https://orcid.org/0000-0002-2747-5618
Figures
Abstract
The immune response to Mycobacterium tuberculosis infection determines tuberculosis disease outcomes, yet we have an incomplete understanding of what immune factors contribute to a protective immune response. Neutrophilic inflammation has been associated with poor disease prognosis in humans and in animal models during M. tuberculosis infection and, therefore, must be tightly regulated. ATG5 is an essential autophagy protein that is required in innate immune cells to control neutrophil-dominated inflammation and promote survival during M. tuberculosis infection; however, the mechanistic basis for how ATG5 regulates neutrophil recruitment is unknown. To interrogate what innate immune cells require ATG5 to control neutrophil recruitment during M. tuberculosis infection, we used different mouse strains that conditionally delete Atg5 in specific cell types. We found that ATG5 is required in CD11c+ cells (lung macrophages and dendritic cells) to control the production of proinflammatory cytokines and chemokines during M. tuberculosis infection, which would otherwise promote neutrophil recruitment. This role for ATG5 is autophagy dependent, but independent of mitophagy, LC3-associated phagocytosis, and inflammasome activation, which are the most well-characterized ways that autophagy proteins regulate inflammation. In addition to the increased proinflammatory cytokine production from macrophages during M. tuberculosis infection, loss of ATG5 in innate immune cells also results in an early induction of TH17 responses. Despite prior published in vitro cell culture experiments supporting a role for autophagy in controlling M. tuberculosis replication in macrophages, the effects of autophagy on inflammatory responses occur without changes in M. tuberculosis burden in macrophages. These findings reveal new roles for autophagy proteins in lung resident macrophages and dendritic cells that are required to suppress inflammatory responses that are associated with poor control of M. tuberculosis infection.
Citation: Kinsella RL, Kimmey JM, Smirnov A, Woodson R, Gaggioli MR, Chavez SM, et al. (2023) Autophagy prevents early proinflammatory responses and neutrophil recruitment during Mycobacterium tuberculosis infection without affecting pathogen burden in macrophages. PLoS Biol 21(6): e3002159. https://doi.org/10.1371/journal.pbio.3002159
Academic Editor: Hans-Uwe Simon, University of Bern, SWITZERLAND
Received: April 12, 2023; Accepted: May 11, 2023; Published: June 15, 2023
Copyright: © 2023 Kinsella 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 paper and its Supporting Information files.
Funding: This work was supported by NIH grants R01 AI132697 and U19 AI142784, a Burroughs Wellcome Fund Investigators in the Pathogenesis of Infectious Disease Award ( 1016714), and the Philip and Sima Needleman Center for Autophagy Therapeutics and Research to C.L.S., a Potts Memorial Foundation postdoctoral fellowship to R.L.K., and a National Science Foundation Graduate Research Fellowship DGE-1143954 and the NIGMS Cell and Molecular Biology Training Grant GM007067 to J.M.K. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. None of the authors received a salary from any of the funders.
Competing interests: The authors have declared that no competing interest exist.
Abbreviations: BMDM, bone marrow-derived macrophage; DC, dendritic cell; dpi, days postinfection; hpi, hours postinfection; LAP, LC3-associated phagocytosis; ROS, reactive oxygen species; TB, tuberculosis; WT, wild-type
Introduction
According to the World Health Organization, 10 million people fell ill with Mycobacterium tuberculosis infection and 1.5 million people died of tuberculosis (TB) in 2020, marking the first increase in TB-associated deaths in over a decade [1]. Whether a person controls the initial M. tuberculosis infection or develops active TB disease is directly impacted by the type of immune response elicited in the infected individual [2]. Therefore, better understanding of what constitutes a protective versus non-protective immune response to M. tuberculosis infection is critical for developing better therapies and prevention measures to fight this deadly disease. Genetic mouse models have provided invaluable insight into the immunological processes that are required for control of M. tuberculosis infection. Infection of mice through the aerosol route leads to phagocytosis of M. tuberculosis by alveolar macrophages, initiating an inflammatory response and recruitment of innate immune cells to the lung [2]. M. tuberculosis replicates within these innate immune cells until antigen specific T cells traffic to the lung where they activate the innate immune cells to restrain M. tuberculosis replication and suppress inflammation. M. tuberculosis establishes a chronic infection in wild-type (WT) mice, which survive for over a year with this infection.
Atg5fl/fl-LysM-Cre mice, which delete the Atg5 gene specifically in macrophages, inflammatory monocytes, some dendritic cells (DCs), and neutrophils, are severely susceptible to M. tuberculosis infection [3–5], highlighting ATG5 as a critical component of a protective immune response to M. tuberculosis. M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice fail to control bacterial replication and succumb to infection by 40 days postinfection (dpi) [3–5]. The uncontrolled M. tuberculosis replication is associated with an early (by 14 dpi) and sustained influx of neutrophils in the M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice. Depletion of neutrophils during M. tuberculosis infection in Atg5fl/fl-LysM-Cre mice rescues the susceptibility and extends their survival [3], demonstrating that the neutrophil-dominated inflammation contributed to their susceptibility. In general, higher abundance of neutrophils during M. tuberculosis infection have been associated with worse disease outcomes in mice [6–13] and humans [12,14–17]. Therefore, understanding the regulatory mechanisms that govern neutrophil recruitment and accumulation during M. tuberculosis infection could be key for manipulating inflammatory responses to better control TB.
ATG5 is required for the intracellular pathway of autophagy, a process by which cytoplasmic contents are targeted to the lysosome for degradation [18,19]. Initiation of autophagy involves phagophore formation from the endoplasmic reticulum, which is mediated by the ULK1 complex (ULK1/ULK2, ATG13, FIP200, and ATG101) and the PI3 kinase complex (ATG14L, BECLIN1, VPS15, and VPS34) [20,21]. Elongation of the autophagosomal double membrane depends on 2 ubiquitin-like conjugation systems. In the first system, ATG12 is activated by ATG7, transferred to ATG10, and covalently attached to ATG5. The second ubiquitin-like component is LC3 (microtubule-associated protein 1 light chain 3), which is conjugated to phosphatidylethanolamine, generating the membrane bound form called LC3-II through the actions of ATG7 and ATG3. ATG5-ATG12 facilitates LC3 lipidation through its interactions with ATG3, while ATG16L1 specifies the localization of LC3 conjugation to the autophagosome membrane [18,19,21,22]. The autophagosome membrane is then completed and targeted for fusion with the lysosome where the autophagosome cargo are degraded. In addition, ATG5 also functions outside of autophagy, including during M. tuberculosis infection [3], although these activities remain poorly understood. Recent work supports an autophagy-dependent role for ATG5 in LysM+ innate immune cells in suppressing neutrophil recruitment to the lungs during M. tuberculosis infection [23]. Using in vitro cell culture experiments, multiple groups have reported that macrophages require autophagy to control M. tuberculosis replication by targeting the pathogen to the lysosome (xenophagy) as well as to prevent necrosis following days of infection in culture [5,23–29]. However, to date there is no evidence that xenophagy functions in this capacity in vivo. Therefore, the mechanistic basis for how loss of ATG5 results in early and exaggerated recruitment of neutrophils during M. tuberculosis infection in vivo remains unknown.
In this manuscript, we dissect the role for ATG5 in regulating neutrophil recruitment and accumulation during M. tuberculosis infection in vivo. We find that ATG5 functions with other autophagy proteins specifically in CD11c+ lung macrophages and DCs to limit the production of cytokines and chemokines that otherwise promote neutrophil influx to the lung early in M. tuberculosis infection. We demonstrate that loss of autophagy in macrophages and DCs does not affect M. tuberculosis burden in these cell types in vivo and instead changes the inflammatory response to the infection. In addition, ATG5 is required in lung macrophages and DCs to limit IL-17A production from CD4+ T cells. Together, our studies reveal new roles for ATG5 and other autophagy proteins in regulating inflammatory responses during infection, which with further dissection could provide insight into pathways that may be targeted to effectively promote protective immune responses during TB.
Results
ATG5 is required in CD11c+ lung macrophages and DCs to control neutrophil recruitment and accumulation early during M. tuberculosis infection in vivo
M. tuberculosis infection of Atg5fl/fl-LysM-Cre mice results in the recruitment of a higher number of neutrophils in the lungs at 14 dpi as compared to Atg5fl/fl controls, despite equivalent bacterial burdens at this time point [3]. There are also no differences in the abundance of non-neutrophil cell types in Atg5fl/fl-LysM-Cre and Atg5fl/fl mice at 14 dpi [3]. This indicates that specifically neutrophils are accumulating in M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice due to a defect in inflammatory responses to infection and not due to higher burden. To determine which LysM+ cells required ATG5 to control the early influx of neutrophils into the lungs during M. tuberculosis infection, we compared bacterial burdens and neutrophil inflammation in Atg5fl/fl-LysM-Cre, Atg5fl/fl-Mrp8-Cre (deletion in neutrophils), Atg5fl/fl-Cd11c-Cre (deletion in lung macrophages and DCs), and Atg5fl/fl controls at 14 dpi. At 14 dpi, the Atg5fl/fl-LysM-Cre and Atg5fl/fl-Cd11c-Cre mice, but not Atg5fl/fl-Mrp8-Cre mice, had higher levels of neutrophil inflammation in the lungs as compared to Atg5fl/fl controls (Fig 1A and 1B). The degree of increased neutrophil frequency was similar in Atg5fl/fl-LysM-Cre and Atg5fl/fl-Cd11c-Cre mice, indicating that loss of Atg5 in CD11c+ cells, but not neutrophils, leads to the early influx of neutrophils into the lungs during M. tuberculosis infection. At 14 dpi, none of the mouse strains harbored increased M. tuberculosis burden in their lungs (Fig 1C), indicating that the increase in neutrophil abundance in Atg5fl/fl-Cd11c-Cre mice is not due to elevated bacterial burden and reflects a dysregulated inflammatory response to infection.
(A) Representative flow cytometry plots of neutrophils (CD45+Ly6G+CD11b+) at 14 dpi from Atg5fl/fl, Atg5fl/fl-LysM-Cre, Atg5fl/fl-CD11c-Cre, and Atg5fl/fl-Mrp8-Cre mice. (B) Proportion of CD45+ cells that are neutrophils in the lung at 14 dpi in Mtb-GFP infected Atg5fl/fl (n = 15), Atg5fl/fl-LysM-Cre (n = 8), Atg5fl/fl-CD11c-Cre (n = 10), and Atg5fl/fl-Mrp8-Cre (n = 6) mice. Neutrophil frequency is reported as a ratio relative to the average neutrophil frequency in Atg5fl/fl control mice at 14 dpi within a given experiment. (C) Lung burden from the right lung at 14 dpi in Mtb-GFP infected Atg5fl/fl (n = 15), Atg5fl/fl-LysM-Cre (n = 8), Atg5fl/fl-CD11c-Cre (n = 10), and Atg5fl/fl-Mrp8-Cre (n = 6) mice. (D) Proportion of CD45+ cells that are neutrophils in the lung at 21 dpi in Mtb-GFP infected Atg5fl/fl (n = 19), Atg5fl/fl-LysM-Cre (n = 6), Atg5fl/fl-CD11c-Cre (n = 6), and Atg5fl/fl-Mrp8-Cre (n = 7) mice. Neutrophil frequency is reported as a ratio relative to the average neutrophil frequency in Atg5fl/fl control mice at 21 dpi within a given experiment. Susceptible and healthy Atg5fl/fl-Mrp8-Cre mice are defined as done previously where susceptible Atg5fl/fl-Mrp8-Cre mice have lost more than 5% of their pre-infection body weight by 20 dpi and healthy Atg5fl/fl-Mrp8-Cre mice have lost less than 5% of their pre-infection body weight at 20 dpi [3]. (E) Lung burden from the right lung at 21 dpi in Mtb-GFP infected Atg5fl/fl (n = 19), Atg5fl/fl-LysM-Cre (n = 6), Atg5fl/fl-CD11c-Cre (n = 6), and Atg5fl/fl-Mrp8-Cre (n = 7) mice. (F) Kaplan–Meier curve of survival proportions during Mtb-GFP infection of Atg5fl/fl (n = 9) and Atg5fl/fl-CD11c-Cre (n = 9) mice. Statistical differences were determined by a log-rank Mantel–Cox test (F) or one-way ANOVA and Šídák multiple comparison test (B–E). * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Differences that are not statistically significant are designated as ns. Pooled data from at least 2 separate experiments is graphed where each data point is from 1 biological replicate. The individual numerical values used to generate the graphed data in Fig 1, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S1 Data. dpi, days postinfection.
