The Multifaceted Antibacterial Mechanisms of the Pioneering Peptide Antibiotics Tyrocidine and Gramicidin S

Cell wall integrity.To assess whether the variable aromatic dipeptide unit of the tyrocidine peptides makes a difference for their in vivo mechanism, we used both tyrocidine A (Phe³-d-Phe⁴) and tyrocidine C (Trp³-d-Trp⁴) (Fig. 1A and B). The tyrocidines have been proposed to bind to fungal cell wall components (27), and gramicidin S was shown to affect cell wall integrity in Bacillus subtilis (37). To study how effectively these antibiotics weaken the bacterial cell wall and cause lysis, we followed the optical density (OD) of B. subtilis cells treated with MICs of the compounds (Fig. 1D). At this concentration, a fraction of the tyrocidine-treated cells lysed after 15 to 20 min of treatment but the culture recovered after approximately 60 min. Gramicidin S treatment did not lead to cell lysis but completely halted further growth. However, at 2× MIC, all three peptides caused lysis and no regrowth could be observed (see Fig. S1 in the supplemental material). Such cell lysis suggests that these antibiotics somehow affect the integrity of the cell wall. Since we wanted to analyze the immediate growth-inhibiting effects of the peptides, instead of pleiotropic lysis effects, we performed further experiments after 10 min of treatment with 1× MIC (unless otherwise noted), when no strong reduction of the optical density was observed (Fig. 1D).

To further examine the effects on cell wall integrity, we employed an organic fixation method that is indicative of holes in the peptidoglycan layer, typically related to impaired synthesis of the cell wall precursor lipid II (45). In line with an earlier study (39), gramicidin S clearly affected cell wall integrity as seen from the protoplast protruding through cell wall breaches (Fig. 2A). However, this phenotype was not observed for tyrocidines A and C, suggesting a clear mechanistic difference from gramicidin S. In fact, transmission electron microscopy revealed that the tyrocidines caused severe cellular damage, intracellular content leakage, and lysis of cells, whereas gramicidin S-treated cells showed only subtle cell shape alterations but no major signs of lysis or physical cell damage (Fig. 2B).

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FIG 2

Effects of tyrocidines and gramicidin S on the cell wall. (A) Impact on cell wall integrity. B. subtilis 168 was treated with peptides for 10 min and subsequently fixed in a 1:3 mixture of acetic acid and methanol. This fixation method leads to extraction of the protoplast through holes in the peptidoglycan layer (arrow) when synthesis of the cell wall precursor lipid II is impaired (37, 45). Since this fixation method can sometimes lead to quick lysis of heavily damaged cells, we examined both 0.5× MIC (nonlytic) and 1× MIC of the tyrocidines but did not find membrane extrusions under both conditions. Bar, 2 µm. (B) Transmission electron microscopy pictures of B. subtilis 168 treated with 1× MIC of the peptides for 10 min. Bar, 0.5 µm.

Membrane permeability.The current belief is that both the tyrocidines and gramicidin S form pores in the cell membrane (26, 31, 33–37). However, for the tyrocidines this has actually never been shown in bacteria, and the pore-forming ability of gramicidin S is heavily disputed (31, 33–37). Therefore, we examined the effects of these peptides on membrane permeability in detail. First, we determined the effects of the peptides on the membrane potential using the membrane potentiometric dye DiSC(3)5. Tyrocidines A and C led to immediate strong depolarization of B. subtilis cells already at 0.5× MIC (Fig. 3A). Inhibitory concentrations of gramicidin S (1× MIC) only gradually and partially depolarized B. subtilis cells. Full depolarization was achieved only at 2× MIC, a concentration causing cell lysis (Fig. S1). These results suggest that the tyrocidines form membrane pores while gramicidin S does not.

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FIG 3

Tyrocidines but not gramicidin S form membrane pores in vivo. (A) Membrane potential measurements of exponentially growing B. subtilis 168 cells using the membrane potential-sensitive fluorescent probe DiSC(3)5. Tyrocidines A and C were applied at 0.5× and 1× MIC, and gramicidin S was applied at 1× and 2× MIC. The red arrow indicates the time point of antibiotic addition. (B) Membrane permeability for propidium iodide. Exponentially growing cells were treated with peptides for 5 min. For the tyrocidines, 0.5× MIC (nonlytic) and 1× MIC were used; gramicidin S was applied at 2× MIC (2 µg/ml, lytic concentration). SDS at 0.01% served as a positive control. Strain, B. subtilis 168. au, arbitrary units.

