Chlamydia psittaci Induces Autophagy in Human Bronchial Epithelial Cells via PERK and IRE1α, but Not ATF6 Pathway

ABSTRACT

Chlamydia psittaci is an important pathogen that causes chronic and atypical pneumonia in humans. Autophagy and the unfolded protein response (UPR) are important mechanisms for regulating the growth of infectious parasitic pathogens in living cells. Here, we explored whether C. psittaci infection induced autophagy via the UPR and the effect of these cellular responses on the survival and replication of C. psittaci in human bronchial epithelial cells (HBEs). Not only were the numbers of autophagosomes and the expression of LC3-II and Beclin1 increased following C. psittaci infection of HBEs, but also the expression of p62 (also called sequestosome-1) was downregulated. Moreover, after C. psittaci infection, the UPR and UPR sensors PERK/eIF2α and IRE1α/XBP1 were activated, but not the ATF6 pathway. When either Bip siRNA was used to block normal initiation of the UPR, or activation of the PERK and IER1α pathways was blocked with specific inhibitors GSK2606414 and 4μ8C, the level of autophagy caused by C. psittaci infection was significantly inhibited. Furthermore, blocking activation of the UPR and associated pathways significantly reduced the number of C. psittaci inclusions. Our research suggests that the UPR, via the PERK and IRE1α, but not ATF6 signaling pathways, regulates HBE-cell autophagy induced by C. psittaci infection and the replication of C. psittaci.

INTRODUCTION

Chlamydia psittaci is an obligate intracellular Gram-negative bacterium. It has a unique developmental cycle involving two different developmental forms, namely, the infectious elementary body (EB) and the metabolic replicative body (RB) (1). Although C. psittaci is the only pathogen responsible for psittacosis in birds, it can also infect a variety of animals, including humans, sheep, cattle, pigs, horses, and cats, causing zoonotic diseases (2–4). Parent birds can transmit C. psittaci to their offspring through regurgitation-feeding to cause transmission between birds, and cause a series of mild or life-threatening diseases, such as pneumonia, conjunctivitis, and enteritis (5–7). Humans can be infected with C. psittaci through indirect contact with respiratory secretions of infected birds, aerosols of dry feces, or feather dust, which can cause atypical pneumonia, accompanied by symptoms such as high fever, chills, myalgia, dyspnea, and general weakness (8–10). Therefore, research on the pathogenesis of C. psittaci is of great significance for the protection of human health and the healthy development of animal husbandry.

Autophagy is the process by which abnormal proteins and damaged organelles are engulfed and degraded to satisfy metabolic needs, renew organelles, and maintain homeostasis in cells (11). It plays an important role in the survival of cells infected with pathogens, including chlamydia. For example, Helicobacter pylori uses cytotoxin-related gene A (CAGA) to upregulate the expression of microtubule-associated protein 1 light chain 3 II (LC3-II), ultimately activating autophagy in gastric epithelial cells and promoting the immune protection of host cells (12). In chlamydia infection, autophagy plays a dual role. One theory is that autophagy is a cellular self-protective mechanism by which Chlamydia infected cells trigger different intracellular mechanisms that interact with autophagy to promote host cells to clear Chlamydia. Yasir et al. (13) proved that, compared with the uninfected group, the ratio of LC3-I to LC3-II is significantly increased in C. trachomatis-infected host cells, and the number of chlamydial inclusions is significantly decreased, indicating that host cells can restrict the growth of C. trachomatis in vivo by autophagy. In mouse embryonic fibroblasts and macrophages infected with C. pneumoniae, the lack of certain components of the autophagy pathway enhances C. pneumoniae replication (14). While some other researchers believe that Chlamydia is a strictly intracellular pathogen that relies on metabolites in host cells for nutrition. As one of the main sources of endogenous metabolites, autophagy can promote the growth and reproduction of Chlamydia in host cells. For example, Al-younes et al. (15) found that autophagy is activated in Hep-2 cells infected with C. trachomatis, and both the maturation of C. trachomatis inclusions and the production of infectious pathogens are inhibited by siRNA interference of LC3.