To determine if the higher levels of neutrophils in the lungs of M. tuberculosis-infected Atg5fl/fl-CD11c-Cre mice was due to elevated neutrophil abundance in circulation prior to or during infection, we monitored neutrophil frequency in the blood in uninfected and 14 dpi Atg5fl/fl and Atg5fl/fl-CD11c-Cre mice. There was no significant difference in the frequency of neutrophils in the blood of uninfected or 14 dpi Atg5fl/fl and Atg5fl/fl-CD11c-Cre mice (S1A Fig), suggesting that the accumulation of neutrophils in the lungs of M. tuberculosis-infected Atg5fl/fl-CD11c-Cre mice was due to specific recruitment of neutrophils to the site of infection or an inability to clear neutrophils from the lung. To begin to investigate this latter possibility, we monitored whether dead neutrophils were accumulating in the lungs of Atg5fl/fl-CD11c-Cre mice during M. tuberculosis infection by analyzing neutrophil viability at 14 dpi by flow cytometry (S1B Fig). We did not observe a significant difference in the frequency of viable neutrophils at 14 dpi in Atg5fl/fl and Atg5fl/fl-CD11c-Cre mice, indicating that increased neutrophil inflammation in the lungs of Atg5fl/fl-CD11c-Cre mice was not due to differences in neutrophil viability.
The higher levels of neutrophils in the lungs of Atg5fl/fl-LysM-Cre and Atg5fl/fl-Cd11c-Cre mice were sustained through 21 dpi (Fig 1D). However, only Atg5fl/fl-LysM-Cre mice, and not Atg5fl/fl-Cd11c-Cre mice, had higher bacterial burdens in the lungs at 21 dpi (Fig 1E), similar to as previously reported [3]. Loss of Atg5 in neutrophils results in increased susceptibility to M. tuberculosis infection in some, but not all, Atg5fl/fl-Mrp8-Cre mice [3]. The susceptible Atg5fl/fl-Mrp8-Cre mice accumulate higher neutrophil numbers and bacterial burdens in their lungs at 21 dpi (Fig 1D and 1E) [3]. Therefore, loss of Atg5 in neutrophils is likely contributing to the higher burdens in the lungs of Atg5fl/fl-LysM-Cre mice at 21 dpi. These data indicate that ATG5 has a role in CD11c+ lung macrophages and DCs to regulate early recruitment of neutrophils, but not the control of M. tuberculosis replication during M. tuberculosis infection at 14 and 21 dpi.
To determine how the loss of Atg5 in CD11c+ cells and the resulting early influx of neutrophils into the lungs affected host susceptibility, we monitored survival in M. tuberculosis-infected Atg5fl/fl-Cd11c-Cre mice as compared to Atg5fl/fl controls. Atg5fl/fl-Cd11c-Cre mice succumbed to M. tuberculosis infection between 100 and 150 dpi, which was significantly earlier than Atg5fl/fl controls (median survival time of 259 dpi) (Fig 1F), but not as early as Atg5fl/fl-LysM-Cre mice (succumb 30 to 40 dpi [3]). These data demonstrate that ATG5 is required in CD11c+ lung macrophages and DCs to control early neutrophil recruitment and promote survival following M. tuberculosis infection.
The role for ATG5 in lung macrophages and DCs in regulating neutrophil recruitment is dependent on other autophagy proteins
Deletion of multiple different autophagy genes in all LysM+ innate immune cells can result in increased neutrophil recruitment to the lung during M. tuberculosis infection [23]. However, we previously showed that at least 1 role for ATG5 in LysM+ innate immune cells in controlling M. tuberculosis infection is autophagy independent [3]. Therefore, it is not known if the role for ATG5 specifically in CD11c+ lung macrophages and DCs is the same as described when broadly deleting Atg5 in all LysM+ cells. To determine whether the regulation of neutrophil recruitment by ATG5 in CD11c+ lung macrophages and DCs was dependent on other autophagy proteins or represented the autophagy-independent role for ATG5 during M. tuberculosis infection, we monitored neutrophil abundance in the lungs of mice lacking expression of another essential autophagy protein, BECLIN 1, in CD11c+ cells (Becn1fl/fl-Cd11c-Cre) at 14 dpi by flow cytometry. Similar to Atg5fl/fl-Cd11c-Cre mice, Becn1fl/fl-Cd11c-Cre mice also exhibited elevated neutrophil frequency in the lung at 14 dpi relative to Becn1fl/fl control mice (Fig 2A), despite no difference in bacterial burden (Fig 2B). In addition, analysis of M. tuberculosis-infected Atg16l1fl/fl-LysM-Cre and Becn1fl/fl-LysM-Cre mice also revealed higher levels of neutrophils in the lungs at 14 dpi relative to controls, without higher bacterial burdens (Fig 2C and 2D).
(A) Proportion of CD45+ cells that are neutrophils (CD45+Ly6G+CD11b+) in the lung at 14 dpi in Mtb-GFP infected Becn1fl/fl (n = 8) or Becn1fl/fl-CD11c-Cre (n = 11) mice. Neutrophil frequency is reported as a ratio relative to the average neutrophil frequency in Becn1fl/fl control mice at 14 dpi. (B) Lung burden from the right lobes of the lung at 14 dpi in Mtb-GFP infected Becn1fl/fl (n = 8) or Becn1fl/fl-CD11c-Cre (n = 11) mice. (C) Neutrophil frequency of CD45+ cells reported as a ratio to the average neutrophil frequency in floxed control mice at 14 dpi in Mtb-GFP infected Atg5fl/fl (n = 6), Atg5fl/fl-LysM-Cre (n = 5), Becn1fl/fl (n = 6), Becn1fl/fl-LysM-Cre (n = 6), Atg16l1fl/fl (n = 7), and Atg16l1fl/fl-LysM-Cre (n = 7) mice. Neutrophil frequencies at 14 dpi in Mtb-GFP infected Rubicon-/- (n = 6) mice were compared to WT C57BL/6J (n = 6) mice. (D) Lung burden from right lobes of the lung at 14 dpi in Mtb-GFP infected Atg5fl/fl (n = 6), Atg5fl/fl-LysM-Cre (n = 5), Becn1fl/fl (n = 6), Becn1fl/fl-LysM-Cre (n = 6), Atg16l1fl/fl (n = 7), Atg16l1fl/fl-LysM-Cre (n = 7), WT C57BL/6J (n = 6), and Rubicon-/- (n = 6) mice. (E, F) The proportion of M. tuberculosis infected (Mtb-GFP+) and the median fluorescence intensity of Mtb-GFP in infected alveolar macrophage (Alv. Mac.), eosinophils (Eos.), neutrophils (Neut.), inflammatory monocytes (Inf. Mono.), CD11b+ DC, CD103+ DC, and non-alveolar macrophages in the lung in Atg5fl/fl (n = 7) and Atg5fl/fl-LysM-Cre (n = 7) mice at 14 dpi. (G) The proportion of viable (Zombie-) alveolar macrophages, eosinophils, neutrophils, inflammatory monocytes, CD11b+ DC, CD103+ DC, and non-alveolar macrophages in the lung in Atg5fl/fl (n = 8) and Atg5fl/fl-CD11c-Cre (n = 7) mice at 14 dpi. Statistical differences were determined by Student t test to compare the LysM-Cre or CD11c-Cre mice to their respective floxed control and Rubicon-/- mice to WT C57BL/6J mice (A–G).* P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Differences that are not statistically significant are designated as ns. Pooled data from at least 2 separate experiments is graphed where each data point is from 1 biological replicate. The individual numerical values used to generate the graphed data in Fig 2, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S2 Data. DC, dendritic cell; dpi, days postinfection; WT, wild-type.
In addition to their role in canonical autophagy, the proteins ATG5, BECLIN 1, and ATG16L1 are also required for the process of LC3-associated phagocytosis (LAP), where LC3 is recruited to the phagosome, resulting in LC3+ single membrane vesicles that traffic to the lysosome for degradation. LAP can dampen inflammatory responses through efferocytosis, pathogen removal, stimulating inhibitory immune-receptor signaling, and reducing auto-antigen levels [30–33]. In contrast to canonical autophagy, LAP uses RUBICON and UVRAG instead of ATG14 in the PI3K complex and does not depend on ULK1 [33,34]. To distinguish between whether ATG5, BECLIN 1, and ATG16L1 were functioning through autophagy or LAP to regulate neutrophil recruitment during M. tuberculosis infection, we infected mice lacking RUBICON expression, a protein specifically required for LAP. Rubicon-/- mice had no difference in neutrophil accumulation or bacterial burdens as compared to WT controls following M. tuberculosis infection (Fig 2C and 2D), indicating that LAP is not required to control neutrophil inflammation during M. tuberculosis infection. Importantly, BECLIN 1 and ATG5 function at different steps of autophagy. Therefore, the requirement of both BECLIN 1 and ATG5 suggests that both the initiation and elongation steps of autophagy are required in CD11c+ cells to control neutrophil recruitment early during M. tuberculosis infection. Targeting of pathogens to the lysosome via autophagy is termed xenophagy, and multiple studies have reported roles for xenophagy in controlling M. tuberculosis replication in macrophages and DCs in cell culture in vitro [5,23–29]. However, in addition to there being no differences in bacterial burden in the lungs at 14 dpi (Figs 1C and 2D) [3], there was no significant difference in the proportion of macrophages, eosinophils, neutrophils, inflammatory monocytes, or DCs that were infected with M. tuberculosis (Fig 2E) and no difference in the M. tuberculosis burden in autophagy-deficient cells in the lungs of Atg5fl/fl-LysM-Cre mice compared to Atg5fl/fl controls at 14 dpi (Fig 2F). Loss of xenophagy in bone marrow-derived macrophages (BMDMs) in vitro has also been associated with increased necrosis during M. tuberculosis infection [23]. However, we did not detect a difference in the viability of macrophages, inflammatory monocytes, eosinophils, neutrophils, or DCs in the lungs of Atg5fl/fl and Atg5fl/fl-Cd11c-Cre mice at 14 dpi (Figs 2G and S1B), although we cannot rule out effects on the balance of different cell death pathways. These data support that the role for autophagy in CD11c+ lung macrophages and DCs early during M. tuberculosis infection in vivo is independent of xenophagy regulating M. tuberculosis replication. In addition, these data show that the elevated neutrophil abundance in M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice is not due to differences in the cell viability or the cell types infected with M. tuberculosis but instead is driven by an imbalanced inflammatory response.
