PNAS PLUS
The autophagic machinery ensures nonlytic transmission of mycobacteria Lilli Gerstenmaiera,1, Rachel Pillaa,1, Lydia Herrmanna, Hendrik Herrmanna,2, Monica Pradoa, Geno J. Villafanoa, Margot Kolonkoa, Rudolph Reimerb, Thierry Soldatic, Jason S. Kingd, and Monica Hagedorna,3 a Section Parasitology, Bernhard Nocht Institute for Tropical Medicine, 20359 Hamburg, Germany; bElectronmicroscopy, Heinrich-Pette-Institute, 20251 Hamburg, Germany; cDepartment of Biochemistry, University of Geneva, 1211-Geneva, Switzerland; and dDepartment of Biomedical Sciences, University of Sheffield, Sheffield S10 2TN, United Kingdom
In contrast to mechanisms mediating uptake of intracellular bacterial pathogens, bacterial egress and cell-to-cell transmission are poorly understood. Previously, we showed that the transmission of pathogenic mycobacteria between phagocytic cells also depends on nonlytic ejection through an F-actin based structure, called the ejectosome. How the host cell maintains integrity of its plasma membrane during the ejection process was unknown. Here, we reveal an unexpected function for the autophagic machinery in nonlytic spreading of bacteria. We show that ejecting mycobacteria are escorted by a distinct polar autophagocytic vacuole. If autophagy is impaired, cell-to-cell transmission is inhibited, the host plasma membrane becomes compromised and the host cells die. These findings highlight a previously unidentified, highly ordered interaction between bacteria and the autophagic pathway and might represent the ancient way to ensure nonlytic egress of bacteria. autophagy
| Dictyostelium discoideum | Mycobacterium marinum | ejection
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n recent years, our understanding of the interactions between the host autophagic machinery and intracellular pathogens has rapidly expanded. These interactions are complex; although, in many cases, the engagement of autophagy protects the host by capturing and destroying the pathogen, some bacteria actively subvert this pathway to promote their own survival (reviewed in ref. 1). Autophagy has also been suggested to promote cell-tocell transmission of Brucella (2, 3), although the molecular mechanisms are unknown. Both Mycobacterium tuberculosis, which causes tuberculosis in humans, and the closely related species M. marinum have been shown to interact with the autophagy machinery of their host cell (4–7). After uptake by immune phagocytes, the bacteria arrest phagosomal maturation and convert their vacuole into a replication-permissive compartment. Both bacteria can translocate into the host cell cytosol dependent on an intact Region-of-Difference1-locus (RD1) (8–11). The genomic RD1-locus encodes a secretion system, ESX-1 (Type-VII secretion system), which has been associated with mycobacterial virulence (ref. 12, reviewed in refs. 13 and 14). Once in the cytosol, M. marinum becomes ubiquitinated (4) likely recruiting adaptor proteins, such as members of the sequestosome-1 family (SQSTM1), which also bind LC3 (microtubule-associated proteins 1A/1B light chains 3A/LC3A and 3B/LC3B), here referred to as Atg8, on autophagosomal membranes. In this way, bacteria are normally targeted to autophagosomes and killed, but M. marinum efficiently escapes this fate, most probably by shedding the ubiquitinated material as a decoy (4). However, infection by M. tuberculosis can be overcome by stimulating the classic autophagic pathway (15) and autophagy can reduce the bacterial burden in vivo (7). It was previously thought that M. marinum and M. tuberculosis leave their host cell by inducing necrotic or apoptotic cell death (16). However, we recently showed that these bacteria also exit their host cell and spread via an F-actin structure, termed the ejectosome (17). This form of egress, which is common to www.pnas.org/cgi/doi/10.1073/pnas.1423318112
M. tuberculosis and M. marinum in the amoeba Dictyostelium, is nonlytic for the host cell, even though its plasma membrane is perforated at the site of ejection. Previously, we showed that ejectosome formation is dependent on ESAT-6 (Early secretory antigenic target 6), a secreted virulence factor encoded in the RD1-locus, and the Dictyostelium small GTPase RacH. However, both the structure and mechanistic details of ejectosome function remain unknown. Using the Dictyostelium–M. marinum system (9, 17, 18) to further dissect the mechanism of ejectosome formation and function, we demonstrate an unexpected role for autophagic membranes in both mycobacteria egress and concomitant cell-tocell transmission. Results Correlative Microscopy Reveals a Vacuolar Structure at the Distal Pole of Ejecting Bacteria. To better understand the mechanism of
