Many bacterial pathogens hijack macrophages to egress from the port of entry to the lymphatic drainage and/or bloodstream, causing dissemination of life-threatening infections. However, the underlying mechanisms are not well understood. Here, we report that Salmonella infection generates directional electric fields (EFs) in the follicle-associated epithelium of mouse cecum. In vitro application of an EF, mimicking the infection-generated electric field (IGEF), induces directional migration of primary mouse macrophages to the anode, which is reversed to the cathode upon Salmonella infection. This infection-dependent directional switch is independent of the Salmonella pathogenicity island 1 (SPI-1) type III secretion system. The switch is accompanied by a reduction of sialic acids on glycosylated surface components during phagocytosis of bacteria, which is absent in macrophages challenged by microspheres. Moreover, enzymatic cleavage of terminally exposed sialic acids reduces macrophage surface negativity and severely impairs directional migration of macrophages in response to an EF. Based on these findings, we propose that macrophages are attracted to the site of infection by a combination of chemotaxis and galvanotaxis; after phagocytosis of bacteria, surface electrical properties of the macrophage change, and galvanotaxis directs the cells away from the site of infection.
Common bacterial pathogens such as Salmonella, Shigella, and Yersinia spp. invade the gut epithelial barrier, preferentially by targeting the relatively small number of M cells located in the follicle-associated epithelium (FAE) [1–3]. Disruption of epithelial integrity releases chemokines that attract immune cells such as neutrophils and macrophages—a process known as chemotaxis [4–7]. Subsequent phagocytosis and clearance of these pathogens by immune cells usually stops the infection. However, some of these bacterial pathogens have developed strategies, such as the type III secretion systems in Salmonella spp. [8–12], to evade macrophage killing and survive inside the macrophage [13–16], an environment in which the pathogen is hidden from the immune system. Survival within the macrophage allows the pathogen to spread from its entry site to the spleen, liver, bone marrow, and other organs via the bloodstream, resulting in life-threatening consequences [17–19]. Although chemotaxis can explain how macrophages reach an infected site, it cannot explain how macrophages harboring pathogens escape from the bacterial entry site to reach the lymphatic drainage and/or bloodstream, a critical initial step in the dissemination process that is understudied and poorly understood. Bioelectric signals have been implicated in development [20–23], wound healing [24–26], and regeneration [27,28]. For example, a wound collapses the transepithelial potential (TEP) difference of an intact epithelial barrier, generating laterally oriented endogenous electric fields (EFs) of up to 1. 4 V cm−1, as well as local electric current densities (JI) of several μA cm−2. These bioelectric phenomena actively control wound healing in the skin and cornea [25,29]; however, they are extremely challenging to study in the gut epithelium due to limited accessibility and have never been characterized in vivo during an active infection. Nonetheless, it is generally appreciated that an EF on this scale can drive directional cell migration—a process known as electrotaxis or galvanotaxis [21]. Many cell types, regardless of their origin, respond to an exogenous EF by directional migration toward the cathode [25,30], whereas others, e. g. , human keratinocytes (HaCat cells) and bone marrow mesenchymal stem cells, migrate toward the anode [31]. Macrophages and lymphocytes also undergo galvanotaxis in vitro and in vivo [32–34]. In the present study, we have developed an ex vivo mouse cecum model of Salmonella infection that enables bioelectricity measurement. We report that Salmonella infection generates a directional EF at the bacterial entry sites that can recruit macrophages by galvanotaxis. We demonstrate that primary macrophages reverse galvanotaxis direction upon Salmonella infection by modifying their surface glycan composition, which reduces the negative electric charge on the surface. This directional switch is independent of the Salmonella pathogenicity island 1 (SPI-1) type III secretion system, a major virulence determinant responsible for Salmonella invasion. Instead, it may require certain glycosidases that are widely conserved in Salmonella spp. , because cleavage of surface-exposed sialic acids with a potent neuraminidase caused severe defects of macrophage galvanotaxis.
