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Autophagy, a eukaryotic cellular activity leading to the degradation of cellular components, serves as a defense mechanism against facultative intracellular bacteria as well as a growth niche for the obligate intracellular bacterium Coxiella burnetii. We here demonstrate that the obligate intracellular bacterial pathogen Chlamydia trachomatis lymphogranuloma venereum strongly induced autophagy in the middle of the chlamydial developmental cycle (24 h after infection), a time point with maximal level of chlamydial replication, but not during the early stages with low overall chlamydial metabolism (before 8 h). No autophagy induction was evident in cells exposed to heat- and ultraviolet-inactivated elementary bodies (EBs, the infectious form of Chlamydia) nor to inocula from which EBs had been removed prior to inoculation. Blocking chlamydial development with chloramphenicol also prevented autophagy induction in cells infected with infectious EBs. It appears that autophagy is activated primarily in response to the metabolic stress consequent to chlamydial replication. However, autophagy-defective ATG5−/− cells supported chlamydial development as efficiently as autophagy-proficient ATG5+/+ cells.
Macroautophagy, frequently referred to as autophagy, is a highly conserved cellular process in eukaryotes, which is characterized by sequestration of cytoplasmic components in a double-membrane vacuole designated autophagosome that eventually fuses with the lysosome in which the sequestrated materials are degraded. Though the primary function of autophagy was once thought to degrade cell organelles and thus to recycle nutrients in response to starvation, recent studies have shown that autophagy is involved in a wide range of physiological processes and may determine the outcome of microbial infection [for review, see (Shintani & Klionsky, 2004, Webster, 2006)]. Thus, autophagy serves as a mechanism for innate immunity against bacterial pathogens. In most cases, invading pathogens are first engulfed into vacuoles commonly referred to as phagosomes, which are then converted into autophagosomes, and are finally destroyed by lysosomal enzymes after the autophagosomes fuse with lysosomes. Even organisms that manage to escape phagosomes can still be taken up by autophagosomes and subsequently be decomposed (Shintani & Klionsky, 2004, Webster, 2006). However, a number of pathogens have developed strategies to circumvent this defense mechanism. For example, Shigella flexneri is able to block the formation of autophagosome and thus to allow the pathogen to replicate in the cytoplasm. S. flexneri mutants defective in this mechanism fail to replicate intracellularly (Ogawa, et al., 2005). Although Mycobacterium tuberculosis enters phagosomes, it blocks the maturation of the bacteria-containing phagosomes into autophagosomes. Manipulations that augment autophagy remove this blockage, leading to intracellular killing of M. tuberculosis (Gutierrez, et al., 2004).
Compared with facultative intracellular bacteria, how obligate intracellular bacteria interact with autophagy in the host cells is not known except in the case of Coxiella burnetii. A proteobacterial pathogen of humans and animals, C. burnetii resides in acidified vacuoles that are characteristic of autophagolysosomes (Heinzen, et al., 1996, Gutierrez, et al., 2005). Stimulation of autophagy leads to increased C. burnetii growth whereas blockage of autophagy results in decreased growth (Gutierrez, et al., 2005). Thus, different from other bacteria studied, C. burnetii has adapted to the autophagic machinery as its growth niche.
Chlamydia trachomatis is a Gram negative bacterium that is responsible for a number of human diseases, including conjunctivitis, respiratory infection and sexually transmitted infection. Like other chlamydiae, C. trachomatis has a biphasic developmental cycle that begins with attachment of the metabolically-inert elementary body (EB) to a host cell that internalizes the bacterium into a vacuole designated as an inclusion. Inside the inclusion, the EB differentiates into the non-infectious, metabolically-active reticulate body (RB) in approximately 1 h. RBs replicate by binary fission during the first half of the developmental cycle and then progressively reorganize back to EBs. When the majority of RBs are converted into EBs (around 40 h), both chlamydial forms are released from the infected cells.