To further characterize the ability of the peptides to generate distinct ion-conducting pores, we performed in vitro conductivity measurements. This allowed us to follow single-ion current events in a planar model lipid system mimicking Gram-positive membranes (3:1 mixture of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoglycerol [POPG] and 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine [POPE]). In line with our in vivo data, we observed a number of conductivity events for tyrocidines A and C at low, nanomolar concentrations (Fig. 4A). Current transitions peaked at 2 pA for tyrocidine A and 3.5 pA for tyrocidine C, corresponding to calculated conductance values of 20 pS and 35 pS, respectively (Fig. 4B). These conductance events are representative of several single-membrane pores that stay open for up to several seconds. Conductance events induced by gramicidin S were observed only at concentrations leading to membrane collapse (5 µM, Fig. 4A), lasted only milliseconds, and exhibited a broad range of amplitudes (Fig. 4A), which is indicative of general bilayer distortion. Similar behavior has been reported for gramicidin S in diphytanoyl phospholipid bilayers (31). These results show that both tyrocidine A and tyrocidine C induce discrete long-lived ion-conducting pores, while gramicidin S did not exhibit pore-forming capacity in our system.

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FIG 4

Conductivity measurements in model membranes. (A) Conductivity measurements in POPG-POPE (3:1) planar model membranes with an artificial transmembrane potential of 100 mV. Representative recordings of single-channel currents of tyrocidine A and tyrocidine C are shown. No ion conductance events were observed for gramicidin S up to concentrations leading to membrane collapse (5 µM). Note the different peptide concentrations and the different y scales. (B) Single-channel current transition amplitude histograms of tyrocidine A (gray) and tyrocidine C (red) in POPG-POPE (3:1) membranes recorded at 100 mV. Normalized frequency refers to the number of times that a pore with a specific current amplitude was observed and is displayed as a fraction of the maximum count. Data from 8 independent membranes and 158 single-channel transitions for tyrocidine A and 12 independent membranes with 560 single-channel transitions for tyrocidine C are shown.

To gain insight into whether tyrocidine pores are small ion channels or large membrane pores that allow big organic molecules to pass through, pore formation was assessed by following the influx of the fluorescent dye propidium iodide, which enters bacterial cells only through membrane lesions large enough for the bulky molecule to pass (46). Both tyrocidine A and tyrocidine C rapidly formed large pores in B. subtilis cells (Fig. 3B). Cells treated with gramicidin S did not show any propidium iodide influx, not even at bacteriolytic concentrations (2× MIC) (Fig. 3B; Fig. S1).

Taken together, both in vivo (Fig. 1 to 3) and in vitro (Fig. 4) experiments strongly indicated that at inhibitory concentrations tyrocidines A and C, but not gramicidin S, form large, ion-conducting pores. The slow and incomplete depolarization by gramicidin S (Fig. 3A) suggests rather a limited increase of passive membrane permeability.

Membrane protein localization.At MICs, gramicidin S did not fully depolarize the cell membrane (Fig. 3A) and even at supra-MICs, it did not lead to large membrane lesions (Fig. 3B). It is therefore plausible that other vital processes must be disturbed that, together with limited depolarization and impairment of the cell wall (Fig. 2A and 3A), will eventually lead to cell death. It is also well known that the membrane potential is required for the localization of peripheral membrane proteins involved in cell division and cell morphology (47), and pore formation by the tyrocidines is likely to affect different essential cellular processes. To investigate whether insertion of gramicidin S and the tyrocidines affects membrane proteins differently, we examined the localization of fluorescent membrane protein fusions involved in a variety of cellular processes. Figure 5A shows peripheral membrane proteins that can be used as reporters for dissipation of the membrane potential. The cell division proteins MinD and DivIVA require the membrane potential for their localization at the membrane (44, 47), while localization of the phospholipid synthase PlsX is independent of the proton motive force (44). In line with membrane depolarization, all three peptides delocalized MinD and DivIVA, whereby the tyrocidines caused a more diffuse cytoplasmic fluorescence signal, while gramicidin S induced stronger clustering of the proteins at the cell periphery (Fig. 5A). Interestingly, the phospholipid synthase PlsX completely lost its membrane localization after treatment with both the tyrocidines and gramicidin S. Since PlsX does not depend on the membrane potential, its delocalization demonstrates that the peptides have other effects on the membrane than only interfering with the barrier function of the cell membrane.