In eukaryotes, endoplasmic reticulum (ER) is the main organelle controling protein synthesis, folding and processing and it plays an important role in maintaining the health of the cell and organism (16). A large number of misfolded or unfolded proteins in the ER can be induced by pathogen infection, drug injury and so on, and this is called ER stress (ERS). Cells experiencing ERS will activate a cellular response called the unfolded protein response (UPR) to ensure the correct folding of proteins and prevent the accumulation of unfolded or misfolded polypeptides, so as to counteract the adverse effects of ERS (17). The UPR consists of three signaling pathways initiated by three transmembrane ER stress sensors: inositol-requiring enzyme 1α (IRE1α), protein kinase RNA-activated (PKR)-like ER kinase (PERK) and activating transcription factor 6 (ATF6). Under normal physiological conditions, these stress sensors are bound to the ER molecular chaperone, glucose-regulated protein (GRP78), which is also called immunoglobulin heavy-chain binding protein (Bip), rendering their luminal domains inactive and Bip is a key sensor of ERS (18). During ERS, however, Bip will dissociate from IRE1α, PERK and ATF6, thereby activating their respective signaling pathways, and then be released into the ER lumen to identify unfolded or misfolded proteins and perform its molecular chaperone function (19, 20). After PERK is activated, it phosphorylates and activates eukaryotic initiation factor 2-α (eIF2α), which inhibits ribosome assembly and the initiation of protein synthesis, thereby halting mRNA translation and downregulating protein synthesis (20). When IRE1α is activated, it acquires endonuclease activity and excises an intron from the XBP1 mRNA, which then encodes a transcription factor, XBP1s, specific to the UPR, which upregulates genes for various ER chaperones and other proteins that relieve ERS (21). Activated ATF6 will migrate from the ER to the Golgi, promoting the expression of ERS-related molecules such as C/EBP-homologous protein (CHOP). This then migrates to the nucleus, where it upregulates transcription of the Bip gene and other genes that help restore protein homeostasis in the ER (22).

Although autophagy and ERS are two independent biological processes, the UPR is not enough on its own to prevent the accumulation of unfolded or misfolded proteins while ERS continues, and it cannot remove damaged organelles. The accumulation of a large number of misfolded proteins during ERS may activate autophagy, and autophagy can help restore ER homeostasis by degrading defective proteins, which in turn limits ERS, and thus reduces cell death and promotes cell survival (23, 24). For example, Chang et al. (25) found that intermittent hypoxia can increase myocardial autophagy as an adaptive response and thereby prevent ERS and apoptosis. Dash and his colleagues (26) reported that hepatitis C virus can induce autophagy as a survival-promoting mechanism for alleviating ERS.

The UPR, a cellular response mechanism that specifically counteracts the adverse effects of ERS, has been reported to be involved in the regulation of the inflammatory response and energy uptake in chlamydia-infected cells. For example, Shima et al. (27) reported that in the early stages of a persistent C. pneumoniae infection induced by interferon-γ, BiP/GRP78 expression can attenuate host cell apoptosis via the regulation of eIF2α phosphorylation, suggesting that the UPR regulates the survival of C. pneumoniae-infected cells. After C. trachomatis infection, the UPR is activated in dendritic cells, and is followed by an increase in the production of inflammatory cytokine IL-23, suggesting that the UPR regulates the inflammatory response in C. trachomatis-infected cells (28). George et al. (29) found that UPR signaling pathways are activated in oviduct epithelial cells of mice infected with C. muridarum, resulting in upregulation of hexokinase II and glucose transporter I, thus promoting host-cell glucose metabolism and ATP synthesis, allowing chlamydia cells to obtain energy from the host.

Whether autophagy is involved in the process of C. psittaci infection has not previously been reported, and the specific role of the UPR and it’s signaling pathways in the process of C. psittaci infection is still unclear. The purpose of this study was to investigate the activation of autophagy and the UPR during C. psittaci infection and the involvement of the UPR in the regulation of autophagy.

RESULTS

C. psittaci induces autophagy in HBE cells.

HBE cells were infected with C. psittaci at different MOIs (0.5, 1, 1.5, 2) for 24 h, and the levels of LC3-II, Beclin1 and p62 were assessed by Western blotting (WB). Compared with the uninfected group, the expression of both LC3-II and Beclin1 were significantly increased, while the level of p62 decreased. This upregulation of LC3-II and Beclin1 was most obvious when the MOI was 1.5 (Fig. 1A) and demonstrates that autophagy systems are activated on infection with chlamydia.

FIG 1.

C. psittaci infection induces autophagy. (A) HBE cells were infected with C. psittaci at MOIs of 0, 0.5, 1, 1.5, 2 for 24 h and the protein expression levels of LC3-II, Beclin1, P62, and GAPDH were assessed by WB. Experiments were carried out three times independently. HBE cells were infected with C. psittaci at an MOI of 1.5 for 24 h and the green fluorescence cluster points representing LC3-II were imaged by IFA (B), while autophagosomes and autophagolysosomes were scanned by TEM (C). NC: negative control, Cps: C. psittaci infected group. Magnification, 8000×; Scale: 1 μm. *, P < 0.05, **, P < 0.01.