Autophagy regulates proinflammatory responses in macrophages during M. tuberculosis infection
We have previously shown that lungs of Atg5fl/fl-LysM-Cre mice at 14 dpi contain higher levels of G-CSF and IL-17A than control mice [3], cytokines that promote neutrophil development and recruitment. At this time point, the primary CD11c+ cell types that are infected by M. tuberculosis are the lung macrophages, encompassing alveolar and interstitial macrophages [35,36]. Therefore, we hypothesized that autophagy could be suppressing the production of these cytokines from M. tuberculosis-infected macrophages. We tested this hypothesis by culturing BMDMs from Atg5fl/fl, Atg5fl/fl-LysM-Cre, Atg16l1fl/fl, Atg16l1fl/fl-LysM-Cre, Becn1fl/fl, and Becn1fl/fl-LysM-Cre mice and infecting with M. tuberculosis in vitro before monitoring cytokine and chemokine production using a cytokine bead array (Bio-Rad) on the supernatants from infected cultures (Figs 3A–3F and S2). Of the 23 cytokines tested, we detected significantly higher levels of IL-1β, G-CSF, KC, TNF-α, and RANTES from the Atg5fl/fl-LysM-Cre macrophage cultures compared to controls at 24 hpi (Fig 3A–3E), despite no difference in bacterial burdens or BMDM viability at this time point (Figs 3F and S2A), indicating that the heightened proinflammatory response of Atg5-/- BMDMs was not in response to increased antigen or macrophage cell death. The levels of these cytokines were only different following M. tuberculosis infection and not in mock-infected cultures, indicating that the increased proinflammatory responses were infection induced. The higher levels of G-CSF and KC, both proinflammatory signals associated with neutrophil inflammation [37–39], produced from Atg5-deficient macrophages in response to M. tuberculosis infection were dose dependent and confirmed by ELISA (Fig 3H and 3I). Similar to Atg5fl/fl-LysM-Cre BMDMs, Atg16l1fl/fl-LysM-Cre BMDMs also produced higher levels of IL-1β, G-CSF, and TNF-α following M. tuberculosis infection in vitro (Fig 3A, 3B and 3D). Becn1fl/fl-LysM-Cre BMDMs also produced more IL-1β and G-CSF following M. tuberculosis infection in vitro compared to control cells (Figs 3A, 3B and S2) despite no difference in M. tuberculosis burden at this time point (Fig 3F). In addition, Becn1fl/fl-LysM-Cre BMDMs produced higher levels of IL-6, MIP-1α, MIP-1β, and MCP-1 following M. tuberculosis infection (Figs 3G and S2). IL-6 in particular is associated with neutrophil recruitment [40–43] and also trended higher in M. tuberculosis-infected Atg5fl/fl-LysM-Cre BMDMs (Fig 3G), so we further analyzed the levels of IL-6 produced by M. tuberculosis-infected Atg5fl/fl-LysM-Cre BMDMs using an ELISA (Fig 3J). These analyses confirmed higher levels of IL-6 secretion from M. tuberculosis-infected Atg5fl/fl-LysM-Cre BMDMs compared to controls. We were able to detect more cytokines and chemokines being differentially produced by M. tuberculosis-infected BMDMs than what was previously detected in the lungs of Atg5fl/fl-LysM-Cre mice at 14 dpi [3]. This is likely due to the dilution of signals produced by macrophages in the context of the total lung homogenate, decreasing our sensitivity to detect macrophage-specific responses. In addition, it is possible that the inflammatory responses of BMDMs differ from lung macrophages and DCs during M. tuberculosis infection. Nonetheless, loss of expression of the autophagy proteins ATG5, ATG16L1, or BECLIN 1 in macrophages both in vivo and in vitro results in higher levels of cytokines and chemokines that are associated with neutrophil recruitment and accumulation following M. tuberculosis infection relative to controls, indicating that canonical autophagy is required in macrophages to control proinflammatory responses during M. tuberculosis infection without affecting M. tuberculosis burden in macrophages.
(A) Cytokine bead array data to quantify cytokines in culture supernatants from Atg5fl/fl, Atg5fl/fl-LysM-Cre, Atg16l1fl/fl, Atg16l1fl/fl-LysM-Cre, Becn1fl/fl, or Becn1fl/fl-LysM-Cre BMDMs mock-treated or infected with Mtb-GFP at an MOI of 10 for 24 h. BMDMs generated from at least 3 mice were tested in duplicate to quantify cytokine production. (A) IL-1β, (B) G-CSF, (C) KC, (D) TNF-α, and (E) RANTES levels at 24 hpi are shown. (F) BMDM CFU counts at 24 hpi. (G) IL-6 levels at 24 hpi are shown. (H) KC, (I) G-CSF, and (J) IL-6 levels at 24 hpi in Atg5fl/fl (n = 8) and Atg5fl/fl-LysM-Cre (n = 8) BMDMs infected with M. tuberculosis at an MOI of 10 or 20 for 24 h determined by ELISA. Statistical differences were determined by Student t test to compare the autophagy gene-deficient cells to their respective floxed control cells (A–J). * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Differences that are not statistically significant are designated as ns. Cytokine levels below detection limits are designated as dbl. Each data point is 1 biological replicate, and the samples were generated from at least 2 separate experiments. The individual numerical values used to generate the graphed data in Fig 3, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S3 Data. BMDM, bone marrow-derived macrophage; hpi, hours postinfection.
Autophagy suppresses neutrophil recruitment early during M. tuberculosis infection independent of mitophagy and inflammasome activation
Autophagy has been shown to suppress proinflammatory responses by negatively regulating inflammasome activation indirectly through regulation of NFκB signaling and directly by degrading pro-IL-1β and clearance of inflammasome components [44–46], which can otherwise promote proinflammatory responses, IL-1β secretion, and neutrophil recruitment [44,45,47–49]. Indeed, Atg5fl/fl-LysM-Cre, Atg16l1fl/fl-LysM-Cre, and Becn1fl/fl-LysM-Cre BMDMs produce significantly more IL-1β in response to M. tuberculosis infection in vitro at 24 hpi compared to control BMDMs (Fig 3A), supporting that loss of autophagy has resulted in increased inflammasome activation. The primary inflammasome activated during M. tuberculosis infection of macrophages is the NLRP3 inflammasome, which consists of the NOD-, LRR-, and pyrin-domain containing protein 3 (NLRP3) sensor, ASC adaptor, and CASPASE 1 [50–53]. TLR engagement and NFκB activation during M. tuberculosis infection constitute the priming step of inflammasome activation, resulting in increased expression of pro-IL-1β and NLRP3 [54,55]. Phagocytosis of M. tuberculosis and subsequent Esx-1-dependent plasma membrane damage leading to potassium efflux is the second signal promoting NLRP3 inflammasome formation, which mediates CASPASE 1 activation followed by IL-1β processing and secretion [26,29].
To determine whether the increased neutrophil inflammation following M. tuberculosis infection in autophagy-deficient mice results from increased inflammasome activation, we crossed Caspase1/11-/- mice to Atg5fl/fl-LysM-Cre and Becn1fl/fl-LysM-Cre mice and monitored neutrophil abundance in the lungs at 14 dpi. Caspase1/11-/-/Atg5fl/fl-LysM-Cre and Caspase1/11-/-/Becn1fl/fl-LysM-Cre mice had similar neutrophil abundances and bacterial burdens in the lungs at 14 dpi as Atg5fl/fl-LysM-Cre mice and Becn1fl/fl-LysM-Cre mice, respectively (Fig 4A and 4B), indicating that increased neutrophil recruitment in the absence of autophagy is occurring independent of CASPASE1/11. Caspase1/11 deletion also did not extend the survival of Atg5fl/fl-LysM-Cre mice, indicating that increased inflammasome activation does not contribute to the early susceptibility of these mice (Fig 4C).
(A) Proportion of CD45+ cells that are neutrophils (CD45+Ly6G+CD11b+) in the lung at 14 dpi in Mtb-GFP infected Becn1fl/fl (n = 5), Becn1fl/fl-LysM-Cre (n = 7), Caspase1/11-/-/Becn1fl/fl (n = 9), Caspase1/11-/-/Becn1fl/fl-LysM-Cre (n = 13), Atg5fl/fl (n = 5), Atg5fl/fl-LysM-Cre (n = 5), Caspase1/11-/-/Atg5fl/fl (n = 4), or Caspase1/11-/-/Atg5fl/fl-LysM-Cre (n = 5) mice reported as a ratio relative to the average neutrophil frequency in corresponding floxed control mice. (B) Lung burden at 14 dpi from right lobes of the lung in Mtb-GFP infected Becn1fl/fl (n = 5), Becn1fl/fl-LysM-Cre (n = 7), Caspase1/11-/-/Becn1fl/fl (n = 9), Caspase1/11-/-/Becn1fl/fl-LysM-Cre (n = 13), Atg5fl/fl (n = 5), Atg5fl/fl-LysM-Cre (n = 5), Caspase1/11-/-/Atg5fl/fl (n = 4), or Caspase1/11-/-/Atg5fl/fl-LysM-Cre (n = 5) mice. The legend in 4A should be used for 4B too. (C) Kaplan–Meier curve of survival proportions during Mtb-GFP infection of Atg5fl/fl, Atg5fl/fl-LysM-Cre, Caspase1/11-/-/Atg5fl/fl, and Caspase1/11-/-/Atg5fl/fl-LysM-Cre mice. (D) Proportion of CD45+ cells that are neutrophils (CD45+Ly6G+CD11b+) in the lung at 14 dpi in Mtb-GFP infected WT (n = 4), Parkin-/- (n = 3), or Pink1-/- (n = 3) mice reported as a ratio relative to the average neutrophil frequency in WT mice. (E) Lung burden from the right lobe of the lung at 14 dpi in Mtb-GFP infected WT (n = 4), Parkin-/- (n = 3), or Pink1-/- (n = 3) mice. Statistical differences were determined by log-rank Mantel–Cox test (C) and one-way ANOVA and Šídák multiple comparison test (A, B, D, and E). * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Differences that are not statistically significant are designated as ns. Pooled data from at least 2 separate experiments is graphed where each data point is from 1 biological replicate. The individual numerical values used to generate the graphed data in Fig 4, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S4 Data. dpi, days postinfection; WT, wild-type.