The mouse is an ideal organism for understanding human infectious diseases and is widely used to study bacterial pathogenesis and mucosal immunity of enteropathogenic bacteria [35–40]. Previously, we have successfully measured bioelectric currents in various tissue and organ cultures using the noninvasive vibrating probe [41–44]; however, measuring bioelectric currents in the mouse small intestinal epithelium is challenging due to limited accessibility [45]. Therefore, we developed a new ex vivo cecum model for measuring bioelectric activity in the gut epithelium (S1 Fig). This model is based on the well-established mouse typhoid model, in which mice orally infected with Salmonella enterica serotype Typhimurium (S. Typhimurium) develop disseminated infection that mimics human typhoid disease [36]. Although the ileum is the most commonly targeted organ to study pathogen–host interactions in vivo, it is too small for the bioelectrical measurements in our ex vivo experimental setting. Instead, the size of the mouse cecum is anatomically optimal. Salmonella invades the cecum in mice and causes acute appendicitis in humans [46]. Furthermore, we can easily identify the FAE under a dissecting microscope, because we found that 90% of C57BL/6 mice have only 1 Peyer’s patch around the blind-end apex, containing 6 to 11 lymphatic follicles. Using microelectrodes [28] in the ex vivo mouse cecum model, we measured a TEP of up to 15 mV, lumen-positive in uninfected mice. Notably, the TEP in the FAE was significantly smaller than that of the surrounding villi (Fig 1A). Next, using a vibrating probe to measure the JI close to the gut surface, we detected outward currents at the FAE and inward currents at the villus epithelium, with a magnitude of around 1 μA cm−2 (Fig 1B). Such currents were not detected in the serosal epithelium, despite the presence of a TEP (Fig 1A), nor in formalin-fixed epithelia (Fig 1B), suggesting the existence of active bioelectricity restricted to the mucosal epithelium. It has been well appreciated that mammalian intestinal mucosa maintains a large transmucosal potential difference [47,48]. In humans, that potential is up to 12 mV, lumen-negative, in the fasting jejunum and ileum [49,50]. Rats, mice, chickens, and fish all maintain a TEP of up to 5 mV in their intestinal epithelium, as measured with Ussing chambers [51–55]; however, these measurements provide no spatiotemporal information [45]. The TEP of control murine ceca (up to 15 mV) that we have measured directly with glass electrodes under microscopic resolution were well within the physiological range described above; however, it differed spatially, i. e. , the TEP of the FAE was significantly lower than the TEP of the surrounding villus epithelium (Fig 1A). Using an ultrasensitive vibrating probe, we further detected ionic currents that run in opposite directions between the FAE and surrounding villi (Fig 1B). Together, these findings unveil lateral voltage gradients and/or constant current loops running between these two structurally and functionally distinct epithelia. In the normal gut scenario, such a bioelectrical landscape may prevent commensals from accidentally entering the FAE and enable the pathogens to specifically target M cells via bacterial galvanotaxis [56]. Salmonella invades the intestinal epithelium, preferentially by targeting M cells located at an FAE [1,57]. If a wound disrupting an epithelium can generate a steady EF, one would envisage that similar EFs must be produced at the Salmonella entry site because of the breakage of epithelial integrity and subsequent short-circuit of the transmucosal potential difference. This is indeed the case in our model. In mice intragastrically challenged with S. Typhimurium, the peak JI remarkably reversed to become inward in FAE compared to control mice, whereas that of the villus epithelium increased nonsignificantly (Fig 1B). Consistent with this finding, we also detected high variation in TEP from individual S. Typhimurium–infected mice (up to 25 mV), still lumen-positive (Fig 1A), which has also been observed in chemically induced colitis in rats [58]. More importantly, penetrating electrodes through disrupted FAE up to 200 μm in depth revealed a stepwise increase in TEP (S2 Fig), suggesting the existence of an electric potential gradient generated by the S. Typhimurium infection. Bacterial invasion and subsequent dissemination to mesenteric lymph nodes (MLNs) and spleen were verified by determination of colony-forming units (CFUs) (S4A Fig), and disruption of FAE was assessed by histological staining (S4B Fig). S. Typhimurium was undetectable in some of the MLNs and spleens, indicating that some of the FAE and surrounding villi were either uninfected or mildly infected, which explains the wide distribution of TEP and JI measured in the S. Typhimurium–challenged mice. Based on these results, we coined the term infection-generated electric field (IGEF) (Fig 1C) to distinguish it from a wound-generated electric field (WGEF) [26]. We demonstrate, for the first time, that Salmonella infection generated a steady EF (up to 5 V cm−1, provided an epithelial thickness of 50 μm) that drives minute directional electric currents, running from the breached FAE into the deep intestinal wall in a stepwise manner. Although how the IGEFs are formed is currently unknown, we speculate on a couple of possible mechanisms. First, in order to establish a steady EF, a potential gradient or a circuit must be formed. Even though there are accumulated charges segregated by the epithelium (positive at the apical side), in a healthy gut a circuit is unlikely to be formed by the epithelium itself due to high resistance of the epithelium, which is sealed by tight junctions. However, given that the intestinal lumen is alkaline (more negative in terms of electrogenesis), a micropotential gradient could be developed