Bacteria elicit a phage tolerance response subsequent to infection of their neighbors
2021; Springer Nature; Volume: 41; Issue: 3 Linguagem: Inglês
10.15252/embj.2021109247
ISSN1460-2075
AutoresElhanan Tzipilevich, Osher Pollak‐Fiyaksel, Bushra Shraiteh, Sigal Ben‐Yehuda,
Tópico(s)Antibiotic Resistance in Bacteria
ResumoArticle8 December 2021free access Source DataTransparent process Bacteria elicit a phage tolerance response subsequent to infection of their neighbors Elhanan Tzipilevich Elhanan Tzipilevich Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Osher Pollak-Fiyaksel Osher Pollak-Fiyaksel Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Bushra Shraiteh Bushra Shraiteh Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Sigal Ben-Yehuda Corresponding Author Sigal Ben-Yehuda [email protected] orcid.org/0000-0001-7023-2163 Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Elhanan Tzipilevich Elhanan Tzipilevich Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Osher Pollak-Fiyaksel Osher Pollak-Fiyaksel Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Bushra Shraiteh Bushra Shraiteh Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Sigal Ben-Yehuda Corresponding Author Sigal Ben-Yehuda [email protected] orcid.org/0000-0001-7023-2163 Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel Search for more papers by this author Author Information Elhanan Tzipilevich1,2,†, Osher Pollak-Fiyaksel1,†, Bushra Shraiteh1 and Sigal Ben-Yehuda *,1 1Department of Microbiology and Molecular Genetics, Institute for Medical Research Israel-Canada (IMRIC), The Hebrew University-Hadassah Medical School, The Hebrew University of Jerusalem, Jerusalem, Israel 2Present address: Department of Biology and Howard Hughes Medical Institute, Duke University, Durham, North Carolina, USA † These authors contributed equally to this work *Corresponding author. Tel: +972 2 6758600; E-mail: [email protected] The EMBO Journal (2022)41:e109247https://doi.org/10.15252/embj.2021109247 See also: EE Maffei & A Harms (February 2022) PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions Figures & Info Abstract Appearance of plaques on a bacterial lawn is a sign of successive rounds of bacteriophage infection. Yet, mechanisms evolved by bacteria to limit plaque spread have been hardly explored. Here, we investigated the dynamics of plaque development by lytic phages infecting the bacterium Bacillus subtilis. We report that plaque expansion is followed by a constriction phase owing to bacterial growth into the plaque zone. This phenomenon exposed an adaptive process, herein termed "phage tolerance response", elicited by non-infected bacteria upon sensing infection of their neighbors. The temporary phage tolerance is executed by the stress-response RNA polymerase sigma factor σX (SigX). Artificial expression of SigX prior to phage attack largely eliminates infection. SigX tolerance is primarily conferred by activation of the dlt operon, encoding enzymes that catalyze D-alanylation of cell wall teichoic acid polymers, the major attachment sites for phages infecting Gram-positive bacteria. D-alanylation impedes phage binding and hence infection, thus enabling the uninfected bacteria to form a protective shield opposing phage spread. Synopsis Plaque spread on a bacterial lawn is limited by a "phage tolerance response", elicited by uninfected bacteria subsequent to infection of their neighbours. This response provides temporary resistance, shielding the non-infected bacterial population. Following a phase of plaque expansion, surprisingly the plaque constricts. During constriction, infected bacteria induce a Sigma X-mediated tolerance response in nearby uninfected neighbours, presumably by releasing a danger signal. Sigma X expresses the dlt operon, encoding enzymes that modify the phage cell surface receptor to inhibit phage binding. Introduction Plaques, reporting bacterial clearance, are the hallmark of bacteriophage (phage) infection since the pioneering discoveries made by Twort and d'Herelle at the beginning of the 20th century (Twort, 1914; d'Hérelles, 1917; Chanishvili, 2012). Plaques are basically visible holes, formed on a lawn of bacteria grown on a solid surface, that report bacterial clearance following successive cycles of infection, including phage adsorption, replication, and spread to nearby hosts. Intriguingly, plaques exhibit a considerable variation in shape according to the host and the infecting phage and are predominantly restricted in size (Abedon & Yin, 2009). It has been proposed that such size limitation is achieved, at least in part, by the entry of bacteria into stationary phase, which