To confirm this, we next used indirect immunofluorescence assay (IFA) and transmission electron microscope (TEM) to examine HBE cells infected with C. psittaci (at an MOI of 1.5). IFA experiments (Fig. 1B) showed green fluorescence cluster points (LC3-II spots) around the nucleus after C. psittaci infection (P < 0.01), while TEM revealed autophagosomes and autophagolysosomes in the perinuclear region of infected cells (Fig. 1C).

C. psittaci infection generates ERS and activates the UPR in HBE cells.

To verify the relationship between C. psittaci infection, ERS and the UPR, HBE cells were infected with C. psittaci at various MOIs (0.5, 1, 1.5, 2) for 24 h and the expression level of Bip was determined by WB. As shown in Fig. 2, Bip expression was upregulated in C. psittaci-infected groups and reached the highest level at an MOI of 1.5, which indicates that C. psittaci infection can trigger ERS and activated the UPR in HBE cells.

FIG 2.

C. psittaci infection triggers ERS and activates the UPR in HBE cells. HBE cells were infected with C. psittaci at MOIs of 0, 0.5, 1, 1.5, 2 for 24 h and the protein expression levels of Bip and GAPDH were assessed by WB. Experiments were repeated three times. *, P < 0.05, **, P < 0.01.

C. psittaci infection activates PERK and IRE1α, but inhibits the ATF6 signaling pathway.

To verify the relationship between C. psittaci infection and the UPR signaling pathways, HBE cells were infected with C. psittaci at different MOIs (0.5, 1, 1.5, 2) for 24 h and the expression levels of PERK, p-PERK, eIF2α, IRE1α, p-IRE1α and ATF6 were assessed by WB, while qRT-PCR was used to measure the XBP1, ATF6 and CHOP mRNA level. We found that PERK, p-PERK, eIF2α, IRE1α and p-IRE1α were all upregulated after C. psittaci infection (Fig. 3A and B), as was XBP1 mRNA (Fig. 3C), reaching the highest levels at an MOI of 1.5. However, either the protein or mRNA level of ATF6 was inversely correlated with MOIs (Fig. 3C). Then the protein and mRNA levels of ATF6 and CHOP (a downstream molecule of the ATF signaling pathway) were detected in HBE cells at 6, 12 and 24h after C. psittaci infection, and showed the same results (Fig. 3D and E).

FIG 3.

C. psittaci activates the PERK and IRE1α signaling pathways, but inhibits the ATF6 pathway, in HBE cells. HBE cells were infected with C. psittaci at MOIs of 0, 0.5, 1, 1.5, 2 for 24 h and the protein expression levels of PERK, p-PERK, eIF2α (A) and IRE1α, p-IRE1α were assessed by WB (B) and expression of XBP1 mRNA determined by qRT-PCR (C). Then the protein level of ATF6 and the mRNA levels of ATF6 and CHOP were detected in HBE cells at 24 h (D), 12 h (E) and 6 h (F) after C. psittaci infection. Experiments were carried out three times. *, P < 0.05, **, P < 0.01.

We also detected the protein and mRNA levels of ATF6 and CHOP in different cells types or infected with different Chlamydia species. Compared with uninfected group, both protein and mRNA levels of ATF6 and CHOP mRNA were decreased after C. psittaci infection in HBE cells, while increased in HeLa cells (Fig. 4). In HBE cells, the expression levels of ATF6 and CHOP were only decreased in C. psittaci infected group, while increased in either C. trachomatis or C. muridarum infected one (Fig. 5). These results indicate that the PERK and IRE1α signaling pathways were activated by C. psittaci infection, while the ATF6 pathway was inhibited in HBE cells.

FIG 4.

The ATF6 pathway was inhibited in HBE cells but activated in HeLa cells after C. psittaci infection. HBE cells or HeLa cells were infected with C. psittaci at MOIs of 1.5 for 24 h, 12 h and 6 h and the expression level of ATF6 (A, B, C) and CHOP (D) were determined by WB or qRT-PCR. Experiments were carried out three times. Notes: NC: negative cotronal, Cps: C. psittaci infection group. *, P < 0.05, **, P < 0.01.

FIG 5.

