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Open Access

Peer-reviewed

Research Article

Role of the type 6 secretion system on apoptosis and macrophage polarization during Burkholderia pseudomallei infection

  • Jacob L. Stockton,

    Roles Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Validation, Writing – original draft, Writing – review & editing

    Current address: USDA APHIS Foreign Animal Disease Diagnostic Laboratory Manhattan, Kansas, United States of America

    Affiliation Department of Microbiology and Immunology, University of Texas Medical Branch Galveston, Texas, United States of America

    ⨯
  • Nittaya Khakhum,

    Roles Conceptualization, Investigation, Methodology, Writing – original draft, Writing – review & editing

    Affiliation Department of Microbiology and Immunology, University of Texas Medical Branch Galveston, Texas, United States of America

    ⨯
  • Alfredo G. Torres

    Roles Conceptualization, Funding acquisition, Project administration, Resources, Supervision, Validation, Writing – original draft, Writing – review & editing

    altorres@utmb.edu

    Affiliations Department of Microbiology and Immunology, University of Texas Medical Branch Galveston, Texas, United States of America, Department of Pathology, University of Texas Medical Branch Galveston, Texas, United States of America

    ⨯

Abstract

Burkholderia pseudomallei (Bpm) is the causative agent of the disease melioidosis. As a facultative intracellular pathogen, Bpm has a complex lifestyle that culminates in cell-to-cell fusion and multinucleated giant cells (MNGCs) formation. The virulence factor responsible for MNGC formation is the type 6 secretion system (T6SS), a contractile nanomachine. MNGC formation is a cell-to-cell spread strategy that allows the bacteria to avoid the extracellular immune system and our previous data highlighted cell death, apoptosis, and inflammation as pathways significantly impacted by T6SS activity. Thusly, we investigated how the T6SS influences these phenotypes within the macrophage and pulmonary models of infection. Here we report that the T6SS is responsible for exacerbating apoptotic cell death during infection in both macrophages and the lungs of infected mice. We also demonstrate that although the T6SS does not influence differential macrophage polarization, the M2 polarization observed is potentially beneficial for Bpm pathogenesis and replication. Finally, we show that the T6SS contributes to the severity of inflammatory nodule formation in the lungs, which might be potentially connected to the amount of apoptosis that is triggered by the bacteria.

Author summary

Burkholderia pseudomallei (Bpm) is an intracellular pathogen and is the etiological agent of melioidosis. This neglected tropical disease results in an estimated 89,000 fatalities a year, however, this number is believed to be severely underreported. The complex intracellular lifestyle and host response to infection is poorly understood, basic cell interactions and responses are critical to defense against infection. Bpm utilizes an array of virulence factors to successfully replicate and disseminate cell-to-cell, one of these is the T6SS which is responsible for MNGC formation and cell-to-cell spread. In this work, we characterized how macrophages respond to infection in the presence or absence of the critical T6SS virulence factor. Macrophages are a primary replicative niche for Bpm but how intracellular replication disrupts macrophages response to infection is poorly understood. By understanding macrophage cell death patterns and polarization, we can dissect responses that are triggered by T6SS activity with the expectation of finding exploitable responses for host directed therapies.

Citation: Stockton JL, Khakhum N, Torres AG (2024) Role of the type 6 secretion system on apoptosis and macrophage polarization during Burkholderia pseudomallei infection. PLoS Negl Trop Dis 18(10): e0012585. https://doi.org/10.1371/journal.pntd.0012585

Editor: Andrew Scott MacDonald, University of Manchester, UNITED KINGDOM OF GREAT BRITAIN AND NORTHERN IRELAND

Received: April 20, 2024; Accepted: September 30, 2024; Published: October 15, 2024

Copyright: © 2024 Stockton et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: All relevant data are within the manuscript and its Supporting Information files.

Funding: This work was supported by U.S. Department of Agriculture APHIS AP20VSD&B000C087 and the U.S. Department of Agriculture APHIS NBAF Scientist Training Program Fellowship to JLS and University of Texas Medical Branch institutional funds to AGT. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Burkholderia pseudomallei (Bpm) is a Gram-negative environmental saprophyte that is the causative agent of melioidosis [1]. Melioidosis is a neglected tropical disease that affects an estimated 165,000 people, with approximately 89,000 deaths, a year [2,3]. Bpm was thought to be restricted to southeast Asia and northern Australia; however, it has been shown to have global distribution [3,4]. This geographical distribution includes the Americas where melioidosis is starting to be recognized and growing public health threat [5–8]. The clinical manifestations of melioidosis are highly variable, which often leads to misdiagnosis, earning Bpm the moniker “The Great Mimicker” [1,9,10]. Bpm is a facultative intracellular pathogen that can successfully infect both phagocytic and non-phagocytic cell types and get distributed to almost every tissue in the host [11,12]. Bpm owes its success as an intracellular pathogen to an arsenal of virulence factors that it utilizes to invade, survive, and spread from cell-to-cell. One critical virulence factor is the type 6 secretion system (T6SS), Bpm utilizes this nanomachine to fuse host cell membranes and generate multinucleated giant cells (MNGCs) [13–16]. MNGC formation is the keystone pathogenesis feature of Bpm and the lack of T6SS activity results in attenuation of the bacterium [13,17]. The mechanism by which host cell membranes are fused by the T6SS and the consequence of MNGC formation, from the host perspective, are both currently unknown.

Previously, we began to interrogate this intracellular pathogen by performing dual RNA-seq using our established in vitro model of gastrointestinal (GI) infection [18,19]. During this analysis, it was found that T6SS activity contributes to modulation of inflammatory responses through NFκB and the differential expression of numerous cell death pathway genes. The primary cell death pathway highlighted was apoptosis, which is significant due to the nature of apoptosis being “immunologically silent” [20]. Cells that undergo apoptosis do not release intracellular contents that would be perceived as damage associated molecular patterns (DAMPs) by the immune system and result in an inflammatory response. Apoptotic corpses are cleared by phagocytic cells to prevent secondary necrosis in a process called efferocytosis [21]. Hijacking this mechanism of cell death to evade the immune system would be advantageous to the bacterium as it can continue to spread from cell-to-cell through efferocytosis under the immune silence of apoptosis. Modulation of phagocytic cell death and evasion of intracellular clearance mechanisms by Bpm [22,23] and apoptosis has been implicated during infection numerous times in a multitude of cell lines [24–28]. Certain virulence factors have been implicated in the activation of apoptosis, including the type 3 secretion system (T3SS) [26] and the BimA protein [24], however, the dual RNA-seq analysis implicated the T6SS apparatus [18]. As the T6SS is downstream of both the T3SS and BimA-mediated actin motility during intracellular pathogenesis, it is likely that the T6SS is the main driver of apoptosis during infection.