frequently restrains phage replication (Abedon & Yin, 2009). However, although plaque employment as a method for monitoring phage infection began decades ago, relatively little is known about the kinetics of plaque development. Furthermore, factors limiting plaque size and expansion are mostly unrevealed. In this study, we investigated the dynamics of plaque formation, utilizing the Gram-positive soil bacterium Bacillus subtilis (B. subtilis) and its lytic phages SPP1 and Phi29. Binding of phages to B. subtilis is commonly mediated by wall teichoic acid (WTA) polymers, a diverse family of cell surface glycopolymers containing phosphodiester-linked glycerol repeat units poly(Gro-P), decorated by glucose and D-alanine moieties, and anchored to peptidoglycan (PG) through an N-acetylmannosaminyl (Brown et al, 2013). WTA polymers were found to be crucial surface components, required for invasion by manifold phages into Gram-positive bacteria such as Bacilli, Listeria, and Staphylococci (Lindberg, 1973; Habusha et al, 2019; Ingmer et al, 2019; Sumrall et al, 2020). SPP1, a double-stranded DNA (dsDNA) phage (44 kb) and a member of the Siphoviridae family, characterized by a long noncontractile tail (Alonso et al, 1997), initiates infection by reversible binding of the tail tip to poly-glycosylated WTA (gWTA). Subsequently, SPP1 binds irreversibly to its membrane receptor protein YueB, resulting in DNA injection into the bacterium cytoplasm (Sao-Jose et al, 2004; Baptista et al, 2008). Phi29 phage, which is significantly smaller (19.3 kb) and belongs to the Podoviridae family, harboring a short noncontractile tail (Salas, 2012), was shown to entirely rely on intact gWTA for infection (Young, 1967). The fact that phages assigned to distinct families utilize gWTA to invade the host strengthens the vital role of these polymers in host recognition by phages and implicates them as major elements for host cell vulnerability. Consistent with this notion, we have shown that a mutant bacteriophage capable of bypassing the need for binding the glucosyl residues, decorating the WTA polymers, gained a broader host range, as it could infect non-host bacterial species presenting dissimilar glycosylation patterns (Habusha et al, 2019). Previously, we demonstrated that phages could occasionally invade resistant cells that acquire phage receptors from their sensitive neighbors, highlighting the importance of understanding infection dynamics in a temporal and spatial fashion (Tzipilevich et al, 2017). Here, we visualized plaques formed on a lawn of B. subtilis bacteria. We revealed that plaque spreading is followed by a phase of constriction mediated by bacterial regrowth into the plaque zone. Characterization of the plaque constriction phase uncovered a temporary immunity mechanism, propelled by a programed transcriptional response, enabling bacteria to tolerate infection by remodeling WTA polymers. This modification reduces phage binding and restricts phage spread. Unlike other mechanisms affording long-term bacterial immunity to phages, namely, restriction enzymes and CRISPR (Labrie et al, 2010), this tolerance mechanism confers a transient adaptive response, providing protection to the uninfected bacterial population subsequent to infection of their neighbors. Results Plaque expansion is followed by a phase of plaque constriction To explore the bacterial population dynamic during phage attack on solid surfaces, we followed the process of plaque formation by SPP1 on a lawn of mCherry-labeled B. subtilis cells at high resolution, using time-lapse confocal microscopy. Maximal SPP1 plaque size was detected approximately eight hours postinfection, but remarkably the spread was counteracted by bacteria growing into the plaque area, limiting plaque expansion (Fig 1A; Movie EV1). Consequently, the final plaque diameter measured after overnight incubation was significantly smaller than the maximal size reached during plaque development (Fig 1A). To further explore this phenomenon, we followed the kinetics of plaque formation on agar plates, the typical methodology used over the years for estimation of plaque-forming units (PFU) [e.g., Abedon and Yin (2009), Ellis and Delbruck (1939)]. Consistent with the confocal microscopy results, plate monitoring revealed a steep expansion phase that was proceeded by a gradual decrease in plaque size, with the final zone being approximately 50% of the maximal plaque area measured during the process (Fig 1B and C; Movie EV2). This plaque constriction occurrence was also evident when bacteria were infected with the distinct lytic phage Phi29 (Fig 1D), suggesting that such a kinetic