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FIG 5

Delocalization of membrane proteins labeled with GFP. (A) Reporter proteins for membrane depolarization. Membrane localization of MinD and DivIVA requires the membrane potential, whereas membrane binding of the peripheral membrane protein PlsX is independent of the membrane potential. (B) Localization of proteins associated with cell wall synthesis. In contrast to MreB, localization of the peripheral MurG and integral MraY proteins is not reliant on the membrane potential. (C) Localization of flotillins involved in the organization of lipid rafts. FloA is an integral membrane protein, and FloT is a peripheral membrane-associated protein. (D) Localization of proteins involved in membrane-bound energy generation. AtpA and SdhA are the peripheral subunits of the ATP synthase and succinate dehydrogenase, respectively. Arrowheads indicate regions from which AtpA and SdhA are fully excluded. B. subtilis strains were grown in LB with appropriate inducer concentrations (see Table S1) and treated with peptides in early exponential growth phase for 10 min. Bar, 2 µm.

The cell wall-damaging effect of gramicidin S has been attributed to detachment of the peripheral peptidoglycan synthesis enzyme MurG, involved in the last steps of lipid II synthesis (39). To test whether the localization of this protein and other cell wall synthesis proteins is affected by the tyrocidines, we examined the localization of MurG, the transmembrane lipid I synthase MraY, and the actin homologue MreB, which coordinates lateral peptidoglycan synthesis (48, 49). As shown in Fig. 5B, MurG completely lost its membrane localization after treatment with all three peptides. However, MraY was clustered into huge foci by tyrocidines A and C, while gramicidin S did not show a clear effect. MreB is known to delocalize after membrane depolarization (47, 50). After treatment with the tyrocidines and, to a lesser extent, also with gramicidin S, MreB localized into discrete clusters distributed all over the cell membrane (Fig. 5B), a peculiar effect not seen before with membrane-targeting antibiotics (44, 47, 50).

The actin homologue MreB has been implicated in the formation of fluid membrane microdomains, so-called regions of increased fluidity (RIFs), which play a role in spatial organization of cell envelope synthesis (44, 50, 51). This made us curious whether the peptides would also affect the distribution of bacterial flotillin proteins involved in the formation of specific rigid membrane domains (known as lipid rafts in eukaryotes) (52, 53). While the integral flotillin FloA was not affected by any of the peptides, the peripheral flotillin FloT was clearly detached from the membrane by tyrocidines A and C but not by gramicidin S (Fig. 5C). FloT interacts with many membrane-associated proteins (54), and the separation from its interaction partners by dissociation from the membrane is likely to affect multiple cellular processes, including membrane organization, fluidity, respiration, protein secretion, membrane transport, signal transduction, and autolysin activity (53, 56–58).

Since gramicidin S was shown to reduce ATP levels in B. subtilis (39) and tyrocidines reduced oxygen consumption in S. aureus (28), we also examined their effects on the localization of AtpA, the peripheral subunit of the ATP synthase, and SdhA, the flavoprotein subunit of the succinate dehydrogenase (complex II) of the respiratory chain. These transmembrane protein complexes are unaffected by membrane potential-dissipating drugs like carbonyl cyanide m-chlorophenylhydrazone (CCCP) and valinomycin, and also the membrane-targeting antibiotic daptomycin (44, 47). Surprisingly, both AtpA and SdhA partially lost their membrane association after treatment with the tyrocidines (Fig. 5D), explaining inhibition of electron transport chain enzymes reported in an early study (29, 59). In contrast, gramicidin S had no marked effect on the localization of these proteins, suggesting that its effect on cellular ATP levels is probably due to delocalization of cytochrome c (39) and partial membrane depolarization.

