et al., 2015). Briefly, 100 μL cell cultures were centrifuged (13000 rpm, 1 min) in Eppendorf tube, with the supernatant being fully removed. The cells were then subjected to hypoionic shock treatment by re-suspending the pellet with pure water (i.e., without the presence of ions; a negative control was set using 0.9% NaCl solution) containing aminoglycoside antibiotic at concentrations as described in Supplementary Table S2. Cell suspension was kept at 25°C for 3 min before washing twice with PBS before subsequent cell survival assay as described above. The effect of proton motive force (PMF) and electron transport was examined by agitating the cell culture in the presence of 20 μM protonophore CCCP or FCCP, 2,4-Dinitrophenol (DNP; 20 μg/mL), rotenone (5 μg/mL) or NaN₃ (200 μg/mL) for 1 h before hypoionic shock treatment.

Tobramycin (gentamicin, kanamycin or streptomycin) extraction coupled with cell growth inhibition was explored for antibiotic uptake assay as follows. Briefly, 1 ml persister cells, after hypoionic shock treatment in the presence of each antibiotic at concentrations as described in Supplementary Table S2, were washed twice with PBS and re-suspended in 100 μL cell wall-digestion buffer (30 mM Tris–HCl, pH 8.0, 1 mM EDTA, 1 mg/mL lysozyme) for further incubation at room temperature for 2 h. Cells were subjected to three cycles of freezing treatment at −80°C, thermally denatured at 90°C for 10 min (Note: the bactericidal activity of each aminoglycoside after heating at 90^oC for 15 min was verified to be almost fully retained; refer to Supplementary Figures S5A, S6A) and centrifuged for removing cell debris and denatured proteins. Afterward, 5 μL supernatant was spotted on E. coli-seeded LB agar dish for further incubation at 37°C for 8–10 h and the diameter of cell growth inhibition zone was measured. In addition, tobramycin or gentamicin uptake by CCCP or FCCP pre-treated persister cells was measured similarly. A standard curve was prepared by directly adding each aminoglycoside at different concentrations (0, 15, 25, 50, 75, and 100 μg/mL) into persister cell suspension with the cell-wall digestion buffer. The tobramycin uptake by S. aureus cells was measured using the same procedure except of applying a different cell wall-digestion buffer (30 mM Tris–HCl, pH 8.0) plus 20 μg/mL lysostaphin [purchased from Sangon Biotech (Shanghai) Co., Ltd.; Cat no.: A609001].

A luciferase-based kit (Beyotime Biotechnology, Shanghai, China; S0026B) was used to measure ATP level according to the manufacturer’s instruction. Briefly, E. coli persister cells, with or without pretreatment of 20 μM CCCP for 1 h, was lysed using the lysis buffer and centrifuged (12000 g, 4°C, 5 min). The supernatant was quickly mixed with the working solution at equal volumes and then transferred into a 96-well plate before light recording on a FLUOstar Omega Microplate Reader using the Luminometer method.

A flow cytometry-based assay was applied to measure the PMF by using the fluorescence probe 3,3′-Diethyloxacarbocyanine Iodide [DiOC2(3); purchased from MaoKang Biotechnology, Inc., Shanghai, China] according to the manufacturer’s instruction. Briefly, E. coli persisters, with or without CCCP pretreatment as described above, were diluted into PBS to a cell density of 10⁶ cells/mL and incubated with DiOC2(3) (at a final concentration of 30 μM) at room temperature for 30 min. Cells were subjected to flow cytometric analysis on FACSymphony^TMA5 (BD Biosciences) with an excitation at 488 nm and emission at both red and green channels.

A carbon source shift from glucose to fumarate was recently reported to induce the formation of E. coli persister cells (Kim et al., 2016; Radzikowski et al., 2016). Here, we examined whether such a nutrient shift from glucose to fumarate could induce persister formation in other bacterial strains, including both gram-negative and gram-positive pathogens, and, if so, whether an aminoglycoside coupled with hypoionic shock could kill those persisters.

