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In this study, we show that the highly pathogenic H5N1 avian influenza virus (AIV) (A/crow/Kyoto/53/04 and A/chicken/Egypt/CL6/07) induced apoptosis in duck embryonic fibroblasts (DEF). In contrast, apoptosis was reduced among cells infected with low-pathogenic AIVs (A/duck/HK/342/78 [H5N2], A/duck/HK/820/80 [H5N3], A/wigeon/Osaka/1/01 [H7N7], and A/turkey/Wisconsin/1/66 [H9N2]). Thus, we investigated the molecular mechanisms of apoptosis induced by H5N1-AIV infection. Caspase-dependent and -independent pathways contributed to the cytopathic effects. We further showed that, in the induction of apoptosis, the hemagglutinin of H5N1-AIV played a major role and its cleavage sequence was not critical. We also observed outer membrane permeabilization and loss of the transmembrane potential of the mitochondria of infected DEF, indicating that mitochondrial dysfunction was caused by the H5N1-AIV infection. We then analyzed Ca2+ dynamics in the infected cells and demonstrated an increase in the concentration of Ca2+ in the cytosol ([Ca2+]i) and mitochondria ([Ca2+]m) after H5N1-AIV infection. Regardless, gene expression important for regulating Ca2+ efflux from the endoplasmic reticulum did not significantly change after H5N1-AIV infection. These results suggest that extracellular Ca2+ may enter H5N1-AIV-infected cells. Indeed, EGTA, which chelates extracellular free Ca2+, significantly reduced the [Ca2+]i, [Ca2+]m, and apoptosis induced by H5N1-AIV infection. In conclusion, we identified a novel mechanism for influenza A virus-mediated cell death, which involved the acceleration of extracellular Ca2+ influx, leading to mitochondrial dysfunction and apoptosis. These findings may be useful for understanding the pathogenesis of H5N1-AIV in avian species as well as the impact of Ca2+ homeostasis on influenza A virus infection.
Avian influenza viruses (AIVs) are classified as highly or low-pathogenic AIVs (HPAIVs or LPAIVs, respectively) based on their pathogenicity in chickens (1). HPAIVs cause systemic infections and high mortality in chickens (28), whereas poultry are asymptomatic or develop mild respiratory problems and/or intestinal illness after LPAIV infection (49). Hemagglutinin (HA) cleavability is a critical determinant of AIV pathogenicity in avian species (61). Other determinants, such as nonstructural (NS) protein and neuraminidase (NA) protein, reportedly regulate the virulence of AIVs (9, 29, 44). However, waterfowl, known as the natural host for AIVs, do not usually have any symptoms during an HPAIV infection (21), whereas they show neurologic symptoms and death after infection with some of the recently emerged HPAIVs, such as the Asian H5N1 virus (11, 46, 62). Thus, the entire mechanism responsible for the pathogenicity of the AIVs is not yet known. Unknown cellular and viral factors probably underlie the pathogenesis of HPAIVs in avian species, especially waterfowl.
The alveolar epithelial cells (66) or vascular endothelial cells (32) of human patients and chickens infected by H5N1-AIV show apoptosis. Other reports suggest that apoptosis of these cells is essential for the development of acute lung injury in mice and acute respiratory distress syndrome in humans (39), which is often observed in H5N1-AIV-infected patients. Therefore, it is necessary to evaluate whether apoptosis is critical for the pathogenesis of H5N1-AIV in vivo and to understand the molecular mechanisms of the apoptotic cell death induced by H5N1-AIV infection.
Ca2+ is a key regulator of cell survival, and the breakdown of Ca2+ homeostasis, due to sustained elevations in Ca2+ inside cells, triggers programmed cell death involving apoptosis (24). Indeed, disruption of Ca2+ homeostasis plays a key role in apoptosis during the pathogenic process of several types of viral infections, including those with human immunodeficiency virus (HIV), hepatitis C virus, and human T-cell leukemia virus type 1 (3, 4, 31, 57). In addition, the HIV gp120 envelope protein induces neuronal cell death through Ca2+ dysregulation, even in the absence of viral particles (25).
In this study, we used duck embryonic fibroblasts (DEF) to elucidate the molecular mechanisms of the apoptotic cell death induced by H5N1-AIV. We show here that H5N1-AIV infection triggered extracellular Ca2+ influx and that this alteration in the concentration of Ca2+ inside the cells subsequently induced mitochondrial dysfunction and led to apoptotic cell death. In addition, we demonstrate that H5N1-HA was a critical viral factor for inducing apoptosis.