The ATF6 pathway in HBE cells was inhibited by C. psittaci but activated by C. muridarum or C. trachomatis. HBE cells were infected with C. psittaci, C. muridarum or C. trachomatis at MOIs of 0 or 1.5 for 24 h, 12 hand 6 h and the protein expression levels of ATF6 were assessed by WB and the expression of ATF6 (A, B, C) and CHOP (D) mRNA determined by qRT-PCR. Experiments were carried out three times. NC: negative cotronal, Cps: C. psittaci infected group. Cm: C. muridarum infected group, Ct: C. trachomatis infected group. *, P < 0.05, **, P < 0.01.

C. psittaci induces autophagy via the UPR in HBE cells.

To verify whether the autophagy observed in HBE cells following C. psittaci infection is regulated by the UPR, we next assessed the expression of LC3-II, Beclin1 and p62 after Bip was downregulated by RNA interference. The results showed that the expression levels of LC3-II and Beclin1 were significantly dowregulated, while p62 was upregulated, and the formation of LC3-II spots around the nucleus was inhibited after Bip siRNA treatment (Fig. 6, P < 0.01). These results indicate that the autophagy induced by C. psittaci infection in HBE cells is regulated by the UPR.

FIG 6.

C. psittaci infection induces autophagy via the UPR. 50 nM Bip siRNA was used to knockdown Bip mRNA for 48 h in HBE cells, which were then infected with C. psittaci at an MOI of 1.5. The expression levels of LC3-II, Beclin1 and p62 were determined by WB (A), and the formation of LC3-II spots was examined by IFA (B). Notes: NC: negative cotronal, Cps: C. psittaci infected group, NC siRNA+Cps: NC siRNA+C. psittaci infected group, Bip siRNA+Cps: Bip siRNA+C. psittaci infected group. Magnification, 400 ×; blue: nucleus; green: LC3-II. *, P < 0.05, **, P < 0.01.

C. psittaci induces autophagy via the PERK and IRE1α signaling pathways.

HBE cells were treated with PERK inhibitor GSK2606414 or IRE1α inhibitor 4μ8C for 2 h before C. psittaci infection at an MOI of 1.5 for 24 h, after which the levels of autophagy proteins were assessed by WB. The results showed that the expression levels of LC3-II and Beclin1 were significantly inhibited, while p62 levels increased in cells treated with either GSK2606414 (Fig. 7A) or 4μ8C (Fig. 8A), compared with untreated infected cells (P < 0.01). IFA analysis showed that the formation of LC3-II spots around the nucleus was significantly inhibited by both GSK2606414 (Fig. 7B) and 4μ8C (Fig. 8B) (P < 0.01). These results indicate that inhibiting either the PERK or the IRE1α signaling pathway can significantly inhibit autophagy induced by C. psittaci infection in HBE cells.

FIG 7.

C. psittaci infection induces autophagy via the PERK signaling pathway. After infection with C. psittaci, HBE cells were treated with 40 μM GSK2606414, and the expression levels of LC3-II, Beclin1 and p62 were determined by WB (A). The formation of LC3-II spots was assessed by IFA (B). Notes: NC: negative cotronal, Cps: C. psittaci infected group, GSK2606414+Cps: C. psittaci infected group was treated with GSK2606414. Magnification, 400 ×; blue: nucleus; green: LC3-II. *, P < 0.05, **, P < 0.01.

FIG 9.

The effects of Bip siRNA, and treatment with GSK2606414 or 4μ8C, on the replication of C. psittaci in HBE cells. C. psittaci IFUs were measured by IFA in HBE cells treated with Bip siRNA (A), GSK2606414 (B) or 4μ8C (C). Notes: Cps: C. psittaci infected group, NC siRNA+Cps: C. psittaci infected group was not treated with Bip siRNA, NC siRNA+Cps: C. psittaci infected group was treated with Bip siRNA, GSK2606414+Cps: C. psittaci infected group was treated with GSK2606414, 4μ8C+Cps: C. psittaci infected group was treated with 4μ8C. Magnification, 200 ×; blue: nucleus; green: C. psittaci inclusions). **, P < 0.01.

FIG 8.

C. psittaci infection induces autophagy via the IRE1α signaling pathway. After infection with C. psittaci, HBE cells were treated with 30 μM 4μ8C, and the expression levels of LC3-II, Beclin1 and p62 were determined by WB (A). The formation of LC3-II spots was assessed by IFA (B). Notes: NC: negative cotronal, Cps: C. psittaci infected group, 4μ8C+Cps: C. psittaci infected group was treated with 4μ8C. Magnification, 400 ×; blue: nucleus; green: LC3-II. *, P < 0.05, **, P < 0.01.