Autophagy has also been shown to suppress inflammatory responses via the process of mitophagy, where autophagy targets old and damaged mitochondria to the lysosome for degradation [56,57]. The build-up of damaged or dysfunctional mitochondria in the absence of autophagy results in loss of mitochondrial membrane potential and the release of reactive oxygen species (ROS), mitochondrial DNA, and ATP to the cytosol where it can lead to oxidative damage, inflammasome activation, and proinflammatory cytokine production [57–61]. Mitophagy requires the canonical autophagy proteins, including ATG5, ATG16L1, and BECLIN 1, as well as PARKIN and PTEN-induced kinase 1 (PINK1) [62]. PINK1 accumulates on damaged mitochondria and activates the mitochondrial E3 ubiquitin ligase, PARKIN, to ubiquitinylate damaged mitochondria [58,61]. Optineurin and NDP52 are the main mitophagy receptors that interact with the ubiquitinylated mitochondria and LC3, leading to autophagosome engulfment of the mitochondria [58,63]. To investigate whether loss of mitophagy could contribute to higher neutrophil accumulation in the lungs following M. tuberculosis infection, we measured neutrophil frequency in the lungs at 14 dpi by flow cytometry in Parkin-/- and Pink1-/- mice relative to WT controls. There was no difference in neutrophil abundance or bacterial burdens in M. tuberculosis-infected Parkin-/- or Pink1-/- mice relative to WT mice at 14 dpi (Fig 4D and 4E), indicating that mitophagy is not required to control neutrophil recruitment early during M. tuberculosis infection. To determine if mitophagy is required in macrophages to control proinflammatory cytokine and chemokine production during M. tuberculosis infection, we generated BMDMs from Parkin-/-, Pink1-/-, and WT mice and infected the macrophages with M. tuberculosis for 24 h. We measured cytokine and chemokine levels from mock and M. tuberculosis-infected cultures using the cytokine bead array (Bio-Rad). Unlike in autophagy-deficient BMDMs, there were no differences in IL-6, IL-1β, G-CSF, KC, TNF-α, or RANTES production by M. tuberculosis-infected Parkin-/- and Pink1-/- macrophages at 24 hpi compared to WT macrophages (S3 Fig), nor any differences in bacterial burden (S3 Fig). Therefore, loss of mitophagy in macrophages does not result in higher levels of inflammation early during M. tuberculosis infection.
ATG5 is required to suppress early TH17 responses in the lungs during M. tuberculosis infection
The higher levels of IL-17A observed in the lungs of Atg5fl/fl-LysM-Cre mice at 14 dpi with M. tuberculosis relative to controls was not reproduced by BMDMs infected with M. tuberculosis for 24 h (S2 Fig). Although there are many reasons to explain this, one possibility is that the macrophages were not the source of IL-17A in vivo. We investigated what cell type was expressing higher levels of IL-17A in the Atg5fl/fl-LysM-Cre mice during M. tuberculosis infection by crossing the Atg5fl/fl and Atg5fl/fl-LysM-Cre mice with an IL-17A reporter mouse that expresses GFP under the IL-17A promoter (Jax # 018472). We infected Il17a-GFP/Atg5fl/fl and Il17a-GFP/Atg5fl/fl-LysM-Cre mice with M. tuberculosis and monitored GFP expression as a proxy of IL-17A expression in immune cells at 14 dpi. The only cell type we reproducibly detected >0.5% of the cells expressing GFP were CD4+ T cells. Similar to previous studies with Atg5fl/fl and Atg5fl/fl-LysM-Cre mice, there was no difference in total CD4+ T cell numbers in the lungs of Il17a-GFP/Atg5fl/fl and Il17a-GFP/Atg5fl/fl-LysM-Cre mice at 14 dpi (Fig 5A) [3]. However, a greater frequency and number of the CD4+ T cells in the lungs of Il17a-GFP/Atg5fl/fl-LysM-Cre mice at 14 dpi were IL-17-GFP+ compared to Il17a-GFP/Atg5fl/fl mice (Fig 5B and 5C). These data indicate that CD4+ T cells contribute to the higher levels of IL-17A in the lungs of M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice and ATG5 is required in innate immune cells to negatively regulate TH17 responses during M. tuberculosis infection.
(A) The number of CD4+ T cells (CD45+ TCRβ+ CD4+) in Atg5fl/fl and Atg5fl/fl-LysM-Cre mice are reported as the total cells per lung in the left lobe at 14 dpi. (B) The frequency of IL-17-GFP+ CD4+ T cells in the lung (CD45+ TCRβ+ CD4+ IL-17-GFP+) of Atg5fl/fl and Atg5fl/fl-LysM-Cre mice are reported as the percentage of CD4+ T cells that are IL-17-GFP positive at 14 dpi. (C) The number IL-17-GFP+ CD4+ T cells in Atg5fl/fl and Atg5fl/fl-LysM-Cre mice are reported as the total cells per lung in the left lobe at 14 dpi. (D) Schematic depicting the timing of CD4-depletion antibody injections. (E) The number of CD4+ T cells (CD45+ TCRβ+ CD4+) in Mtb-GFP infected Atg5fl/fl and Atg5fl/fl-LysM-Cre mice are reported as the total cells per lung in the left lobe at 14 dpi following antibody treatment. (F) Lung burden from the right lobes of the lung at 14 dpi in Mtb-GFP infected Atg5fl/fl or Atg5fl/fl-LysM-Cre mice that received isotype or CD4-depletion antibodies. (G) Proportion of CD45+ cells that are neutrophils (CD45+Ly6G+CD11b+) in the lung at 14 dpi in Mtb-GFP infected Atg5fl/fl or Atg5fl/fl-LysM-Cre mice that received CD4-depletion or isotype treatment. Neutrophil frequency is reported as a ratio relative to the average neutrophil frequency in Atg5fl/fl control mice at 14 dpi. (H) Schematic depicting delivery of the IL-17 neutralizing antibody treatments. (I) Lung burden from the right lobes of the lung at 14 dpi in Mtb-GFP infected Atg5fl/fl or Atg5fl/fl-LysM-Cre mice that received isotype or IL-17 neutralizing antibodies. (J) Proportion of CD45+ cells that are neutrophils (CD45+Ly6G+CD11b+) in the lung at 14 dpi in Mtb-GFP infected Atg5fl/fl or Atg5fl/fl-LysM-Cre mice that received isotype or IL-17 neutralizing antibodies. Neutrophil frequency is reported as a ratio relative to the average neutrophil frequency in Atg5fl/fl control mice at 14 dpi. Statistical differences were determined by Student t test (A) or one-way ANOVA and Šídák multiple comparison test (A–C, E–G, I, and J). * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Differences that are not statistically significant are designated as ns. Pooled data from at least 2 separate experiments is graphed where each data point is from 1 biological replicate. The individual numerical values used to generate the graphed data in Fig 5, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S5 Data. The mouse and needle images used in Fig 5 were made with an academic account on BioRender.com where BioRender’s academic license (UU25CW6WGU) terms enable the use of these graphics in journal publications. dpi, days postinfection.
IL-17A drives neutrophil influx by promoting the production of neutrophil chemokines MIP-1α and KC and through activation of endothelial cells [64–66]. Therefore, the increased TH17 responses could be responsible for the early influx and accumulation of neutrophils in the lungs of M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice. To determine if the increased expression of IL-17A by T cells was responsible for the enhanced influx of neutrophils at 14 dpi in Atg5fl/fl-LysM-Cre mice, we depleted CD4+ T cells by administering antibodies specific for CD4 from day −2 to 14 dpi (Fig 5D and 5E). At 14 dpi, we harvested the lungs for enumeration of M. tuberculosis burden and neutrophil abundance and found that there was no effect of CD4+ T cell depletion on either readout (Fig 5F and 5G). In addition, blocking IL-17A signaling by administering an anti-IL-17A antibody from day −1 to 14 dpi (Fig 5H) did not affect M. tuberculosis burden (Fig 5I) or neutrophil abundance (Fig 5J) in Atg5fl/fl-LysM-Cre mice and Atg5fl/fl mice at 14 dpi. Therefore, although ATG5 is required in innate immune cells to suppress IL-17A expression in T cells, this role does not contribute to regulating neutrophil accumulation early during M. tuberculosis infection.
Discussion
Atg5fl/fl-LysM-Cre mice are extremely susceptible to M. tuberculosis infection, where neutrophils accumulate in the lungs of infected Atg5fl/fl-LysM-Cre mice by 14 dpi and are sustained at high levels until the mice succumb to the infection between 30 and 40 dpi [3]. It was previously unknown how ATG5 imparted control of neutrophil recruitment to the lungs during M. tuberculosis infection. We have discovered that ATG5 is required in CD11c+ lung macrophages and DCs to regulate proinflammatory cytokine production and neutrophil accumulation in the lungs early during M. tuberculosis infection. This role for ATG5 is shared with ATG16L1 and BECLIN 1, but not RUBICON, indicating it is autophagy dependent and does not involve LAP. We were able to reproduce the heightened proinflammatory responses in M. tuberculosis-infected autophagy-deficient BMDMs in vitro, suggesting that autophagy specifically suppresses inflammatory responses from macrophages during M. tuberculosis infection, although this does not rule out a similar role in DCs in vivo. Alveolar macrophages are among the first cells to encounter M. tuberculosis in the airways and orchestrate the initial response to infection, recruiting other innate immune cells to the lung [35,36]. We postulate that similar to the BMDMs, autophagy-deficient alveolar macrophages overproduce proinflammatory signals during M. tuberculosis infection, leading to increased neutrophil recruitment early during infection.
The increased levels of cytokines and chemokines produced by autophagy-deficient macrophages was dependent on M. tuberculosis infection, demonstrating that pathogen detection was required. However, the heightened proinflammatory responses in autophagy-deficient macrophages occurred in the absence of differences in bacterial burden, indicating that the enhanced inflammatory response is not due to increased antigen. Although multiple studies have shown effects of loss of autophagy on M. tuberculosis burden in macrophages and DCs in vitro in cell culture [5,23–29], there was no difference in M. tuberculosis burden in these cell types in vivo when we detected higher levels of proinflammatory cytokine production and neutrophil influx. Therefore, the role for autophagy in regulating proinflammatory cytokine production from CD11c+ lung macrophages and DCs during M. tuberculosis infection is independent of xenophagy controlling M. tuberculosis replication. Atg5fl/fl-CD11c-Cre mice succumb to M. tuberculosis infection at a similar time as Atg16l1fl/fl-LysM-Cre and Atg7fl/fl-LysM-Cre mice [23], suggesting that loss of the role for autophagy that we have identified in CD11c+ lung macrophages and DCs is responsible for the susceptibility of Atg16l1fl/fl-LysM-Cre and Atg7fl/fl-LysM-Cre mice to M. tuberculosis. These data demonstrate that autophagy is not only required in CD11c+ cells to control early inflammatory responses during M. tuberculosis infection, but also to promote survival through the chronic phase of infection. Although the cause of this later susceptibility of Atg5fl/fl-CD11c-Cre to M. tuberculosis infection is still unknown, previous studies showed that the lungs of M. tuberculosis-infected Atg16l1fl/fl-LysM-Cre and Atg7fl/fl-LysM-Cre mice harbor higher numbers of neutrophils despite equivalent bacterial burdens compared to controls at 70 dpi [23], indicating an association between continued neutrophil accumulation and susceptibility.
We ruled out the involvement of CASPASE1/11-dependent inflammasome activity and mitophagy in the autophagy-dependent regulation of neutrophil accumulation during M. tuberculosis infection, leaving open the question of how autophagy regulates macrophage proinflammatory responses during infection. Our understanding of the number of cellular pathways regulated by autophagy continues to grow, and there are many possible mechanisms by which autophagy could regulate inflammatory responses in macrophages [18]. One possibility is that targeting of autophagy components to intracellular M. tuberculosis prevents sensing of M. tuberculosis by cytosolic pattern recognition receptors and subsequent proinflammatory cytokine and chemokine production, without impacting M. tuberculosis replication. Autophagy can also regulate levels of IL-1β [44,62,67], IL-6 [32,68,69], and TNF-α [68,69] downstream of pathogen sensing, in part by negatively regulating NFκB activation [70–72]. Autophagy can both directly promote autophagic cell death and apoptosis as well as negatively regulate RIPK3-dependent necrotic cell death, all which can impact inflammatory responses [23,73–77]. Although we did not detect overall differences in the viability of autophagy-deficient cells at the time points we observed effects on cytokine and chemokine production in vivo or in vitro, the mechanism of cell death in Atg5fl/fl and Atg5-/- cells during M. tuberculosis infection may vary and could impact the inflammatory responses [73,74,78]. Autophagy may also be required in macrophages for the efficient removal of dead or infected neutrophils from the lung and resolution of inflammation. The best-studied process for removal of dead cells by phagocytes is efferocytosis [79]. LAP has specifically been shown to contribute to efferocytosis [80], and although we rule out a role for LAP in regulating neutrophil accumulation during M. tuberculosis infection, in remains possible that processing of dead cells may be attenuated in the absence of autophagy. Another possible mechanism for how autophagy regulates inflammatory responses from macrophages during M. tuberculosis infection involves the process of ER-phagy. ER-phagy is induced under conditions of ER stress, accumulation of unfolded proteins, and during infection [81,82]. ER-phagy restrains ER stress responses by targeting excess or damaged endoplasmic reticulum to autophagosomes for degradation [81], but in the absence of autophagy, ER stress activates NFκB-dependent transcription of inflammatory cytokines, such as IL-1β, IL-6, IL-18, and TNF-α [82].