DEF and chicken embryonic fibroblasts (CEF) were prepared using 14-day-old duck eggs and 11-day-old chicken eggs, respectively, as described previously (5), and cultured in Dulbecco's modified Eagle's medium (DMEM)-low glucose (1.0%) supplemented with 10% fetal bovine serum (FBS) and antibiotics. Human embryonic kidney 293T cells were also cultured in DMEM-low glucose supplemented with 10% FBS and antibiotics. Madin-Darby canine kidney (MDCK) cells were cultured in minimal essential medium supplemented with 10% FBS and antibiotics. The HPAIVs (A/crow/Kyoto/53/04 [Cw/Kyoto] [H5N1] and A/chicken/Egypt/CL6/07 [H5N1]) were isolated from embryonated chicken eggs inoculated with tracheal or lung homogenates from dead crow or dead chicken, respectively (18). The LPAIVs (A/duck/HK/342/78 [Dk/HK] [H5N2], A/duck/HK/820/80 [H5N3], A/wigeon/Osaka/1/01 [H7N7], and A/turkey/Wisconsin/1/66 [H9N2]) were kindly provided by Yoshinobu Okuno, Osaka Prefectural Institute of Public Health. Cw/Kyoto, A/chicken/Egypt/CL6/07, Dk/HK, A/duck/HK/820/80, A/wigeon/Osaka/1/01, and A/turkey/Wisconsin/1/66 were grown in 9-day-old embryonated chicken eggs (60) and titrated by a focus-forming assay with DEF (17). DEF were infected with each virus at a multiplicity of infection (MOI) of 3 for 1 h at 37°C and then cultured in DMEM-F-12 containing 3% FBS and antibiotics.
The transcription plasmids and protein expression plasmids were constructed as described previously (20) but with slight modifications. Briefly, the pCAGGS/MCSII set of protein expression plasmids was constructed by inserting the open reading frames of the polymerase proteins (PB2, PB1, and PA) and nucleoprotein (NP) into the multicloning site of the pCAGGS/MCSII plasmid. Those genes were derived from influenza A/WSN/33 virus (14). To generate the plasmids that directed the synthesis of the eight vRNA segments (PB2, PB1, PA, HA, NP, NA, matrix [M], and NS), each of the full-length positive-strand segments of Cw/Kyoto and Dk/HK was cloned into pUC18-based plasmids, between the human RNA polymerase I promoter and the hepatitis delta virus ribozyme (pPOLI). Mutant HA genes, replacing N′-RRKKR-C′ with N′-TR-C′ at the cleavage site, were generated by site-directed mutagenic PCR (GeneTailor site-directed mutagenesis system; Invitrogen, Carlsbad, CA) using Cw/Kyoto cDNA that had been inserted into the pGEM-T Easy vector (Promega, Madison, WI) and subcloned into the transcription plasmid pPOLI. A mutant HA gene, replacing N′-TR-C′ with N′-RRKKR-C′ at the cleavage site in A/duck/HK/820/80, was generated as described previously (14).
We transfected 1.75 × 105 293T cells that had been cocultured with 0.75 × 105 CEF on 24-well plates by TransIT-LT1 (Mirus, Madison, WI) according to the manufacturer's instructions. Briefly, 1.5 μl of transfection reagent was diluted in 50 μl of Opti-MEM (Invitrogen) and incubated for 10 min at room temperature. Then, the mixture was added to the premixed plasmids: 31.25 ng of each of the eight vRNA coding transcription plasmids (pPOLI-PB2, -PB1, -PA, -HA, -NP, -NA, -M, and -NS) and 62.5 ng of each of the protein expression plasmids (pCAGGS/MCSII-PB2, -PB1, -PA, and -NP). After incubation for 20 min at room temperature, the plasmid mixtures were added to the cells. During production of the recombinant viruses with the monobasic amino acids in the HA cleavage site, 5 μg/ml of acetylated trypsin (Sigma, St. Louis, MO) was added to the plates at 1 and 4 days posttransfection. At 7 days posttransfection, the supernatants of the cultures were collected and injected into 9-day-old chicken eggs. The allantoic fluid was collected by 3 days postinfection (p.i.) and titrated by the number of focus-forming units (FFU). The recombinant Dk/HK virus that included the HA gene of Cw/Kyoto was generated by the helper virus method as described previously (51). Briefly, pPOLI-HA derived from Cw/Kyoto was transfected with pCAGGS/MCSII protein expression plasmids into 293T cells cocultured with CEF. At 24 h posttransfection, cells were infected with Dk/HK at an MOI of 1, as a helper virus. At 72 h p.i., recombinant viruses in the supernatants were plaque purified from MDCK cells in the absence of trypsin. The presence of the HA gene in the recombinant viruses was confirmed by sequence analysis.