in close proximity to the apical side of the gut epithelia, which can drive outward ionic currents (Fig 1C). Second, differential expression and asymmetric distribution of ion pumps and channels essential for selective absorption and/or secretion of electrogenic ions by enterocytes and M cells [47,48] are likely to be critical for the generation of the aforementioned bioelectricity. Future studies using specific channel blockers in combination with our bioelectrical experimental model and techniques will help to pinpoint the molecular mechanism of an IGEF by identifying the channel (s) or pump (s) involved. Third, as in the formation of a WGEF [21,59], Salmonella preferentially invades and destroys M cells and collapses the epithelial barrier at the FAE (S4B Fig), which short-circuits the TEP. Subsequently, the short-circuited and augmented TEP could then drive inward ionic currents as supported by the measurements of stepwise increase of the TEP (S2B Fig). Fourth, in contrast to the healthy alkaline gut, the microenvironment at the Salmonella entry site (i. e. , the FAE) is likely to be more acidic (more positive in terms of electrogenesis) because of the local inflammatory responses [60] and metabolic changes [61] induced by Salmonella infection. Such a microenvironmental pH switch could be related or attributed to the reversal of ionic current flow as we detected with vibrating probes (Fig 1B). Like the injury currents reported by Sawyer and colleagues in early publications [62–64], IGEF-driving ionic currents could affect small blood vessels in the intestinal wall and mesentery to cause a transvascular potential drop or reversal, resulting in 2 possible consequences: an intravascular occlusion that may benefit transendothelial penetration of immune cells (e. g. , monocytes, leukocytes) and/or the creation of a galvanotactic route between the infected epithelium and the electrically impacted vessels [65]. Although IGEFs may provide a guidance cue for the enterocytes or even local stem cells, contributing to the repair process of damaged epithelium, the major focus of this work is, rather, to investigate its biopathological role in the systemic Salmonella infection, specifically during the initiation of macrophage-driven dissemination. Previous studies have shown that applying an EF in vitro can direct macrophage galvanotaxis to the anode [32–34]. We confirmed this phenotype by demonstrating that the primary mouse peritoneal macrophages (PMs), in response to an exogenous EF tuned to mimic the in vivo IGEF (mathematics in Materials and methods), underwent robust unidirectional migration to the anode (S1 Movie). This unidirectional migration was verified by immunostaining showing that nearly 100% of macrophages were polarized to the anode with a distinct morphology characterized by a leading pseudopodium of dense actin meshwork and a rearward uropod. However, upon challenge with S. Typhimurium IR715, the average directionality of EF-induced galvanotaxis decreased significantly, with approximately 41% of the macrophages reversing their migratory direction from the anode to the cathode (Fig 2 and S2 Movie). Although PMs have been widely used in bacterial infection studies [66], likely due to the ease of harvesting, the most commonly used primary murine macrophages are the bone marrow-derived macrophages (BMDMs) because of their high yield and less heterogeneous nature, which makes them phenotypically and functionally different from PMs [67]. To exclude the possibility that the observed bidirectional migration is restricted to PMs, we generated and tested galvanotaxis of BMDMs as we did with PMs. Although both types of macrophages exhibited similar unidirectional migration (mean directionality: −0. 98 versus −0. 88) to the anode, reversal in BMDMs infected by Salmonella IR715 was more robust than in PMs infected by the same Salmonella strain (mean directionality: 0. 36 versus −0. 18) (Fig 2C and S5B Fig). Notably, this phenotype can be reproduced in BMDMs challenged by another 2 virulent S. Typhimurium strains, LT2 and SL1344 (S5 Fig), suggesting that directional migration in response to electrical stimuli is an intrinsic hallmark of the macrophages regardless of their origins and that the ability to manipulate and subvert galvanotaxis in these macrophages is conserved in virulent Salmonella spp. If the cells had simply stopped sensing the EF, then we would have observed random migration, as was the case for control macrophages not subjected to an EF (Fig 2E). The observed change in migration pattern can be attributed to S. Typhimurium infection for a variety of reasons. First, a gentamycin protection assay confirmed the presence of intracellular bacteria (Fig 2D); second, flow cytometry demonstrated a high S. Typhimurium infection rate (approximately 53%) (S6 Fig); and third, high-resolution confocal microscopy revealed that macrophages containing intracellular S. Typhimurium switched polarity to the cathode (Fig 2G). Based on these findings, we conclude that Salmonella-containing macrophages respond to galvanotaxis stimuli by reversing their primary directional migration. Macrophages are professional phagocytes that uptake a broad range of substances; meanwhile, pathogenic Salmonella has developed several virulence means to evade macrophage killing, with SPI-1 being the major virulence factor responsible for colonization and invasion [10,68]. To better understand whether the directional reversal is phagocytosis dependent or SPI-1 specific, we monitored galvanotaxis under identical conditions in BMDMs challenged with (i) fluorescently labeled microspheres similar in size to bacteria, (ii) S. Typhimurium constitutively expressing mCherry, and (iii) a green fluorescent protein (GFP) -expressing ΔinvA mutant that lacks a functional SPI-1 (unable to inject its effectors into cells) [69] (Fig 3A). Macrophages