Cell death can have a profound impact on the immune microenvironment; different modes of death can vastly change how the immune system responds. For example, pyroptosis and necroptosis release pro-inflammatory DAMPs and cytokines, which is in contrast to the silent death occurring during apoptosis [29]. Both processes can impact the behavior of macrophages through the mechanism of polarization. Macrophage polarization is a phenomenon during which these immune cells get activated and skew towards pro-inflammatory (M1) or alternatively activated (M2). The M2 macrophages commonly take on anti-inflammatory and homeostatic characteristics, while classically activated M1 macrophages are primarily involved in pathogen clearance and tissue damage [30]. We have recently reported that Bpm elicits differential pulmonary macrophage polarization during infection with a Bpm ΔbicA strain, a T3SS mutant. While wild-type (WT) infected mice resulted in both M1 and M2 polarization, the ΔbicA failed to generate M2 polarization [31]. In this work, we evaluate the contribution of the T6SS system as an inducer of apoptosis and macrophage polarization to understand the consequences of MNGC formation and start elucidating its role in pathogenesis.

Materials and methods

Ethics statement

All manipulations of B. pseudomallei were conducted in CDC/USDA-approved and registered BSL3 facilities at the University of Texas Medical Branch (UTMB) in accordance with approved BSL3 standard operating practices. The animal studies at UTMB were carried out humanely in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals by the National Institutes of Health. The protocol (IACUC no. 0503014E) was approved by the Animal Care and Use Committee of UTMB.

Bacterial strains and growth conditions

All experiments were conducted with the prototypical wild-type B. pseudomallei strain K96243 or derivative strains (Δhcp1 (ΔBPSS1498) [32], Δhcp1::hcp1). All Bpm strains were routinely grown at 37°C on LB agar plates and in LB broth with shaking. Escherichia coli S17-1 λpir were grown in LB agar plates and broth at 37°C, and kanamycin was added for plasmid selection. For the counter selection, co-integrants were grown in YT medium supplemented with 15% sucrose.

Construction of hcp1 strain complementation

The in cis complementation of the Bpm hcp1 mutant was performed by inserting the hcp1 gene back into Bpm Δhcp1 strain via allelic exchange using Burkholderia optimized vector pMo130 [32]. Purified PCR amplicon of upstream-BPSS1498-downstream and pMo130 vector were digested by NheI and HindIII restriction enzyme followed by ligation. The ligated DNA was transformed to E. coli S17-1 λpir donor strain. The upstream-BPSS1498-downstream/pMo130 plasmid was introduced into Bpm Δhcp1 strain by biparental mating as described elsewhere [18]. The clonal selection of complemented Bpm hcp1 mutant was confirmed by PCR and sequencing at GENEWIZ.

Macrophage culture conditions and infection assays

RAW 264.7 cells (ATCC TIB-71) were grown in Gibco Dulbecco’s Modified Eagle Medium (DMEM) plus 10% heat-inactivated fetal bovine serum (Gibco), 100 U/mL penicillin, and 100 μg/mL streptomycin (Gibco) at 37°C with 5% CO2. The RAW 264.7 cells were maintained in T-75 flasks (Corning), treated using Accutase cell detachment solution (Biolegend) and seeded into 12 or 24 well plates (Corning). Bone marrow was collected from the femur and tibia of female BALB/c mice (Jackson Laboratories), RBCs lysed (Invitrogen 10x RBC Lysis buffer), and cells were added to polystyrene petri dishes (Sigma, 100mm x 20mm) containing RPMI 1640 w/ L-glutamine and HEPES (Gibco) plus 5 μM sodium pyruvate (Sigma), 100 U/mL penicillin, 100 μg/mL streptomycin (Gibco), 10% heat-inactivated fetal bovine serum (Gibco), and 25 ng/mL M-CSF (Biolegend). Cells were incubated at 37°C with 5% CO2 for 5 days with media changes on days 3 and 5. The resulting adherent cells were detached from the petri dishes using Accutase cell detachment solution (Biolegend) and seeded into 12 or 24 well plates (Corning) for further use.

RAW 264.7 cells or BMDMs were seeded at 5 × 105/well in complete DMEM or RPMI without antibiotics into 24 well-plates and allowed to adhere overnight. Bpm strains were streaked on LB agar plates, grown at 37°C for 48 h, LB broth was inoculated and grown at 37°C with shaking for 12 h. Bacterial culture was diluted to 5 × 106 CFU/mL in antibiotic free complete DMEM or RPMI and added to the cells for a multiplicity of infection (MOI) of 10. Cells were incubated with inoculum for 1 h for internalization, washed with PBS, and then media containing 500 μg/mL kanamycin was added to kill off extracellular bacteria. For bacterial enumeration, cells were washed twice with PBS to remove any extracellular bacteria, lysed with 0.1% TritonX-100, serially diluted in PBS, and plated on LB agar plates.

In vitro evaluation of apoptosis

RAW 264.7 cells were seeded at 2 x 106/well in six well plates and infected as described above and the infection was allowed to progress for 3, 6, 8, or 12 h. At the defined timepoint, cells were washed with PBS, removed from the well using Accutase, and pelleted in a PBS wash at 500 xg for 5 min. Cells were resuspended in 400 nM Apotracker Green (Biolegend) and incubated for 15 min before adding 1 mL of Zombie NIR (1/10,000 in PBS) for 5 min. Stained cells were washed in FACS buffer and fixed with 4% ultrapure formaldehyde for 48 h at 4°C before removal from Biosafety Level 3 (BSL3). Analysis was done on a BD Symphony full spectrum flow cytometer. Data were analyzed using FlowJo software.

Intranasal challenge and survival studies

Female 6–8-week-old BALB/c mice (n = 5/group) (Jackson Laboratories) were intranasally (i.n.) challenged with 3–5 LD50 Bpm K96243, Δhcp1, or Δhcp1::hcp1 in 50 μL (25 μL/nare). One LD50 is equal to 312 CFU. Infected mice were monitored for survival and weight loss for 21 days post-infection and euthanized if the animal reached the threshold for humane endpoint. On day 21 post-infection, survivors were humanely euthanized, and lungs, liver, and spleen were collected for bacterial enumeration.