Gel-phase domains.Interestingly, many of the tyrocidine-treated cells displayed large areas from which AtpA and SdhA were excluded (Fig. 5D, arrowheads; Fig. S2, left panels; and Fig. S3A). It seems as if these empty areas might have been caused by plasmolysis of the cell membrane due to partial release of cytoplasmic content. However, the corresponding phase-contrast images did not indicate that these were partially empty cells (Fig. 5D and Fig. S2, right panels). To examine the effects of the peptides on the cell membrane in more detail, we employed superresolution structured illumination microscopy (SIM) and stained the cell membrane with MitoTracker green, a very bright fluorescent membrane stain that provides excellent SIM contrast (60). As shown in Fig. 6A, a 10-min incubation with the tyrocidines resulted in strongly fluorescent foci (open arrows) but also regions that were completely unstained (closed arrows). Gramicidin S-treated cells did not cause gaps in the fluorescent membrane stain but some small fluorescent membrane foci. When we expressed green fluorescent protein (GFP), it became apparent that many of the tyrocidine-treated cells lost their cytosolic GFP signal (Fig. 6B), indicating leakage of cellular content. Cells were counterstained with the red fluorescent membrane dye Nile red, which showed similar gaps in its fluorescent stain (Fig. 6B) as observed with MitoTracker green (Fig. 6A). Importantly, these gaps were also observed in cells that still contained their cytoplasmic GFP (Fig. 6B and Fig. S3B), indicating that these gaps are not caused by the absence of cell membrane due to lysis. The only remaining explanation for the absence of both fluorescent membrane dyes as well as membrane proteins is that these membrane areas are in a rigid, gel-phase state that excludes most dyes and proteins (61). Although stabilization of gel-phase domains by antimicrobial peptides has been observed in vitro (62, 63), to the best of our knowledge, such domains have not been documented before in bacterial cells.

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FIG 6

Tyrocidines and gramicidin S cause aberrant membrane staining and plasmolysis. (A) SIM images of B. subtilis 168 stained with MitoTracker green. Note the unstained membrane areas (arrowheads) and bright lipid accumulations (arrows) in cells treated with tyrocidines A and C. See also Fig. S5A for quantification of membrane patches. (B) Tyrocidines induce membrane invaginations. SIM images of B. subtilis bSS82 expressing cytosolic GFP from the strong ribosomal PrpsD promoter were stained with Nile red. Sucrose (20%) was used to induce osmotic upshock and used as a control for plasmolysis (bottom panel). (C) Tyrocidines cause plasmolysis. Time-lapse images of B. subtilis bSS82 cells treated with tyrocidines A and C. Membranes were stained with the nontoxic membrane dye FM5-95. Arrows indicate membrane patches and corresponding areas with reduced intracellular GFP. Bars, 2 µm. Experiments were performed in early exponential log phase using 1× MIC of the peptides.

Accumulation of membrane material.Cells stained with membrane dyes and treated with gramicidin S, and especially with tyrocidines A and C, showed membrane patches that were highly fluorescent (Fig. 6A and B and Fig. S3B and C). In the case of the tyrocidines, these membrane patches seemed to partially displace the cytoplasmic GFP signal (Fig. 6B, arrowheads), suggesting that the increased fluorescence signal is caused by an accumulation of membrane material (44, 50, 64). Formation of this extra membrane material can be caused by overactivation of fatty acid synthesis (65). However, when either fatty acid synthesis or protein synthesis was blocked with triclosan or chloramphenicol, respectively, bright membrane patches were still observed upon peptide treatment (Fig. S3D). Alternatively, these membrane accumulations can be caused by plasmolysis due to a loss of turgor pressure and cell shrinkage, as shown by plunging cells in a sucrose solution (Fig. 6B, bottom panel) (66). To confirm that tyrocidines A and C cause cell shrinkage, we performed a time-lapse experiment using cells stained with the membrane dye FM5-95. As shown in Fig. 6C, membrane patches are formed within minutes, rapidly followed by reduction of the GFP signal in their immediate proximity and cell shrinkage. Thus, tyrocidines induce large fluorescent membrane invaginations (patches) by causing a loss of turgor and subsequent cell shrinkage.