Of the seven gram-negative bacterial strains (refer to Supplementary Table S1), we found that the tolerance of S. typhimurium, S. flexneri, and E. coli to typical bactericidal antibiotics (ampicillin [Amp], ofloxacin [Ofl], and Tom) were significantly increased by different degrees after antibiotic-sensitive exponential-phase cells grown in LB medium were transferred to fumarate-containing M9 medium 4 h prior to antibiotic treatment (Figure 1A). Meanwhile, cell densities were largely held constant before and after the nutrient shift (as indicated in the “untreated” column in Figure 1A), i.e., exponential-phase cells in a growing state were switched to a non/slowly growing state, which is a prerequisite for persister formation. A. baumanii Ab6 and K. pneumonia KP-D367 were not tested because of their antibiotic resistance (Supplementary Figures S1A,B). Both A. hydrophila and P. aeruginosa PAO1, despite tolerance to antibiotics after culture in fumarate-containing M9 medium for 4 h (Supplementary Figures S1C,D), were able to grow substantially (refer to the colony density in the red frames of figures). These strains were not used in further studies because of this growth.

Of the four gram-positive bacterial strains (refer to Supplementary Table S1), we found that the antibiotic tolerance of S. aureus, S. epidermidis, B. thuringiensis, and B. Subtilis to Amp, Ofl, and Tom were all increased after the nutrient shift to fumarate (Figure 1B; for S. aureus, also refer to Figure 1C). Meanwhile, their cell densities were largely held constant (as indicated in the “untreated” column in Figure 1B).

Then, prompted by our earlier observations on stationary-phase E. coli persisters (Jiafeng et al., 2015), we examined whether hypoionic shock could facilitate eradication of nutrient shift-induced persisters by aminoglycoside antibiotics. For this purpose, fumarate-induced E. coli and S. aureus persisters (representing gram-negative and gram-positive bacteria, respectively) and four aminoglycoside antibiotics (Tom, gentamicin [Genta], kanamycin [Kana], and streptomycin [Strep]) were tested.

Cell survival assay revealed that E. coli persisters could be killed by Tom- or Genta-containing pure water after the cells were resuspended in the solution and incubated for only 3 min (Figure 2A). By contrast, Tom and Genta had little effect if they were dissolved in 0.9% NaCl solution (right panel, Figure 2A), which is consistent with our early observation that the presence of ions abolished aminoglycoside potentiation (Jiafeng et al., 2015). Notably, such nutrient shift-induced E. coli persisters were not killed by Kana and Strep upon hypoionic shock (Figure 2A), whereas cells before the nutrient shift were killed (Supplementary Figure S2A). Time-dependent analysis revealed that the minimal time for hypoionic shock-enabled eradication of E. coli persisters was approximately 2 min (Supplementary Figure S2B). In contrast, fumarate-induced S. aureus persisters were not killed by combined treatment with aminoglycoside antibiotics and hypoionic shock (upper panel, Supplementary Figure S2C), although the same treatment enabled Tom to kill normally growing exponential-phase S. aureus cells [lower panel, Supplementary Figure S2C, and as reported earlier (Jiafeng et al., 2015)].

We sought to examine whether CCCP, an uncoupler of the proton gradient, could suppress hypoionic shock-potentiated aminoglycoside killing of persisters, given that the bacterial uptake of aminoglycosides requires a PMF across cytoplasmic membranes of bacteria [reviewed in Taber et al. (1987)]. For this purpose, we pretreated fumarate-induced E. coli persisters with CCCP for 1 h and then subjected the cells to combined treatment with Tom and hypoionic shock. Cell survival assay revealed that CCCP, as well as its functional analogs FCCP and DNP, efficiently suppressed hypoionic shock-induced Tom potentiation that could kill E. coli persisters (Figure 2B). We confirmed that such CCCP pretreatment decreased intracellular ATP levels (Figure 2C) and also the PMF (Figure 2D) in E. coli persisters, as monitored by luciferase assay and membrane potential probe-based flow cytometric analysis, respectively. In line with the results from CCCP pretreatment, rotenone and NaN₃, two electron transport inhibitors that inhibit the transfer of electrons from iron-sulfur centers in complex I to ubiquinone and cytochrome c oxidase, respectively, were found to significantly suppress hypoionic shock-induced Tom potentiation (Figure 2B). Similarly, we found that all of these uncouplers or inhibitors significantly suppressed hypoionic shock-induced Genta potentiation that could kill nutrient shift-induced E. coli persisters (Supplementary Figure S2D).

We next examined whether hypoionic shock could facilitate aminoglycoside antibiotic killing of other persisters. Experimentally, we adopted the starvation-induced persister model described in an earlier report (Eng et al., 1991), in which stationary-phase E. coli and S. aureus cells were centrifuged and resuspended by 10-fold dilution in new medium without any nutrients, thus ruling out effects of the old medium and high cell density on antibiotic killing (Gutierrez et al., 2017).