We also discovered that ATG5 was required in innate immune cells to suppress early TH17 responses during M. tuberculosis infection. At this point, we do not know if the increased abundance of IL-17+ CD4+ T cells in M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice is due to an autophagy-dependent or independent role for ATG5, but there are multiple other reports of autophagy regulating TH17 responses. The effect of loss of Atg5 in innate immune cells on IL-17A expression from T cells may be explained by the requirement for autophagy in DCs to negatively regulate surface expression of disintegrin and metalloproteinase domain-containing protein 10 (ADAM10). ADAM10 cleaves its substrate ICOSL and lower levels of ICOSL leads to decreased ICOSL-ICOS interactions between DCs and T cells, resulting in less CD25hi CD4+ T regulatory cells and more IL-17+ CD4+ T cells [83]. In addition, autophagy negatively regulates TH17 differentiation by reducing IL-23 and IL-1β levels, which promote TH17 differentiation and IL-17A secretion [4,84–86]. Nonetheless, depleting CD4+ T cells or blocking IL-17A did not rescue the increased neutrophil accumulation in the lungs of M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice at 14 dpi, suggesting that the hyper-inflammatory responses from infected autophagy-deficient macrophages and DCs is sufficient to recruit excessive neutrophils early during infection. However, it is possible that the heightened TH17 responses in M. tuberculosis-infected Atg5fl/fl-LysM-Cre mice have a longer-term impact on the increased susceptibility of these mice and could contribute to the susceptibility of Atg5fl/fl-Cd11c-Cre mice during the chronic phase of infection.
Our data support a role for autophagy in restricting neutrophil accumulation in the lung but does not rule out additional contributions from autophagy-independent roles for ATG5, BECLIN 1, and ATG16L1. In particular, loss of Atg5, Becn1, or Atg16l1 in LysM+ cells led to different degrees of elevated neutrophil frequency in the lung at 14 dpi relative to control mice, where loss of Atg5 results in the greatest increase in neutrophil abundance. This could in part be due to differences in efficiency of gene deletion in the different lines [23]. In addition, we have previously identified an autophagy-independent role for ATG5 in neutrophils, where loss of ATG5 in neutrophils can result in early lethality during M. tuberculosis infection [3]. Therefore, loss of Atg5 in LysM+ cells, which includes neutrophils, may result in higher neutrophil frequency in the lung at 14 dpi than loss of Becn1 or Atg16l1 because of this autophagy-independent role for ATG5 in neutrophils. We hypothesize that the combination of the newly discovered role for autophagy in CD11c+ lung macrophages and DCs to regulate inflammatory responses and an autophagy-independent role for ATG5 in neutrophils collectively allow for control of M. tuberculosis infection, where loss of both functions results in the extreme susceptibility of Atg5fl/fl-LysM-Cre mice to M. tuberculosis infection. Higher abundances of neutrophils have been associated with poor disease prognosis and treatment outcomes in TB patients [14–16]. Therefore, our new findings and future dissection of the ATG5-dependent mechanisms of regulating neutrophil recruitment to the lungs during M. tuberculosis infection will provide critical insight into how to promote protective immune responses to TB.
Materials and methods
Mice
All flox mice (Atg5fl/lfl, Atg16l1fl/fl, and Becn1fl/fl) used in this study have been described previously [3,87–89] and colonies are maintained in an enhanced barrier facility. LysM-Cre (Jax #004781), Cd11c-Cre (Jax #007567), Mrp8-Cre (Jax #021614) from the Jackson Laboratory were crossed to specific flox mice. Il17a-IRES-GFP-KI (Jax # 018472) reporter mice were bred to Atg5fl/fl-LysM-Cre and Atg5fl/fl mice to generate the Il17a-GFP/Atg5fl/fl-LysM-Cre and Il17a-GFP/Atg5fl/fl lines. Rubicon-/- (Jax # 032581) and WT littermates were provided by D. Green and J. Martinez [90]. Caspase 1/11-/- (Jax #016621) were bred to Atg5fl/fl-LysM-Cre and Becn1fl/fl-LysM-Cre mice. Parkin-/- (Jax # 006582) [91], Pink1-/- (Jax # 017946) [56], and WT control mice were a gift from Dr. Jonathan Brestoff at Washington University School of Medicine. Male and female littermates (aged 6 to 12 weeks) were used and were subject to randomization. A minimum of 3 mice were used per experiment and each experiment was performed twice. Statistical consideration was not used to determine mouse sample sizes. The mice were housed and bred at Washington University in St. Louis in specific pathogen-free conditions in accordance with federal and university guidelines, and protocols were approved by the Animal Studies Committee of Washington University.
Infection of mice with M. tuberculosis and measurement of bacterial burden in the lungs
M. tuberculosis Erdman expressing GFP (Mtb-GFP [10,92]) was used in all experiments except experiments with the Il-17a-GFP/Atg5fl/fl-LysM-Cre reporter mice when WT Erdman was used. M. tuberculosis was cultured at 37°C in 7H9 (broth) or 7H11 (agar) (Difco) medium supplemented with 10% oleic acid/albumin/dextrose/catalase (OADC), 0.5% glycerol, and 0.05% Tween 80 (broth). Cultures of GFP expressing M. tuberculosis were grown in the presence of kanamycin (20 μg/mL) to ensure plasmid retention. M. tuberculosis cultures in logarithmic growth phase (OD600 = 0.5–0.8) were washed with PBS + 0.05% Tween-80, sonicated to disperse clumps, and diluted in sterile water before delivering 100 CFUs of aerosolized M. tuberculosis per lung using an Inhalation Exposure System (Glas-Col). Within 2 h of each infection, lungs were harvested from at least 2 control mice, homogenized, and plated on 7H11 agar to determine the input CFU dose. At 14 dpi, M. tuberculosis titers were determined by homogenizing the superior, middle, and inferior lobes of the right lung and plating serial dilutions on 7H11 agar. Colonies were counted after 3 weeks of incubation at 37°C in 5% CO2.
Flow cytometry from blood and infected lungs
Blood was collected by cardiac puncture into K2EDTA anticoagulant tubes (BD, 365974). Red blood cell lysis was performed by adding 2 mL of ACK lysis buffer (Gibco, A10492-01) per 100 μL of EDTA-treated blood for 5 min at room temperature. Cells were pelleted at 500×g for 5 min and then resuspended in 50 μL of PBS + 2% HI-FBS + 2 mM EDTA in the presence of Fc receptor blocking antibody (BioLegend, 101302). Cells were labeled with antibodies as described below. Lungs were perfused with sterile PBS and digested for 1 h with 625 μg/mL collagenase D (Roche 11088875103) and 75 U/mL DNase I (Sigma D4527). Cells were quenched with PBS + 2% heat-inactivated (HI)-FBS, + 2 mM EDTA and passed through a 70 μm filter. Cells were stained with Zombie-violet or Zombie-NIR in PBS at 1:2,000 dilution for 5 min at room temperature prior to resuspending in PBS + 2% HI-FBS + 2 mM EDTA in the presence of Fc receptor blocking antibody (BioLegend, 101302) for blocking. Cells were labeled with antibodies at a 1:200 dilution using the following mouse markers: CD11b_BV605 or PerCP-Cy5.5 (clone M1/70), CD45_AF700 (BioLegend, 103259), Ly6G_PE-Cy7 or AF647 (clone 1A8), MHCII_Spark blue 550 (BioLegend, 107662), CD62L_Pe/Cy5 (BioLegend, 104410), CD44_BV510 (BioLegend, 103044), CD11c_PerCP (BioLegend, 117325), Ly6C_BV605 (BioLegend, 128036), CD4_BV570 (clone RM4-5), TCRb_BV421 (clone H57-597), CD19_Pacific blue (BioLegend, 115526), Zombie-violet or -NIR (BioLegend, 423113 or 423105), SiglecF_BV480 (BD Biosciences, 746668), MHC-II_Sparkblue550 (BioLegend, 107662), CD64_PerCP-eFluor 710 (eBiosciences, 46061482) and MerTK_PE/Cy7 (eBiosciences, 25575182). Cells were incubated for 20 min at 4°C with antibodies, washed and then fixed in 4% paraformaldehyde (Electron Microscope Sciences) for 20 min at room temperature. Flow cytometry was performed on an LSR-Fortessa (BD Bioscience) or an Aurora (Cytek Biosciences, with 4 laser 16V-14B-10YG-8R configuration) and analyzed using FlowJo software (Tree Star). Absolute cell counts were determined using Precision count beads (BioLegend) or volumetric-based counting on the Aurora. Gating strategies to identify Mtb-GFP+ myeloid cell populations, viable lung neutrophils, CD4+ T cells and IL-17-GFP+ CD4+ T cells are in S4A–S4C Fig.
Culturing and infection of bone marrow-derived macrophages (BMDMs)
BMDMs were generated by flushing femurs and tibias of mice and culturing the cells in DMEM, 20% HI-FBS, 10% supernatant from 3T3 cells overexpressing M-CSF + 1% MEM non-essential amino acids (Cellgro 25-025-CI), 2 mM L-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin (Sigma P4333) at 37°C in 5% CO2 in non-TC treated plates. After 6 days, non-adherent cells were removed and 1 × 106 adherent macrophages were seeded per well in 6-well non-TC treated plates in DMEM, 10% HI-FBS, 1% MEM non-essential amino acids, and 2 mM L-glutamine. BMDMs were rested overnight at 37°C in 5% CO2. M. tuberculosis was grown to an OD of 0.6 to 0.8, washed with PBS twice, sonicated to disperse clumps, centrifuged at 55×g for 10 min to remove clumps and resuspended in antibiotic-free BMDM media. Macrophages were infected at an MOI of 10 by centrifuging the cells at 200×g for 10 min. BMDMs were washed with PBS twice to remove free M. tuberculosis and fresh BMDM media was added to each well. Cells were incubated at 37°C and 5% CO2 for 24 h. To determine CFU counts, the cells were lysed with 0.05% triton X-100, serially diluted, and plated onto 7H11 agar and incubated for 21 days when bacterial colonies were counted. At 24 hours postinfection (hpi), supernatants were stored at −80°C for cytokine analysis. To assess BMDM viability, cells were harvested at 24 hpi by gently scrapping, washed with PBS twice, stained with Zombie-violet in PBS at 1:2,000 dilution for 5 min at room temperature, washed once with PBS, and fixed in 4% PFA for 20 min at room temperature before performing flow cytometry. Flow cytometry was performed on an LSR-Fortessa (BD Bioscience) and analyzed using FlowJo software (Tree Star). The gating strategy to identify viable BMDMs at 24 hpi is depicted in S1D Fig.