The 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) cell survival assays were performed according to the manufacturer's instructions (Cell Titer 96 Aqueous One Solution cell proliferation assay; Promega). Briefly, 20 μl of MTT was added to the culture medium. After incubation at 37°C for 90 min, 25 μl of 10% sodium dodecyl sulfate was added to the cells and then the absorbance of each sample was measured at 492 nm with an automated plate reader (Multiskan MS-UV; Labsystems, Helsinki, Finland). The results were calculated as the absorbance ratio of infected to mock-infected cells.
The RNA of the progeny particles was extracted from the infected culture medium with a QIAamp viral RNA mini kit (Qiagen, Hilden, Germany) in order to quantify the number of viral genomic RNA copies. Total RNA was extracted from the infected cells using TRIzol (Invitrogen) so that the number of copies of the viral mRNA could be quantified. Next, cDNA was synthesized using RevaTra Ace reverse transcriptase (Toyobo, Osaka, Japan) with an M gene-specific primer (5′-GGTACCCGGGAGCGAAAGCAGGTAGATATT-3′) for genomic RNA (27) or an oligo(dT) primer for mRNA. Synthesized cDNA was subjected to a SYBR green real-time PCR assay (Applied Biosystems, Foster City, CA) with the Applied Biosystems StepOne real-time PCR system. pPOL-M was used for the standard curve. All reactions were performed in triplicate. The primer sequences were as follows: M forward primer, 5′-TTCTAACCGAGGTCGAAACG-3′, and M reverse primer, 5′-ACAAAGCGTCTACGCTGCAG-3′ (30).
The infected cells were stained with 10 μM Hoechst 33342 (Sigma) at 37°C for 60 min. Apoptotic cells were characterized by chromatin condensation or nuclear fragmentation, and the number of apoptotic cells was counted from a population of more than 500 cells.
The activity of caspases 3/7, 8, and 9 was measured using the Caspase Glo assay kit (Promega) according to the manufacturer's instructions. Briefly, 100 μl of Caspase Glo reagent was added to the culture medium of the infected cells in a 96-well white-wall plate. After incubation at room temperature in the dark for 30 min, the luminescence of the samples was measured with a luminometer (Multi-Detection Microplate Reader PowerscanHT; DS Pharma Biomedical, Osaka, Japan). Luciferase activity was corrected against the amount of total protein (bicinchoninic acid [BCA] protein assay kit; Thermo Fisher Scientific, Waltham, MA), and the n-fold increase in protease activity was determined by comparing the luciferase activity of the infected cells with that of the mock-infected cells.
Cellular fractionation was performed according to the procedures of Wang et al. (67). Briefly, 4 × 106 cells were washed with phosphate-buffered saline and collected with a scraper. The cell pellets were collected after centrifugation and resuspended with 200 μl of the homogenate buffer (250 mM sucrose, 10 mM Tris-HCl [pH 7.4], 2 mM EDTA, and protease inhibitor cocktail [Nacalai Tesque, Kyoto, Japan]). The cells were homogenized with a 1-ml Dounce homogenizer via 40 strokes on ice and were centrifuged at 1,000 × g for 10 min. The supernatants were recentrifuged at 13,000 × g for 60 min. The pellets were considered the mitochondrial (Mit) fraction and were resuspended with 50 μl of the homogenate buffer. The supernatants were considered the cytosolic (Cyt) fraction.
The protein concentrations of the mitochondrial and cytosolic samples were quantified with the BCA protein assay kit (Thermo Fisher Scientific). Next, 20 μg of protein or 15 μg of protein was used with Western blotting to quantify the amount of cytochrome c or apoptosis-inducing factor (AIF), respectively, in the samples. Blots were probed with an anti-cytochrome c (Zymed, South San Francisco, CA) or anti-AIF (Cell Signaling, Danvers, MA) primary monoclonal antibody (MAb), followed by a horseradish peroxidase-conjugated secondary antibody. Blots were visualized using an enhanced chemiluminescence system (Amersham ECL Plus Western blotting detection system; GE Healthcare, Buckinghamshire, United Kingdom).
The culture medium of the infected cells was changed to DMEM-F-12 that did not include phenol red and that was supplemented with 2 μM JC-1 dye (Sigma). After incubation at 37°C for 30 min, the cells were observed with a fluorescence microscope.
The culture medium of the infected cells was changed to DMEM-F-12 that did not include phenol red, and 5 μM Fluo-4 AM (Invitrogen) or 10 μM Rhod-2 AM (Invitrogen) was added as an indicator of Ca2+ in the cytosol ([Ca2+]i) or mitochondria ([Ca2+]m), respectively. After incubation at 37°C for 30 min, the cells were observed with a fluorescence microscope.