Impact on membrane fluidity.No signs of plasmolysis were observed with gramicidin S (Fig. 6B), in line with the fact that this antibiotic does not cause rapid cell lysis at inhibitory concentrations (Fig. 1D). However, gramicidin S also caused highly fluorescent membrane patches. Irregular membrane staining of fluorescent membrane dyes has been observed before when cells were treated with the membrane antibiotic daptomycin and the proton ionophore CCCP, and the irregularly stained areas were identified as highly fluid membrane domains, which increase the fluorescence of membrane dyes and can also accommodate more dye (50, 64). To examine whether these bright fluorescent membrane patches are fluid lipid domains, we used the membrane dye DiIC12, which displays a strong affinity for fluid membranes, including RIFs, due to its short hydrocarbon tail (50, 67, 68). Indeed, after treatment with gramicidin S the natural distribution of RIFs was severely disturbed, and the dye accumulated in one or two large patches per cell (Fig. 7A). The same patches were observed in cells treated with the tyrocidines (Fig. 7A), and they seem to correspond to the large membrane invaginations caused by plasmolysis (Fig. 6). Presumably, fluid membrane domains are more likely to invaginate due to the higher flexibility of fluid lipids.

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FIG 7

Effect of tyrocidines and gramicidin S on membrane fluidity. (A) Fluid lipid domain (RIF) staining with DiIC12. (B) Laurdan GP microscopy indicating local fluidity differences. Filled arrows indicate the cell membrane; open arrows indicate fluid membrane patches. (C) Quantification of membrane fluidity from laurdan microscopy pictures. The minimum GP within the membrane patch and the average GP in the lateral membrane were measured in individual cells. (D) Changes in overall membrane fluidity measured by laurdan GP in microtiter plate cultures. The membrane fluidizer benzyl alcohol served as a positive control. The arrow indicates the time point of antibiotic addition. Error bars represent standard deviations of the mean of three replicate experiments.

To confirm that these large DiIC12-stained membrane patches are indeed more fluid, we employed laurdan generalized polarization (GP). The fluorescence membrane dye laurdan shifts its fluorescence spectrum depending on the amount of water molecules in the bilayer, and laurdan GP can therefore be used as a measure of phospholipid head group spreading and fatty acid chain flexibility (69, 70). As shown in Fig. 7B, this assay confirmed that the membrane patches (open arrows) have an increased fluidity compared to the bulk membrane (filled arrows). When we quantified laurdan GP in the patches and the remaining membrane, it became apparent that not only did the peptides cause fluid membrane patches but this also led to a rigidification of the rest of the membrane (Fig. 7C).

To gain insight into how fast these fluidity changes happen, laurdan GP was measured spectroscopically in batch culture. We found that there was a rapid (2-min) reduction of membrane fluidity when the peptide antibiotics were added (Fig. 7D), indicating that this is a direct effect of peptide-membrane interaction and not a bacterial adaptation strategy, which would require the induction of specific enzymes and thus take more time. In line with our laurdan microscopy data, the overall membrane rigidification was stronger for tyrocidines A and C than for gramicidin S (Fig. 7D), supporting our notion that the tyrocidines change the physical parameters of the lipid bilayer in a different way than gramicidin S.

Protein delocalization is independent of MreB.Previous work suggested that RIFs are formed by MreB polymers, both in B. subtilis and in E. coli (50, 51). All three peptides caused clustering of MreB (Fig. 5B); therefore, we examined whether these MreB clusters are responsible for the large DiIC12-stained RIFs that emerge upon addition of the peptide antibiotics (Fig. 7A). Interestingly, the GFP-MreB clusters did not overlap DiIC12 clusters, both in the case of gramicidin S and in the case of the tyrocidines (Fig. 8A). Thus, the peptide antibiotics are able to form artificial RIFs without the help of MreB. To confirm this, we tested a triple deletion strain lacking MreB and its homologues MreBH and Mbl. Indeed, in such mutant cells the addition of gramicidin S and the tyrocidines still resulted in large DiIC12-stained membrane patches (Fig. S4).

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FIG 8

Overlap of GFP-tagged proteins with fluid membrane domains. (A to C) Colocalization of MreB (A), MraY (B), and AtpA (C) with DiIC12 was tested after a 10-min treatment with tyrocidine A, tyrocidine C, and gramicidin S. Open arrows indicate fluid membrane patches induced by the peptides. Closed arrows indicate membrane areas from which both DiIC12 and AtpA are excluded. (D) Effects of the unrelated antimicrobial peptides daptomycin and MP196. Cells were treated with 1 µg/ml daptomycin (in LB supplemented with 1.25 mM CaCl₂) or 10 µg/ml MP196. B. subtilis strains were grown in LB supplemented with appropriate inducer concentrations (Table S1) and treated with peptides in early exponential growth phase for 10 min. Bar, 2 µm.