The chlamydial inclusion is not acidified and does not fuse with lysosomes or autophagosomes through out the chlamydial developmental cycle (Heinzen, et al., 1996, Wyrick, 2000). These findings indicate that Chlamydia either inhibits or resists autophagy. In this work, we analyzed the expression ratio of LC3-II/LC3-I, a specific biochemical marker for autophagy, and the localization of LC3 and the lysosomal marker LAMP-1, following infection with C. trachomatis lymphogranuloma venereum. We also compared chlamydial growth efficiencies in cells with various levels of autophagic activity. Our results suggest that this organism has developed a unique strategy to interact with the autophagic pathway of host cells.
HeLa cells were obtained from American Type Culture Collections (ATCC, Manassas, VA). The HeLa-HA-LC3 cell line was derived by transfecting the human cervical epithelial HeLa cells with an expression vector for human LC3 carrying dual amino-terminal hemagglutinin (HA) and Flag epitope tags and selection with G418. ATG5−/− mouse embryonic fibroblasts (MEFs) and the control ATG5+/+ MEFs were kindly provided by Dr. Noboru Mizushima (Tokyo Metropolitan Institute of Medical Science). All cell lines were maintained as adherent cultures using Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum (FBS).
Strain 434/bu of C. trachomatis serovar L2 (L2), a lymphogranuloma venereum pathogen, was purchased from ATCC and was amplified using HeLa cells (Balakrishnan, et al., 2006). HeLa-HA-LC3, ATG5+/+ or ATG5−/− cells were seeded on 6 or 24 well plates at the density that resulted in 20–30% confluence after overnight incubation. To infect the cells, the overnight culture medium was replaced with fresh medium containing EBs. In selected experiments, cells were pretreated with the autophagy activator Hank’s balanced salt solution (HBSS) or with full culture medium containing rapamycin (3S,6R,7E,9R,10R,12R,14S,15E,17E,19E,21S,23S,26R,27R,34aS)-9,10,12,13,14,21,22,23,24,25,26,27,32,33,34,34a-hexadecahydro -9,27-dihydroxy-3-[(1R)-2-[(1S,3R,4R)-4-hydroxy-3-methoxycyc lohexyl]-1-methylethyl]-10,21-dimethoxy-6,8,12,14,20,26-hexa methyl-23,27-epoxy-3H-pyrido[2,1-c][1,4]oxaazacyclohentriaco ntine-1,5,11,28,29(4H,6H,31H)-pentone, final concentration: 50 nM), another autophagy activator, or the control rapamycin vehicle (0.0005% Tween-20, 0.0045% ethanol and 0.095% dimethyl sulfoxide) for 2 h prior to infection (Gutierrez, et al., 2004). Unless indicated otherwise, the multiplicity of infection (MOI) was 5 or 10 inclusion-forming units (IFUs) per cell, which results in infection of >90% of cells (Balakrishnan, et al., 2006). The time that the EB stock was added onto cells was defined as t0. After incubation at 37 °C for 3 h (i.e. 3 h after infection), the remaining free EBs were removed by three washes, and cells were cultured with medium supplemented with FBS. In selected wells, chloramphenicol was added to culture medium (final concentration: 50 μM) at the end of 3 h attachment/entry period to block chlamydial protein synthesis and consequently chlamydial growth. Alternatively, some EB stocks were subjected to filtration through a syringe filter with a pore size of 200 nm (Millipore, Billerica, MA) or heat inactivation (56 °C, 10 min) or irradiation with ultraviolet light (UV) (1.4 J/cm2).
Chlamydial inclusions in live cultures were visualized with an IX-51 Olympus phase-contrast microscope and imaged with a monochrome DCC camera.