TUNEL staining

Female 6–8-week-old BALB/c mice (n = 5/group) (Jackson Laboratories) were intranasally (i.n.) challenged with 3–5 LD50 Bpm K96243, Δhcp1, or Δhcp1::hcp1 in 50 μL (25 μL/nare). At 48 hours post-infection (hpi), lungs were removed and fixed in 10% buffered formalin for 48 h before removal from BSL3. Lungs were sent to the UTMB Anatomical Pathology core for paraffin embedding and mounting on slides. Mounted lung sections were deparaffinized, TUNEL stained according to the included assay protocol in the Click-iT Plus TUNEL assay Alexa Fluor 594 kit (Invitrogen), and then stained with Hoechst 33342 (ThermoFisher) to highlight nuclei. Stained slides were imaged using an Echo Revolve microscope.

In vitro polarization assay

RAW 264.7 cells were seeded at 2 x 106/well in six well plates and infected as previously described, with and the infection progressing for 8 h. At the end point, the cells were removed from the well via Accutase and washed in PBS before staining for flow cytometry. Briefly, cells were incubated with Zombie NIR (Biolegend) for 5 min in PBS, washed, and incubated with TruStain X plus (Biolegend) for 30 min followed by the extracellular antibodies (CD80, CD86, and CD163). Cells were fixed and permeabilized using Cytofix/Cytoperm (BD Biosciences) and stained for intracellular arginase-1. Stained cells were fixed with 4% ultrapure formaldehyde for 48 h at 4°C before removal from BSL3. Analysis was done on a BD Symphony full spectrum flow cytometer. Data were analyzed using FlowJo software.

Pre-polarization of RAW 264.7 cells

RAW 264.7 cells were seeded at 1 x 106 cells/well in twelve well plates and allowed to adhere overnight. After adherence, polarization media containing either 50 ng/mL IFNγ (MilliporeSigma) + 50 ng/mL LPS (MilliporeSigma) for M1 or 40 ng/mL IL-4 (MilliporeSigma) for M2 was added for 24 h. M0 cells were treated with mock polarization media containing an equitable amount of DMSO for 24 h. This protocol was validated via flow cytometry before progressing to infection assays.

Flow cytometry

Female 6–8-week-old BALB/c mice (n = 5/group) (Jackson Laboratories) were i.n. challenged with 3–5 LD50 Bpm K96243, Δhcp1, or Δhcp1::hcp1; at 48 hpi, animals were euthanized, and lungs harvested for processing. Lung tissue was cut into small pieces and dissociated via incubation for 30 min at 37°C with slight rocking in RPMI plus 0.5 mg/mL collagenase IV and 30 μg/mL DNase I. The dissociated tissue was homogenized through a 100 μm cell strainer and fibroblasts and debris was pelleted via a 60 xg centrifugation for 1 min. Supernatant was collected and RBCs were lysed for 5 min at RT. Following washes, pulmonary cells were adjusted to 1 x 106 cells and stained using the reagents in Table 1. Briefly, cells were incubated with Zombie NIR (Biolegend) for 5 min in PBS, washed, and incubated with TruStain X plus (Biolegend) for 30 min followed by the extracellular antibodies (Table 1). Cells were fixed and permeabilized using Cytofix/Cytoperm (BD Biosciences) and stained for intracellular markers. Fully stained cells were resuspended in 4% ultrapure formaldehyde in PBS for 48 h in accordance with the inactivation protocol approved by UTMB Department of Biosafety before removal from BSL3 laboratory for analysis via BD Symphony full spectrum flow cytometer. Data were analyzed using FlowJo software.

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Table 1. Flow cytometry antibodies and reagents.

https://doi.org/10.1371/journal.pntd.0012585.t001

Lung pathology

Lungs were collected from mice after humane euthanasia 48 h post-infection and fixed in 10% formalin for 48 h. Formalin fixed lung samples were submitted to the UTMB Anatomical Pathology core for paraffin embedding, mounting, and H&E staining. Slides were imaged using an Olympus BX51 microscope.

Statistical analysis

All statistical analysis was done using GraphPad Prism software (v9.0). P-values of < 0.05 are considered statistically significant. Survival differences were assessed via Kaplan-Meier survival curve followed by a log-rank test. An ordinary one-way ANOVA followed by Tukey’s post hoc test was used to analyze differences in intracellular replication and flow cytometry populations.

Results

The T6SS is dispensable for survival inside of macrophages

To begin understanding how T6SS activity affects apoptosis and polarization, we first established how our T6SS mutant, Δhcp1 (BPSS1498), replicated inside of macrophages. Hcp1 is the most prevalent structural protein of the T6SS and hexamerizes to form the inner sheath of the injectosome, deletion of hcp1 ablates MNGC formation while attenuating the bacterium during in vivo infections [13,15]. We chose to evaluate intracellular survival in two macrophage models: RAW 264.7 cells and BALB/c bone marrow-derived macrophages (BMDMs). As RAW 264.7 cells were initially derived from BALB/c mice, we chose the same background for the primary BMDMs. Unlike the previously characterized regulatory mutant ΔbicA [31], the Δhcp1 strain did not display an intracellular survival defect in either macrophage model (Fig 1A and 1B). The Δhcp1 strain does appear to survive significantly better than the WT or complemented Δhcp1::hcp1 strains in the RAW 264.7 model but the mechanism behind this phenotype remains unclear (Fig 1A). Together, these data suggest that although the T6SS is critical for virulence, it is dispensable for replication within macrophages.

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Fig 1. Intracellular survival of Bpm in macrophages is not T6SS-dependent.

RAW 264.7 cells (A) or BALB/c BMDMs (B), were infected at an MOI of 10 with Bpm K96243 WT, Δhcp1, or Δhcp1::hcp1 and bacteria enumerated at 3, 6, and 12 hpi to assess intracellular replication. Bars represent an average of two independent experiments performed in triplicate ± SD. Significant differences were assessed via one-way ANOVA followed by Tukey’s multiple comparison test. p < 0.05 *, p < 0.01**, p < 0.005***, p < 0.0001****.

https://doi.org/10.1371/journal.pntd.0012585.g001

T6SS activity exacerbates apoptosis in macrophages and during in vivo infection

To evaluate apoptosis, we utilized flow cytometry to measure the externalization of phosphatidylserine (PS) via Apotracker dye paired with a live/dead (L/D) viability dye. This allows for the differentiation between apoptotic death (Apotracker +, L/D +/-) and necrotic forms of cell death (Apotracker -, L/D+). We performed an infection time course in RAW 264.7 cells and measured the percentage of macrophages that were apoptotic (Apotracker +) at 3-, 6-, 8-, and 12-hours post infection (hpi) (Fig 2A–2E). Beginning at 6 hpi, WT infection results in significantly more apoptotic cells (Fig 2B and 2E), and by 8 and 12 hpi all infection groups had elevated levels of apoptosis events (Fig 2C–2E). Although Δhcp1 had increased apoptosis compared to mock infected cells, WT and Δhcp1::hcp1 demonstrated a dramatic increase over Δhcp1 at both 8 and 12 hpi. Considerable amounts of intracellular replication results in a robust apoptotic response, however, T6SS activity exacerbates apoptosis in macrophages.