pattern is widespread. Figure 1. Plaque formation dynamics reveals phases of expansion and constriction A. BDR2637 (Pveg-mCherry) cells were infected with low concentrations (10-8 PFU/ml) of SPP1, placed on an agarose pad, and plaque formation was followed by time-lapse confocal microscopy. Shown are overlay images from mCherry signal (purple) and phase contrast (gray) of the bacterial lawn captured at the indicated time points (h). The plaque is seen as a hole formed on the bacterial lawn. Scale bar, 150 µm. Corresponds to Movie EV1. B. Plaque formation was monitored by automated scanning (Levin-Reisman et al, 2010) on a lawn of infected PY79 (WT) (10-6 PFU/ml) cells spread over an MB agar plate. Shown are images captured at the indicated time points (h). Scale bar, 2 mm. Corresponds to Movie EV2. C, D. The dynamic of SPP1 (C) and Phi29 (D) plaque formation following PY79 (WT) infection was monitored as described in (B). Shown is the diameter of individual plaques for each phage (n ≥ 12), with the average highlighted in red. Source data are available online for this figure. Source Data for Figure 1 [embj2021109247-sup-0007-SDataFig1.xlsx] Download figure Download PowerPoint To concomitantly track plaque development and phage localization, we followed plaque formation by SPP1 harboring its lysin gene (gp51) fused to a yellow fluorescent protein (YFP) as a sole copy (SPP1-lysin-yfp), marking active infection (Tzipilevich et al, 2017). Fluorescence from YFP demarcated the plaque periphery even during the constriction phase, signifying the presence of actively infected cells that release phage particles (Fig EV1A–C; Movie EV3). This observation implies that bacteria at the rim could withstand the presence of phages. Isolating bacteria from the edge of 30 different plaques subsequent to the constriction phase and re-plating them over plate-containing phages revealed all tested bacteria to remain phage sensitive (Fig EV1D). We refer to the phenomenon of phage-sensitive bacteria that can confront phages at the plaque circumference as “phage tolerance”. Click here to expand this figure. Figure EV1. Evidence for the presence of phages at the plaque periphery during constriction PY79 (WT) cells were infected with low concentrations (10-8 PFU/ml) of SPP1-lysin-yfp, placed on an agarose pad, and plaque formation was followed by time-lapse confocal microscopy. Shown are overlay images of phase contrast (gray) and signal from Lysin-SPP1-YFP (cyan) captured at the indicated time points (h) postinfection (left panels). Corresponding signal from Lysin-SPP1-YFP (cyan) is shown separately (right panels). Scale bars, 50 µm. Corresponds to Movie EV3. Quantification of the SPP1-lysin-yfp fluorescence intensity (AU) at the indicated time points. Fluorescence from Z sections that include the plaque region and flanking area was measured. Corresponds to EV1A. Quantification of the diameter of the YFP fluorescence (AU) ring, derived from SPP1-lysin-yfp. Corresponds to EV1A. Screening for phage-resistant bacteria at the plaque rim. PY79 (WT) cells were infected with SPP1 (10-6 PFU/ml) and plated for plaque formation. At t = 18 h, similar numbers of bacteria were collected from 30 "non-plaque" and 30 "plaque rim" regions, and bacterial smears were plated over plates with (10-4 PFU/ml) or without phages. No phage-resistant colonies were detected in both populations. Source data are available online for this figure. Download figure Download PowerPoint SigX is necessary for plaque constriction The phenomenon of plaque constriction directed by phage-sensitive bacteria prompted us to postulate that bacteria residing at the plaque periphery could mount a transient phage tolerance response. We further reasoned that such a response could be orchestrated by one of the B. subtilis extracytoplasmic function (ECF) RNA polymerase sigma (σ) factors. These factors are activated in response to stress imposed on the cell envelope (Helmann, 2016) that could stem from remains of surrounding lysed cells. To examine this premise, we deleted each of the seven known ECF sigma factors of B. subtilis and assayed their impact on the final plaque size. Intriguingly, ∆sigX strain exhibited significantly larger plaques in comparison with wild-type (WT) when challenged with SPP1 or Phi29 phages (Fig 2A; Appendix Fig S1A). Furthermore, monitoring plaque dynamics revealed that this size difference is due to the substantial attenuation of the plaque constriction phase (Fig 2B and C). Importantly, when grown in liquid cultures, ∆sigX cells propagated with kinetics similar to that of WT cells (Appendix Fig S1B), indicating that the observed phenotype