Cytokine analysis
BMDM supernatants were filtered through a 0.22 μm filter twice to remove M. tuberculosis and analyzed using the BioPlex-Pro Mouse Cytokine 23-Plex Immunoassay (Bio-Rad) as per the manufacturer’s instructions. ELISAs were performed according to the manufacturer’s instructions (R&D systems): KC/CXCL1 (DY453), IL-6 (DY406), and G-CSF (DY414).
IL-17A blocking and T cell depletion
To neutralize IL-17A, 100 μg of InVivo monoclonal anti-IL-17A (Bio X Cell, BE0173) neutralizing antibody was administered to Atg5fl/fl and Atg5fl/fl-LysM-Cre mice by intraperitoneal (i.p.) injection every other day starting at 1 day prior to infection, with the final dose delivered at 13 dpi, similar to published protocols [93]. Control mice received 100 μg of IgG from mouse serum (Sigma, I5381) by i.p. injection every other day starting 1 day prior to infection and finishing on 13 dpi. To deplete CD4+ T cells from mice, 250 μg of anti-mouse CD4 (Leinco Technologies, C1333) was administered by i.p. injection at 2 days prior to infection, 5 dpi, and 12 dpi. Control mice received 250 μg of IgG from rat serum (Sigma, 18015) i.p. on 2 days prior to infection, 5 dpi, and 12 dpi.
Data and statistics
All experiments were performed at least twice. When shown, multiple samples represent biological (not technical) replicates of mice randomly sorted into each experimental group. No blinding was performed during animal experiments. Animals were only excluded when pathology unrelated to M. tuberculosis infection was present (i.e., bad teeth leading to weight loss). Determination of statistical differences was performed with Prism (GraphPad Software) using log-rank Mantel–Cox test (survival), unpaired two-tailed t test (to compare 2 groups with similar variances), or one-way ANOVA with Šídák multiple comparison test (to compare more than 2 groups). When used, center values and error bars represent the mean +/- SEM. In all figures, all significant differences are indicated by asterisks: * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Nonsignificant comparisons of particular interest are noted as ns.
Supporting information
S1 Fig. ATG5 is not required in CD11c+ cells to regulate neutrophil viability in the lung or accumulation of neutrophils in the blood during M. tuberculosis infection.
(A) The proportion of lung neutrophils (CD45+ Ly6G+ CD11b+) that are viable (Zombie-) in M. tuberculosis-infected Atg5fl/fl (n = 8) and Atg5fl/fl-CD11c-Cre (n = 7) mice at 14 dpi. (B) The proportion of CD45+ cells in the blood that are neutrophils in Atg5fl/fl (n = 4 in naïve and n = 3 in 14 dpi) and Atg5fl/fl-CD11c-Cre (n = 4) mice at 14 dpi and uninfected (naïve) mice. Statistical differences were determined by Student t test comparing the genotypes within a particular cell type or treatment group. * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Differences that are not statistically significant are designated as ns. Pooled data from at least 2 separate experiments are graphed where each data point is from 1 biological replicate. The individual numerical values used to generate the graphed data in S1 Fig, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S6 Data.
https://doi.org/10.1371/journal.pbio.3002159.s001
(TIF)
S2 Fig. Levels of viability, cytokines, and chemokines in autophagy-deficient macrophages during M. tuberculosis infection.
(A) The proportion of Atg5fl/fl (n = 5) and Atg5fl/fl-LysM-Cre (n = 5) BMDMs that are viable (Zombie-) at 24 hpi in mock and Mtb-GFP treated groups. Cytokine bead array data from mock treated and Mtb-GFP infected BMDMs. Atg5fl/fl, Atg5fl/fl-LysM-Cre, Atg16l1fl/fl, Atg16l1fl/fl-LysM-Cre, Becn1fl/fl, and Becn1fl/fl-LysM-Cre BMDMs were cultured for 24 hpi, and cytokine levels were measured in the spent media from mock treated or infected macrophages. BMDMs from at least 3 mice were tested in duplicate to quantify the cytokines in the bead array. All cytokine and chemokine data that are not significantly different between Atg5fl/fl-LysM-Cre and Atg5fl/fl mice are reported here. (B) IL-1α, (C) IL-4, (D) IL-10, (E) IL-12(p40), (F) IL-12(p70), (G) IFN-γ, (H) MIP1β, G (I) M-CSF, (J) MIP1α, (K) MCP-1, (L) Eotaxin, and (M) IL-17 levels at 24 hpi. Statistical differences were determined by one-way ANOVA and Šídák multiple comparison test (A) and Student t test comparing the autophagy-deficient macrophage with its floxed control within a treatment condition (B–M). * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Cytokine levels below detection limits are designated as dbl. Differences that are not statistically significant are designated as ns. Each data point is 1 biological replicate, and the samples were generated from at least 2 separate experiments. The individual numerical values used to generate the graphed data in S2 Fig, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S7 Data.
https://doi.org/10.1371/journal.pbio.3002159.s002
(TIF)
S3 Fig. Mitophagy is not required in macrophages to regulate proinflammatory responses during M. tuberculosis infection.
WT, Parkin-/- and Pink1-/-, BMDMs were cultured for 24 hpi and cytokine levels were measured by cytokine bead array in the media from mock treated or Mtb-GFP infected macrophages. (A) IL-1β, (B) G-CSF, (C) IL-6, (D) KC, (E) RANTES, and (F) TNF-α levels at 24 hpi (G) BMDM CFU counts from 24 hpi. BMDMs from at least 3 mice were tested in duplicate to quantify cytokines in the bead array. Each point is 1 biological replicate. Statistical differences were determined by one-way ANOVA and Šídák multiple comparison test (A–G). * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001. Cytokine levels below detection limits are designated as dbl. Statistical differences that are not significant are designated as ns. Each data point is 1 biological replicate, and the samples were generated from at least 2 separate experiments. The individual numerical values used to generate the graphed data in S3 Fig, the statistical analyses performed to analyze these data, and the p values from these statistical tests are in S8 Data.
https://doi.org/10.1371/journal.pbio.3002159.s003
(TIF)
S4 Fig. Gating strategies for flow cytometry identification of myeloid cells and IL-17 expressing T cells in the lung or viable BMDMs.
Representative flow cytometry plots depicting the gating strategy used to identify Mtb-GFP+ myeloid cells (A), viable lung neutrophils (B), and IL-17-GFP+ CD4+ T cells (C) in the lung at 14 dpi. (D) Representative flow cytometry plots showing the gating strategy to identify viable BMDMs at 24 hpi. FSC-A, forward scatter area; SSC-A, side scatter area; FSC-H, forward scatter height; FSC-W, forward scatter width, Alv. Mac., alveolar macrophages; Eos., eosinophils; DC, dendritic cell, and Inf. Mono., inflammatory monocytes.
https://doi.org/10.1371/journal.pbio.3002159.s004
(TIF)
S1 Data. Supporting data files for Fig 1.
These supporting data files include the individual numerical values used to generate the graphed data in Fig 1, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s005
(XLSX)
S2 Data. Supporting data files for Fig 2.
These supporting data files include the individual numerical values used to generate the graphed data in Fig 2, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s006
(XLSX)
S3 Data. Supporting data files for Fig 3.
These supporting data files include the individual numerical values used to generate the graphed data in Fig 3, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s007
(XLSX)
S4 Data. Supporting data files for Fig 4.
These supporting data files include the individual numerical values used to generate the graphed data in Fig 4, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s008
(XLSX)
S5 Data. Supporting data files for Fig 5.
These supporting data files include the individual numerical values used to generate the graphed data in Fig 5, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s009
(XLSX)
S6 Data. Supporting data files for S1 Fig.
These supporting data files include the individual numerical values used to generate the graphed data in S1 Fig, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s010
(XLSX)
S7 Data. Supporting data files for S2 Fig.
These supporting data files include the individual numerical values used to generate the graphed data in S2 Fig, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s011
(XLSX)
S8 Data. Supporting data files for S3 Fig.
These supporting data files include the individual numerical values used to generate the graphed data in S3 Fig, the statistical analyses performed to analyze these data, and the p value from these statistical tests.
https://doi.org/10.1371/journal.pbio.3002159.s012
(XLSX)
Acknowledgments
The authors thank Dr. Jonathan Brestoff at Washington University School of Medicine for generously providing the Parkin-/- and Pink1-/- mice.
References
- 1.
World Health Organization. Regional Office for South-East Asia. WHO global Tuberculosis report 2021 [Internet]. New Delhi: World Health Organization. Regional Office for South-East Asia; 2022 [cited 2022 Sep 9]. Available from: https://apps.who.int/iris/handle/10665/361162.
- 2. Kinsella R, Zhu D, Harrison G, Mayer Bridwell A, Prusa J, Chavez S, et al. Perspectives and Advances in the Understanding of Tuberculosis. Annu Rev Pathol. 2021 Jan 24;16:377–408. pmid:33497258
- 3. Kimmey JM, Huynh JP, Weiss LA, Park S, Kambal A, Debnath J, et al. Unique role for ATG5 in neutrophil-mediated immunopathology during M. tuberculosis infection. Nature. 2015 Dec;528(7583):565–9. pmid:26649827
- 4. Castillo EF, Dekonenko A, Arko-Mensah J, Mandell MA, Dupont N, Jiang S, et al. Autophagy protects against active tuberculosis by suppressing bacterial burden and inflammation. Proc Natl Acad Sci U S A. 2012 Nov 13;109(46):E3168–76. pmid:23093667
- 5. Watson RO, Manzanillo PS, Cox JS. Extracellular M. tuberculosis DNA Targets Bacteria for Autophagy by Activating the Host DNA-Sensing Pathway. Cell. 2012 Aug;150(4):803–15. pmid:22901810
- 6. Mishra BB, Lovewell RR, Olive AJ, Zhang G, Wang W, Eugenin E, et al. Nitric oxide prevents a pathogen-permissive granulocytic inflammation during tuberculosis. Nat Microbiol. 2017 Jul;2(7):17072. pmid:28504669
- 7. Mishra BB, Rathinam VAK, Martens GW, Martinot AJ, Kornfeld H, Fitzgerald KA, et al. Nitric oxide controls the immunopathology of tuberculosis by inhibiting NLRP3 inflammasome–dependent processing of IL-1β. Nat Immunol. 2013 Jan;14(1):52–60.
- 8. Nandi B, Behar SM. Regulation of neutrophils by interferon-γ limits lung inflammation during tuberculosis infection. J Exp Med. 2011 Oct 24;208(11):2251–62.
- 9. Niazi MKK, Dhulekar N, Schmidt D, Major S, Cooper R, Abeijon C, et al. Lung necrosis and neutrophils reflect common pathways of susceptibility to Mycobacterium tuberculosis in genetically diverse, immune competent mice. Dis Model Mech. 2015 Jan 1;dmm.020867.