First, we evaluated the antibiotic tolerance of starvation-adapted E. coli and S. aureus stationary-phase cells by agitating them in the presence of each aminoglycoside antibiotic for 3 h. Cell survival assay revealed that starvation adaptation caused E. coli to be highly tolerant to Kana and Strep and moderately tolerant to Tom and Genta (Supplementary Figure S3A) and S. aureus to be highly tolerant to Genta/Kana/Strep and moderately tolerant to Tom (Supplementary Figure S3B), results that are in line with those presented in an earlier report (Eng et al., 1991).

Next, we determined the efficacy of each aminoglycoside antibiotic coupled with hypoionic shock in killing starvation-induced persister cells. Cell survival assay revealed that starvation-induced E. coli persisters were killed by each aminoglycoside antibiotic upon hypoionic shock (Figure 3A), with this efficacy lost in the presence of 0.9% NaCl. Time-dependent analysis revealed that the cell survival ratio was constant with combined treatment from 1 min to 5 min (Supplementary Figure S3C), indicating that the effect of hypoionic shock on aminoglycoside antibiotics occurs as early as 1 min. Again, starvation-induced S. aureus persisters showed little killing after the combined treatment (Figure 3B and upper panel, Supplementary Figure S3D). However, stationary-phase S. aureus cells before starvation adaptation were killed by each aminoglycoside antibiotic upon hypoionic shock (lower panel, Supplementary Figure S3D).

We further examined the effect of proton gradient uncouplers (CCCP, FCCP, and DNP) and electron transport inhibitors (rotenone and NaN₃) on hypoionic shock-induced aminoglycoside potentiation that could kill starvation-induced E. coli persisters. We found that both CCCP and FCCP abolished Tom potentiation by hypoionic shock and that rotenone and NaN₃ exhibited a smaller but still significant suppressive effect (Figure 3C). We confirmed that CCCP pretreatment reduced intracellular ATP levels in E. coli persisters (Figure 3D) but had no significant effect on the PMF (Supplementary Figure S4), presumably because the basal PMF in starvation-induced E. coli persisters is quite low, as reported previously (Allison et al., 2011). Intriguingly, DNP showed a weak suppressive effect in this assay. Similarly, we found that all of these uncouplers or inhibitors significantly suppressed hypoionic shock-induced Genta potentiation for starvation-induced E. coli persister cell killing (Supplementary Figure S3E).

In view of the fact that pretreatment with CCCP or FCCP can abolish potentiation (Figures 2B, 3C) and that aminoglycoside uptake is dependent on a PMF [reviewed in Taber et al. (1987)], we hypothesized that hypoionic shock-induced aminoglycoside potentiation is accomplished by enhancing bacterial uptake of antibiotics. Taking advantage of the high thermal stability of Tom (Supplementary Figure S5A) and the irreversible nature of aminoglycoside uptake by E. coli cells (Nichols and Young, 1985), we explored a protocol to measure the bacterial uptake of Tom. To this end, Tom taken up by E. coli cells was extracted by cell wall digestion coupled with cycled freezing/thawing and thermal denaturation, and then the relative level of Tom in the lysate was measured by bacterial cell growth inhibition assay (for details, refer to the section “Materials and Methods”).

Cell growth inhibition assay revealed that Tom extracted from nutrient shift-induced E. coli persisters upon hypoionic shock significantly suppressed bacterial cell growth on LB agar plates (red frame, Figure 4A). In contrast, no significant cell growth inhibition was observed if Tom was extracted from the persister cells not in hypoionic shock or pretreated with CCCP or FCCP (Figure 4A). A regression analysis (Supplementary Figure S5B) based on standards (upper panel, Figure 4A) showed that the concentration of Tom extracted from nutrient shift-induced E. coli persister cells in hypoionic shock was up to 57 ± 4 μg/mL, whereas that extracted from the cells treated in NaCl-containing solution or pre-treated with CCCP was less than 15 μg/mL. Similarly, whereas hypoionic shock dramatically enhanced the uptake of Tom by starvation-induced E. coli persister cells (red frame, Figure 4B), this enhancement was diminished by the presence of 0.9% NaCl, CCCP, or FCCP pretreatment.