ATG5+/+ and ATG5−/− cells were inoculated with an EB stock and incubated on ice for 2 h. At the end of the attachment period, cells were washed 3 times with cold medium and were collected in sucrose-phosphate-glutamate buffer (SPG) (Balakrishnan, et al., 2006) immediately or, after they were cultured at 37 °C for 40 h. The cell harvests were frozen at −80 °C. To quantify input and progeny EBs, the harvests were thawed, sonicated, 1:10 serially diluted and inoculated on to HeLa monolayers on 96-well plates. The inclusions that formed on the 96-well plates were stained with a monoclonal antibody against the chlamydial lipopolysacharide (LPS) (a generous gift from Dr. Guangming Zhong) and FITC-conjugated goat anti-mouse IgG and scored under an IX51 fluorescence microscope (Balakrishnan, et al., 2006). The production of EBs from HeLa-HA-LC3 cells with different autophagic levels was determined in the same fashion excepting that the infection was carried out at 37 °C and that only the progeny EBs but not the input EBs were quantified.
Cells were harvested in sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer. The proteins in the cell extracts were resolved by electrophoresis in a 15% gel. The endogenous or HA-tagged LC3-I and LC3-II were visualized by sequential reactions with a polyclonal rabbit anti-LC3 or a monoclonal anti-HA (clone 16B2) (BAbCO, Richmond, CA) and an appropriate secondary antibody conjugated with horseradish peroxidase and an ECL plus kit (Amersham, Piscataway, NJ). The intensities of the LC3 bands were determined by densitometry.
The following primary antibodies were used for immunofluorescence assays: monoclonal mouse anti-chlamydial LPS described above, polyclonal rabbit anti-sera against LC3 (Sigma-Aldrich) and the inclusion membrane protein IncA (Rockey, et al., 1997), monoclonal mouse anti-Flag (Sigma-Aldrich), monoclonal mouse anti-human LAMP-1 and monoclonal rat anti-mouse LAMP-1 (Santa Cruz Biotechnology). Sample preparation and immunostaining procedures have been reported previously (Balakrishnan, et al., 2006). All images presented were acquired with an Olympus IX51 epifluorescence microscope except those in Fig 5B, which were acquired with a Zeiss LSM510 META confocal microscope.
Two-tailed analysis of variation was used to analyze densitometry and infectivity (inclusion-forming units) data. A significant difference was defined as p-value of <0.05.
Since C. trachomatis inclusions do not fuse with lysosomes, we hypothesized that successful C. trachomatis infection required the inhibition of autophagy. Therefore, we determined whether rapamycin and HBSS pretreatment would affect the growth of C. trachomatis. We measured the LC3-II/LC3-I ratio to confirm the induction of autophagy by rapamycin and HBSS. LC3-II is derived from LC3-I by post-translational modifications that require the activation of autophagy (Kabeya, et al., 2000, Mizushima, 2004). Compared with LC3-I, which is a cytosolic protein, LC3-II is associated with autophagosome as well as isolation membranes, which are precursors of autophagosomes (Kabeya, et al., 2000, Mizushima, 2004). To facilitate the detection of LC3 forms in the cervical carcinoma HeLa cells, which are broadly used to study the interaction of C. trachomatis with host cells, we derived HeLa-HA-LC3 cells. As expected, the HA-LC3-II/HA-LC3-I ratio increased in cells cultured with either rapamycin or HBSS but not in those cultured with the rapamycin vehicle control (Figure 1A and B). The abilities of cells to support C. trachomatis infection following different pretreatments were judged by inclusion formation (Figure 1C) and by quantifying the infectious EBs produced (Figure 1D). Evidently, induction of autophagy prior to infection had no effects on C. trachomatis infection.