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Fig 2. Functional T6SS exacerbates apoptosis in macrophages during infection.

RAW 264.7 cells were infected at an MOI of 10 with Bpm K96243 WT, Δhcp1, Δhcp1::hcp1, or mock infected and collected at 3 (A), 6 (B), 8 (C), or 12 (D) hpi. Cells were evaluated for apoptosis via staining with Apotracker Green and Zombie NIR (Live/Dead). Percentage of apoptotic cells were counted as Apotracker+ and L/D+/- (Q2 & Q3) (E). Bars represent an average of three independent experiments performed in duplicate ± SD. Significant differences were assessed via one-way ANOVA followed by Tukey’s multiple comparison test. p < 0.05 *, p < 0.01**, p < 0.005***, p < 0.0001****.

https://doi.org/10.1371/journal.pntd.0012585.g002

With the T6SS-exacerbated apoptosis phenotype established in vitro, we wanted to assess apoptosis in murine lungs during pulmonary melioidosis. BALB/c mice intranasally challenged with Δhcp1 demonstrated 100% survival, confirming what has previously been reported [13]. As expected, WT and Δhcp1::hcp1 challenged groups saw complete lethality (Fig 3A). The Δhcp1 infected mice exhibited minimal weight loss and were mostly clear of persistent infection on day 21 post infection (Fig 3B and 3C). We selected 48 h post infection to assess pulmonary apoptosis due to the disparity in disease severity observed between WT/Δhcp1::hcp1 and Δhcp1 at this time point. As such, another set of BALB/c mice were challenged, lungs were removed and TUNEL staining was performed to detect apoptosis. Representative images of WT and Δhcp1::hcp1 infected lungs exhibited intense TUNEL signal, while Δhcp1 infected lungs displayed intermediate amounts of staining (Fig 4). This non-quantitative assay recapitulates what was seen in vitro (Fig 2A–2E) with the Δhcp1 strain eliciting a small to moderate amount of apoptosis, while an active T6SS triggers large scale apoptosis. However, it should be noted that the in vivo analysis was performed with a single replicate which limits the veracity until it can be replicated.

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Fig 3. The Δhcp1 strain is attenuated in the intranasal melioidosis model.

BALB/c mice (n = 5/group) were intranasally challenged with 3–5 LD50 of Bpm K96243 WT, Δhcp1, or Δhcp1::hcp1 (1 LD50 ~ 312 CFU) and monitored for 21 days post infection for survival (A) and weight loss (B). Animals were euthanized once the humane endpoint threshold was reached. On day 21 post infection, Δhcp1 survivors were euthanized and lungs, liver, and spleen were homogenized for bacterial enumeration (C). Error bars in (B) represent SEM and lines in (C) represent median value.

https://doi.org/10.1371/journal.pntd.0012585.g003

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Fig 4. Pulmonary apoptosis mirrors in vitro Bpm T6SS-mediated exacerbation.

BALB/c mice (n = 5/group) were intranasally challenged with 3–5 LD50 of Bpm K96243 WT, Δhcp1, or Δhcp1::hcp1 (1 LD50 ~ 312 CFU) and at 48 h post-infection lungs were harvested, formalin fixed, and mounted on slides. Sections were stained with TUNEL (red) and Hoechst 33342 (blue) to evaluate apoptosis in the lungs.

https://doi.org/10.1371/journal.pntd.0012585.g004

Bpm infection triggers in vitro macrophage polarization independent of T6SS

We previously established that Bpm elicits both M1 and M2 polarization in vivo but the M2 population was BicA-dependent [31]. As BicA is involved in T3SS-mediated virulence and thus the bicA mutant has an intracellular survival defect, we sought to understand if the differential polarization is dependent on just intracellular survival or requires a functional T6SS. Therefore, RAW 264.7 cells were infected, and at 8 hpi, were stained with markers for polarization: CD80, CD86 (M1) and CD163, Arginase-1 (M2). In this assay, cell populations that are CD80+ are being considered M1 while populations that are Arg-1+ are M2. Surprisingly, there was no significant difference in M2 polarization across the infection groups (Fig 5A and 5C) with only WT Bpm demonstrating an increase over mock infected cells. WT did trend higher than Δhcp1 on average but due to variability in the samples, this was not significant. All infection groups generated a consistent and robust M1 response (Fig 5B and 5D). The mock infected group did exhibit a sizable amount of residual CD80 staining, however, there was a distinct shift in intensity upon infection. This result suggests that infection with an intracellular survival competent strain is enough to trigger both pro-inflammatory and alternative effector functions in macrophages and does not require a functional T6SS.

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Fig 5. The Bpm T6SS does not contribute to differential macrophage polarization in vitro.

RAW 264.7 cells were infected at an MOI of 10 with Bpm K96243 WT, Δhcp1, Δhcp1::hcp1, or mock infected and collected at 8 hpi. Cells were processed, stained, and evaluated for expression of M2 (A & C) and M1 (B & D) markers. Cells that were Arg-1+ are denoted as M2 while CD80+ cells are M1. Bars represent an average of three independent experiments performed in duplicate ± SD. Significant differences were assessed via one-way ANOVA followed by Tukey’s multiple comparison test. ns; non-significance, p < 0.05 *, p < 0.01**, p < 0.005***, p < 0.0001****.

https://doi.org/10.1371/journal.pntd.0012585.g005

M2 polarization promotes intracellular survival of Bpm

The advantages and disadvantages of macrophage polarization for Bpm are unclear, as different bacterial pathogens skew polarization one way or the other to promote infection [33]. To address this dichotomy, RAW 264.7 cells were pre-polarized to M1 (IFNγ + LPS) or M2 (IL-4) prior to infection and intracellular survival was assessed and compared to an M0 control (Fig 6A and 6B). M2 macrophages had decreased relative phagocytic capacity compared to M0 (Fig 6A) and increased intracellular survival at 3 hpi compared to both M1- and M0-polarized cells (Fig 6B). Interestingly, M1 polarization showed no significant advantage on bacterial clearance as compared to M0. These data further suggest that M2 skewing by Bpm might be offering an advantage during infection and is potentially an example of hijacking the host response to promote pathogenesis and replication.