We also examined the bacterial uptake of Genta, Strep, and Kana during hypoionic shock. First, we verified the high thermal stability of these aminoglycosides (Supplementary Figure S6A). Then, we extracted each and measured their inhibitory effects on cell growth. Data presented in Supplementary Figure S6B revealed that Genta extracted from nutrient shift-induced E. coli persisters in hypoionic shock dramatically suppressed bacterial cell growth on LB agar plates (as indicated by the red frame). In contrast, the extracted Kana only slightly inhibited cell growth (Supplementary Figure S6C), and the extracted Strep hardly showed any inhibitory effect (Supplementary Figure S6D). These results agreed with the weak bactericidal actions of these antibiotics in persister cells (Figure 2A and Supplementary Figure S2A, respectively). Similarly, Genta extracted from starvation-induced E. coli persisters significantly suppressed cell growth (Supplementary Figure S7A), whereas extracted Kana and Strep exhibited hardly any inhibitory effects (Supplementary Figures S7B,C). These results were in accordance with the strong bactericidal action of Genta and relatively weak killing action of Kana and Strep (Figure 3A).

An intriguing observation in our study is that neither nutrient shift- nor starvation-induced S. aureus persisters were sensitive to hypoionic shock-induced Tom potentiation, whereas exponential- and stationary-phase S. aureus cells were sensitive (Supplementary Figures S2C, S3D). To clarify the reason for this, we measured Tom uptake by these different S. aureus cells. We observed that Tom extracted from exponential-phase S. aureus cells (i.e., cells obtained before the nutrient shift) dramatically inhibited cell growth on LB agar plates (red frame, left panel, Figure 4C), whereas Tom extracted from nutrient shift-induced S. aureus persister cells showed little inhibitory effect (right panel, Figure 4C). Similarly, we observed significant cell growth inhibition by Tom extracted from stationary-phase S. aureus cells (i.e., cells before starvation induction) (red frame, Supplementary Figure S8A), but not by Tom from starvation-induced S. aureus persister cells (Supplementary Figure S8B). These results suggest that the insensitivity of both types of S. aureus persisters to hypoionic shock-induced Tom potentiation is most likely due to the failure of hypoionic shock to enhance bacterial uptake of Tom.

We recently reported that hypoionic shock enabled aminoglycosides to kill stationary-phase E. coli persisters (Jiafeng et al., 2015). Here, we found that aminoglycoside antibiotics exhibited different actions against E. coli and S. aureus cells with different growth statuses upon hypoionic shock. These actions can be summarized as follows. (1) Kana and Strep eradicated exponential-phase E. coli cells but not nutrient-shifted E. coli cells (Supplementary Figure S2A), whereas Tom and Genta killed both (Figure 2A); (2) Tom killed exponential-phase S. aureus cells but not nutrient-shifted S. aureus cells, whereas the other three aminoglycoside antibiotics had little effect on either of these cell types (Supplementary Figure S2C); and (3) each aminoglycoside antibiotic eradicated stationary-phase S. aureus cells but not starvation-induced S. aureus cells (Supplementary Figure S3D). Notably, these distinct bactericidal actions of aminoglycoside antibiotics induced by hypoionic shock (Figures 2A, 3A,B and Supplementary Figures S2A,C, S3D) agreed well with the amount of aminoglycoside taken up by persister cells (Figure 4 and Supplementary Figures S6–S8), which strongly suggests that hypoionic shock-induced aminoglycoside potentiation is achieved by enhancing aminoglycoside uptake.

Hypoionic shock-induced aminoglycoside potentiation that can kill E. coli persisters appears to depend on the cellular respiration of bacteria based on the following evidence. First, it is well-known that aminoglycoside uptake depends on a PMF, which is generated through respiration (Taber et al., 1987). Second, recent studies have suggested that the downstream lethal action of aminoglycosides depends on the respiration of bacterial cells (Lobritz et al., 2015; Meylan et al., 2017). In our study, CCCP or FCCP alone was able to abolish hypoionic shock-induced aminoglycoside potentiation (Figures 2B, 3C and Supplementary Figures S2D, S3E) and uptake (Figure 4 and Supplementary Figures S6B, S7A). In addition, sodium azide and rotenone (inhibitors of the electron transport chain) significantly suppressed such potentiation (Figures 2B, 3C and Supplementary Figures S2D, S3E). These observations indicate that hypoionic shock, although lasting for only a couple of minutes, dramatically enhances bacterial uptake of aminoglycosides in a respiration-dependent manner.

Based on these observations, we hypothesize that certain channels on the cytoplasmic membrane of bacterial cells may be responsible for hypoionic shock-induced aminoglycoside potentiation. These channels can be activated for aminoglycoside uptake in response to hypoionic shock and may exhibit selectivity in transporting structurally different aminoglycosides, as demonstrated by the potentiation of some aminoglycosides and not others in this study. In addition, the protein level and/or transportation activity of these channels could be tightly regulated by growth conditions; therefore, they are functionally dependent on cellular respiration and physiological status. As such, an aminoglycoside (e.g., Tom) coupled with hypoionic shock can kill exponential-phase S. aureus cells but not these same cells after a nutrient shift to fumarate (Supplementary Figure S2C). In addition, it should be pointed out that ribosome is still the acting target of aminoglycoside during such hypoionic shock as revealed in our earlier study using the streptomycin-resistant E. coli strain MC4100 (Jiafeng et al., 2015).