The UPR and its signaling pathways regulate the replication of C. psittaci in HBE cells.

We next verified whether there is any association between the UPR or its signaling pathways and the replication of C. psittaci in HBE cells. As shown in Fig. 9A, the number of C. psittaci inclusions was 1.13 × 10⁷ IFU/ml in cells subjected to Bip siRNA, which was significantly lower than that of the untreated C. psittaci infection group (2.33 × 10⁷ IFU/mL) and NC siRNA-infected cells (2.41 × 10⁷ IFU/mL). Similar results were obtained in GSK26026414- (1.46 × 10⁷ IFU/mL versus 2.50 × 10⁷ IFU/mL in the control, Fig. 9B) or 4μ8C-treated (1.59 × 10⁷ IFU/mL versus 2.50 × 10⁷ IFU/mL in the control, Fig. 9C) groups. These data indicate that the UPR and its PERK and IRE1α signaling pathways play a key role in promoting intracellular replication of C. psittaci in HBE cells.

DISCUSSION

C. psittaci is an obligate intracellular parasitic pathogen; in humans, an opportunistic host, it can cause atypical pneumonia and other diseases (1). However, the mechanism of pathogenicity remains unclear.

The ability of cells to adapt to stress, such as pathogen infection, requires diverse changes in cellular metabolism that include autophagy, a critical catabolic process, which maintains cell stability and survival by removing and recycling unwanted cellular material (30). The binding of microtubule-associated protein 1A/1b-light chain 3-I (LC3-I) to phosphatidylethanolamine promotes the formation of LC3-phosphatidylethanolamine (LC3-II), which contributes to the development of the autophagosome bilayer and the recruitment of proteins during autophagy; thus, LC3-II is considered an autophagy marker protein (31). Beclin1, a mammalian homologue of yeast ATG6, binds with PI3KC3/VPS34 to regulate the formation of the autophagosome, and is a core component of autophagosome membrane nucleation (32). p62 is an adapter protein that binds ubiquitinated proteins and mediates their destruction via the proteasome. It is also an autophagy receptor protein, participating in selective autophagy where it recognizes an N-terminal protein degradation domain and combines with LC3 to form a complex, which is often used as an indicator of autophagic flux. The degradation of p62 is usually consistent with an increase in LC3-II when autophagic flux is not hindered (33, 34). In this study, the expression of LC3-II and Beclin1 increased, while p62 levels decreased in C. psittaci-infected HBE cells. This coincided with the appearance of LC3A/B puncta and an increased number of autophagosomes and autophagolysosomes around the nucleus. These phenomena suggest that autophagy is induced in HBE cells during C. psittaci infection.

The ER is an important site for protein synthesis and is involved in the correct folding and modification of proteins (35). As bacterial pathogens infect their host cells, the increased demand for protein may lead to an excessive number of proteins or misfolded proteins in the ER lumen, resulting in ERS and activation of the UPR response (36). Previously, the relationship between ERS and chlamydia was mainly known from research on C. trachomatis and C. pneumoniae. Studies have shown that C. pneumoniae can activate the UPR response by upregulating the expression of Bip, ATF4 and CHOP mRNA, and induce adipocytes to secrete fatty acid binding protein 4. This promotes the release of glycerol and fatty acids and the production of ATP, which in turn is exploited by the pathogen for its survival (37). C. pneumoniae has been shown to induce the UPR by activating IRE1α RNase via the TLR4 pathway (38). The upregulation of Bip is not only an indicator of ERS in mammalian cells, but also of UPR activation (18). Our results confirm that C. psittaci infection can stimulate Bip expression in HBE cells, and thus induce ERS and activate the UPR response.

The UPR has three signaling pathways, namely, PERK, IRE1α, and ATF6. Our results show that C. psittaci can affect both PERK-eIF2α and IRE1α-XBP1 but inhibit the ATF6 signaling pathway in HBE cells. We detected the levels of ATF6 and one of its downstream molecule CHOP at different times after C. psittaci infection, and found the level of both of them were decreased. We then tried this experiment using HeLa cell or two different Chlamydia species, and it is interesting to note that ATF6 and CHOP were decreased only in HBE cells infected with C. psittaci. This phenomenon is different from C. pneumoniae and C. trachomatis which can simultaneously activate all three signaling pathways (28, 39).