nonlytic bacterial ejection, we examined the ultrastructure of the ejectosome. Using a correlative approach, we were able to identify ejectosomes by fluorescence microscopy (Fig. 1A and Fig. S1 A and E), before ultrastructural analysis of serial thin sections by transmission electron microscopy (TEM) (Fig. 1 B–D and Movie S1). Fig. 1C shows a representative section of a bacterium at a very late stage of ejection with the proximal pole Significance Pathogenic mycobacteria can be transmitted by direct ejection from one host cell to another. However, the mechanism of ejection, and how lysing the host cell is prevented are unknown. This study explains how the host cell remains intact and alive while Mycobacterium marinum breaks through its plasma membrane during ejection. We show that a membraneous cup is specifically recruited to the distal pole of ejecting M. marinum. We demonstrate that these membranes are formed by the canonical autophagic pathway, though they do not mature to autophagolysosomes. Disruption of autophagy causes the host cells to become leaky and die during ejection. This dramatically reduces cell-to-cell transmission of the infection, demonstrating an important and unexpected role for autophagy in maintaining plasma membrane integrity during mycobacterial infection. Author contributions: L.G., R.P., R.R., T.S., and M.H. designed research; L.G., R.P., L.H., H.H., M.P., G.J.V., M.K., J.S.K., and M.H. performed research; L.G., R.P., H.H., M.P., G.J.V., M.K., and M.H. analyzed data; R.R., T.S., J.S.K., and M.H. contributed new reagents/analytic tools; and J.S.K. and M.H. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. 1
L.G. and R.P. contributed equally to this work.
2
Deceased July 28, 2014.
3
To whom correspondence should be addressed. Email:
[email protected].
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. 1073/pnas.1423318112/-/DCSupplemental.
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Edited by Ralph R. Isberg, Howard Hughes Medical Institute, Tufts University School of Medicine, Boston, MA, and approved January 7, 2015 (received for review December 9, 2014)
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The presence of Atg8-containing membranes at bacteria during ejection indicates the specific recruitment of the autophagic machinery. Selective autophagy is mediated by ubiquitination of the target and recruitment of adaptor proteins such as SQSTM1 that contain both ubiquitin- and Atg8-binding domains (reviewed in ref. 20). Consistent with this pathway, both ubiquitin and DdSQSTM1 (GFP-SQSTM1), the single Dictyostelium ortholog of SQSTM1 accumulated in a pocket around the distal pole of ejecting bacteria, similarly to Atg8 (Fig. 2 E–G). Extracellular M. marinum was never associated with Atg8 or GFPSQSTM1, indicating that the autophagic membrane is retained inside the host. Importantly, the bacterial localization of both ubiquitin and Atg8 was largely restricted to ejecting bacteria. Although less than 15% of total cytoplasmic bacteria were associated with ubiquitin (Fig. 2 H and I) or Atg8 (Fig. 2 H and J)
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(white arrowhead) being extracellular and the distal pole (white arrow) lagging behind within the cell. The plasma membrane, which is ruptured at the site of ejection, is tightly apposed to the bacterium (Fig. 1C’, arrows). Strikingly, in every electron micrograph, the distal pole of the bacterium in the course of ejection was tightly enclosed by a vacuolar structure. Both spacious (with an electron lucent lumen, Fig. 1D and Fig. S1 F and G), and flat cisternal structures were observed (Fig. S1 B–D), but 3D-reconstruction using serial thin sections revealed these cisternal structure always formed a vacuole around the distal bacterial pole (Fig. 1E, Fig. S2, and Movie S2).
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Movie S2) is reminiscent of an expanding autophagic membrane. We therefore looked for the presence of autophagosomal markers around bacteria in the course of ejection. Consistent with the TEM images, immunofluorescence imaging showed that Atg8, a protein that specifically incorporates into expanding autophagosomal membranes (19), strongly localized around the distal pole of ejecting M. marinum (Fig. 2A and Movie S3). This Atg8-positive cup was associated with M. marinum at all stages of ejection, from the very early stages when the bacterium is mainly inside the cell, until the bacterium is completely ejected, where an Atg8-positive cup still caps the extremity of the almost fully extracellular bacterium (Fig. 2 B–D). 2 of 6 | www.pnas.org/cgi/doi/10.1073/pnas.1423318112
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Fig. 1. A vacuole caps the distal pole of ejecting M. marinum. (A) A Dictyostelium cell ejecting a M. marinum bacterium was localized by confocal fluorescence microscopy. The bacterium is shown in red, actin is shown in green. (B) Overlay of the corresponding brightfield and transmission electron microscopy (TEM) image after processing. (Scale bar: 2 μm.) (C) TEM image of the ejecting bacterium. The white arrow indicates the distal pole of the ejecting bacterium, the white arrowhead points toward the proximal pole. (Scale bar: 2 μm.) (C’) shows a magnification of the region where the bacterium perforates the plasma membrane. Typical for ejection, the plasma membrane is protruding and tightly apposed to the ejecting bacterium (indicated by black arrows). (D) High magnification of the distal pole of the ejecting bacterium. (Scale bar: 500 nm.) Black arrowheads indicate the vacuolar membrane apposed to the bacterium, black arrows point to the membrane exposed to the host cell cytosol. The actin-rich ejectosome is indicated with an asterisk in all images. (E) 3D-model of the vacuolar pocket around the pole of an ejecting bacterium. Serial thin sections were imaged by TEM and surface rendered. Bacteria are depicted in red, polar vacuole is depicted in yellow, and plasma membrane is depicted in green.