Considering that the clinical application of aminoglycosides has dropped substantially in recent decades due to their toxicity and the rise of antibiotic resistance (Mingeot-Leclercq and Tulkens, 1999; Mingeot-Leclercq et al., 1999), improving the efficacy of aminoglycosides by hypoionic shock while limiting their side effects would be a clinically valuable approach. An antibiotic potentiation strategy would entail exposing subjects to aminoglycosides for only 1 or 2 min, therefore reducing the toxicity associated with aminoglycosides use. Nevertheless, this approach cannot be directly applied to curing infections in animals and humans, largely because of the ubiquity of ions throughout the animal body (e.g., Na⁺, K⁺, Cl^–, and charged amino acids) that could abolish the potentiation effect (Jiafeng et al., 2015). If the biochemical mechanism underlying hypoionic shock-induced aminoglycoside potentiation can be discovered (e.g., if the membrane channels for aminoglycoside uptake during hypoionic shock can be identified and fully characterized), however, this would be helpful for developing new anti-persister strategies that are based on the mechanism rather than on hypoionic shock. Studies to identify this mechanism are currently underway in our laboratory.

Metabolite-stimulated aminoglycoside potentiation has recently been shown to kill stationary-phase E. coli, P. aeruginosa, and Edwardsiella tarda persister cells (Allison et al., 2011; Barraud et al., 2013; Peng et al., 2015; Meylan et al., 2017; Su et al., 2018). This approach has even been validated for eradication of persisters in animal models (Allison et al., 2011; Peng et al., 2015). Apparently, metabolic stimulation dramatically changes the physiological states of persister cells, boosting their respiration and reprogramming their metabolic network. It follows that persister cells might regrow during a lengthy period of metabolic stimulation (usually a couple of hours), and, if this occurs, cell regrowth would reduce the benefits of aminoglycoside potentiation to kill persisters. In comparison, our method of hypoionic shock-induced aminoglycoside potentiation requires only 1 or 2 min of stimulation [Supplementary Figures S2B, S3C; or refer to our earlier report (Jiafeng et al., 2015)]. Another advantage is that hypoionic shock enables aminoglycosides to eradicate normally growing bacterial cells [Supplementary Figures S2A,C; or refer to our earlier report (Jiafeng et al., 2015)].

Nutrient shift-stimulated bacterial persistence has been widely reported for E. coli cells that are grown in a batch culture containing two carbon sources and exhibiting diauxic growth phases (Amato et al., 2013; Amato and Brynildsen, 2014, 2015; Kotte et al., 2014). Among the carbon sources, fumarate is highly potent in increasing the formation of persisters in exponential-phase E. coli cells (Amato et al., 2013; Amato and Brynildsen, 2014, 2015; Kim et al., 2016; Radzikowski et al., 2016). Nevertheless, fumarate was found to impair persister formation in P. aeruginosa stationary-phase cells exposed to Tom by activating cellular respiration and generating a PMF through stimulation of the tricarboxylic acid (TCA) cycle (Meylan et al., 2017). These actions might be linked to the role of fumarate as a metabolite of the TCA cycle and/or as an electron acceptor (Unden et al., 2014, 2016).

Here, we have shown that a nutrient shift to fumarate is able to increase persister formation in exponential-phase gram-negative (as represented by E. coli, S. typhimurium, and S. flexneri) and gram-negative (as represented by S. aureus, S. epidermidis, B. thuringiensis, and B. subtilis) bacterial cells. According to the recent definition by Balaban et al. (2019), these fumarate-induced persisters should be considered type I/triggered persisters. On the other hand, the concentration of intracellular fumarate was shown to be proportional to the frequency of persisters among exponential-phase E. coli cells (Kim et al., 2016), illustrating its critical role in the formation of type II/spontaneous persisters [according to the definition in Balaban et al. (2019)]. Based on our findings, we propose that intracellular fumarate may converge on both external and intrinsic signals in bacterial cells and together, these signals determine persister formation, conceivably by finely tuning the electron transport chain and/or TCA cycle (Unden et al., 2014, 2016).