Similar results have been reported for other pathogen infections. For example, Chen et al. found that infection with porcine reproductive and respiratory syndrome virus activates PERK and IRE1α, but inhibits the ATF6 signaling pathway (40). The PERK-eIF2α and IRE1α-XBP1 signaling pathways are activated in human cytomegalovirus-infected human foreskin fibroblasts, but the ATF6 pathway is blocked after glycosylation of ATF6 (41). Increasing evidence has shown that the three UPR signaling pathways are coregulated, but they can produce different transcriptional outputs, resulting in diverse signal transduction effects (42). For example, Lin et al. (43) confirmed that a lethal dose of thapsigargin or tunicamycin, both ERS inducers, causes long-term PERK activation, but only transient activation of IRE1α and ATF6. Therefore, it is possible that C. psittaci infection causes only a transient activation of ATF6 in HBE cells, which was undetected in our experiments, or alternatively blocks the ATF6 pathway by glycosylation of ATF6.

Autophagy is known to be associated with the UPR. For example, coxsackievirus B3 induces autophagy in host cells by activating the PERK, IRE1α and ATF6 signaling pathways (44). To clarify the regulatory role of the C. psittaci-induced UPR response in host cell autophagy, we examined the effect on autophagy of knockdown of the key UPR protein, Bip, or treatment with PERK inhibitor GSK2606414 or IRE1α inhibitor 4μ8C. In all three experiments, expression levels of LC3-II and Beclin1 were significantly downregulated, while p62 was upregulated, and the formation of LC3-II spots around the nucleus was inhibited. These results indicate that autophagy in HBE cells induced by C. psittaci infection is regulated by the UPR, in particular the PERK and IRE1α signaling pathways. Similar results have been reported for other pathogens. Seneca Valley virus can increase the phosphorylation of PERK by stimulating Bip expression in BHK-21 cells, thereby promoting the conversion of LC3-I to LC3-II and inducing autophagy (45). Porcine epidemic virus activates the PERK, IRE1α and ATF6 signaling pathways in Vero cells to induce autophagy by upregulating the expression of LC3-II. The same authors showed that both PERK and IRE1α pathway inhibitors limit the expression level of LC3-II and cause a reduction in virus titer in infected cells (46).

The UPR also plays an important role in the replication of pathogens in the host. For example, Guimarães et al. (42) confirmed that Brucella abortus infection induces the expression of UPR target genes Bip and XBP1 in mouse macrophages. After treatment with the UPR inhibitor TUDCA, Brucella abortus levels in the cells decreased. Xue et al. (47) proved that transmissible gastroenteritis coronavirus (TGEV) infection can activate the UPR via the PERK pathway both in vitro and in vivo, and in vitro this negatively regulates TGEV replication. In our study, we found that the number of C. psittaci inclusions in cells is lower than that in the normal infection group, either following Bip siRNA interference or after GSK2606414 and 4μ8C inhibitor treatment. Thus, replication of C. psittaci in HBE cells is likely regulated by the UPR, PERK and IRE1α signaling pathways. Because these same pathways regulate autophagy in C. psittaci-infected cells, we hypothesize that C. psittaci may use autophagy to provide itself with the building blocks (i.e., amino acids, etc.) to promote its reproduction.

In conclusion, our results demonstrate for the first time that C. psittaci infection induces ERS and a complete autophagic process in HBE cells. The UPR and its associated PERK and IRE1α signaling pathways regulate the host-cell autophagy induced by C. psittaci infection and replication. These results provide insights into the interactions between C. psittaci and host cells, and may provide useful evidence for further research into the prevention and treatment of C. psittaci infection.

MATERIALS AND METHODS

Cell culture.

Human bronchial epithelial cells (HBE; ATCC 135-E6E7) and human cervical epithelial HeLa 229 cells (ATCC CCL-2.1) were cultured in 6-well, 24-well or 96-well flat-bottom plates (Corning, NY, USA) to a suitable density, and maintained in Dulbecco modified eagle medium (DMEM) (HyClone, Logan, USA) containing 10% (vol/vol) fetal bovine serum (FBS) (Gibco BRL, Gaithersburg, USA) at 37°C in an incubator supplied with 5% CO₂.

Chlamydias propagation and titration.