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Fig 6. M2 polarization promotes Bpm intracellular survival.

RAW 264.7 cells were pre-polarized with IFNγ + LPS (M1), IL-4 (M2), or media control (M0) and infected at an MOI of 10 with Bpm K96243. Phagocytic capacity of M1 and M2 macrophages was compared to M0, and relative phagocytic capacity was measured after 1 h (A). Intracellular survival was evaluated at 3 hpi (B). Percent intracellular survival was calculated by dividing the output at 3 hpi by the mean number of bacteria phagocytosed after 1 h for each group. Bars represent an average of two independent experiments performed in triplicate ± SD. Significant differences were assessed via one-way ANOVA followed by Tukey’s multiple comparison test. ns; non-significance, p < 0.05 *., p < 0.01**, p < 0.005***, p < 0.0001****.

https://doi.org/10.1371/journal.pntd.0012585.g006

Bpm infection triggers in vivo macrophage polarization independent of T6SS

After examining the relationship between the T6SS and macrophage polarization in vitro, we assessed the role of the T6SS in macrophage polarization in vivo. BALB/c mice were intranasally challenged with WT, Δhcp1, or Δhcp1::hcp1 and at 48 hpi lungs were removed and processed for flow cytometry. We devised a comprehensive panel (Table 1) and gating strategy adapted from [34] and previously utilized in [31] (S1 Fig) to interrogate macrophage activity within the lungs. We are denoting macrophages as cells that are MHCII+ and F4/80+ after being filtered through gating and cells within that population as M1-like (CD80+ and CD86+/-) or M2-like (Arginase-1+ and CD163+). We found that although Δhcp1 is drastically attenuated in vivo, there was no difference in macrophage recruitment to the lungs during infection (Fig 7A and 7B). When examining the activation states of pulmonary macrophages, we found no difference in M1-like or M2-like macrophages (Fig 7C and 7D), however, there was a distinct downward shift in the intensity of CD80 staining in Δhcp1 (Fig 7A).

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Fig 7. M2 polarization is not T6SS-dependent in vivo.

BALB/c mice (n = 5/group) were intranasally challenged with 3–5 LD50 of Bpm K96243 WT, Δhcp1, or Δhcp1::hcp1 (1 LD50 ~ 312 CFU) and at 48 hpi lungs were harvested and processed for flow cytometry. A comprehensive gating strategy (S1 Fig) was used to filter and evaluate macrophages within the lungs (A). Total pulmonary macrophages: MHCII+ F4/80+ (B), M1: CD80+ CD86+/- (C), and M2: Arg-1+ CD163+ (D) were assessed. Significant differences were assessed via one-way ANOVA followed by Tukey’s multiple comparison test. ns; non-significance. p < 0.05 *, p < 0.01**, p < 0.005***, p < 0.0001****.

https://doi.org/10.1371/journal.pntd.0012585.g007

Inflammatory nodules predictive of M2 polarization but not T6SS-dependent

Previously, we observed that the presence of an M2 macrophage population in the lungs correlated with distinct inflammatory nodules [31]. We examined H & E-stained lungs from infected BALB/c mice infected at 48 hpi for the presence or absence of this pathological feature. We found that all strains generated inflammatory nodules (Fig 8). However, even though the Δhcp1 strain generated these nodules, a non-quantitative analysis indicated that they were smaller and less numerous compared to WT and Δhcp1::hcp1 strains. The WT and Δhcp1::hcp1-associated nodules appear to have more cellular debris compared to Δhcp1 but the cellular content of each nodule is currently unknown. Although this phenotype was consistent across all animals evaluated, this study was only performed one time. We hypothesize that the nodules are likely primary replication hot spots for Bpm within the lungs of infected animals.

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Fig 8. Inflammatory nodule formation is not contingent on T6SS.

BALB/c mice (n = 5/group) were intranasally challenged with 3–5 LD50 of Bpm K96243 WT, Δhcp1, or Δhcp1::hcp1 (1 LD50 ~ 312 CFU) and at 48 hpi lungs were harvested, formalin fixed, and mounted on slides before hematoxylin and eosin staining. Representative images were taken using a 10x microscope objective.

https://doi.org/10.1371/journal.pntd.0012585.g008

Discussion

Melioidosis is a neglected tropical disease that is a looming global public health threat [2,3]. As a facultative intracellular pathogen, Bpm deploys an arsenal of virulence factors to successfully survive and replicate within the host cells [1,4]. One critical virulence factor is the T6SS, an injectosome apparatus that Bpm utilizes to fuse host membranes and generate MNGCs. Formation of MNGC is the keystone pathogenesis event and T6SS mutants are highly attenuated in vivo [13]. The mechanisms of T6SS-mediated pathogenesis and the impact that MNGC formation has on the host response is currently poorly understood. Previously, our laboratory sought to establish the impact of the T6SS on host response via dual RNA-seq in our in vitro model of GI infection [18]. This analysis revealed that in the absence of the T6SS, there is substantial differential expression in pathways that are involved in inflammation, cell death, and apoptosis. The differential expression of inflammation pathways was validated by demonstrating that there is a T6SS-dependent blockage of NFκB activation, even after priming with TNFα. This work was done in primary murine intestinal epithelial cells, and it is currently unclear how this model translates to other models of infection. Therefore, to investigate how the T6SS participates in the inflammation process and cell death, we chose the in vitro macrophage system and the intranasal infection model to perform in vivo studies. Respiratory involvement is one of the most common clinical presentations of melioidosis and can progress into necrotizing pneumonia, making the intranasal model of infection particularly useful and relevant to study [1]. Macrophages are a primary replicative niche for Bpm and are omnipresent in all host tissues, circulating or as tissue resident sentinels [11,35]. They are an attractive target for Bpm to manipulate as they are integral in the immune response to infection, and we have previously shown that Bpm is capable of differentially activating macrophages [31].

We first needed to establish how the T6SS affects intracellular survival within macrophages, as other secretion system mutants exhibit intracellular defects [18,31,36]. We chose two different macrophage models to evaluate intracellular survival: RAW 264.7 cells, an immortalized murine macrophage cell line, and bone marrow-derived macrophages harvested from BALB/c mice. The RAW 264.7 cells were initially collected from a BALB/c background, so we selected the same genetic background for our primary model. We found that, unlike the T3SS, the T6SS is dispensable for replication within both immortalized and primary macrophages (Fig 1A and 1B). There was a significant increase in intracellular survival of Δhcp1 in RAW 264.7 cells that was not observed in the primary BMDMs. One possibility for this result is that RAW 264.7 cells lack the inflammasome adapter protein, ASC, which limits the ability of the NLRP3 inflammasome. However, it has been demonstrated that BMDMs lacking ASC do not facilitate increased replication of Bpm [37].