The infected samples were subjected to Western blotting and probed with an anti-AIV (Dk/HK) polyclonal antibody (18), or an anti-glucose-regulated protein 78 (GRP78) (Santa Cruz Biotechnology, Santa Cruz, CA), anticalnexin (Affinity BioReagents, Golden CO), anti-phosphorylated protein kinase R-like endoplasmic reticulum (ER) protein (P-PERK) (Cell Signaling), anti-Bcl-2 (BD Biosciences, San Jose, CA), or anti-Bax (Santa Cruz Biotechnology) MAb, followed by a horseradish peroxidase-conjugated secondary antibody (65). Blots were visualized as described above. The amount of α-tubulin (protein loading control) in each sample was assessed with an anti-α-tubulin primary antibody (Sigma).
The infected samples were dissolved in TRIzol reagent (Invitrogen), and the total RNA was extracted, reverse transcribed with an oligo(dT) primer, and subjected to PCR using Go Taq DNA polymerase (Promega) with primers for the viral M gene, ER degradation-enhanced α-mannosidase-like protein 1 (EDEM1) (F, 5′-CATCCTCTTTGGAGAGAAGG-3′; R, 5′-CCATGTGTTCATCAGCTGTCC-3′), and β-actin (control) (F, 5′-CACAGATCATGTTTGAGACCTT-3′; R, 5′-CATCACAATACCAGTGGTACG-3′) (15).
Data are expressed as means ± standard errors of the means (SEM). Statistical analysis was performed by Student's t test. A P value of <0.01 was considered significant.
We used the H5N1-AIVs, a group of recently emerged HPAIVs (Cw/Kyoto and A/chicken/Egypt/CL6/07), and compared their cytotoxicity to those of several LPAIVs (Dk/HK, A/duck/HK/820/80, A/wigeon/Osaka/1/01, and A/turkey/Wisconsin/1/66). The domestic ducks infected with Cw/Kyoto belonging to subclade 2.5 would not show any clinical signs (37), whereas those infected with A/chicken/Egypt/CL6/07 belonging to subclade 2.2 showed neurological symptoms and died suddenly (2).
DEF infected with these AIVs were incubated at 37°C, and cell viability was measured with the MTT assay at 24, 48, and 72 h p.i. Viability decreased among all AIV-infected DEF at 24 and 48 h p.i., indicating AIV-induced cytotoxicity (Fig. (Fig.1A).1A). At 72 h p.i., cell viability continued to deteriorate among the H5N1-AIV-infected cells but not among the LPAIV-infected cells (Fig. (Fig.1A).1A). This finding suggests that infection with the H5N1-AIVs induced progressive, irreversible cytotoxicity in DEF, whereas infection with the LPAIVs induced moderate cytopathicity. In addition, the percentages of apoptotic cells induced by each type of viral infection were similar at 24 h p.i. (Fig. (Fig.1B).1B). However, infection with the H5N1-AIVs resulted in higher numbers of apoptotic cells at 48 h p.i. than did infection with the LPAIVs (Fig. (Fig.1B).1B). In contrast, necrotic cell death was not detected in the DEF infected with the H5N1-AIVs or LPAIVs (data not shown). At 48 h p.i., the MTT assay showed that viabilities of cells infected with A/chicken/Egypt/CL6/07 and LPAIVs were similar (Fig. (Fig.1A),1A), whereas the apoptotic cell numbers of A/chicken/Egypt/CL6/07-infected cells were significantly higher (Fig. (Fig.1B).1B). This discrepancy is probably due to the property of the MTT assay. As the assay is used for measuring the activity of mitochondrial enzymes, it can sometimes be influenced by metabolic activity, resulting in large differences in the results even if the number of viable cells is kept constant.
In order to elucidate the molecular mechanisms underlying the H5N1-AIV-induced apoptosis, we utilized Cw/Kyoto (H5N1) and Dk/HK (H5N2) as a control. First, we measured viral mRNA expression and progeny virion production from the DEF infected by each virus. At 0, 4, 8, 12, 24, 36, and 48 h p.i., infected cells were collected and subjected to quantitative real-time PCR with an M gene-specific primer set. The amounts of viral mRNA obtained per total RNA were similar for the two viruses at each time point (Fig. (Fig.2A).2A). Also, the culture medium of the infected cells was replaced at 6, 12, 24, 36, and 48 h p.i. so that the number of progeny viruses could be determined via quantitative real-time PCR and the focus-forming assay. As found for viral mRNA expression in the infected cells, the amounts of viral genome (copies per ml) in the culture medium were similar between the two viruses (Fig. (Fig.2B).2B). However, FFU were somewhat higher following Cw/Kyoto infection at 24 h p.i. (Fig. (Fig.2B),2B), suggesting that infectious viral particles continue to be produced in the late phase of Cw/Kyoto infection, whereas defective particles would increase during Dk/HK infection. Thus, we conclude that the differences observed in the amounts of apoptosis induced by Cw/Kyoto and Dk/HK were not due to viral replication.