challenged with microspheres exhibited migratory behavior similar to that of controls, i. e. , unidirectional migration to the anode (Fig 3B and S3 Movie). Macrophages challenged with ΔinvA migrated with a significantly decreased overall directionality close to that of macrophages infected with wild-type (WT) Salmonella (Fig 3B), even though the number of intracellular mutants was indeed lower than that of the WT (Fig 3C). Time-lapse recording captured a marked opposing directional migration pattern: macrophages containing microspheres migrated to the anode, and macrophages with either WT or ΔinvA bacteria inside migrated to the cathode (Fig 3D and 3E, and S3 Movie), further confirming that the observed directional switch was Salmonella infection dependent. There were similar phagocytosis and/or infection rates between the macrophages challenged with microspheres or Salmonella in the given multiplicity of infection (MOI; S7A and S7B Fig), as verified by flow cytometry (S6 Fig). Cells containing no or variable microspheres migrated to the anode exclusively (S7C and S7D Fig), ruling out mechanisms solely based on phagocytosis. The fact that cells containing either WT or ΔinvA migrated to the cathode suggests that the SPI-1 type III secretion system is not required or, at least, is insufficient for the directional switch. These data are in accordance with previous studies that identified an SPI-1-independent pathway contributing to early dissemination of S. Typhimurium in the mouse typhoid model [17]. We therefore argue for the existence of a general infection-dependent mechanism that involves phagocytosis and subsequent interplay between the macrophages and internalized bacterial pathogens. Charged cell-surface components are critical for EF-induced motility in 3H3 cells [70] and have been implicated in electro-osmosis of concanavalin A (Con A) binding receptors on the surface of myotomal spheres [71]. We hypothesized that the directional switch of macrophage galvanotaxis could result from bacteria-induced modifications to the charged components of the cell surface, which would not occur following microsphere challenge (Fig 4A). To this end, we screened Salmonella-infected and control macrophages against a panel of fluorescently labeled lectins (glycan-binding proteins) capable of detecting charged and uncharged cell surface glycans (S1 Table). The normalized mean fluorescent intensity of Maackia Amurensis Lectin II (MAL-2), a lectin that recognizes pathogen-binding sialic acids, was significantly decreased in macrophages infected by Salmonella but not in those carrying microspheres (Fig 4B–4D). Marked Galanthus Nivalis Lectin (GNL) - and Con A–binding aggregates were visible within macrophages infected by Salmonella (S8 and S9 Figs), raising the possibility that the decrease in MAL-2-binding sialic acids could be the result of bacterial internalization and subsequent degradation. Sialylated cell surface molecules are negatively charged, creating an electronegative zeta potential [72]. Using an electrophoretic light-scattering technique, we determined the zeta potential of BMDMs under various conditions. The negative zeta potential of macrophages infected by S. Typhimurium was significantly reduced, i. e. , less negative than that of control macrophages (P < 0. 05) (Fig 4E). By contrast, macrophages challenged with microspheres showed a nonsignificant zeta potential change compared to that of control macrophages (Fig 4E), suggesting that the decrease in surface-exposed sialic acids and subsequent reduction of surface negativity is mediated by active bacterial product (s) [73]. If a decrease of the negatively charged sialic acids of the surface glycoproteins is critical for the directional switch in macrophage galvanotaxis, then cells with their sialic acids enzymatically removed should exhibit a switch or at least a defect in directional migration when exposed to an EF. To test this, we treated freshly isolated mouse BMDMs with a potent neuraminidase (an enzyme that cleaves terminal sialic acid residues from surface-exposed glycoproteins) [74]. Cleavage of sialic acids following enzymatic treatment was confirmed by flow cytometry and confocal microscopy (Fig 5A–5C). As predicted, the zeta potentials of neuraminidase-treated macrophages were significantly reduced (Fig 5D). These cells also lost anodal migration compared to control macrophages monitored in parallel (Fig 5E and 5F, and S4 Movie). Further inspection and morphological quantification of macrophages stained with fluorescently labeled actin and/or lectin revealed that 71% of the macrophages treated with neuraminidase failed to establish a polarity and notably, 12% of the cells were polarized to the cathode whereas nearly all the control macrophages (97%) were polarized to the anode in response to the EF (Fig 5G). It is well known that a change in the environmental pH can dramatically influence the growth and virulence of Salmonella [75,76]. Moreover, by exposure to acidic pH, a condition that is required for activation of several Salmonella virulence factors [77–79], it is possible to modify macrophage surface electrical properties [30]. Therefore, to further determine the importance of surface negativity, we incubated BMDMs in medium at pH 5. 