We then evaluated the contribution of the T6SS to apoptosis of RAW 264.7 cells during infection. We found that although Δhcp1 triggered increased apoptosis as compared to the mock infected macrophages, WT and Δhcp1::hcp1 infected RAW cells exhibited remarkably high proportions of apoptotic cells (Fig 2A–2E). The increased viability of Δhcp1 at 12 hpi (Fig 2D and 2E) helps explain the increased intracellular survival in Fig 1A as dead and dying cells release the bacteria into the media containing kanamycin. This increase in apoptosis between 6 and 8 hpi correlates with the historical timeline of MNGC formation and thus is likely the driving force behind the rise in apoptosis. As this is an in vitro system, we wanted to evaluate the consistency of this phenotype in vivo using an intranasal challenge model using BALB/c mice. The attenuation of Δhcp1 has been previously documented [13], however, we needed to confirm the restoration of Δhcp1::hcp1 in vivo. We found that Δhcp1::hcp1 recapitulates WT virulence during intranasal challenge (Fig 3A and 3B) while Δhcp1 remained attenuated. At day 21 post infection, the Δhcp1 survivors had predominantly cleared the infection, with only a couple of animals harboring small numbers of bacteria (Fig 3C). We chose to evaluate pulmonary apoptosis at 48 hpi as there is a distinct disparity in weight loss (a predictor of disease severity) between Δhcp1 and WT/Δhcp1::hcp1 (Fig 3B). TUNEL staining was used to evaluate apoptosis, and paraffin embedded lung sections from 48 hpi were probed with TUNEL and a DNA counter stain and imaged to visualize relative amounts of apoptosis within the lungs (Fig 4). Much like the in vitro assay, Δhcp1 elicits lesser amounts of apoptosis but WT and Δhcp1::hcp1 trigger much higher and more widely distributed TUNEL signal (Fig 4). The in vitro and in vivo apoptosis phenotypes being nearly identical suggests that T6SS-mediated exacerbation of apoptosis is not a macrophage specific phenomenon but is a common mechanism across phagocytic and non-phagocytic cell types. There are two signaling pathways that converge on caspase-3 activation and apoptosis; the extrinsic pathway, that is initiated through an external death receptor, and the intrinsic pathway, that is triggered by internal cellular damage and release of specific mitochondrial molecules [20]. Intracellular damage, like that caused by cell fusion and ineffective ROS, is an intrinsic lethal stimulus that triggers caspase-9 mediated apoptosis. Common ligands for the extrinsic pathway are TNFα, Fas-L, and TRAIL, Bpm has been shown to shut down NFκB which is a driver of TNFα production. The expression of these specific molecules is unknown. However, it has also been demonstrated that splenic monocytes/macrophages from heavily colonized mice produce increased levels of TNFα and that correlated with increasing severity of pyogranulomatous lesions on the spleen [38]. There is established crosstalk between the two pathways, specifically, caspase-8 cleavage of BID leading to cytochrome C release from the mitochondria and activation of the intrinsic “apoptosome”, a multimeric structure that acts as a scaffold for caspase-9 activity [39,40]. Future studies are needed to determine the signaling cascade that is involved in Bpm-mediated apoptosis and if there is a cell type specific contribution to the microenvironment that influences this phenotype.

We next evaluated how the T6SS affects the base inflammation state of macrophages in vitro via assessing the expression of polarization markers on infected RAW 264.7 cells. For this assay, an M1 macrophage is denoted as CD80+ and CD86+/- while an M2 macrophage is Arg-1+ and CD163+/-. The 8 hpi time point was selected due to the difference in the apoptosis phenotype between Δhcp1 and WT/Δhcp1::hcp1 and, although not directly measured, comparable intracellular survival. Although WT exhibited a significant increase in M2 macrophages over mock, there was no difference across the infection groups (Fig 5A and 5C). This phenotype was highly variable in the infection groups, especially within cells infected with Δhcp1. On the other hand, M1 polarization was highly consistent across infection groups and all strains elicited a highly significant increase over mock (Fig 5B and 5D). Mock infected RAW 264.7 cells exhibited moderate basal CD80 expression, however, upon infection there was a distinct shift in intensity that is indicative of M1 polarization. The near complete M1 polarization tells us that the pro-inflammatory activation is the primary response to infection and that is not dependent on T6SS activity. The M2 response appears to be less evident and a secondary reaction to infection. Such activation state is highly variable and potentially is a response to the apoptosis that is occurring during infection. Traditionally, apoptotic corpses are cleared by phagocytes and anti-inflammatory molecules are released to avoid unnecessary inflammatory damage and to maintain homeostasis, a process that is called efferocytosis [21,41]. M2 polarization by Bpm might be incidental, an indirect response to apoptosis, but it still could be beneficial to the pathogen by creating a more permissive environment for replication. To address the question of whether macrophage polarization is beneficial to Bpm, we pre-polarized RAW 264.7 cells and infected with WT Bpm to evaluate intracellular survival. The phagocytic capacity of M1 and M2 macrophages was compared to mock treated M0 macrophages, and we found that M2s have a decreased phagocytic capacity compared to M0s (Fig 6A). Intracellular survival was assessed at 3 hpi and although M2s had a decreased phagocytic capacity, they facilitated increased intracellular survival compared to M1s and M0s (Fig 6B). This suggests that M2 polarization is a beneficial replication environment for Bpm and M1 polarization is inconsequential during infection. It should be noted that this phenotype is within the context of an in vitro infection of monocultured cells, and within the in vivo environment, this is more complex with multiple cell types contributing to the immune landscape during infection.

To incorporate the complexities of a multicellular immune system, we evaluated macrophage polarization in the lungs of infected mice at 48 hpi. Lungs were collected, processed, and total pulmonary macrophages and M1/M2 polarization within that population of pulmonary macrophages were evaluated (Fig 7A). We found that there was no difference in total macrophages present in the lungs at 48 hpi (Fig 7B). When we evaluated the expression of our polarization markers, we found no differences in M1 (Fig 7C) or M2 (Fig 7D) macrophages across the infection groups. This matches what was observed in vitro (Fig 5A–5D). We previously reported that WT Bpm elicited an M2 population along with inflammatory nodules within the lungs [31] so we evaluated lung pathology at 48 hpi to determine if these inflammatory nodules were T6SS-depedendent. We found that, although they were smaller and less numerous, Δhcp1 infection resulted in the formation of inflammatory nodules (Fig 8). The structure of Δhcp1-associated nodules was distinct from WT/Δhcp1::hcp1, lacking the cellular debris and segmented appearance that has been observed numerous times in WT infections, both in murine and human infections [31,38,42]. This confirms that M2 macrophages are associated with these inflammatory nodules; however, a functional T6SS might contribute to the complexity of these nodules.