In order to determine which gene is essential for the apoptosis, we generated a set of Cw/Kyoto-derived recombinant viruses; each of 8 genes of Cw/Kyoto virus was replaced by those of Dk/HK virus. Infection with the recombinant virus containing the HA gene of Dk/HK (rCw/Kyoto-HADk) improved cell viability only in comparison with the parent Cw/Kyoto virus (Fig. (Fig.3A),3A), although it still resulted in lower cell viability than did the parental Dk/HK infection (Fig. (Fig.3B).3B). These results were consistent with our previous report showing that H5N1-HA was critical for apoptosis of mammalian primary cells (14). Subsequently, another recombinant virus, rDk/HK-HACw, was generated in which the HA gene of Cw/Kyoto was placed into the Dk/HK virus. Figure Figure3B3B showed that infection with rDk/HK-HACw induced significantly lower cell viability at 48 and 72 h p.i. as did Cw/Kyoto infection. Furthermore, like Cw/Kyoto, rDk/HK-HACw induced severe levels of apoptotic death of the infected cells (data not shown). Therefore, the HA gene of Cw/Kyoto could be involved in the process of apoptosis induced by H5N1-AIV infection, although other viral genes may also be involved. Next, in order to determine whether the multibasic amino acids were involved in the apoptosis induced by H5N1-AIV infection, we used two recombinant viruses: (i) recombinant Cw/Kyoto in which the multibasic cleavage site of the parent virus was changed to a monobasic cleavage site (rCw/Kyoto-HACwmono) and (ii) recombinant A/duck/HK/820/80 in which the monobasic cleavage site of the parent virus was changed to a multibasic cleavage site (rA/duck/HK/820/80-HAN3multi). These parental and recombinant viruses showed similar mRNA expression kinetics until 48 h p.i. (data not shown). Cell viability of the DEF infected with the parent or recombinant viruses was assessed with the MTT assay at 24, 48, and 72 h p.i. At every time point, the viability of the cells infected with each recombinant virus was similar to the viability of the cells infected with each parent virus (Fig. (Fig.3C),3C), suggesting that HA cleavability is not important for inducing apoptosis of DEF infected with H5N1-AIVs.
We assessed the involvement of caspases in the cell death process induced by infection because caspases participate in several biochemical cascades that mediate apoptosis (50). Caspase activity was assayed in the culture medium as well as in cells infected with Cw/Kyoto or Dk/HK by a luminometer at 12, 24, 48, and 72 h p.i. The activity of caspases 3/7, 8, and 9 was significantly elevated at 48 h p.i. with Cw/Kyoto compared to that with Dk/HK (Fig. (Fig.4A).4A). Next, DEF were treated with 100 μM pan-caspase inhibitor Z-VAD-fmk. Twelve hours later, the cells were infected with Cw/Kyoto, and then caspase activity and cell viability were measured at 0, 24, 48, and 72 h p.i. Caspase 3/7 activities were inhibited in the Z-VAD-fmk-treated DEF at every time point (Fig. (Fig.4B,4B, left panel); however, the treated cells were as viable as the untreated cells (Fig. (Fig.4B,4B, right panel), indicating that apoptosis of avian cells following AIV infection was primarily through a caspase-independent pathway(s). However, the possible contribution of a caspase-dependent pathway(s) to the cell death process could not be eliminated.