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Fig. 2. The autophagic machinery is specifically recruited to the distal pole of ejecting M. marinum. (A–D) Fluorescence microscopy of infected cells fixed and stained with anti-Atg8 showing polar accumulation of Atg8 at different stages of ejection. Early in ejection, Atg8 (green) is found as a large pocket around the bacterium (outlined with a broken white line) (B), which gets more focused as the bacterium becomes more extracellular (C and D). The outline of the cell is depicted with a red line. (E–G) Fluorescence micrograph of an infected cell expressing GFP (E), GFP-SQSTM1 (F), or stained with anti-ubiquitin (UB; G). (A and E–G) Bacteria are shown in blue, actin is shown in red, and the respective marker is shown in green. Ejectosomes are indicated by a white arrow, the distal poles of bacteria are highlighted with a white arrowhead. (H) Quantification of association of ubiquitin and Atg8 with cytoplasmic and ejecting bacteria. n = 3. (I and J) Cytoplasmic M. marinum associated with ubiquitin (UB, I) and Atg8 (J). The bacteria are shown in blue, the respective marker is shown in green. (Scale bars: 2 μm.)
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Fig. 3. The atg1− mutant forms nonfunctional ejectosomes. (A) Ejectosomes are present in atg1− cells (white arrowheads). Actin is shown in red, bacterium in blue. (B) Ejectosome frequency is unaffected in atg1− cells compared with wild-type cells (WT), n = 3. Infected atg1− cells stained for Atg8 (C) illustrate the absence of Atg8 recruitment. Actin is in red, Atg8 antibody staining in green. (D and E) Transmission electron micrograph of an ejectosome in atg1− cells confirms absence of the distal vacuole. (D) The ejectosome region is indicated with white arrowheads. (Scale bar: 1 μm.) (E) The distal pole of the ejecting bacterium is cytosolic and not capped by a vacuole. (Scale bar: 500 nm.) Material spilling from the ejectosome region is indicated with black arrows (D and E). (F) A transmission electron micrograph of the distal pole of an ejecting bacterium from wild-type cells (WT). The vacuole engulfing the bacterium is indicated with black arrows. (Scale bar: 500 nm.)
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Fig. 4. Elimination of autophagy reduces cell-to-cell transmission and causes cell leakage. (A) Dictyostelium knockout mutants were infected with M. marinum and the recruitment of Atg8 to the distal poles of ejectosomes was scored in three independent experiments. Statistical significance was calculated using the one way ANOVA. (***P ≤ 0.001; n.s., not significant). (B) Wild-type cells expressing GFP-Atg18 and GFP-2xFYVE were infected with M. marinum and the accumulation of the respective marker at the distal pole of ejecting bacteria was scored (n = 3). (C and D) Dictyostelium wild-type cells were coinfected (Fig. S5) with M. marinum wild-type and M. marinum ΔRD1 (C) and M. smegmatis (D), respectively. Ejectosome structures (white arrowheads) are formed by both, M. marinum ΔRD1 (C, blue) and M. smegmatis (D, blue). Distinct Atg8 (green) localization at the distal pole (white arrows) is observed for both, M. marinum ΔRD1 (C) and M. smegmatis (D) alike M. marinum wild-type. (E) Cell-to-cell transmission was measured by flow cytometry. Acceptor cells (wild-type) were mixed with infected donor cells (wild-type, racH−, and atg1−) at 6 hpi and transmission measured at 6, 24, 28 and 32 hpi. Both racH− and atg1− cells are significantly impaired in transmitting bacteria to acceptor cells (n = 3, mean ± SEM; *P ≤ 0.1; **P ≤ 0.01). Two-way ANOVA analysis). (F and G) Ejectosome structures (white arrowhead) formed by atg1− cells showed spilling of cytosolic Atg8 (green) on the extracellular side of the ejecting bacterium (indicated by white arrow). (Scale bar: 2 μm.) (H) In contrast to wild-type (WT) Dictyostelium cells, atg1− cells show a significant increase in mean PI accumulation upon M. marinum infection (24 hpi), n = 3. Similarly, cell death was significantly increased (I) when atg1− cells were infected with M. marinum, whereas wild-type cells were unaffected (24 hpi). Statistical analysis was performed using one-way ANOVA analysis (n = 3–4; mean ± SEM; ***P ≤ 0.001). (Scale bars: 2 μm.)