The failure of rapamycin and HBSS pretreatment to inhibit chlamydial growth suggests that C. trachomatis might suppress autophagy. Therefore, we determined the LC3-II/LC3-I ratio following chlamydial infection. In uninfected cells, the HA-LC3-II/HA-LC3-I ratio increased to some degree as they were kept in culture (Figure 2A, B). The rise in autophagic activity is likely due to the increase in cell confluence as previously reported (Fuertes, et al., 2003, Wu, et al., 2006) as well as the infection procedures (e.g. medium and/or temperature changes). At both 3 and 8 h after infection, the HA-LC3-II/HA-LC3-I ratios were similar between uninfected and chlamydia-infected cells. Surprisingly, at 24 h after infection, there was a marked increase in the autophagic index in the infected cells compared with the uninfected cells (Figure 2A, B). Separate experiments showed that there was no significant further increase in the index at 30 and 40 h after infection (data not shown). Representative images of the uninfected cells and infected cells containing chlamydial inclusions, which were obtained immediately before the cells were harvested at 24 h post-infection, are shown in Figure 2C. As expected, inclusions were too small to be detected by light microscopy at 3 and 8 h after infection (data not shown). These results suggest that autophagy is induced around the midpoint of the chlamydial developmental cycle.
To determine whether induction of autophagy requires chlamydial replication, we subjected the EB stock to heat and UV irradiation before inoculation. We also inoculated cells with EB stocks from which EBs had been filtered out. Finally, for selected wells infected with live EBs, we incorporated chloramphenicol, which inhibits bacterial but not eukaryotic protein synthesis, in the culture medium to block chlamydial growth. As expected, no chlamydial inclusions were detected in cultures subjected to these treatments (Figure 2C). The LC3-II/LC3-I ratios in these cultures were comparable to that in the uninfected cells at the respective time points (Figure 2A, B).
Chlamydiae have a limited biosynthetic capacity, and therefore rely on host cells for the supply of many types of nutrients, including amino acids, nucleotides and lipids from host cells. At 24 h after infection, there are significant decreases in the sizes of nutrient pools in host cells, as a result of active bacterial growth. However, during the early stages (e.g., within the first 8 h) of the developmental cycle, the overall demand on host cells for nutrients is relatively low because only small numbers of RBs exist in cells (McClarty, 1999). Therefore, data presented in Figure 2 may suggest that autophagy is induced in response to metabolic stress as a result of chlamydial growth. Alternatively, it is also plausible that autophagy is induced when there are significantly more intracellular bacteria by 24 h after infection compared to earlier time points. However, in cells inoculated with UV- or heat-inactivated EBs at doses equivalent to 100 inclusion-forming units (IFUs) per cell, the LC3-II/LC3-I ratios were still comparable to those in the control uninfected cells (Figure 3A and 3B). Taken together, the data presented in Figs. 2 and and33 indicate that autophagy may be induced in response to the decline in the nutrient pools of host cells, similar to the response to nutrient starvation.
Whereas LC3-I has a diffused distribution pattern, LC3-II is located in punctate structures including autophagosomes and autophagolysosomes (Mizushima, 2004). Therefore, we performed LC3 immunostaining to corroborate the LC3-immunoblotting data. Also, since the conversion of LC3-I to LC3-II is a relatively early step in the autophagic pathway, we determined if LC3 colocalizes with LAMP1, a lysosomal marker, to analyze autophagy flux in chlamydia-infected cells. For these analyses, we used rabbit anti-LC3 and mouse anti-LAMP1 as primary antibodies, and Rhodamine-conjugated anti-rabbit IgG and FITC-conjugated anti-mouse IgG as secondary antibodies. As expected, cells treated with rapamycin displayed stronger punctate LC3 distribution than untreated cells; they also showed increased colocalization of LC3 with LAMP1 (Figure 4). The increases in punctate LC3 distribution and LC3/LAMP1 colocalization are even stronger in chlamydia-infected cells than rapamycin-treated cells. These results are consistent with data from LC3 Western blotting analyses presented in Figure 1 and Figure 2, and suggest that LC3-II-containing autophagosomes fuse with lysosomes in chlamydia-infected cells.