Apoptotic cell death has been described as “immunologically silent” because it does not trigger a pro-inflammatory response from the phagocytic cells tasked with clearing the corpse. It has long been hypothesized that Bpm uses MNGC formation as a mechanism of cell-to-cell spread to prevent interacting with the external milieu and apoptosis is simply a response to MNGC formation. Our data suggested that apoptosis is a purposeful immune evasion mechanism that Bpm uses to avoid triggering an effective pathogen clearing response. The robust M1 response, which occurs both in vitro and in vivo, is not effective at bacterial clearance at first glance. There is a possibility that in the absence of cell-to-cell spread and apoptosis, this M1 response is productive and can eliminate bacteria that have limited cell-to-cell mobility (Δhcp1). The correlation of M2 macrophages and the inflammatory nodule pathology might suggest that the M2 polarization is a response to tissue damage caused by the pro-inflammatory response and bacterial replication. More work needs to be done understanding which M2 subtype(s) are present inside the lung as that can shed light on their activity, as well as positioning where both M1 and M2 macrophages are in relation to the inflammatory nodules which might be working as replication hotspots.

In summary, we explored the contribution of the Bpm T6SS to inflammation and cell death during infection. We demonstrated that the T6SS participates as the driver of apoptosis in macrophages and within the lung but does not result in differential macrophage polarization compared to WT. The increase in inflammatory nodule severity correlated with apoptosis in the lungs, suggesting that triggering apoptosis is advantageous for pathogenesis. There are limitations to these results, namely, only performing the pulmonary TUNEL staining and pathology observations in a single replicate. The results were consistent across all animals evaluated; however, this will need to be replicated to fully support our hypothesis.

Supporting information

S1 Fig. Flow cytometry gating strategy.

Gating strategy used to filter macrophages.

https://doi.org/10.1371/journal.pntd.0012585.s001

(PDF)

Acknowledgments

We would like to thank Meredith Weglarz in the UTMB Flow Cytometry Core for the expertise and help in designing and implementing the flow cytometry experiments. We would also like to thank Alex Badten for his help during animal experiments, Dr. Alison Coady for allowing us to utilize her Echo Revolve microscope, and Paige Diaz for training and troubleshooting on the Echo microscope.