8 that markedly reduced the zeta potential (S11A Fig), presumably through protonation of the sialylated surface molecules. Similar to the neuraminidase treatment, galvanotaxis of BMDMs under acidic conditions was significantly impaired, resulting in nearly half of the macrophages (47%) losing their directional migration and 14% of the macrophages reversing their polarity to the cathode (S11B-S11D Fig and S5 Movie). Although these data are consistent with our previous studies, showing that low pH abrogates directional migration of epithelial cells in response to an EF [80,81], it also suggests that the acidic environment is not only required for the activation of Salmonella but also contributes to triggering dissemination. How does Salmonella infection instruct macrophages to reverse EF-directed migration? Firstly, it is important to note that infecting macrophages with a SPI-1 mutant resulted in a directional switch similar to that of WT, with nearly all the macrophages containing live fluorescent protein-expressing bacteria migrating to the cathode (Fig 3 and S3 Movie). This suggests a general mechanism, independent of this major virulence factor, even though other specific factor (s) may still be involved [82,83]. Previously reported effects have suggested that negatively charged surface glycan moieties are critical for EF-induced cell motility and polarization, which are consistent with our data, and provide a long sought-after mechanism of action. Because macrophages challenged with microspheres did not show a significantly reduced zeta potential (Fig 4E), it is likely that the decrease in surface-exposed sialic acids and reduction of surface negativity is mediated by active bacterial product (s) rather than by metabolic changes in the host itself. It is also important to note that although both Salmonella infection and neuraminidase treatment decreased the surface-exposed sialic acids, the latter caused a serious defect in macrophage galvanotaxis without reversing the overall directionality (Fig 5E), in contrast to Salmonella infection (Fig 3B). There may be multiple glycosidases involved in surface glycan modification to reverse directionality (Fig 5H) because Salmonella possesses at least 51 putative glycosidases that likely function in glycan degradation [84]. In fact, a recent study identified several glycosidases, including a putative neuraminidase, as new virulence factors essential for Salmonella infection of epithelial cells, which is again independent of the SPI-1 [85]. We are in the process of performing genetic knockouts to identify the factors involved. It is also possible that modification of surface glycan and reduction of zeta potential were mediated by internalization during phagocytosis of the bacterium itself rather than by bacterial enzymatic activities. For example, macrophages express Toll-like receptors (TLRs) that recognize structurally conserved molecules derived from Salmonella and other pathogens. All TLRs contain N-linked glycosylation consensus sites, and both TLR2 and TLR4 require glycosylation for surface translocation and function [86,87]. Binding of Salmonella to these glycosylated receptors and subsequent internalization may reduce surface negativity of macrophages, leading to directional switch under an EF. This idea is supported by our observations in which the accumulation of certain lectin binding aggregates within macrophages infected by Salmonella but not in cells challenged by microspheres (S8 and S9 Figs). It is also possible that a macrophage can uptake microspheres without significantly changing its zeta potential (e. g. , through a neutralized receptor) and thus still migrate to the anode. Disseminated Salmonella infection is a major health problem of developing countries, responsible for approximately 433,000 deaths annually [88]. Understanding the mechanisms that trigger dissemination is critical for efforts to target this key process for preventive and therapeutic purposes. We propose that macrophages are attracted to the site of infection by a combination of chemotaxis and galvanotaxis, driving the cells in the same direction. After phagocytosis of bacteria, surface electrical properties of the macrophage change, and galvanotaxis directs the cells away from the site of infection (Fig 6). Our study represents a new perspective for the initiating mechanisms, suggesting that Salmonella disseminates through infection-generated bioelectrical control of macrophage trafficking. It is important to emphasize that the demonstrated bidirectional migration of macrophages to the IGEF-like EFs is not a physical electrophoresis (movement of charged particles under direct-current EF) but rather a complex yet poorly understood biological process that requires phosphoinositide 3-kinases and other critical signaling activities, as well as the cellular motility machinery [25,81]. It is also worth emphasizing that the model proposed in this work is not mutually exclusive with respect to chemotaxis but offers an alternative and/or complementary mechanism of directional migration. Both directional cues can coexist and play equally important roles in orchestrating the initial stage of the innate immune response against bacterial infection (Fig 6), although a chemical gradient could be overridden by a strong electrical stimulus [59,89]. Both chemotaxis and galvanotaxis likely share critical signaling pathways as suggested in studies of macrophage-like Dictyostelium, which are highly sensitive to cyclic adenosine monophosphate gradients, as well as to electrical potential gradients [90–92]. Future work utilizing transgenic animals and pharmacological perturbations to target specific pathways (known or unknown) will help to pinpoint key molecules mediating the infection-dependent directional reversal (i. e. , the molecular mechanisms to initiate disseminated infection) and response to bioelectric signaling. Triggering bacterial dissemination mediated by macrophage galvanotaxis might be a common strategy, not only for pathogenic Salmonella but also for other bacterial pathogens that are able to invade macrophages and survive intracellularly. Although the present work deals primarily with gut epithelium and enteric bacteria, the general mechanism that emerged from this work could also apply to other mucous epithelia such as the respiratory tract and its associated pathogens, e. g. , Mycobacterium tuberculosis (the causative agent of tuberculosis) [93], which is another major public health concern.