To characterize this compartment further, we also examined the localization of both GFP-Atg18 (ortholog of the mammalian proteins WIPI1/2), and the PI(3)P reporter GFP-2xFYVE. Canonically, both proteins are only recruited to the early, expanding PNAS Early Edition | 3 of 6
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The Canonical Autophagy Machinery Is Required for Cup Formation.
To determine whether the recruitment of Atg8-positive membrane to ejecting bacteria required the canonical autophagy pathway, we infected Dictyostelium mutants with a defective atg1 gene (atg1−). Atg1 (Ulk1/2) is a key component of the canonical autophagy initiation complex and the Dictyostelium atg1− mutant is strictly impaired in autophagy (21). atg1− null cells were however still able to form ejectosomes closely resembling the actin-rich structures generated in wild-type cells, with the same frequency (Fig. 3 A and B). In atg1− cells, none of the ejecting bacteria were associated with Atg8 (Fig. 3C), although half of them (46.4% ± 3.6) retained the association with ubiquitin at their distal pole (Fig. S4). Consistent with this, TEM analysis in atg1− cells showed a total loss of the vacuolar cup structures around the distal pole of ejecting bacteria (Fig. 3 D and E). To test the requirement for other components of the canonical autophagic pathway, we also tested for the recruitment of Atg8 to ejecting bacteria in mutants of the ubiquitin-like conjugation machinery (Atg5 and Atg7) and the phosphatidyl inositol 3-kinase (PI3K) complex (Atg6/Beclin1−) (Fig. 4A). We found that atg5 and atg7 were strictly required for association of Atg8 with the ejectosome, whereas disruption of one of the two Dictyostelium atg6 orthologs gave a significant, although less penetrant defect, consistent with the partial block in autophagy reported by others (22) (Fig. 4A). We therefore conclude that the core autophagic machinery is required for the formation of the ejectosome-associated vacuole.
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at 24 h postinfection (hpi), all ejecting bacteria were labeled at their intracellular pole (Fig. 2H). Furthermore, although the nonejecting cytosolic bacteria recruited ubiquitin, atg8, and GFP-SQSTM1 to patches along their entire surface (Fig. 2 I and J and Fig. S3), this was restricted to the distal pole during ejection. These observations indicate that the recruitment of the autophagy machinery during ejection is by a specialized pathway with a higher level of regulation and organization.
isolation membrane, in contrast to Atg8, which remains associated with the autophagosome throughout (23, 24). We observed that Atg18 localized to 38 ± 2.9% and 2xFYVE was present at 8.9 ± 4.4% of ejecting bacteria (Fig. 4B). This indicates that the pocket is formed at the site of ejection, by the canonical autophagic pathway. Furthermore, as Atg8 could be observed on >90% of ejecting bacteria (Fig. 2H), this pocket must also be undergoing maturation to some extent. However, when we labeled the lysosomal system by feeding cells fluorescent dextran, we never observed an accumulation at ejecting bacteria, indicating that fusion with endolysosomes does not occur (Fig. S6). To examine how the autophagic machinery is recruited, we also disrupted the recently identified Dictyostelium ortholog of SQSTM1, the sole known autophagic adaptor in this organism (22). Surprisingly however, loss of SQSTM1 had no affect on Atg8 recruitment. The alternative autophagy adaptors identified in mammals are poorly conserved through evolution and no clear orthologs can be identified in Dictyostelium. Therefore, either additional unknown adaptors must exist, or an alternative mechanism of recruitment is responsible for the recruitment of the autophagy machinery. Recruitment of the Autophagic Machinery Is ESX-1 Independent. The specific recruitment of the autophagy machinery to the distal pole of ejecting bacteria raises the question whether polar mycobacterial virulence factors are involved in this recruitment and whether this is specific for pathogenic M. marinum. The virulence secretion system ESX-1 is enriched at the poles of M. marinum (25) and is encoded in the RD1-region. Involvement of this secretion system in ejection has been demonstrated before (17). We therefore asked whether local secretion activity of ESX-1 at the pole might be involved in the observed activation of autophagy. As ESX-1 is required for the bacteria to escape from phagosomes into the cytosol before ejection, we established a coinfection protocol that used wild-type M. marinum to break phagosomes that contained M. marinum mutants lacking the RD1-locus (M. marinum ΔRD1) at the same time (assay depicted in Fig. S5). Immunofluorescence microscopy showed that both wild-type and ΔRD1 M. marinum, were able to form ejectosomes and recruit Atg8 to their distal poles (Fig. 4C). Therefore, ESX-1 –mediated ESAT-6 secretion is required for ejectosome formation (17), the polarization of this secretion system and its local activity does not mediate recruitment of the autophagy machinery. To monitor whether the observed ejection is restricted to pathogenic mycobacteria we performed coinfection with nonpathogenic M. smegmatis. Immunofluorescence micrographs showed that M. smegmatis was indeed able to form an ejectosome and the autophagy machinery was recruited to its distal pole (Fig. 4D). Autophagy Is Required for Efficient Nonlytic, Cell-to-Cell Transmission.