C. psittaci 6BC (ATCC VR-125), C. trachomatis Serovar E (ATCC VR-348B) or C. muridarum (Nigg strain, presented by Zhong, UTHSCSA Laboratory, USA) were propagated in confluent HBE cell monolayers in complete growth medium with 10% FBS as described previously (48). Chlamydial elementary bodies (EBs) were harvested 48 h postinfection (pi.). Cells containing mature EBs were centrifuged at 1,000 rpm for 10 min and then washed twice with phosphate-buffered saline (PBS) [pH 7.4] (Sigma-Aldrich, Munich, Germany). Cell pellets were resuspended in sucrose-phosphate-glutamate (SPG) buffer (0.2 moL/L sucrose, 3.8 mmoL/L KH₂PO₄, 6.7 mmoL/L Na₂HPO₄, 5 mmoL/L l-glutamic acid, pH 7.4), then disrupted by sonication and stored at −80°C until use. C. psittaci was used in all experiments at a multiplicity of infection (MOI) of 0.5∼2. The MOI was determined using standard serial dilution methods by counting the number of Chlamydial inclusion-forming units (IFU) in HBE cells using rabbit anti-C. psittaci antiserum.

Chlamydias infection and cell treatment.

HBE or HeLa229 cells were grown in 6-well or 24-well flat-bottom plates and maintained in DMEM supplemented with 10% FBS. After culturing overnight at 37°C in an incubator supplied with 5% CO₂ to a suitable density, the cells were infected with C. psittaci C. trachomatis or C. muridarum at an MOI of 0∼2, with an uninfected cell group as blank control. After 2 h adsorption, inocula were removed and the medium was replaced with complete DMEM containing 10% FBS. After 24 h, 12 h or 6 h of infection, proteins were extracted for WB analysis, mRNA was extracted for qRT-PCR analysis, and the autophagic structures of the cells were scanned by electron microscope, and the inclusions of C. psittaci were counted by immunofluorescence.

For the RNA interference experiment, prior to C. psittaci infection, 5 μL Lipofectamine 2000 (Invitrogen; Thermo Fisher Scientific, Inc.) and a suitable volume of Bip or negative-control (NC) small interfering RNA (siRNA) (Table 1) (GenePharma, China; 97% purity) were added into 500 μL antibiotic- and serum-free DMEM to obtain a final concentration of 50 nM for each siRNA and then incubated at room temperature for 20 min to allow the siRNA-Lipofectamine complex to form. Then, the mixture was added to HBE cells, and after incubating for 6 h, the medium was replaced with fresh DMEM. After incubation for 48 h, the cells were infected with C. psittaci for 24 h.

TABLE 1.

siRNA sequences of Bip and GAPDH

siRNA senseantisense	Sense	Antisense
BiP	5′-GAGGUGUCAUGACCAAACUTT-3′	5′-AGUUUGGUCAUGACACCUCTT-3′
GAPDH	5′-UUCUCCGAACG GUCACGUTT-3′	5′-ACGUGACACGUUCGGAGAATT-3′

For the pathway inhibition experiment, prior to chlamydia infection, the cells were treated with or without 40 μM PERK inhibitor GSK2606414 (MedChemexpress, FL, USA) or 30 μM IRE1α inhibitor 4μ8C (MedChemexpress) for 2 h. The outcome of both gene silencing and pathway inhibition experiments was measured by Western blotting.

Western blot analysis.

Cells were collected and lysed for 30 min with RIPA buffer (10 mM phosphate buffer pH 7.4, 150 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% sodium deoxycholate, 1% Triton X-100) containing 1 mM sodium orthovanadate and protease inhibitors. Lysates were collected and boiled at 100°C for 8 min. An equal amount of total protein was separated on 10% ∼15% SDS-PAGE gels and transferred to polyvinylidene difluoride (PVDF) membranes (Millipore, Billerica, USA). After blocking with 5% skimmed milk dissolved in TBST (0.02% Tween 20 and 1 M Tris buffer, pH 8.0) for 2 h at room temperature, the membranes were incubated with specific primary antibodies overnight at 4°C. Primary antibodies used: LC3A/B, Beclin1, SQSTM1/p62 rabbit MAbs (from Affinity, USA); Bip, PERK, p-PERK, eIF2α, IRE1α, p-IRE1α, ATF6 rabbit MAbs (from CST, USA); a GAPDH rabbit MAb (Proteintech, USA) was used as internal reference antibody. After washing three times with TBST, the membranes were incubated with horseradish peroxidase (HRP)-conjugated Affinipure Goat Anti-Rabbit IgG (H+L; Abcam) with a dilution ratio of 1:3000 in 5% skimmed milk for 1 h at room temperature and then analyzed using an Azure C300 (Azure Biosystems, California, USC) imaging system. Image J software was used for band density analysis.

Indirect immunofluorescence assay.