A number of facultative intracellular bacteria become associated with LC3 as they are targeted by autophagy after entering cells (Gutierrez, et al., 2004, Nakagawa, et al., 2004, Shintani & Klionsky, 2004, Ogawa, et al., 2005, Birmingham, et al., 2006, Webster, 2006). We also determined whether LC3 colocalizes with chlamydiae or the chlamydial inclusion membrane. For this purpose, LC3 was stained with either polyclonal rabbit anti-LC3 or monoclonal mouse anti-Flag, whereas chlamydiae and the chlamydial inclusion membrane were stained with monoclonal mouse anti-chlamydial LPS and polyclonal rabbit anti-IncA, respectively. Immunofluorescence microscopy did not detect any colocalization of LC3 with chlamydiae (Figure 5A). No colocalization was observed between LC3 and the vast majority of the inclusion membranes (Figure 5B) although single loci (mostly only one locus) of fluorescence copatching were detected occasionally (~5% of infected cells). These results indicate that the LC3-containing autophagosomes rarely fuse with the chlamydial inclusion.
To determine the role of autophagy in chlamydial infection, we compared the efficiencies of chlamydial infection in two isogenic fibroblastic cell lines derived from wild-type (Atg5+/+) and ATG5 knockout (Atg5−/−) mice, respectively. Since ATG5 is essential for autophagy, Atg5−/− cells are deficient in autophagy while Atg5+/+ cells are proficient (Kuma, et al., 2004). Chlamydial inclusions in Atg5+/+ and Atg5−/− cells were indistinguishable in both the number and size at any of the time points (16, 24 and 40 h after infection) that the experiments were conducted (Figure 6A). Quantitative analyses of EBs presented in Figure 6B clearly indicate that Atg5+/+ and Atg5−/− cells supported chlamydial infection to generate infectious EBs equally efficiently.
Analyses were performed to confirm the statuses of autophagy in Atg5+/+ and Atg5−/− cells following chlamydial infection. L2 infection resulted in an elevated level of endogenous LC3-II protein in Atg5+/+ cells (Figure 7A). Densitometry analysis showed that the LC3-II/LC3-I ratio increased from 5.7 in the uninfected Atg5+/+ cells to 11.4 in L2-infected Atg5+/+ cells (Figure 7B). These results are consistent with those obtained with the HeLa-HA-LC3 cells (Figure 2) although the fibroblasts appear to have a higher basal level of autophagy compared to HeLa cells. Immunofluorescence microscopy also revealed increases in punctate LC3 distribution and colocalization of LC3 with LAMP1 in chlamydia-infected Atg5+/+ cells, as compared to uninfected cells (Figure 7C). On the contrary, in the autophagy-deficient Atg5−/− cells, the LC3-II bands (Figure 7A, B) and punctate LC3 distribution (Figure 7C) were completely undetectable under both uninfected and chlamydia-infected conditions. Thus, autophagy induction in response to chlamydial infection depends on the function of ATG5. Overall, data presented in Figure 6 and Figure 7 suggest that autophagy is not required for chlamydial infection.
In this work we have investigated how C. trachomatis lymphogranuloma venereum interacts with autophagy. Surprisingly, we discovered that autophagy is strongly induced in L2-infected cells. Experimentation under a variety of conditions suggests that the induction of autophagy depends on active chlamydial growth. Thus, removal of EBs from the inocula, exposure of the inocula to heat and UV to inactivate the infectivity of EBs prior to infection, and culturing cells infected with live EBs with chloramphenicol all prevented the induction of autophagy.
As obligate intracellular parasites with a limited biosynthetic capacity, chlamydiae obtain a variety of nutrients, including amino acids, nucleotides and lipids from host cells (McClarty, 1999). At 24 h after infection, the sizes of nutrient pools in host cells are significantly reduced. However, during the early stages of the developmental cycle, the overall demand on host cells for nutrients is relatively low because only small numbers of RBs exist in the cells (McClarty, 1999). Interestingly, C. trachomatis infection-induced autophagy is prominent at 24 h after infection but not at 3 and 8 h. This suggests that autophagy may be induced in response to the declines in the nutrient pools of host cells, similar to the response to nutrient starvation caused by culturing cells in medium poor in nutrients. Nevertheless, autophagy-deficient ATG5−/− cells support chlamydial infection as efficiently as ATG5+/+ cells. A possible explanation for these “negative” data relies on the recently reported upregulation of chaperone-mediated lysosome-dependent protein degradation pathway (Kaushik, et al., 2008) and additional yet-to-be defined compensatory mechanisms in ATG5−/− cells.