Although not required for their formation, we next asked whether autophagy was required for ejectosome function. Ejection is the major mechanism of cell-to-cell transmission in Dictyostelium (17). Therefore, to determine whether ejectosomes generated in atg1deficient cells are functional, we used a flow cytometry-based assay to monitor the transmission of GFP-expressing M. marinum from mutant donor cells to mRFP-expressing wild-type acceptor cells (Fig. S7). The acceptor cells were added to infected donor cells (wild-type, atg1−, or racH−) at 6 hpi and transmission was measured starting from 24 hpi onwards. Whereas wild-type cells efficiently transmitted the bacteria to the acceptor cells, transmission was reduced by 50% in atg1− mutants (Fig. 4E). This decrease in transmission is comparable to the defect in racH− cells that we previously showed to be deficient in nonlytic cell-to-cell transmission due to their complete inability to form ejectosomes (17). Therefore, although not required for ejectosome biogenesis, autophagy is essential for efficient cell-to-cell transmission. 4 of 6 | www.pnas.org/cgi/doi/10.1073/pnas.1423318112
Although atg1− cells still form a tightly localized actin ring around ejecting bacteria, we frequently observed by light-microscopy cytoplasmic material spilling out on the extracellular side of the ejectosome (Fig. 4 F and G). Extracellular material around the ejecting bacterium was also observed by EM (Fig. 3 D and E, black arrows). This was never observed in wild-type cells, suggesting that the ejectosomes in atg1− cells may not be closing as tightly around the ejecting bacteria. We therefore used the nuclear accumulation of the membrane impermeable dye propidium iodide (PI) to determine the membrane integrity of infected autophagy-deficient cells. After a 10 min exposure of infected cells (24 hpi) to PI, 40 ± 3.2% of live atg1− cells showed a significantly higher mean fluorescence than noninfected atg1− cells indicating that the cells become partially permeable to the dye (Fig. 4H and Fig. S8). In contrast, the mean fluorescence of wild-type cells increased only moderately (10 ± 6%) upon infection. Cell leakiness will also eventually result in cell death. Although other ejectosome-independent roles of autophagy may also contribute to this, when we quantified the number of dead cells at 24 hpi using the PI-staining and the characteristic forward/side scatter of dead cells (Fig. S8) we found significantly more dead cells when autophagy is blocked (Fig. 4I). Discussion In this paper, we describe a previously unidentified interaction between pathogenic M. marinum and the autophagic machinery. In contrast to previously described host-pathogen interactions, this is specific to ejecting bacteria and is required for the transmission of bacteria to a naïve host cell. Although this mechanism of transmission clearly benefits the bacteria, it might be considered to also benefit the host, as the polar autophagic vacuole apparently prevents plasma membrane leakage and host cell death. Recently, a number of studies have demonstrated autophagyindependent roles for several autophagy-related proteins. We show that the recruitment of the double-membrane cup to ejecting bacteria requires proteins from the canonical autophagy pathway. Importantly, this includes the Atg1 initiation complex, which is dispensible for the autophagy-independent recruitment of Atg8/LC3 to the single membranes of phagosomes and macropinosomes in mammalian cells (26). In addition, the requirement for the PI3K complex subunit Atg6 and the ubiquitin-like conjugation machinery components Atg5 and Atg7, as well as the presence of PI(3)P and Atg18 at the bacterial pole indicate that the ejectosome-associated vesicle is formed at the site of ejection by the canonical autophagy machinery. However, although the presence of GFP-Atg18 on only a subset of these vesicles indicates that they undergo some maturation, the lack of dextran delivery to this compartment implies that it does not undergo fusion with the endolysosomal system and therefore is unlikely to become degradative. The autophagic membrane is tightly restricted to the distal pole of ejectosome-associated bacteria, however, it is unclear how this polar recruitment is orchestrated. It has previously been reported that the autophagy machinery can be recruited to damaged membranes via galectins (27) and the protrusion of a bacterium through the plasma membrane may well be interpreted by the cell as plasma membrane wounding. However, at early stages of ejection, the distal pole of these rod-shaped bacteria is several microns away from the plasma membrane (Fig. S1C), and it is therefore possible that the recruitment of this vacuole is directed by the bacteria themselves in some way. The virulence secretion system ESX-1 localizes to the poles of M. marinum (25) and is encoded in the RD1-region. Involvement of this secretion system in ejection has been demonstrated before (17) in particular the secreted factor ESAT-6. However, the results of our coinfection assay show that neither the RD1 region nor M. marinum-specific virulence factors are responsible for Gerstenmaier et al.