The mouse strains used were in a C57BL/6 background (both male and female mice were used in experiments). Mice were purchased from Jackson lab and maintained under a strict 12-h light cycle and given a regular chow diet in a specific pathogen-free facility at University of California (UC), Davis. All animal experiments were performed in accordance with regulatory guidelines and standards set by the Institutional Animal Care and Use Committee of UC Davis. In brief, we dissected mouse cecum following euthanasia and opened longitudinally along the mesenteric attachment remnant to avoid incision damage to the single Peyer’s patch located under the antimesenteric mucosa near the apex (S1A Fig). After thorough washing in mouse Ringer’s solution (154 mM NaCl, 5. 6 mM KCl, 1 mM MgCl2,2. 2 mM CaCl2,10 mM glucose, and 20 mM HEPES [pH 7. 4]) to remove the luminal contents, we placed the cecum with mucous side facing up, on a 30° slope of silicone gel prepared from polydimethylsiloxane (PDMS) in custom-made measuring chambers. The cecum was aligned and immobilized with fine metal (tungsten) pins prior to taking measurements (S1B Fig). This process was usually completed within 5 min. We used glass microelectrodes to directly measure the TEP of intestinal epithelium as previously described [28,94]. TEPs were recorded by microelectrode impalement through the epithelial layers. Microelectrodes (1–2 μm tip diameter; NaCl 3 M electrolyte) had resistances of approximately 1 to 2 MΩ, and the potentials were offset to 0 mV prior to impalement. Cecal FAE and adjacent villus epithelium were discriminated under a dissecting microscope within a Faraday cage on an antivibration table. In some cases, the TEP were measured as follows: first at the epithelial surface (0 μm), then stepwise at 50,100, and 200 μm in depth, controlled by a micromanipulator (S2A Fig). The potential typically returns to the baseline of 0 mV after microelectrode withdrawal. If the reference baseline was > ±1 and ≤ ±5 mV, the value was subtracted from the TEP recorded as shown in the equation (S2B Fig); if > ± 5 mV, the trace was rejected. As a control, we measured the TEP of serosa epithelium and the TEP of formalin-fixed mucous epithelium. Measurements were performed at room temperature in mouse Ringer' s solution. Data were acquired (saturated sampling at 100 Hz) and extracted using pClamp 10 (Molecular Devices) and analyzed using Excel (Microsoft, Redmond, WA). We used noninvasive vibrating probes to measure the electric current density (JI) in μA cm−2 of mouse cecum epithelium as previously described [28,41,42]. The probes, platinum-electroplated at the tip (approximately 30 μm ball diameter), vibrated at a frequency between 100 and 200 Hz. Prior to measurements, the probe was calibrated to the experimental conditions by an applied JI of 1. 5 μA cm−2 (S3C Fig). Under a dissecting microscope, mounted mouse ceca were positioned in the nonconductive measuring chamber (S3A Fig). The plane of probe vibration was perpendicular to the epithelial surface at a distance as close as possible (S3B Fig). JI was recorded until the plateau peak was reached (<1 min) (S3C Fig). Reference values were recorded with the probe away from the epithelium surface (≫1 mm) (S3A Fig). Measurements were taken at room temperature in mouse Ringer' s solution. During calibrations and measurements, a Faraday “wall” (grounded aluminum-wrapped cardboard) covered the microscope. As a control, we measured JI near the surface of serosal epithelium and formalin-fixed mucous epithelium. Data were acquired and extracted using WinWCP V4 (Strathclyde Electrophysiology Software, University of Strathclyde, Glasgow, United Kindom) and analyzed using Excel (Microsoft, Redmond, WA). Special reagents used in this work are listed in S2 Table. Plasmids and Salmonella strains used in this work are listed in S3 Table. Cultures of Escherichia coli (for plasmid extraction) and S. Typhimurium were incubated aerobically at 37°C in Luria-Bertani (LB) broth (per liter: 10 g tryptone, 5 g yeast extract, 10 g NaCl) or on LB agar plates (1. 5% Difco agar) overnight. Antibiotics were used at the following concentrations unless stated otherwise: 30 μg ml−1 chloramphenicol, 50 μg ml−1 nalidixic acid, 100 μg ml−1 ampicillin, 50 μg ml−1 kanamycin, and 10 μg ml−1 tetracycline. In an effort to monitor live intracellular Salmonella during macrophage galvanotaxis we constructed an S. Typhimurium strain derived from IR715 that constitutively expresses mCherry coded in its genome [95]. In brief, the glmS: : mCherry allele was transferred to IR715 via P22 transduction from the donor strain S. Typhimurium SL1344St (a gift from Leigh Knodler) [96] and selected by chloramphenicol. To remove the FRT-flanked chloramphenicol cassette, the transductant was transformed with pCP20 encoding FLP recombinase. The resulting strain constitutively expressing mCherry encoded by glmS: : mCherry: : FRT was designated KLL18. The GFP-expressing SPI-1 mutant ΔinvA was generated by electroporating pGFT/RalFc into AJB75 [69]. The plasmid pGFT/RalFc was constructed in 2 steps. First, a fragment of a gfp-mut3 gene under the control of the constitutively active kanamycin-resistance gene aphA3 promoter was amplified from pJC43 [97] with the primers GFPmut3-F (5ʹ- AGAGCTCCAGCGAACCATTTAAGGTGATAG -3ʹ) and GFPmut3-R (5ʹ- ACTGCAGTTATTTGTATAGTTCATCCATGCC -3ʹ). This fragment was then digested with SacI/PstI and cloned into pFT/RalFc, a low–copy-number plasmid based on pBBR1-MCS4 [98]. The mouse is a well-established animal model for studying Salmonella pathogenesis [35]. C57BL/6 and other mice carrying a mutation in nramp1 develop disseminated infections when challenged by S. Typhimurium, which mimics human typhoid fever [99]. For the mouse infection experiment, we used the hyper-disseminative D23580 strain [100,101]. In brief, D23580 was used to inoculate LB broth and incubated overnight at 37°C. C57BL/6 mice (6 to 10 wk old, mixed sexes) were intragastrically infected with 109 bacteria (actual inoculum was determined by plating) in 0. 