Because mitochondria are a central component of the apoptotic signaling pathways (36), DEF infected with AIVs were evaluated for mitochondrial outer membrane permeabilization (MOMP). DEF were mock infected or infected with Cw/Kyoto or Dk/HK, as was performed for the experiments shown in Fig. Fig.1.1. The infected cells were collected at 42 h p.i. and processed in order to obtain the Mit and Cyt fractions. Mitochondrial intermembrane space proteins such as cytochrome c and AIF were detected in the Mit and Cyt fractions by Western blotting. DEF infected with Cw/Kyoto showed marked liberation of these proteins into the Cyt fraction (Fig. (Fig.5A).5A). Although these proteins were also liberated in DEF infected with Dk/HK, the amounts of these proteins that leaked from the Mit to the Cyt compartment were smaller than those for the Cw/Kyoto-infected cells (Fig. (Fig.5A).5A). Indeed, the Cyt/Mit ratio for AIF was significantly higher for the Cw/Kyoto-infected cells than for the Dk/HK-infected cells, as assessed with Image-J software (Fig. (Fig.5B).5B). We also investigated whether MMP was retained in the infected cells, utilizing JC-1 dye staining. At 24 h p.i., most of the cells infected with Cw/Kyoto or Dk/HK retained MMP (Fig. (Fig.5C,5C, upper panels). However, at 48 h p.i., MMP was severely lost by cells infected with Cw/Kyoto but retained by most cells infected with Dk/HK (Fig. (Fig.5C,5C, lower panels). These results suggest that Cw/Kyoto infection induced more severe MOMP and loss of MMP than did Dk/HK infection, resulting in mitochondrial dysfunction.
Previous reports indicate that one of the factors causing mitochondrial dysfunction is an elevated concentration of [Ca2+]m (22, 33, 48, 64). Thus, we investigated Ca2+ dynamics in DEF infected with AIVs. As shown in Fig. Fig.6,6, infected cells were stained with Fluo-4 AM and Rhod-2 AM dyes as an indicator of [Ca2+]i and [Ca2+]m, respectively, at 24, 36, and 48 h p.i. DEF infected with each virus showed similar, moderate increases in [Ca2+]i and [Ca2+]m at 24 h p.i. (Fig. (Fig.6).6). Marked elevations in [Ca2+]i and [Ca2+]m were detected in the Cw/Kyoto-infected cells at 36 and 48 h p.i., whereas moderate increases were observed for the Dk/HK-infected cells (Fig. (Fig.6).6). These results suggest that Cw/Kyoto infection induced a marked elevation in [Ca2+]i and [Ca2+]m in the infected cells.
Elevation of [Ca2+]i is caused by the efflux of Ca2+ from the ER ([Ca2+]ER), the influx of extracellular Ca2+, or both (19, 45). Because efflux of [Ca2+]ER is closely related to the ER stress response, we analyzed the expression levels of ER stress-mediated proteins and mRNA. As shown in Fig. Fig.7A,7A, the protein expression levels of GRP78 and calnexin increased to similar extents at 12 h p.i. in Cw/Kyoto- and Dk/HK-infected cells as in the positive control, 10 μM thapsigargin-treated cells. P-PERK was not detected in cells infected with either virus during our observation period, which is consistent with previous findings (34). The mRNA expression levels for EDEM1 were similarly elevated at 4 h p.i. for the two types of viral infections (Fig. (Fig.7B).7B). The expression levels of EDEM1 and the internal control (β-actin) decreased in Cw/Kyoto-infected cells, probably due to severe cell toxicity, at 8 h p.i. (Fig. (Fig.7B).7B). The expression levels of the Bcl-2 and Bax proteins, which regulate Ca2+ efflux from the ER (54, 59), were comparable at every time point for cells infected with Cw/Kyoto and those infected with Dk/HK (Fig. (Fig.7C).7C). These results suggest that AIV infection induced ER stress; however, ER stress did not directly contribute to the differential increase in [Ca2+]i between the Cw/Kyoto- and Dk/HK-infected cells. Thus, we shifted our focus toward understanding the pathway mediating the extracellular influx of Ca2+.
DEF infected with Cw/Kyoto were treated with 2.5 mM EGTA, a chelator of extracellular Ca2+, at 24 h p.i., and the concentrations of [Ca2+]i and [Ca2+]m were analyzed as discussed for the experiments shown in Fig. Fig.6.6. Reduced levels of [Ca2+]i and [Ca2+]m were found in the presence of EGTA at 48 h p.i. (Fig. (Fig.8A).8A). Especially, EGTA treatment inhibited elevation of [Ca2+]i, resulting in the reduced [Ca2+]i levels of mock-infected cells (Fig. (Fig.8A).8A). These results suggest that the influx of extracellular Ca2+ caused an increase in [Ca2+]i and [Ca2+]m in the Cw/Kyoto-infected cells. In addition, the cytopathic effects of Cw/Kyoto infection were attenuated in the EGTA-treated cells (Fig. (Fig.8A).8A). As shown in Fig. Fig.8B,8B, viral titers in the supernatants were similar in the presence and absence of EGTA at 36 h p.i. The viral titer was a little higher in the absence of EGTA at 48 h p.i., indicating that apoptosis might promote viral release. Moreover, the number of apoptotic cells was significantly reduced at 36 and 48 h p.i. in EGTA-treated DEF that had been infected with Cw/Kyoto (Fig. (Fig.8C),8C), which is consistent with the observations shown in Fig. Fig.8A.8A. Finally, the number of apoptotic cells was counted at 48 h p.i. in DEF infected with the recombinant viruses. In rCw/Kyoto-HACwmono- and rDk/HK-HACw-infected cells, the numbers of apoptotic cells markedly decreased after EGTA treatment as in Cw/Kyoto-infected cells (Fig. (Fig.8D).8D). As expected, the number of apoptotic cells was not significantly altered by the presence of EGTA after infection with Dk/HK or rCw/Kyoto-HADk (Fig. (Fig.8D).8D). Taken together, these findings suggest that the influx of extracellular Ca2+ was a critical factor leading to apoptosis following H5N1-AIV infection. In addition, the HA gene of H5N1-AIV might be involved in the apoptosis signaling pathway.