Materials and Methods Dictyostelium Cell Culture. Wild-type Dictyostelium cells (Ax2) were axenically cultivated in HL5c medium (Formedium) at 22 °C. The Dictyostelium atg1− cells were described (34) and racH− cells from F. Rivero (University of Hull, Hull, United Kingdom). Dictyostelium mutants atg5−, atg6−, and atg7− were received from dictyBase (www.dictybase.org). Dictyostelium mutant SQSTM1- was generated using the pKOSG-IBA-Dicty1 system from Stargate.
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Mycobacteria Cultivation, Strains, and Plasmids. Mycobacteria were cultivated with agitation at 32 °C in Middlebrook 7H9 (Difco), supplemented with 10% (vol/vol) OADC (oleic acid, albumin, dextrose, catalase) (Becton Dickinson), 5% (vol/vol) glycerol and 0.2% Tween80 (Sigma Aldrich). M. marinum wild type (M-strain) and the msp12::GFP plasmid were kindly provided by L. Ramakrishnan (Washington University, Seattle) and the pCHERRY3 plasmid (No. 24659; ref. 35; Tanya Parish, Infectious Disease Research Institute, Queen Mary’s School of Medicine, University of Washington, Seattle) was supplied by Addgene. Transformed bacteria were selected either with hygromycin (pCHERRY) or kanamycin (msp12::GFP) at 50 μg/mL. Phalloidin and Antibodies. Rabbit polyclonal antibody was raised against fulllength recombinant Atg8, FK2H (directed against ubiquitinylated conjugates) was bought from Enzo (BML-PW0150), and anti-RFP from Chromotek (5F8). Secondary antibodies were goat anti-rabbit or goat anti-mouse/chicken IgG coupled to Alexa 488, Alexa 568, or Alexa 647 (Invitrogen). Phalloidin staining was performed with Alexa Fluor488-, Alexa Fluor568-, or Alexa Fluor647phalloidin (Molecular Probes). Infection Assay. Mycobacteria were grown to an OD600 =1 (∼5 × 108 cells per mL) and clumps disrupted by passing through 50 μm CellTrics-filters (Partec) and blunt 25-gauge needles (Dispomed, Neoject). Adherent Dictyostelium wild type or mutant cells were grown overnight without antibiotics to a density of 80–100%. Bacteria were added at a multiplicity of infection (MOI) of 50–100 and centrifuged on the Dictyostelium cells (500 × g, four times for 5 min each). The cells were left to phagocytose for additional 10–30 min before free bacteria were washed off with HL5c. To inhibit extracellular proliferation of bacteria the cells were subsequently resuspended in HL5c supplemented with 5 μg/mL streptomycin and maintained at 25 °C. The moment when the bacteria were added to the attached Dictyostelium cells was defined as 0 hpi. Immunofluorescence and Phalloidin Staining. Dictyostelium cells were infected as above with GFP or mCHERRY expressing M. marinum. At 24 hpi, cells were centrifuged on poly-lysine coated coverslips (500 × g, 5 min) and fixed either in methanol (−80 °C, 1 h) or with Soerensen buffer (14.7 mM KH2PO4, 2.5 mM NaHPO4, pH 6.3) containing 4% paraformaldehyde and stained as described (9, 36). The fluorescence images were documented using an Olympus IX81 confocal microscope with a 60× 1.35 NA or 100× oil immersion objective and Fluoview software v1.7b. Recording parameters for fields of 1024 × 1024 pixels with appropriate electronic zoom (6–12×) were 3× line averaging (Kalman). To adjust the brightness and contrast of complete images ImageJ (imagej.nih.gov/ij/) was used. Correlative Light and Electron Microscopy (CLEM). The combination of light and electron microscopy was used to analyze the ultrastructure of ejectosomes. Adherent Dictyostelium cells expressing Lifeact-GFP were infected as described above with mCHERRY expressing M. marinum. Infected cells were plated in a sterile 35 mm μ-dish with an imprinted grid (Ibidi). After 24 h post infection (hpi), the cells were fixed for two hours with 2% paraformaldehyde (Electron Microscopy Sciences) and 0.25% glutaraldehyde (Electron Microscopy Sciences) in HL5c medium. Fixed cells were washed and stained for 30 min with Alexa Fluor488-phalloidin. Localization and fluorescence images of ejectosome structures were documented using the Olympus IX81 confocal microscope as described above. Subsequently, the cells were fixed with 2.5% glutaraldehyde in 1× PBS (137 mM NaCl, 2.7 mM KCl, pH 7.4) overnight. After washing the samples twice, they were incubated with OsO4 (1% in PBS, 30 min), washed again and finally incubated with gallic acid (1% in 0.5× PBS, 30 min). After sample dehydration in a rising ethanol series they were embedded in 50% Epoxy resin (Roth, 8619.2) in 100% Ethanol overnight