It has been reported that certain amino acids and 3-methyladenine, which inhibit autophagy, reduce the growth of C. trachomatis L2, the same strain used in this study (Al-Younes, et al., 2004). However, a more recent study has revealed that those amino acids inhibit chlamydial development by competitive inhibition of the transport of other amino acids into chlamydiae (Braun, et al., 2007). The adverse effect of 3-methyladenine on chlamydial infection may not be due to the inhibition of autophagy since a number of studies have suggested that this compound has other cellular targets (Mizushima, 2004), including MAP kinase-mediated signaling pathways, which are critical for chlamydial infection (Su, et al., 2004).
Autophagy acts as primary defense in a number of cell types, including epithelial cells, which constitute the first line of natural guard against numerous microbial pathogens, macrophages, which are crucial in both innate and acquired immunities, and fibroblasts (Gutierrez, et al., 2004, Nakagawa, et al., 2004, Shintani & Klionsky, 2004, Ogawa, et al., 2005, Birmingham, et al., 2006, Webster, 2006). For example, a majority of group A Streptococcus is encased in autophagosomes within 1 h after entering HeLa cells (Nakagawa, et al., 2004). Efficient survival and proliferation of this human pathogen occur only in Atg5−/− cells but not in Atg5+/+ cells (Nakagawa, et al., 2004). Autophagy also plays a pivotal role in defense against mycobacteria and legionellae in macrophages (Gutierrez, et al., 2004, Amer & Swanson, 2005). Mucosal epithelial cells in the urogenital tract are the initial targets for C. trachomatis. Macrophages are recruited to the site of infection in response to cytokines and chemokines released from the infected epithelial cells. C. trachomatis can survive and replicate in macrophages, although inefficiently compared with epithelial cells. Findings from this study suggest that in epithelial cells and fibroblasts autophagy might not be an innate immune mechanism against C. trachomatis infection. First, immunostaining analysis indicates that chlamydiae or the chlamydial inclusion is not targeted by autophagosomes marked by LC3-II. This is consistent with well-established observations that chlamydial inclusions are not acidified and do not fuse with lysosomes (Heinzen, et al., 1996, Wyrick, 2000). Second, chlamydial infection-induced autophagy is a delayed response as demonstrated by our time point analyses. Third, induction of autophagy by HBSS or rapamycin prior to inoculation fails to inhibit chlamydial infection. Finally, inactivation of autophagy due to the knockout of ATG5 does not increase the efficiency of chlamydial infection. However, if and how autophagy regulates chlamydial infection in macrophages and other non-epithelial and non-fibroblast cell types remains to be examined.
In summary, we have demonstrated that autophagy is strongly upregulated upon active growth of C. trachomatis lymphogranuloma venereum. However, neither the augmentation of nor the deficiency in autophagy has a detectable effect on chlamydial infection. Thus, it appears that this parasite has developed a unique way to interact with autophagy, which differs from both facultative intracellular bacterial pathogens that require the inhibition of autophagy for successful infection and the obligate intracellular C. burnetii, an extreme case that requires autophagy for growth. This seemly undemanding relation between C. trachomatis and autophagy may have contributed to the evolution of C. trachomatis into arguably one of the most successful parasites of mankind.
We thank Dr. Guangming Zhong for the anti-LPS and anti-MOMP, Dr. Ted Hackstadt for the anti-IncA, Dr. Noboru Mizushima for the ATG5+/+ and ATG5−/− cells, and Dr. Estela Jacinto for rapamycin. This work was supported in part by grants from the National Institutes of Health (AI064441 and AI071954).