An indirect immunofluorescence assay (IFA) was used to analyze the formation of LC3 spots in the perinuclear region and to count the number of chlamydia inclusions. HBE cells in a 24-well plate with or without UPR pathway inhibition were infected with C. psittaci at an optimal MOI (1.5) for 24 h. Next, the cells were fixed with 4% paraformaldehyde for 30 min, and then permeabilized with 0.3% (vol/vol) Triton X-100 for 15 min and blocked with DMEM containing 10% FBS at 37°C for 1 h. After washing twice with PBS, the cells were immunostained overnight with rabbit anti-LC3A/B or anti-C. psittaci 6BC at 4°C. Following three washes with PBS, the cells were incubated with cy2-conjugated goat anti-rabbit IgG (1:200; Green, Jackson ImmunoResearch Laboratories, USA) containing Hoechst 33258 (1:4000; Sigma, USA) at room temperature for 1 h. After washing with PBS five times, the cells were viewed and photographed with an inverted fluorescence microscope (TS 2R; NIKON). The fluorescence of LC3 spots was analyzed using Image pro plus 6.0 software and the number of inclusions was counted using ImageJ.

Transmission electron microscopy.

HBE cells were plated in 96 mm cell culture dishes, and infected with C. psittaci at an MOI of 1.5 for 24 h. After immobilization with electron microscopy solution at 4°C for 2∼4 h, the cells were collected and centrifuged at 4°C for 2000 rpm for 5 min. The cells were pre-embedded in agar and fixed with 1% osmium acid (prepared with 0.1 M phosphate buffer) at room temperature for 2 h. Next, a series of 30%, 50%, 70%, 80%, 95%, 100%, and 100% alcohol solutions were used for dehydration at room temperature and then the cells were embedded in epoxy resin mixture (45% epon812, 30% DDSA, 23% MNA, 2% DMP30). After treatment for 48 h at 60°C, the embedded samples were ultrathin-sectioned (60–80 nm) with a microtome and stained with a 2% uranyl acetate-saturated alcohol solution in the dark for 8 min. After washing the slices, autophagosomes and autolysosomes were imaged under a transmission electron microscope (Hitachi, Ltd., Tokyo, Japan).

Real-time fluorescence quantitative PCR (qRT-PCR) analysis.

After infection with C. psittaci, C. trachomatis or C. muridarum, total RNA was extracted from HBE or HeLa cells with TRIzol reagent (Qiagen, Hilden, Germany) according to the manufacturer’s protocol. Reverse transcription was performed using Servicebio®RT First Strand cDNA Synthesis kit (Servicebio, China). The expression level of XBP1, ATF6, CHOP mRNA were measured using a fluorescent quantitative PCR system (Applied Biosystems, Inc., USA): predenaturation at 95°C for 10 min, then 40 cycles of denaturation at 95°C for 15 s, annealing and elongation at 60°C for 1 min. The primers used in this study are shown in Table 2. Data analysis was performed using the ΔΔCt method, with the Ct value of GAPDH as the standard ΔCt=the Ct value for XBP1/ATF6/CHOP – Ct value for GAPDH. Expression of the XBP1/ATF6/CHOP gene was reported as 2^-ΔΔCt.

TABLE 2.

Primer sequences of XBP1, ATF6, CHOP, and GAPDH gene

Gene	Primer sequence
XBP1	5′-GGAAGCTTGTCATCAATGGAAATC-3′ (sense primer)
	5′-TGATGACCCTTTTGGCTCCC-3′ (antisense primer)
ATF6	5′-CTGATGGCTGTTCAATACACAG -3′ (sense primer)
	5′-GATCCCTTCGAAATGACACAAC -3′ (antisense primer)
CHOP	5′-GAGAATGAAAGGAAAGTGGCAC-3′ (sense primer)
	5′-ATTCACCATTCGGTCAATCAGA-3′ (antisense primer)
GAPDH	5′-ATGGATTCTGGCGGTATTGACT-3′ (sense primer)
	5′-AGAGAAAGGGAGGCTGGTAAGG-3′ (antisense primer)

Statistical analysis.

Data are expressed as the mean ± standard deviation of at least three independent experiments and statistical analysis of the data were performed using SPSS version 23.0 software (SPSS, Inc., Chicago, IL, USA). One-way analysis of variance (ANOVA) was used for the comparison of multiple groups, and an independent sample t test was used to analyze the difference between two groups. Statistical images were analyzed and produced by OriginPro 9.0 software. A difference was considered to be statistically significant (*) at P < 0.05 or highly significant (**) at P < 0.01.