- 10. Nair S, Huynh JP, Lampropoulou V, Loginicheva E, Esaulova E, Gounder AP, et al. Irg1 expression in myeloid cells prevents immunopathology during M. tuberculosis infection. J Exp Med. 2018 Apr 2;215(4):1035–45. pmid:29511063
- 11. Dorhoi A, Yeremeev V, Nouailles G, Weiner J, Jörg S, Heinemann E, et al. Type I IFN signaling triggers immunopathology in tuberculosis-susceptible mice by modulating lung phagocyte dynamics. Eur J Immunol. 2014 Aug;44(8):2380–93. pmid:24782112
- 12. Moreira-Teixeira L, Stimpson PJ, Stavropoulos E, Hadebe S, Chakravarty P, Ioannou M, et al. Type I IFN exacerbates disease in tuberculosis-susceptible mice by inducing neutrophil-mediated lung inflammation and NETosis. Nat Commun. 2020 Dec;11(1):5566. pmid:33149141
- 13. Moreira-Teixeira L, Tabone O, Graham CM, Singhania A, Stavropoulos E, Redford PS, et al. Mouse transcriptome reveals potential signatures of protection and pathogenesis in human tuberculosis. Nat Immunol. 2020 Apr;21(4):464–76. pmid:32205882
- 14. Carvalho ACC, Amorim G, Melo MGM, Silveira AKA, Vargas PHL, Moreira ASR, et al. Pre-Treatment Neutrophil Count as a Predictor of Antituberculosis Therapy Outcomes: A Multicenter Prospective Cohort Study. Front Immunol. 2021 Jul 2;12:661934. pmid:34276654
- 15. Han Y, Kim SJ, Lee SH, Sim YS, Ryu YJ, Chang JH, et al. High blood neutrophil-lymphocyte ratio associated with poor outcomes in miliary tuberculosis. J Thorac Dis. 2018 Jan;10(1):339–46. pmid:29600065
- 16. Lowe DM, Bandara AK, Packe GE, Barker RD, Wilkinson RJ, Griffiths CJ, et al. Neutrophilia independently predicts death in tuberculosis: Table 1–. Eur Respir J. 2013 Dec;42(6):1752–7.
- 17. Berry MPR, Graham CM, McNab FW, Xu Z, Bloch SAA, Oni T, et al. An interferon-inducible neutrophil-driven blood transcriptional signature in human tuberculosis. Nature. 2010 Aug;466(7309):973–7. pmid:20725040
- 18. Kinsella RL, Nehls EM, Stallings CL. Roles for Autophagy Proteins in Immunity and Host Defense. Vet Pathol. 2018 May;55(3):366–73. pmid:29433400
- 19. Kimmey JM, Stallings CL. Bacterial Pathogens versus Autophagy: Implications for Therapeutic Interventions. Trends Mol Med. 2016 Dec;22(12):1060–76. pmid:27866924
- 20. Lamb CA, Yoshimori T, Tooze SA. The autophagosome: origins unknown, biogenesis complex. Nat Rev Mol Cell Biol. 2013 Dec;14(12):759–74. pmid:24201109
- 21. Mizushima N, Yoshimori T, Ohsumi Y. The Role of Atg Proteins in Autophagosome Formation. Annu Rev Cell Dev Biol. 2011 Nov 10;27(1):107–32. pmid:21801009
- 22. Mizushima N, Noda T, Yoshimori T, Tanaka Y, Ishii T, George MD, et al. A protein conjugation system essential for autophagy. Nature. 1998 Sep;395(6700):395–8. pmid:9759731
- 23. Golovkine GR, Roberts AW, Morrison HM, Rivera-Lugo R, McCall RM, Nilsson H, et al. Autophagy restricts Mycobacterium tuberculosis during acute infection in mice. Nat Microbiol [Internet]. 2023 Apr 10 [cited 2023 Apr 11]. Available from: https://www.nature.com/articles/s41564-023-01354-6. pmid:37037941
- 24. Manzanillo PS, Ayres JS, Watson RO, Collins AC, Souza G, Rae CS, et al. The ubiquitin ligase parkin mediates resistance to intracellular pathogens. Nature. 2013 Sep;501(7468):512–6. pmid:24005326
- 25. Köster S, Upadhyay S, Chandra P, Papavinasasundaram K, Yang G, Hassan A, et al. Mycobacterium tuberculosis is protected from NADPH oxidase and LC3-associated phagocytosis by the LCP protein CpsA. Proc Natl Acad Sci U S A [Internet]. 2017 Oct 10 [cited 2023 Apr 4];114(41). Available from: https://pnas.org/doi/full/ pmid:28973896
- 26. Aylan B, Bernard EM, Pellegrino E, Botella L, Fearns A, Athanasiadi N, et al. ATG7 and ATG14 restrict cytosolic and phagosomal Mycobacterium tuberculosis replication in human macrophages. Nat Microbiol [Internet]. 2023 Mar 23 [cited 2023 Apr 4]. Available from: https://www.nature.com/articles/s41564-023-01335-9. pmid:36959508
- 27. Dutta RK, Kathania M, Raje M, Majumdar S. IL-6 inhibits IFN-γ induced autophagy in Mycobacterium tuberculosis H37Rv infected macrophages. Int J Biochem Cell Biol. 2012 Jun;44(6):942–54.
- 28. Deretic V. Autophagy in Tuberculosis. Cold Spring Harb Perspect Med. 2014 Nov 1;4(11):a018481–a018481. pmid:25167980
- 29. Sakowski ET, Koster S, Portal Celhay C, Park HS, Shrestha E, Hetzenecker SE, et al. Ubiquilin 1 Promotes IFN-γ-Induced Xenophagy of Mycobacterium tuberculosis. Lewinsohn DM, editor. PLoS Pathog. 2015 Jul 30;11(7):e1005076.
- 30. Heckmann BL, Green DR. LC3-associated phagocytosis at a glance. J Cell Sci. 2019 Mar 1;132(5):jcs231472.
- 31. Wan J, Weiss E, Ben Mkaddem S, Mabire M, Choinier PM, Picq O, et al. LC3-associated phagocytosis protects against inflammation and liver fibrosis via immunoreceptor inhibitory signaling. Sci Transl Med. 2020 Apr 15;12(539):eaaw8523. pmid:32295902
- 32. Inomata M, Xu S, Chandra P, Meydani SN, Takemura G, Philips JA, et al. Macrophage LC3-associated phagocytosis is an immune defense against Streptococcus pneumoniae that diminishes with host aging. Proc Natl Acad Sci U S A. 2020 Dec 29;117(52):33561–9.
- 33. Martinez J, Almendinger J, Oberst A, Ness R, Dillon CP, Fitzgerald P, et al. Microtubule-associated protein 1 light chain 3 alpha (LC3)-associated phagocytosis is required for the efficient clearance of dead cells. Proc Natl Acad Sci U S A. 2011 Oct 18;108(42):17396–401. pmid:21969579
- 34. Sanjuan MA, Dillon CP, Tait SWG, Moshiach S, Dorsey F, Connell S, et al. Toll-like receptor signalling in macrophages links the autophagy pathway to phagocytosis. Nature. 2007 Dec;450(7173):1253–7. pmid:18097414
- 35. Cohen SB, Gern BH, Delahaye JL, Adams KN, Plumlee CR, Winkler JK, et al. Alveolar Macrophages Provide an Early Mycobacterium tuberculosis Niche and Initiate Dissemination. Cell Host Microbe. 2018 Sep;24(3):439–446.e4. pmid:30146391
- 36. Rothchild AC, Olson GS, Nemeth J, Amon LM, Mai D, Gold ES, et al. Alveolar macrophages generate a noncanonical NRF2-driven transcriptional response to Mycobacterium tuberculosis in vivo. Sci Immunol. 2019 Jul 5;4(37):eaaw6693.
- 37. Vieira S, Lemos H, Grespan R, Napimoga M, Dal-Secco D, Freitas A, et al. A crucial role for TNF-a in mediating neutrophil influx induced by endogenously generated or exogenous chemokines, KC/CXCL1 and LIX/CXCL5. Br J Pharmacol. 2009;11.
- 38. Bendall LJ, Bradstock KF. G-CSF: From granulopoietic stimulant to bone marrow stem cell mobilizing agent. Cytokine Growth Factor Rev. 2014 Aug;25(4):355–67. pmid:25131807
- 39. Semerad CL, Liu F, Gregory AD, Stumpf K, Link DC. G-CSF Is an Essential Regulator of Neutrophil Trafficking from the Bone Marrow to the Blood. Immunity. 2002 Oct;17(4):413–23. pmid:12387736
- 40. Pollara G, Turner CT, Rosenheim J, Chandran A, Bell LCK, Khan A, et al. Exaggerated IL-17A activity in human in vivo recall responses discriminates active tuberculosis from latent infection and cured disease. Sci Transl Med. 2021 May 5;13(592):eabg7673. pmid:33952677
- 41. Sasaki Y, Guo YM, Goto T, Ubukawa K, Asanuma K, Kobayashi I, et al. IL-6 Generated from Human Hematopoietic Stem and Progenitor Cells through TLR4 Signaling Promotes Emergency Granulopoiesis by Regulating Transcription Factor Expression. J Immunol. 2021 Aug 2;ji2100168. pmid:34341172
- 42. Hu S, He W, Du X, Yang J, Wen Q, Zhong XP, et al. IL-17 Production of Neutrophils Enhances Antibacteria Ability but Promotes Arthritis Development During Mycobacterium tuberculosis Infection. EBioMedicine. 2017 Sep;23:88–99. pmid:28821374
- 43. Fielding CA, McLoughlin RM, McLeod L, Colmont CS, Najdovska M, Grail D, et al. IL-6 Regulates Neutrophil Trafficking during Acute Inflammation via STAT3. J Immunol. 2008 Aug 1;181(3):2189–95. pmid:18641358
- 44. Saitoh T, Akira S. Regulation of inflammasomes by autophagy. J Allergy Clin Immunol. 2016 Jul;138(1):28–36. pmid:27373323
- 45. Harris J, Hartman M, Roche C, Zeng SG, O’Shea A, Sharp FA, et al. Autophagy Controls IL-1β Secretion by Targeting Pro-IL-1β for Degradation. J Biol Chem. 2011 Mar;286(11):9587–97.
- 46. Pang Y, Wu L, Tang C, Wang H, Wei Y. Autophagy-Inflammation Interplay During Infection: Balancing Pathogen Clearance and Host Inflammation. Front Pharmacol. 2022 Feb 22;13:832750. pmid:35273506
- 47. Ko JH, Yoon SO, Lee HJ, Oh JY. Rapamycin regulates macrophage activation by inhibiting NLRP3 inflammasome-p38 MAPK-NFκB pathways in autophagy- and p62-dependent manners. Oncotarget. 2017 Jun 20;8(25):40817–31.
- 48. Shi CS, Shenderov K, Huang NN, Kabat J, Abu-Asab M, Fitzgerald KA, et al. Activation of autophagy by inflammatory signals limits IL-1β production by targeting ubiquitinated inflammasomes for destruction. Nat Immunol. 2012 Mar;13(3):255–63.
- 49. Pyrillou K, Burzynski LC, Clarke MCH. Alternative Pathways of IL-1 Activation, and Its Role in Health and Disease. Front Immunol. 2020 Dec 18;11:613170. pmid:33391283
- 50. McElvania TeKippe E, Allen IC, Hulseberg PD, Sullivan JT, McCann JR, Sandor M, et al. Granuloma Formation and Host Defense in Chronic Mycobacterium tuberculosis Infection Requires PYCARD/ASC but Not NLRP3 or Caspase-1. Tyagi AK, editor. PLoS ONE. 2010 Aug 20;5(8):e12320. pmid:20808838
- 51. Beckwith KS, Beckwith MS, Ullmann S, Sætra RS, Kim H, Marstad A, et al. Plasma membrane damage causes NLRP3 activation and pyroptosis during Mycobacterium tuberculosis infection. Nat Commun. 2020 Dec;11(1):2270. pmid:32385301
- 52. Abdalla H, Srinivasan L, Shah S, Mayer-Barber KD, Sher A, Sutterwala FS, et al. Mycobacterium tuberculosis Infection of Dendritic Cells Leads to Partially Caspase-1/11-Independent IL-1β and IL-18 Secretion but Not to Pyroptosis. Kwaik YA, editor. PLoS ONE. 2012 Jul 24;7(7):e40722.