at room temperature and polymerized at 60 °C with fresh 100% Epoxy resin for 6 h. Consecutive ultrathin sectioning was performed with a Leica EM UC7 μL tramicrotome and the samples were examined with a Tecnai Spirit transmission electron microscope at 80 kV (FEI). For 3D reconstruction the image stack was aligned with ImageJ (stackreg; imagej.nih.gov/ij/) and reconstructed in Imaris (6.2). Movies were exported from Imaris and rendered in AdobeAfterEffects and AdobeMediaEncoder. Transmission Assay. All Dictyostelium strains (Ax2, racH−, and atg1−, donor cells) were infected as described above with GFP-expressing M. marinum. Infection rates were measured by flow cytometry (Accuri C6, BD Biosciences), and normalized to each other by dilution with the appropriate strain. At 6 hpi the infected donor cells were mixed with RFP-expressing acceptor Dictyostelium cells at a 1:5 ratio. At each timepoint (6, 24, 28, and 32 hpi), cells were
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distal vacuole generation, which would rather favor a host-driven process. At a molecular level, the accumulation of ubiquitin and SQSTM1 suggest recruitment of the autophagy machinery by factors which are directly present on the bacterial surface. The observation that bacteria ubiquitination is partly reduced when atg1 is disrupted (Fig. S4) also implies that the autophagic membrane somehow helps stabilize the ubiquitination. The recruitment of Atg8 even in the absence of SQSTM1, suggests the involvement of additional autophagy adaptors. In mammalian cells, multiple adaptors have been shown to mediate xenophagy including NDP52 and optineurin (28, 29). Although orthologs of these proteins have yet to be found in lower eukaryotes, given the universal challenge of containing intracellular pathogens (particularly if you are a professional phagocyte) it seems highly likely that additional adaptors will exist. Previously identified roles for autophagy during infection are aimed at capturing the bacteria within a vacuole, either to kill it, or allow it to replicate. In contrast, during ejection the autophagic membrane is required both to maintain host cell plasma membrane integrity and promote cell-to-cell transmission. Plasma membrane damage can be sealed by lysosomal fusion in a calciumdependent fashion (30). Recently, a new calcium-dependent wound healing mechanism has been discovered. The endosomal sorting complexes required for transport (ESCRT) machinery has been demonstrated to catalyze the shedding of small vesicles carrying membrane wounds (31). Even though no molecular mechanism has been proposed, autophagosome interactions with the plasma membrane have been suggested to allow the release of virus particles (32) and the cell-to-cell transmission of Brucella (2). Recently, using a fluorescent timer approach, it was shown that the autophagic pathway contributes to the shedding of microvesicles that harbor infectious coxsackievirusB and contribute to the spreading of the virus from cell to cell (33). Autophagy has also been shown to promote the cell-to-cell transmission of other pathogens, although the molecular mechanisms are unknown (2, 3). In contrast to our observations, in which the ejecting bacteria are cytosolic, Starr and colleagues showed that the Brucella replicating vacuole converts into a vacuole with autophagic features, and that this is independent of Atg5 and Atg7. Interestingly, they also show that this conversion is necessary for efficient bacterial transmission. Furthermore, they also report no increase in cell death upon Brucella release, suggesting a nonlytic form of egress, reminiscent of ejectosomemediated transmission. Here, we show that the autophagic machinery is necessary for nonlytic cell-to-cell transmission of M. marinum. We propose that the membrane generated by the autophagic machinery at the distal pole of ejecting bacteria helps seal the membrane wound generated by the ejection (Fig. S9). The loss of the polar autophagic membrane leads to increased permeability at the ejectosome and death of the host cell. The range of interactions between intracellular pathogens and the autophagy machinery continues to expand. The previously unidentified and unexpected role for autophagy highlighted in our study indicates the complexity of host-pathogen interactions that have evolved over millions of years, and how pathogen virulence and host defense pathways have reached a form of equilibrium that benefits both organisms.