To our surprise, untreated MreB triple mutant cells also showed some small RIFs (Fig. S4). This is in disagreement with what we have reported previously (50). An in-depth analysis revealed that DiIC12-stained RIFs become visible only during logarithmic growth and disappear when cells enter the stationary phase of growth, despite the fact that MreB is still present in the stationary phase (Fig. S4 to S6 and Text S1). This growth-phase-dependent occurrence of RIFs explained the differences from our previous finding. Based on this and previous studies, we propose that MreB is not involved in establishing RIFs but rather helps to organize them along the cell axis.

Peptides induce fluid lipid domains by stabilizing the liquid crystalline phase.The previous results suggested that both tyrocidines and gramicidin S induce fluid lipid domains independently from the MreB cytoskeleton. To confirm that the observed fluidity changes are a direct cause of peptide-lipid interaction, we tested whether the peptides influence phase separation behavior of model lipid membranes using ³¹P solid-state nuclear magnetic resonance (NMR). As before, we used 3:1 POPG-POPE model membranes to mimic Gram-positive bacterial cell membranes. To exclude the possibility that the observed effects correspond to head group demixing rather than transition from liquid crystalline to gel phase, we additionally tested pure POPE bilayers. All three peptides stabilized the liquid crystalline (fluid) phase in both model membrane systems (Fig. S7). For the tyrocidines, similar effects have been described before in model membranes made of dielaidoylphosphatidylethanolamine (DEPE) (26), and gramicidin S has been shown to have a preference for more-fluid membranes (32). The NMR data indicate that the generation of fluid lipid domains is a direct cause of peptide-lipid interaction and suggest a model in which the peptides preferentially insert into fluid lipid domains like RIFs and increase their local fluidity. The DiIC12 and laurdan data suggest that these domains also attract more flexible (fluidizing) lipid species, further increasing local fluidization.

Membrane proteins localize to fluid domains.The relatively bulky membrane-targeting domains of most peripheral membrane proteins need a certain flexibility of the lipid bilayer for their binding and insertion into the membrane (71). Similarly, the hydrophobic surface area of transmembrane proteins causes an inherent preference of these proteins for areas of a particular membrane thickness and fluidity (72). It is therefore likely that the segregation of lipids into fluid and rigid (membrane) domains by the tyrocidines and gramicidin S (Fig. 7B) is responsible for the observed delocalization of membrane proteins (Fig. 5). To investigate this, we examined the localization of MraY (transmembrane) and AtpA (peripheral subunit of a transmembrane complex) in relation to fluid lipid domains stained with DiIC12. MraY clearly overlapped DiIC12 clusters caused by the tyrocidines and gramicidin S, suggesting that this protein preferentially partitions from the rigidified membrane regions into fluid membrane patches (Fig. 8B). AtpA has been used as a reporter for cell membrane invagination (44, 50), and it does not accumulate in fluid lipid domains caused by antibiotics (50) (also MP196, Fig. 8B). In line with membrane invaginations due to plasmolysis caused by the tyrocidines but not by gramicidin S (Fig. 6), AtpA showed a substantial overlap with the tyrocidine-induced DiIC12 clusters but not with clusters induced by gramicidin S (Fig. 8C).

To compare the effects of the tyrocidines and gramicidin S with those of other membrane-targeting peptides, we performed the same costaining experiment with two well-studied antimicrobial peptides, the lipopeptide daptomycin, which selectively inserts into RIFs and tightly aggregates them into rigid clusters (44), and the small cationic peptide MP196, which disturbs membrane architecture and depolarizes the membrane by inhibiting the respiratory chain (39). In line with an earlier study (44), daptomycin only slightly affected the localization of MreB and had no effect on MraY and AtpA localization (Fig. 8B). On the other hand, MP196 caused MreB clusters that overlapped aggregated RIFs, which is typical for depolarizing compounds (50). MraY also accumulated in the DiIC12-stained membrane patches, whereas AtpA localization was unaffected. These effects are quite different from what was observed with the tyrocidines and gramicidin S, indicating that these old peptide antibiotics disrupt the bacterial cell membrane by thus-far unique mechanisms.