References

  1. 1. Wiersinga WJ, Virk HS, Torres AG, Currie BJ, Peacock SJ, Dance DAB, et al. Melioidosis. Nat Rev Dis Primers. 2018;4:17107. pmid:29388572
  2. 2. Savelkoel J, Dance DAB, Currie BJ, Limmathurotsakul D, Wiersinga WJ. A call to action: time to recognise melioidosis as a neglected tropical disease. Lancet Infect Dis. 2022;22:e176–e82. pmid:34953519
  3. 3. Limmathurotsakul D, Golding N, Dance DA, Messina JP, Pigott D M, Moyes CL, et al. Predicted global distribution of Burkholderia pseudomallei and burden of melioidosis. Nat Microbiol. 2016;1:15008.
  4. 4. Meumann EM, Limmathurotsakul D, Dunachie SJ, Wiersinga WJ, Currie BJ. Burkholderia pseudomallei and melioidosis. Nat Rev Microbiol. 2024;22:155–69.
  5. 5. Rolim DB, Lima RXR, Ribeiro AKC, Colares RM, Lima LDQ, Rodríguez-Morales AJ, et al. Melioidosis in South America. Trop Med Infect Dis. 2018;3:60. pmid:30274456
  6. 6. Hall CM, Jaramillo S, Jimenez R, Stone NE, Centner H, Busch JD, et al. Burkholderia pseudomallei, the causative agent of melioidosis, is rare but ecologically established and widely dispersed in the environment in Puerto Rico. PLoS Negl Trop Dis. 2019;13:e0007727.
  7. 7. Cossaboom CM, Marinova-Petkova A, Strysko J, Rodriguez G, Maness T, Ocampo J, et al. Melioidosis in a Resident of Texas with No Recent Travel History, United States. Emerg Infect Dis. 2020;26:1295–9. pmid:32442394
  8. 8. Torres AG. The public health significance of finding autochthonous melioidosis cases in the continental United States. PLoS Negl Trop Dis. 2023;17:e0011550. pmid:37619236
  9. 9. Garg R, et al., Shaw T, Vandana KE, Magazine R, Mukhopadhyay C. Melioidosis In Suspected Recurrent Tuberculosis: A disease in disguise. J Infect Dev Ctries. 2020;14:312–6. pmid:32235093
  10. 10. Ninan F, Mishra AK, John AO, Iyadurai R. Splenic granuloma: Melioidosis or Tuberculosis? J Family Med Prim Care. 2018;7:271–3. pmid:29915776
  11. 11. Jones AL, Beveridge TJ, Woods DE. Intracellular survival of Burkholderia pseudomallei. Infect Immun. 1996;64:782–90.
  12. 12. Whiteley L, Meffert T, Haug M, Weidenmaier C, Hopf V, Bitschar K, et al. Entry, Intracellular Survival, and Multinucleated-Giant-Cell-Forming Activity of Burkholderia pseudomallei in Human Primary Phagocytic and Nonphagocytic Cells. Infect Immun. 2017;85:e00468–17.
  13. 13. Burtnick M.N., et al. The cluster 1 type VI secretion system is a major virulence determinant in Burkholderia pseudomallei. Infect Immun. 2011;79:1512–25. pmid:21300775
  14. 14. Toesca IJ, French CT, Miller JF. The Type VI secretion system spike protein VgrG5 mediates membrane fusion during intercellular spread by pseudomallei group Burkholderia species. Infect Immun. 2014;82:1436–44.
  15. 15. Lennings J, West TE, Schwarz S. The Burkholderia Type VI Secretion System 5: Composition, Regulation and Role in Virulence. Frontiers Microbiol. 2019;10:3339.
  16. 16. Coulthurst S. The Type VI secretion system: a versatile bacterial weapon. Microbiology (Reading). 2019;165:503–15. pmid:30893029
  17. 17. Hopf V, Göhler A, Eske-Pogodda K, Bast A, Steinmetz I, Breitbach K. BPSS1504, a cluster 1 type VI secretion gene, is involved in intracellular survival and virulence of Burkholderia pseudomallei. Infect Immun. 2014;82:2006–15.
  18. 18. Sanchez-Villamil JI, Tapia D, Khakhum N, Widen SG, Torres AG. Dual RNA-seq reveals a type 6 secretion system-dependent blockage of TNF-α signaling and BicA as a Burkholderia pseudomallei virulence factor important during gastrointestinal infection. Gut Microbes. 2022;14:2111950.
  19. 19. Sanchez-Villamil JI, Tapia D, Borlee GI, Borlee BR, Walker DH, Torres AG. Burkholderia pseudomallei as an Enteric Pathogen: Identification of Virulence Factors Mediating Gastrointestinal Infection. Infect Immun. 2020;89:e00654–20. pmid:33106293
  20. 20. Elmore S. Apoptosis: a review of programmed cell death. Toxicol Pathol. 2007;35:495–516. pmid:17562483
  21. 21. Mohammad-Rafiei F, Moadab F, Mahmoudi A, Navashenaq JG, Gheibihayat SM. Efferocytosis: a double-edged sword in microbial immunity. Arch Microbiol. 2023;205:370. pmid:37925389
  22. 22. Krakauer T. Living dangerously: Burkholderia pseudomallei modulates phagocyte cell death to survive. Med Hypotheses. 2018;121:64–9.
  23. 23. Mariappan V, Vellasamy KM, Barathan M, Girija ASS, Shankar EM, Vadivelu J. Hijacking of the Host’s Immune Surveillance Radars by Burkholderia pseudomallei. Front Immunol. 2021;12:718719.
  24. 24. Jitprasutwit N, Rungruengkitkun A, Lohitthai S, Reamtong O, Indrawattana N, Sookrung N, et al. In Vitro Roles of Burkholderia Intracellular Motility A (BimA) in Infection of Human Neuroblastoma Cell Line. Microbiol Spectr 2023;11:e01320–23.
  25. 25. Place DE, Christgen S, Tuladhar S, Vogel P, Malireddi RKS, Kanneganti TD. Hierarchical Cell Death Program Disrupts the Intracellular Niche Required for Burkholderia thailandensis Pathogenesis. mBio. 2021;12:e0105921.
  26. 26. Suparak S, Kespichayawattana W, Haque A, Easton A, Damnin S, Lertmemongkolchai G, et al. Multinucleated giant cell formation and apoptosis in infected host cells is mediated by Burkholderia pseudomallei type III secretion protein BipB. J Bacteriol. 2005;187:6556–60.
  27. 27. Kespichayawattana W, Rattanachetkul S, Wanun T, Utaisincharoen P, Sirisinha S. Burkholderia pseudomallei induces cell fusion and actin-associated membrane protrusion: a possible mechanism for cell-to-cell spreading. Infect Immun. 2000;68:5377–84.
  28. 28. Vellasamy KM, Mariappan V, Shankar EM, Vadivelu J. Burkholderia pseudomallei Differentially Regulates Host Innate Immune Response Genes for Intracellular Survival in Lung Epithelial Cells. PLoS Negl Trop Dis. 2016;10:e0004730.
  29. 29. Wang Y, Kanneganti T-D. From pyroptosis, apoptosis and necroptosis to PANoptosis: A mechanistic compendium of programmed cell death pathways. Comput Struct Biotechnol J. 2021;19:4641–57. pmid:34504660
  30. 30. Murray PJ. Macrophage Polarization. Annu Rev Physiol. 2017;79:541–66. pmid:27813830
  31. 31. Stockton JL, Khakhum N, Stevenson HL, Torres AG. Burkholderia pseudomallei BicA protein promotes pathogenicity in macrophages by regulating invasion, intracellular survival, and virulence. mSphere. 2023;8:e0037823.
  32. 32. Khakhum N, Bharaj P, Myers JN, Tapia D, Kilgore PB, Ross BN, et al. Burkholderia pseudomallei ΔtonB Δhcp1 Live Attenuated Vaccine Strain Elicits Full Protective Immunity against Aerosolized Melioidosis Infection. mSphere. 2019;4:e00570–18.
  33. 33. Thiriot JD, Martinez-Martinez YB, Endsley JJ, Torres AG. Hacking the host: exploitation of macrophage polarization by intracellular bacterial pathogens. Pathog Dis. 2020;78:ftaa009. pmid:32068828
  34. 34. Misharin AV, Morales-Nebreda L, Mutlu GM, Budinger GR, Perlman H. Flow cytometric analysis of macrophages and dendritic cell subsets in the mouse lung. Am J Respir Cell Mol Biol. 2013;49:503–10. pmid:23672262
  35. 35. Zhang C, Yang M, Ericsson AC. Function of Macrophages in Disease: Current Understanding on Molecular Mechanisms. Front Immunol. 2021;12:620510. pmid:33763066
  36. 36. Burtnick MN, Brett PJ, Nair V, Warawa JM, Woods DE, Gherardini FC. Burkholderia pseudomallei type III secretion system mutants exhibit delayed vacuolar escape phenotypes in RAW 264.7 murine macrophages. Infect Immun. 2008;76:2991–3000. pmid:18443088
  37. 37. Ceballos-Olvera I, Sahoo M, Miller MA, Del Barrio L, Re F. Inflammasome-dependent pyroptosis and IL-18 protect against Burkholderia pseudomallei lung infection while IL-1β is deleterious. PLoS Pathog. 2011;7:e1002452. pmid:22241982
  38. 38. Amemiya K, Dankmeyer JL, Bearss JJ, Zeng X, Stonier SW, Soffler C, et al. Dysregulation of TNF-α and IFN-γ expression is a common host immune response in a chronically infected mouse model of melioidosis when comparing multiple human strains of Burkholderia pseudomallei. BMC Immunol. 2020;21:5.
  39. 39. Li H, Zhu H, Xu CJ, Yuan J. Cleavage of BID by caspase 8 mediates the mitochondrial damage in the Fas pathway of apoptosis. Cell. 1998;94:491–501. pmid:9727492
  40. 40. Luo X, Budihardjo I, Zou H, Slaughter C, Wang X. Bid, a Bcl2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of cell surface death receptors. Cell. 1998;94:481–90. pmid:9727491
  41. 41. Doran AC, Yurdagul A, Tabas I. Efferocytosis in health and disease. Nat Rev Immunol. 2020;20:254–67. pmid:31822793
  42. 42. Savelkoel J, Tiemensma M, Birnie E, Wiersinga WJ, Currie BJ, Roelofs JJTH. A Graphical Overview of the Histopathology of Human Melioidosis: A Case Series. Open Forum Infect Dis. 2023;10:ofad367. pmid:37547853