fixed with 4% paraformaldehyde in Soerensen buffer and immunostained with an anti-RFP antibody to enhance the red signal. Subsequently, 250,000 cells were measured by flow cytometry. The results were analyzed and plotted with Flowjo 7.6.3 (TreeStar). To quantify transmission of M. marinum from donor to acceptor cells the ratio of infected acceptor versus infected donor cells was calculated. Permeability Test with Propidium Iodide (PI). Dictyostelium cells were infected with GFP-expressing bacteria as described above. At 24 hpi 2 × 106 cells were centrifuged (5 min, 500 × g) and resuspended in 500 μL Soerensen sorbitol buffer (120 mM sorbitol). PI (Sigma Aldrich, stock solution: 0.5 mg/mL) was added at a dilution of 1:10 and fluorescence (FL3) of 250,000 cells was measured (530 nm excitation, 610 nm emission) by flow cytometry (Accuri C6, BD Biosciences). As a positive control for dead and leaky cells, Dictyostelium cells were heat killed (60 °C, 10 min), stained, and measured as described above. To obtain the percentages of permeable cells which are right-shifted the shown, data were normalized between compared data sets and areas were computed using the density estimate of channel FL3 (PI). Area estimates for the overlap region between two curves (noninfected and infected) was computed and used to obtain the fraction of the total area shifted right of the overlap. All computations were preformed in R (R Core Team (2014). R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria).
added (ratio 10:1). The live cells were imaged at 25 °C using a Zeiss 510 Meta confocal microscope equipped with a 63× Europlan apochromat oil immersion objective (N.A. 1.4) and the Zeiss confocal microscope software release 3.2. Two different laser-lines were used (Argon-Laser 488 nm and HeNeLaser 543 nm). The imaging was performed in a multitrack-mode with a pinhole of 2 Airy units. Cells were imaged for 10 min with time intervals of 6 s between each scan. Brightness and contrast adjustments to images, as well as annotations, were performed using ImageJ. Coinfection Assay. ABD-GFP expressing Dictyostelium cells were simultaneously infected with both green fluorescent (GFP-expressing) M. marinum and red fluorescent (mCHERRY-expressing) M. smegmatis or M. marinum ΔRD1 as described under Infection assay. The bacteria were grown separately in shaking culture and added to the adherent Dictyostelium cells at the same time (defined as 0 hpi) with MOIs of 100 for M. marinum-GFP and 30 for mCHERRY-expressing M. smegmatis or M. marinum ΔRD1. Phagocytosis and washing was performed as described (infection assay) and cells prepared for immunofluorescence (as described) at 24 phi.
Live-Cell Imaging. Lifeact-RFP/Atg8-GFP–expressing Dictyostelium cells were infected with mCHERRY-expressing M. marinum and at 4 hpi transferred to a 35 mm μ-dish (Ibidi). At 6 hpi, nonfluorescent Dictyostelium cells were
ACKNOWLEDGMENTS. We gratefully acknowledge Francisco Rivero for providing Dictyostelium mutant racH− and Lalita Ramakrishnan for providing M. marinum and plasmids; and Ulrike Fröhlke and Carola Schneider for technical help. This work was supported by The Leibniz Society, Deutsche Forschungsgemeinschaft (DFG; HA3474/3-1, M.H.), Ciencia sem fronteiras (CsF, BEX2607-13-1, postdoctoral fellowship to R.P.), Cancer Research-UK and The Royal Society Research Grant RG130655 (to J.S.K.), and a grant from the Swiss National Science Foundation (to the T.S. laboratory). This work would not have been possible without dictyBase (37).
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