1 ml LB broth. Uninfected mice used as a control were given 0. 1 ml sterile LB broth in place of Salmonella. Mice were euthanized at 16 h and 40 h post infection (PI) by CO2 asphyxiation followed by cervical dislocation as the secondary method of euthanasia. The 16 h PI was chosen based on previous studies showing that the best-characterized route for the phagocytes harboring Salmonella to reach the bloodstream normally requires between 12 and 20 h [17,102]. MLNs and spleens were collected aseptically and homogenized in phosphate-buffered saline (PBS) for CFU enumeration. Ceca were dissected, cleaned, and then either mounted for bioelectrical measurement, prepared for histopathological fixation, or homogenized in PBS for CFU enumeration. Ceca were fixed in 10% neutral buffered formalin. After fixation, tissues were routinely processed, embedded in paraffin, sectioned, and stained with hematoxylin–eosin. Both PMs and BMDMs were isolated from C57BL/6 mice (6 to 10 wk old, mixed sex) following standard procedures as previously described [103]. PMs were seeded onto 6-well plates and allowed to adhere to the plastic for 1 to 2 d in DMEM (Invitrogen) with 10% fetal bovine serum (Invitrogen) and 1× antibiotic-antimycotic solution (Invitrogen). BMDMs were cultured in the same medium as described above but supplemented with 20% L-929 conditioned medium for 6 d (plus an extra feed at day 3), followed by 1-d culture without the conditioned medium. Adherent macrophages were then harvested by gently scraping with a “policeman” cell scraper and used for subsequent experiments accordingly. Cell viability was determined by trypan blue staining and counting. Our initial galvanotaxis experiments were carried out in PMs. Because we observed better directional switch in BMDMs infected by Salmonella (S5 Fig), subsequent experiments were done in BMDMs, unless stated otherwise. The gentamycin protection assays were carried out as previously described [104]. In 24-well tissue culture-treated plates, 2 × 105 cells were seeded per well for 5 to 6 h in culture medium (DMEM with 10% fetal bovine serum and no antibiotics). Salmonella were grown overnight and used to infect macrophages at an MOI of 20. After 60 min of incubation, cells were gently washed 3× with PBS and further incubated in gentamycin-containing culture media at a final concentration of 50 μg ml−1 for an additional 60 min. Afterwards, media were replaced with culture media containing 10 μg ml−1 gentamycin for the duration of the experiment. Intracellular CFU was measured at 16 h PI. To measure intracellular CFU, macrophages were lysed using 0. 5% Tween 20 for 5 min at room temperature and released by scraping with 1 ml pipette tips. CFUs were enumerated by plating. Typically, 4 × 104 primary mouse macrophages were seeded per well of engineered silicon stencils sealed in custom-made EF chambers (see “Engineering silicone stencil and EF chamber design” for details) or 96-well glass bottom plates (Nunc) or 2 × 105 cells per well in 24-well tissue culture-treated plates depending on different experiment needs for 5 to 6 h in culture medium. Overnight cultures of Salmonella or fluorescently labeled microspheres were used to infect and/or challenge macrophages at an MOI of 20. The rest of the procedures were similar to the gentamycin protection assay. Cells were cultured in medium containing 10 μg ml−1 gentamycin for 16 h, and subsequent galvanotaxis experiments were carried out in the same medium containing gentamycin. For the neuraminidase treatment, cells were incubated in culture medium containing 100 mU ml−1 neuraminidase from Vibrio cholerae (Sigma-Aldrich) for 30 min at 37°C [70]. Cells were then washed with culture medium, and subsequent galvanotaxis experiments were carried out in the culture medium containing no neuraminidase. For the low pH experiments, cells were incubated in culture medium of pH 5. 8 buffered with 15 mM MES (Sigma-Aldrich) for 60 min, and subsequent galvanotaxis experiments were carried out in the same media of pH 5. 8 [81]. Control experiments were carried out always in parallel in culture medium of pH 7. 4 buffered with 14. 