was not due to growth perturbation. To further elucidate the role of SigX in counteracting phage spread, we challenged ∆sigX cells with SPP1 or Phi29 phages in liquid cultures. No difference in lysis kinetics was observed when bacteria were infected at 1:1 (phage:bacteria) multiplicity of infection (MOI). However, while infected with low MOI (phage:bacteria, 1:20), ∆sigX cells lysed significantly faster than WT cells (Fig 2D; Appendix Fig S1C), a phenotype that could be reversed by ectopic expression of sigX (Appendix Fig S1D). The low MOI might be equivalent to the bacteria:phage ratio reached during the phase of plaque constriction. These results are consistent with the view that uninfected bacteria induce a SigX-regulated defense response, capable of tempering future phage infections. Figure 2. SigX is required for plaque constriction A. The indicated strains were infected with SPP1 (10-6 PFU/ml), spread over MB plates, and plaque diameter was monitored after 20 h of incubation. Shown is average plaque diameter and SD for each strain (n ≥ 54). B, C. Plaque formation dynamic of SPP1 (B) and Phi29 (C) was monitored by automated scanning (Levin-Reisman et al, 2010) on a lawn of infected (10-6 PFU/ml) PY79 (WT) or ET19 (∆sigX) cells spread over an MB agar plate. Shown are average values and SD from kinetic of random plaques for each strain (n ≥ 7). D. PY79 (WT) and ET19 (∆sigX) cells were infected with SPP1 at either high (phages:bacteria 1:1) or low (1:20) MOI, and OD600nm was followed at 2-min intervals. Shown is a representative experiment out of 6 biological repeats, and the average values and SD of 8 technical repeats. E. BDR2637 (Pveg-mCherry) (WT, purple) and ET191 (PrrnE-gfp, ∆sigX) (∆sigX, cyan) cells were mixed, infected with low concentrations (10-8 PFU/ml) of SPP1, placed on an agarose pad, and plaque formation was followed by time-lapse confocal microscopy. Shown are overlay images from mCherry (purple) and GFP (cyan) signals of the bacterial lawn captured at the indicated time points (h). The plaque is seen as a hole formed on the bacterial lawn. Scale bar, 100 µm. Corresponds to Movie EV4. F. ET27 (PsigX-gfp) cells were infected with low concentrations (10-8 PFU/ml) of SPP1, placed on an agarose pad, and plaque formation was followed by time-lapse confocal microscopy. Shown are fluorescence from GFP (upper panels) and corresponding phase contrast images (lower panels), captured at the indicated time points (h). The plaque is seen as a hole formed on the bacterial lawn. Scale bar, 100 µm. Source data are available online for this figure. Source Data for Figure 2 [embj2021109247-sup-0008-SDataFig2.zip] Download figure Download PowerPoint To compare the response to phage attack of WT and ∆sigX cells in real time, we mixed mCherry-labeled WT cells with GFP-labeled ∆sigX cells and monitored plaque dynamics following infection with SPP1, utilizing time-lapse confocal microscopy. At the initial stages of phage spreading, both strains appeared to be infected and to be lysed equally (Fig 2E, t = 5, 9 h; Appendix Fig S2A; Movie EV4). However, during the constriction phase, when the bacteria re-grew into the plaque zone, WT cells outcompeted the ∆sigX cells, as signified by the dominant colonization of the mCherry-labeled WT cells at the plaque rim (Fig 2E, t = 12, 16 h; Fig EV2; Appendix Fig S2A, Movie EV4). When GFP- and mCherry-labeled WT cells were mixed as a control, both strains were evenly distributed at the plaque edge even 16 hours postinfection (Fig EV2). Moreover, WT and ∆sigX cells equally occupied regions located remotely from visible plaque sites (Fig EV2), and, in accord, the growth rate of ∆sigX cells was not significantly affected by the presence of WT cells in a co-culture (Appendix Fig S2C). Consistent with these results, fusion of the sigX promoter to gfp (PsigX-gfp) specified that cells located at the plaque rim produced GFP chiefly during the constriction phase (Fig 2F and Appendix Fig S2B). Interestingly, infection at 48°C, a temperature shown to activate sigX expression (Huang et al, 1997), led to a significant reduction in plaque size in a sigX-dependent manner (Fig EV3A). Of note, although no measurable difference between ∆sigX and WT growth kinetics was seen at 48°C (Fig EV3B), phage manufacture could be alleviated at high temperatures (Schachtele et al, 1970). In sum, we conclude that ∆sigX cells are deficient in inducing a defense mechanism that enables bacteria to tolerate the presence of phages and invade into the plaque zone. Click here to expand this figure. Figure EV2. ∆sigX cells are excluded from the plaque rim during constriction BDR2637 (Pveg-mCherry) (WT) (purple) cells were mixed with AR16 (PrrnE-gfp) (WT) (cyan) (1) or with ET191 (∆sigX, PrrnE-gfp) (cyan) (2-5) cells. The mixtures were infected with low concentrations (10-8 PFU/ml) of SPP1, placed on an agarose pad, and plaque formation was followed by time-lapse confocal microscopy. Shown are overlay images of mCherry (purple) and GFP (cyan) signals captured 16 h postinfection. (1-3) show plaque regions, whereas (4-5) show regions remote from any visible plaque site. Scale bar, 100 µm. Quantification of images 1, 3, and 4 presented in EV2A. Fluorescence intensity (AU) of the plaques formed by phages infecting the corresponding cells is shown. Fluorescence from Z sections that include the plaque region and flanking area or control areas was measured. Source data are available online for this figure. Download figure Download PowerPoint Click here to expand this figure. Figure EV3. Monitoring SigX activation during phage infection PY79 (WT) and ET19 (∆sigX) cells were infected with SPP1 or Phi29 (10-6 PFU/ml), spread over MB agar plates, and incubated at either 37°C or 48°C. Plaque diameter was monitored after 20 h of incubation (n ≥ 50). Shown are average values and SD of 3 independent repeats. PY79 (WT) and ET19 (∆sigX) were grown in liquid LB medium at 48°C and OD600nm monitored. Shown are average values and SD of 3 biological repeats. Corresponds to the experiment presented in Fig 3A. Phage-sensitive PY79 (WT) cells were mixed with phage-resistant BS12 (ΔyueB, PsigX-gfp) cells and the mixture was infected with SPP1 at 2:1 (phages:bacteria) MOI, and OD600nm was followed at 2.5-min intervals. Uninfected mixed population served as a control (-SPP1). Shown is a representative experiment out of 3 biological repeats, and the average values and SD of n ≥ 3 technical repeats. BS12 (PsigX-gfp, ∆yueB) cells were infected with SPP1 at 2:1 (phages:bacteria) MOI, and fluorescence intensity from PsigX-gfp (AU) was followed at 2.5-min intervals. Uninfected BS4 (PsigX-gfp) cells served as a control. Shown is a representative experiment out of 3 biological repeats, and the average values and SD of 3 technical repeats. ET26 (PsigX-sigX-gfp) cells were infected with SPP1 at 5:1 (phages:bacteria) MOI, placed on an agarose pad, and followed by time-lapse fluorescence microscopy. Shown are signal from SigX-GFP (upper panels), and corresponding phase contrast images (lower panels), captured at the indicated time points postinfection. Scale bar, 1 µm. Source data are available online for this figure. Download figure Download PowerPoint SigX is activated in non-infected bacteria following infection of their neighbors The results so far raised the prospect that bacteria could sense a danger signal, emanating from nearby infected bacteria, and in turn activate SigX-dependent phage tolerance response. To assay sigX induction in uninfected bacteria, SPP1 was added to WT cells that were co-cultured with SPP1-resistant bacteria, lacking the phage receptor (ΔyueB), and harboring the PsigX-gfp reporter. Indeed, monitoring GFP fluorescence showed a continuous increase in sigX expression that reached the maximal level approximately 35 min postinfection and started to decline at t = 60 min (Figs 3A and EV3C and D). A similar fluorescence profile was obtained when the two strains were separated by a membrane that allows the passage of small molecules while compartmentalizing the cells (Fig 3B), suggesting that the sigX-inducing factor is secreted into the shared medium. To further substantiate the ability of bacteria to activate SigX in response to nearby infected cells, we followed the production and localization of SigX protein during infection at the cellular level. In the absence of phages, a functional SigX-GFP fusion (PsigX-sigX-gfp) mainly localized onto the membrane, frequently forming focal assemblies in proximity to the cell circumference and at septal positions (Fig 3C and Appendix Fig S1E). This localization pattern is consistent with previous reports, showing that SigX is sequestered to the plasma membrane by its anti-sigma factor as a way to halt its action (Ho & Ellermeier, 2012). To assay SigX activity in uninfected bacteria, we added SPP1 phage to mCherry-labeled WT bacteria mixed with ΔyueB phage-resistant bacteria, harboring sigX-gfp. Time-lapse microscopy revealed repositioning of SigX-GFP from membrane and foci locations to massive nucleoid deployment in the resistant bacteria (Fig 3D; t = 35 min), indicating a switch from an inactive to an active mode. Noticeably, this shift in localization occurred prior to lysis of nearby infected sensitive bacteria and corresponded to the increase in