- 53. Mayer-Barber KD, Barber DL, Shenderov K, White SD, Wilson MS, Cheever A, et al. Cutting Edge: Caspase-1 Independent IL-1β Production Is Critical for Host Resistance to Mycobacterium tuberculosis and Does Not Require TLR Signaling In Vivo. J Immunol. 2010 Apr 1;184(7):3326–30.
- 54. Lin KM, Hu W, Troutman TD, Jennings M, Brewer T, Li X, et al. IRAK-1 bypasses priming and directly links TLRs to rapid NLRP3 inflammasome activation. Proc Natl Acad Sci U S A. 2014 Jan 14;111(2):775–80. pmid:24379360
- 55. Qiao Yu, Wang Peng, Qi Jianni, Zhang Lei, Gao Chengjiang. TLR-induced NF-jB activation regulates NLRP3 expression in murine macrophages. 2012;586:1022–6.
- 56. Goldberg MS, Fleming SM, Palacino JJ, Cepeda C, Lam HA, Bhatnagar A, et al. Parkin-deficient Mice Exhibit Nigrostriatal Deficits but Not Loss of Dopaminergic Neurons. J Biol Chem. 2003 Oct;278(44):43628–35. pmid:12930822
- 57. Lazarou M, Sliter DA, Kane LA, Sarraf SA, Wang C, Burman JL, et al. The ubiquitin kinase PINK1 recruits autophagy receptors to induce mitophagy. Nature. 2015 Aug;524(7565):309–14. pmid:26266977
- 58. Killackey SA, Philpott DJ, Girardin SE. Mitophagy pathways in health and disease. J Cell Biol. 2020 Nov 2;219(11):e202004029. pmid:32926082
- 59. Zhang W, Ren H, Xu C, Zhu C, Wu H, Liu D, et al. Hypoxic mitophagy regulates mitochondrial quality and platelet activation and determines severity of I/R heart injury. elife. 2016 Dec 20;5:e21407. pmid:27995894
- 60. Wang SH, Martin SM, Harris PS, Knudson CM. Caspase Inhibition Blocks Cell Death and Enhances Mitophagy but Fails to Promote T-Cell Lymphoma. Chandra D, editor. PLoS ONE. 2011 May 17;6(5):e19786. pmid:21611191
- 61. Nguyen TN, Padman BS, Lazarou M. Deciphering the Molecular Signals of PINK1/Parkin Mitophagy. Trends Cell Biol. 2016 Oct;26(10):733–44. pmid:27291334
- 62. Harris J, Deen N, Zamani S, Hasnat MA. Mitophagy and the release of inflammatory cytokines. Mitochondrion. 2018 Jul;41:2–8. pmid:29107116
- 63. Song Y, Zhou Y, Zhou X. The role of mitophagy in innate immune responses triggered by mitochondrial stress. Cell Commun Signal. 2020 Dec;18(1):186. pmid:33239048
- 64. Griffin GK, Newton G, Tarrio ML, Bu DV, Maganto-Garcia E, Azcutia V, et al. IL-17 and TNF-α Sustain Neutrophil Recruitment during Inflammation through Synergistic Effects on Endothelial Activation. J Immunol. 2012 Jun 15;188(12):6287–99.
- 65. Roussel L, Houle F, Chan C, Yao Y, Bérubé J, Olivenstein R, et al. IL-17 Promotes p38 MAPK-Dependent Endothelial Activation Enhancing Neutrophil Recruitment to Sites of Inflammation. J Immunol. 2010 Apr 15;184(8):4531–7. pmid:20228195
- 66. Halwani R, Al-Muhsen S, Hamid Q. T Helper 17 Cells in Airway Diseases. Chest. 2013 Feb;143(2):494–501.
- 67. Ding WX, Jaeschke H. Autophagy in macrophages regulates the inflammasome and protects against liver injury. J Hepatol. 2016 Jan;64(1):16–8. pmid:26456339
- 68. Bussi C, Peralta Ramos JM, Arroyo DS, Gaviglio EA, Gallea JI, Wang JM, et al. Autophagy down regulates pro-inflammatory mediators in BV2 microglial cells and rescues both LPS and alpha-synuclein induced neuronal cell death. Sci Rep. 2017 Mar 3;7(1):43153. pmid:28256519
- 69. Peng X, Wang Y, Li H, Fan J, Shen J, Yu X, et al. ATG5-mediated autophagy suppresses NF-κB signaling to limit epithelial inflammatory response to kidney injury. Cell Death Dis. 2019 Mar 15;10(4):253.
- 70. Chang CP, Su YC, Hu CW, Lei HY. TLR2-dependent selective autophagy regulates NF-κB lysosomal degradation in hepatoma-derived M2 macrophage differentiation. Cell Death Differ. 2013 Mar;20(3):515–23.
- 71. Liu K, Zhang L, Zhao Q, Zhao Z, Zhi F, Qin Y, et al. SKP2 attenuates NF-κB signaling by mediating IKKβ degradation through autophagy. Su B, editor. J Mol Cell Biol. 2018 Jun 1;10(3):205–215.
- 72. Yang Q, Liu TT, Lin H, Zhang M, Wei J, Luo WW, et al. TRIM32-TAX1BP1-dependent selective autophagic degradation of TRIF negatively regulates TLR3/4-mediated innate immune responses. Philpott DJ, editor. PLoS Pathog. 2017 Sep 12;13(9):e1006600. pmid:28898289
- 73. Lim J, Park H, Heisler J, Maculins T, Roose-Girma M, Xu M, et al. Autophagy regulates inflammatory programmed cell death via turnover of RHIM-domain proteins. elife. 2019 Jul 9;8:e44452. pmid:31287416
- 74. Bock FJ, Riley JS. When cell death goes wrong: inflammatory outcomes of failed apoptosis and mitotic cell death. Cell Death Differ. 2023 Feb;30(2):293–303. pmid:36376381
- 75. Tang D, Kang R, Coyne CB, Zeh HJ, Lotze MT. PAMPs and DAMPs: signal 0s that spur autophagy and immunity. Immunol Rev. 2012 Sep;249(1):158–75. pmid:22889221
- 76. Zhao X, Khan N, Gan H, Tzelepis F, Nishimura T, Park SY, et al. Bcl-xL mediates RIPK3-dependent necrosis in M. tuberculosis-infected macrophages. Mucosal Immunol. 2017;10(6):1553–1568. pmid:28401933
- 77. Yousefi S, Perozzo R, Schmid I, Ziemiecki A, Schaffner T, Scapozza L, et al. Calpain-mediated cleavage of Atg5 switches autophagy to apoptosis. Nat Cell Biol. 2006 Oct;8(10):1124–32. pmid:16998475
- 78. Mázló A, Jenei V, Burai S, Molnár T, Bácsi A, Koncz G. Types of necroinflammation, the effect of cell death modalities on sterile inflammation. Cell Death Dis. 2022 May 2;13(5):423. pmid:35501340
- 79. Doran AC, Yurdagul A, Tabas I. Efferocytosis in health and disease. Nat Rev Immunol. 2020 Apr;20(4):254–67. pmid:31822793
- 80. Asare PF, Roscioli E, Hurtado PR, Tran HB, Mah CY, Hodge S. LC3-Associated Phagocytosis (LAP): A Potentially Influential Mediator of Efferocytosis-Related Tumor Progression and Aggressiveness. Front Oncol. 2020 Aug 5;10:1298. pmid:32850405
- 81. Moretti J, Roy S, Bozec D, Martinez J, Chapman JR, Ueberheide B, et al. STING Senses Microbial Viability to Orchestrate Stress-Mediated Autophagy of the Endoplasmic Reticulum. Cell. 2017;171(4):809–823.e13. pmid:29056340
- 82. Chipurupalli S, Samavedam U, Robinson N. Crosstalk Between ER Stress, Autophagy and Inflammation. Front Med. 2021;5(8):758311. pmid:34805224
- 83. Niven J, Madelon N, Page N, Caruso A, Harlé G, Lemeille S, et al. Macroautophagy in Dendritic Cells Controls the Homeostasis and Stability of Regulatory T Cells. Cell Rep. 2019 Jul;28(1):21–29.e6. pmid:31269441
- 84. Peral de Castro C, Jones SA, Ní Cheallaigh C, Hearnden CA, Williams L, Winter J, et al. Autophagy Regulates IL-23 Secretion and Innate T Cell Responses through Effects on IL-1 Secretion. J Immunol. 2012 Oct 15;189(8):4144–53.
- 85. Cho ML, Kang JW, Moon YM, Nam HJ, Jhun JY, Heo SB, et al. STAT3 and NF-κB Signal Pathway Is Required for IL-23-Mediated IL-17 Production in Spontaneous Arthritis Animal Model IL-1 Receptor Antagonist-Deficient Mice. J Immunol. 2006 May 1;176(9):5652–61.
- 86. Sutton C, Brereton C, Keogh B, Mills KHG, Lavelle EC. A crucial role for interleukin (IL)-1 in the induction of IL-17–producing T cells that mediate autoimmune encephalomyelitis. J Exp Med. 2006 Jul 10;203(7):1685–91. pmid:16818675
- 87. Gawriluk TR, Hale AN, Flaws JA, Dillon CP, Green DR, Rucker EB. Autophagy is a cell survival program for female germ cells in the murine ovary. Reproduction. 2011 Jun;141(6):759–65. pmid:21464117
- 88. Hara T, Nakamura K, Matsui M, Yamamoto A, Nakahara Y, Suzuki-Migishima R, et al. Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature. 2006 Jun;441(7095):885–9. pmid:16625204
- 89. Hwang S, Maloney NS, Bruinsma MW, Goel G, Duan E, Zhang L, et al. Nondegradative Role of Atg5-Atg12/ Atg16L1 Autophagy Protein Complex in Antiviral Activity of Interferon Gamma. Cell Host Microbe. 2012 Apr;11(4):397–409. pmid:22520467
- 90. Martinez J, Malireddi RKS, Lu Q, Cunha LD, Pelletier S, Gingras S, et al. Molecular characterization of LC3-associated phagocytosis reveals distinct roles for Rubicon, NOX2 and autophagy proteins. Nat Cell Biol. 2015 Jul;17(7):893–906. pmid:26098576
- 91. Kitada T, Pisani A, Porter DR, Yamaguchi H, Tscherter A, Martella G, et al. Impaired dopamine release and synaptic plasticity in the striatum of PINK1 -deficient mice. Proc Natl Acad Sci U S A. 2007 Jul 3;104(27):11441–6.
- 92. Huynh JP, Lin CC, Kimmey JM, Jarjour NN, Schwarzkopf EA, Bradstreet TR, et al. Bhlhe40 is an essential repressor of IL-10 during Mycobacterium tuberculosis infection. J Exp Med. 2018 Jul 2;215(7):1823–38. pmid:29773644
- 93. Domingo-Gonzalez R, Das S, Griffiths KL, Ahmed M, Bambouskova M, Gopal R, et al. Interleukin-17 limits hypoxia-inducible factor 1α and development of hypoxic granulomas during tuberculosis. JCI Insight. 2017 Oct 5;2(19):e92973.