In this study, we showed that Cw/Kyoto (HPAIV) induced significantly higher levels of apoptosis than did Dk/HK (LPAIV) in avian cells and that the HA glycoprotein had a critical function in viral cytotoxicity. We also demonstrated that Cw/Kyoto infection induced the influx of extracellular Ca2+ which elevated [Ca2+]i and [Ca2+]m and led to MOMP and the loss of MMP. Additional investigations revealed that mitochondrial dysfunction led to apoptosis via caspase-dependent and -independent mechanisms. The proposed mechanism of H5N1-AIV-induced apoptosis is shown in Fig. Fig.99.
Ca2+ functions as a universal second messenger in virtually all eukaryotic cells (23, 24). Cells maintain a very low [Ca2+]i by actively pumping Ca2+ out of the cell, into the ER, and by binding Ca2+ to various host molecules (19). Temporally and spatially organized increases in [Ca2+]i, [Ca2+]m, or nuclear Ca2+ ([Ca2+]n) serve as common intracellular signaling mechanisms (24). However, prolonged changes in Ca2+ distribution, including elevations in [Ca2+]i, [Ca2+]m, or [Ca2+]n or a decrease in [Ca2+]ER, trigger a variety of cellular cascades that lead to cell death (24). So far, several reports have shown that Ca2+-dependent apoptosis occurs following infections with viruses (10) such as hepatitis C virus (3), human T-cell leukemia virus type 1 (57), HIV (25, 26, 31), and rotavirus (56). In addition to these findings, we show here that H5N1-AIV infection induced an influx of extracellular Ca2+, leading to apoptosis of DEF. We also found that EGTA treatment significantly reduced apoptosis of CEF infected with Cw/Kyoto but not cells infected with Dk/HK (data not shown), although both viruses induced apoptosis in CEF via both caspase-dependent and -independent pathways (data not shown). These findings are consistent with the results shown for DEF in Fig. 8C and D. To our knowledge, this is the first report showing that the influx of extracellular Ca2+ induces apoptosis following infection with influenza A virus (55).
The mechanisms underlying H5N1-AIV infection-mediated changes in Ca2+ influx and the subsequent elevation in [Ca2+]i are still unknown. Several factors are known to cause elevations in [Ca2+]i: opening of Ca2+ channels on the plasma membrane, downregulation of the Ca2+ pumps in the ER and the plasma membrane, and disruption of the ability of Ca2+-binding proteins to bind free [Ca2+]i (19, 24). In this study, we confirmed by a reverse-genetics approach that the HA of Cw/Kyoto was involved in the apoptosis signaling pathway (Fig. (Fig.33 and and8).8). There are 49 amino acids, out of a total of 567 amino acids, that differ between the HA proteins of Cw/Kyoto and Dk/HK. Thus, recombinant AIVs expressing a series of chimeric HA proteins could provide information regarding the functional domain(s) and residue(s) that contribute to the ability of these viruses to induce apoptosis. As shown in Fig. Fig.3C,3C, the cleavage sequence within HA is not involved in the ability of these viruses to induce apoptosis. Therefore, another domain within HA could contribute to the cytotoxicity of the H5N1-AIV in avian species. Also, inactivated virus showed neither cytopathicity nor elevation in [Ca2+]i or [Ca2+]m (see Fig. S1 in the supplemental material), suggesting that de novo viral replication is essential for this apoptosis pathway. HA glycoprotein could alter Ca2+ influx and elevate [Ca2+]i through mechanisms as described above. Further investigations using inhibitors or blockers of the Ca2+ channels and pumps (43), immunoprecipitation (31), or small interfering RNA (siRNA) knockdown (16) approaches may reveal the key factor(s) important for disrupting Ca2+ homeostasis in H5N1-AIV-infected cells.