Influence on the chromosome.Although the bacterial cell membrane is considered the key target of both the tyrocidines and gramicidin S, in vitro experiments have shown that these antibiotics also interact with DNA (40–43). However, it has never been shown whether this DNA-binding activity plays a role in vivo. To investigate this, we stained the bacterial chromosome with the DNA-binding dye 4′,6-diamidino-2-phenylindole (DAPI). Cells treated with tyrocidines A and C showed a clear condensation of the nucleoid (Fig. 9A). As a control, we treated cells with CCCP but did not observe any nucleoid condensation, indicating that this is not a mere effect of membrane depolarization (Fig. 9A). Since nucleoid condensation occurred rapidly (<2 min) and coincided with shrinking of the cells, it is possible that loss of turgor pressure plays a role in tyrocidine-induced nucleoid condensation. In fact, an osmotic upshock can lead to plasmolysis, nucleoid compaction, and dissociation of RNA polymerase from the DNA (73). Therefore, we examined the localization of RNA polymerase by tagging its subunit RpoC with GFP. As shown in Fig. 8B, RNA polymerase remained attached to the nucleoid but seemed to cluster. To test whether the peptides influence DNA replication, we examined the localization of the DNA replication regulator ParB (Spo0J) (74, 75) and the DNA polymerase subunit DnaN. The ribosome subunit RpsB was included as a cytosolic control protein. Both tyrocidines caused a complete detachment of ParB from the origin of replication (Fig. 9B). DnaN remained attached to the chromosome but exhibited some clustering toward the center of the cells after treatment with tyrocidines A and C (Fig. 9B). The same effect is seen with the gyrase inhibitor ciprofloxacin (60), which causes arrest of the replication fork (76). RpsB, which is not associated with DNA, was not affected by the peptides. To examine whether the peptides cause DNA damage, we looked at the localization of RecA. Upon DNA damage, this repair protein forms large nucleoprotein filaments (77, 78). As shown in Fig. 8B, RecA formed clear foci after treatment with tyrocidines A and C. In contrast to the tyrocidines, gramicidin S had no effect on the localization of any of these proteins. These results indicate that the tyrocidines also target the bacterial DNA, resulting in inhibition of DNA replication and DNA damage, whereas gramicidin S does not show any DNA-targeting activity in vivo.

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FIG 9

Influence of tyrocidines and gramicidin S on the bacterial chromosome. (A) Tyrocidines cause condensation of the nucleoid within 2 min. CCCP was included as a control for depolarization. DNA was stained with DAPI. Strain, B. subtilis 168. (B) Localization of DNA-associated proteins (see main text for details). B. subtilis strains were grown in LB with appropriate inducer concentrations (see Table S1) and treated with peptides in early exponential growth phase for 10 min. The ribosomal subunit protein RpsB was chosen as control for a cytosolic protein not associated with the nucleoid. Bars, 2 µm.

Activity against persister cells.Antibiotic tolerance caused by nongrowing persister cells is an important problem in antibacterial therapy as it leads to chronic and recurrent bacterial infections (79). Membrane-active compounds have been suggested as treatment options for such persistent infections (80). Since the tyrocidines and gramicidin S affect multiple cellular processes, including the cell membrane, we tested the potential of the peptides to kill nongrowing overnight cultures of B. subtilis and S. aureus that have been used as simple model systems for persister cells (81, 82). As shown in Fig. 9A, ampicillin and chloramphenicol were highly active against growing B. subtilis and S. aureus cultures (Fig. 10A). However, overnight cultures were completely resistant to 9-h treatment with these antibiotics. In contrast, both the tyrocidines and gramicidin S were highly efficient in killing nongrowing B. subtilis cells (Fig. 10B). A 9-h incubation with the tyrocidines killed 90% to 99% of nongrowing S. aureus cells, whereas gramicidin S achieved a CFU reduction of as much as 5 log units (Fig. 10C), underlining the clinical potential of the peptides.

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FIG 10

Activity against growing and nongrowing cultures of B. subtilis 168 and S. aureus 8325. (A) MIC(s) against B. subtilis, the model organism used in this study, and the pathogenic Gram-positive bacterium S. aureus. MICs for B. subtilis were additionally determined in a test tube assay to exactly match the conditions for mode-of-action experiments. (B) Log₁₀ CFU reduction of nongrowing B. subtilis and S. aureus cultures treated with ampicillin and chloramphenicol. (C) Log₁₀ CFU reduction of nongrowing B. subtilis and S. aureus cultures (stationary overnight cultures).