4 mM HEPES (Invitrogen). Macrophages were seeded in either 96-well glass bottom plates or custom-made EF chambers and infected, challenged, and/or treated by following procedures as described above. The cells were fixed with 4% paraformaldehyde immediately or after EF exposure for 3 h with field orientation marked. Salmonella were detected with a polyclonal antibody specific to Salmonella spp. (Mybiosource, San Diego, CA) stained by an Alexa Fluor 488-conjugated secondary antibody. F-actin was labeled by Alexa Fluor 555 Phalloidin. Nuclei were labeled by Hoechst 33342. In the cases of lectin staining, fixed cells were incubated with FITC-labeled lectin (S1 Table) overnight at 4°C, washed extensively, and then stained with DAPI for 10 min on ice. Cells were photographed using either an inverted (for cells on cover glass with no EF) or an upright (for cells on plastic EF chambers) Leica TCS SP8 confocal microscope (Leica microsystem). Images were processed using ImageJ. Quantification and comparison of fluorescent intensity were done in images taken in the same batch with the same optical setup and parameters. Lectin binding aggregates stained after permeabilization were quantified by thresholding. Cells were counted using particle analysis function. Infected, challenged, and/or treated macrophages, handled according to the procedures described above, were then incubated with Fc-block (BD, Franklin Lakes, NJ) on ice for 15 min, stained with FITC-labeled lectin (S1 Table) for 1 h on ice, and then stained with Aqua-LIVE/DEAD (Invitrogen, Carlsbad, CA) for 30 min at room temperature. Cells were washed after each step and before being analyzed on a BD Fortessa flow cytometer. Data were analyzed using FlowJo software (Tree Star Inc. Ashland, OR). After gating single cells and live cells, the geometric mean fluorescence intensity and standard error (SE) were collected for each FITC lectin in each condition in addition to Fluorescence Minus One (FMO) for FITC-lectin (no FITC-lectin staining). The geometric mean fluorescence intensity was then standardized across experiments using the following equation: xFMO (MFI) =x (MFI) −FMO (MFI) FMO (SD), (2) where xFMO (MFI) is the standardized geometric mean fluorescence intensity of a specific lectin for a specific experiment, x (MFI) is the geometric mean fluorescence intensity of a specific lectin for a specific experiment, FMO (MFI) is the geometric mean fluorescence intensity of the FMO for a specific experiment, and FMO (SD) is the standard deviation of the FMO for a specific experiment. Standardized geometric mean fluorescent intensities were then plotted and tested for statistical significance (S10 Fig). Macrophages were seeded onto 24-well tissue culture plates and infected, challenged, and/or treated following procedures as described above. Cells were fixed in 2% paraformaldehyde and washed with motility buffer (10−4 M potassium phosphate buffer at pH 7. 0, with 10−4 M EDTA) [56]. Cells were then gently collected by scraping with a “policeman” cell scraper, and subsequent measurements were done in motility buffer, except for the macrophages tested for acidic treatment, which were measured in either pH 5. 8 medium or pH 7. 4 medium as a control. Zeta potential was determined by electrophoretic light scattering at 25°C with a Zetasizer (Malvern Panalytical Ltd, Malvern, United Kingdom). Zeta potential was calculated in mV, and differences between groups were analyzed by Student t test. Galvanotaxis data from representatives of at least 4 independent experiments were routinely presented as mean ± SE, unless stated otherwise. Distributions of macrophage polarity between control and neuraminidase treated or between neutral and acidic conditions were analyzed using χ2 test. Student t test and one-way ANOVA analysis followed by post hoc Tukey HSD test were used for paired or unpaired comparisons among 2 groups or multiple groups (more than 2), respectively.
Bacterial pathogens can invade and survive within macrophages and use them as a vehicle to reach important organs of a human body, resulting in life-threatening infections, but the underlying mechanisms are not well understood. Our current understanding is that macrophages are recruited to the infected site by sensing gradients of chemokines and/or cytokines released by damaged cells—a process known as chemotaxis. However, this mechanism does not explain how macrophages containing the pathogens escape the site to reach the bloodstream. Here, we use a disseminated Salmonella infection model in mice and detect electric fields (EFs) generated by Salmonella infection at the gut epithelium, which can drive unidirectional migration of macrophages towards the anode—a biological process also known as galvanotaxis. We further demonstrate that macrophage galvanotaxis can be reversed to the cathode by phagocytosis of the bacteria. Based on these findings, we propose that macrophages are attracted to the site of infection by a combination of chemotaxis and galvanotaxis; after phagocytosis of bacteria, the electrical properties of the macrophage change, and galvanotaxis directs the cells away from the site of infection allowing the escape of the macrophages that contain pathogens.
blood cells
medicine and health sciences
immune cells
pathology and laboratory medicine
engineering and technology
electronics
cathodes
pathogens
immunology
electricity
microbiology
salmonellosis
bacterial diseases
enterobacteriaceae
electric field
bacteria
bacterial pathogens
salmonella typhimurium
infectious diseases
white blood cells
animal cells
medical microbiology
microbial pathogens
biological tissue
salmonella
physics
anodes
cell biology
anatomy
epithelium
biology and life sciences
cellular types
physical sciences
macrophages
electrodes
organisms
Infection-generated electric field in gut epithelium drives bidirectional migration of macrophages