In addition to HA, another viral factor(s) may participate in apoptotic cell death, as shown in Fig. Fig.3B.3B. Reportedly, expression of the H5N1-NS1 protein induced apoptosis in human airway epithelial cells (40), and double-stranded RNA (dsRNA), NA, PB1-F2, and M proteins induced apoptosis following infection with the influenza A virus (8, 12, 58). PB1-F2 is known to facilitate release of proteins housed within the mitochondrial intermembrane space that trigger apoptosis (68). In our study, infection with the recombinant Cw/Kyoto virus, with its PB1 gene replaced by the Dk/HK PB1 gene, resulted in severe cytotoxicity and loss of MMP, as observed with the parent Cw/Kyoto (Fig. (Fig.3A3A and data not shown). These results suggest that the PB1 genes of Cw/Kyoto and Dk/HK played either equal roles or no role in the induction of apoptosis. Indeed, both viruses possess Asn at position 66 in PB1-F2, which confers a less virulent genotype than that of the viruses with Ser at position 66 (13). Additional investigations with reverse-genetics methods are necessary to elucidate the functional role of the viral factors that induce apoptosis after H5N1-AIV infection.
We have shown that [Ca2+]i and [Ca2+]m increased after H5N1-AIV infection and that mitochondrial dysfunction via elevated [Ca2+]m must be essential for activating the apoptotic signaling pathway. On the other hand, reportedly, elevations in [Ca2+]i lead to increased [Ca2+]n, resulting in the cleavage of nuclear DNA by nucleases, disruption of cytoskeletal organization, and mitochondrial dysfunction during apoptosis (47, 52, 53). Therefore, DNA damage induced by increased [Ca2+]n may also contribute to apoptosis. We were unable to directly determine whether the efflux of [Ca2+]ER contributed to the induction of apoptosis. Previous reports indicate that ER stress is a major reason for the efflux of [Ca2+]ER and, conversely, that efflux of [Ca2+]ER induced ER stress (3). In this study, although ER stress temporarily occurred after AIV infection, ER stress-mediated genes were expressed at similar levels in cells infected with Cw/Kyoto and those infected with Dk/HK (Fig. 7A and B). These findings suggest that the efflux of [Ca2+]ER is not critical for H5N1-AIV-induced apoptosis.
In Fig. Fig.4B,4B, we show that the cytotoxicity of an AIV infection did not change with Z-VAD-fmk treatment. Previous reports suggest that the most probable reason that the caspase inhibitors failed to prevent cell death was that these inhibitors were unable to prevent MOMP, which occurs upstream of caspase activation in many pathways that culminate in cell death (6, 38). In this study, marked MOMP was detected after AIV infection; therefore, the efficacy of Z-VAD-fmk may have been diminished. On the other hand, we previously reported that apoptotic cell death was markedly reduced with Z-VAD-fmk treatment of human and swine airway epithelial cells infected with H5N1-AIV (14). Inconsistencies in the results may be due to the animal species of the host cells; the caspase-independent pathway plays a predominant role in apoptotic cell death in avian cells infected with H5N1-AIV, whereas the caspase-dependent pathway is critical for mammalian cells.
In this study, we did not elucidate whether the disruption of Ca2+ homeostasis was essential for the pathogenesis of H5N1-AIV in vivo. However, Ca2+ imbalance is thought to be related to the pathogenesis of several types of viral infection (3, 7, 25) and various pathophysiologies (35, 52, 63), including acute lung injury (41, 42). Therefore, extracellular Ca2+ influx and elevations in [Ca2+]i may be important precursors to the pathophysiology caused by H5N1-AIV infection. Further investigations into Ca2+ regulation and balance with H5N1-AIV infection are required.
We thank Yoshinobu Okuno, Osaka Prefectural Institute of Public Health/Research Foundation for Microbial Diseases of Osaka University, for helpful advice and Yohei Watanabe, Ritsuko Kubota-Koketsu, and Atsuyo Yoshioka, RIMD, Osaka University, for helpful advice and technical assistance.
This work was supported in part by a Grant-in-Aid for Scientific Research from the Ministry of Education, Science, Sports, Culture, and Technology (MEXT) to T.N.; a Grant-in-Aid for Young Scientists from the Japan Society for the Promotion of Science (JSPS) to T.D.; and a grant from the Kato Memorial Bioscience Foundation to T.D. M.U. is a research fellow of the JSPS Research Fellowships for Young Scientists. M.S.I. is the recipient of a JSPS postdoctoral fellowship for foreign researchers.
Published ahead of print on 6 January 2010.
†Supplemental material for this article may be found at http://jvi.asm.org/.