PsigX-GFP signal observed (Fig 3A and B). SigX-GFP level was dropped, and its localization into foci was largely restored in the resistant bacteria 95 min postinfection (Fig 3D), in line with the decline in sigX expression (Fig 3A and B), presumably corresponding to conclusion of the phage-sensing response. Taken together, SigX appears to be activated in phage-resistant bacteria upon sensing a danger signal from nearby infected sensitive cells. Notably, infecting sigX-gfp-sensitive cells with SPP1 showed that SigX-GFP largely displaces its position form the membrane to the nucleoid in the course of infection (Fig EV3E), denoting that also infected cells activate the SigX response. Figure 3. SigX is activated in non-infected cells upon infection of their neighbors Phage-sensitive PY79 (WT) cells were mixed with phage-resistant BS12 (ΔyueB, PsigX-gfp) cells and the mixture was infected with SPP1, as illustrated. Infection was conducted at 2:1 (phages:bacteria) MOI, and fluorescence intensity from PsigX-gfp (AU) was followed at 2.5-min intervals. Uninfected mixed population served as a control, and its fluorescence was subtracted from the overall GFP signal of the infected culture. Shown is a representative experiment out of 3 biological repeats, and the average values and SD of 8 technical repeats. Phage-sensitive PY79 (WT) cells were infected with SPP1 at 2:1 (phages:bacteria) MOI, and placed in the outer ring of a transwell, as illustrated. Phage-resistant BS12 (ΔyueB, PsigX-gfp) were placed in the inner ring. Fluorescence intensity from PsigX-gfp (AU) of the inner compartment was followed at 2.5-min intervals. Uninfected population served as a control, and its fluorescence was subtracted from the overall GFP signal of the infected culture. Shown is a representative experiment out of 3 biological repeats, and the average values and SD of 8 technical repeats. ET26 (PsigX-sigX-gfp) cells were visualized by fluorescence microscopy. Shown are signal from SigX-GFP (cyan) (left panel), and an overlay image of phase contrast (gray) and signal from SigX-GFP (cyan) (right panel). Scale bar, 1 μm. BDR2637 (Pveg-mCherry) (WT, purple) and ET261 (∆yueB, PsigX-sigX-gfp) (cyan) cells were mixed, infected with SPP1 at 5:1 (phages:bacteria) MOI, placed on an agarose pad, and followed by time-lapse fluorescence microscopy. Shown are overlay images from mCherry (purple), SigX-GFP (cyan), and phase contrast (gray), captured at the indicated time points postinfection. Scale bar, 1 μm. Source data are available online for this figure. Source Data for Figure 3 [embj2021109247-sup-0009-SDataFig3.xlsx] Download figure Download PowerPoint Expression of SigX protects from phage infection The impact of SigX on phage infection was further explored by constructing bacteria artificially expressing SigX under an IPTG-inducible promoter. Remarkably, expressing sigX prior to phage addition markedly attenuated both SPP1 and Phi29 infections, with the cells being capable of extending the infection process (Fig 4A). Next, mCherry-labeled cells, over-expressing SigX (PIPTG-sigX), were incubated with non-labeled WT cells, and the mixture was infected with SPP1-lysin-yfp. Consistent with the above observations, WT cells were rapidly infected and lysed, while cells over-expressing SigX appeared to be infected at slower kinetics and to a lesser extent (Fig 4B and C), a phenomenon that was also observed during infection with Phi29 (Fig EV4A). Figure 4. SigX expression confers phage tolerance PY79 (WT) and ET28 (PIPTG-sigX) cells were infected with SPP1 or Phi29 (t = 60 min) at 1:20 (phages:bacteria) MOI, and OD600nm was followed at 2-min intervals. IPTG was added 30 min before infection (t = 30 min). Shown is a representative experiment out of 3 biological repeats, and the average values and SD of 4 technical repeats. ET29 (Pveg-mCherry, PIPTG-sigX) (purple) cells were grown in the presence of IPTG and mixed with PY79 (WT) cells. The mixture was infected with SPP1-lysin-yfp 5:1 (phages:bacteria) MOI, placed on an IPTG-containing agarose pad, and followed by time-lapse fluorescence microscopy. Shown are overlay images of phase contrast (gray), signal from mCherry-labeled cells (purple), and signal from Lysin-SPP1-YFP (cyan), captured at the indicated time points postinfection (upper panels). Corresponding signal from Lysin-SPP1-YFP (cyan) is shown separately (lower panels). Arrows highlight the delayed infection of ET29 cells. Scale bar, 1 μm. Quantification of the experiment described in (B). Shown is the percentage of phage infected PY79 (WT) and ET29 (Pveg-mCherry, PIPTG-sigX) ce
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