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Correspondence Guangyu Li, Department of Internal Medicine, Division of Infectious Diseases, University of Texas Medical Branch, 301 University Boulevard, Galveston, TX 77555-0435, USA. Tel: 409-747-0275; fax: 409-772-6527; email: email@example.com
Punta Toro virus (PTV; family Bunyaviridae, genus Phlebovirus) causes severe hepatic damage through brisk apoptosis of hepatocytes. In the present study, two viral proteins encoded by the S segment of the viral genome, non-structural (NSs) and nucleocapsid protein (N), were examined for their roles in apoptosis. Expression of NSs in HepG2 cells led to apoptosis in 45% of transfected cells, and with N, 28%, on average. These levels represent a four- to an eightfold increase over cells transfected with the mutated protein vectors. Caspase-3, -8 and -9 activities were increased by N protein when compared with the control NC (P < 0.05), and by NSsA and NSsB, as compared to control NSsC (P < 0.01). Treatment of the transfected cells with caspase-8 or -9 inhibitors markedly decreased apoptosis. Neutralization of TNF-α or Fas ligand had no effect on apoptosis. These results indicate that both NSs and N are responsible for causing hepatocyte apoptosis by triggering the extrinsic caspase-8 and intrinsic caspase-9 pathways.
terminal deoxynucleotidyl transferase-mediated dUTP-nick end labeling
Punta Toro virus, a member of the genus Phlebovirus, family Bunyaviridae, has been isolated repeatedly in Panama and Colombia, from phlebotomous sandflies and humans. Like most other members of the sandfly fever group of viruses, it produces an acute febrile illness lasting 2 to 5 days in infected people (1–3). Although there are fragmentary observations suggesting that RVFV attacks endothelium and initiates disseminated intravascular coagulation with concomitant hepatic damage (4), the pathogenesis of the phleboviruses is largely unknown.
In previous studies (5, 6), a hamster model of PTV-induced hemorrhagic fever was established and validated; this model had many similarities to the severe disease produced in animals and humans by RVFV (Phlebovirus genus). The brisk apoptosis of hepatocytes associated with the acute illness was one of the main characteristics (6). It was independently shown that PTV infection in vitro, without a secondary inflammatory cellular reaction, was responsible for hepatocytic apoptosis (7, 8). Cusi et al. have reported that TOSV, another phlebovirus, induced apoptotic death in neurons of experimentally infected mice (9). These observations suggest that inhibition of cellular apoptotic pathways during the acute infection may be a target for future therapeutics for diseases caused by phleboviruses. Thus, further examining the roles of individual viral components and their relationship with host cell response are important.
Previous studies (5) have also found that two geographically distinct strains of PTV (Adames and Balliet) produce a differential pathogenesis in the Syrian hamster, with PTV-Adames (PTVA) causing a RVFV-like illness with death, whereas animals infected with the PTV-Balliet (PTVB) strain survived. Genetic reassortments between PTVA and PTVB were used to investigate viral genetic determinants for pathogenesis and lethality in the hamster model; these studies concluded that the S segment of PTV was a critical determinant of lethality in the hamster (10). Similarly, the S segment of RVFV carries determinants for attenuation and virulence in mice (11). The S segment of PTV encodes the nucleocapsid protein (N) and non-structural protein (NSs). Amino acid sequences available to date show considerable homology between the N proteins of different phleboviruses, and at least five regions of the protein appear to be highly conserved (12). The NSs of RVFV contributes to viral pathogenesis by promoting cell death and/or inhibiting the early innate immune response in mice (13).
To expand our knowledge of phlebovirus pathogenesis, we cloned the N and NSs of PTVA and PTVB to assess the potential of these proteins in causing apoptosis of liver cells. The data obtained, using the constructed vectors, demonstrated that N and NSs were sufficient to induce apoptosis in transfected HepG2 cells. N- and NSs-induced apoptosis proceeded through caspase-8- and caspase-9-dependent pathways, as is the case with liver cells infected with PTV (8), and were independent of TNF-α or Fas signaling.
MATERIALS AND METHODS
Virus and cells
PTV strains (Adames and Balliet) were obtained from the World Reference Center of Emerging and Arboviruses Collection at University of Texas Medical Branch (UTMB). Both virus strains were originally isolated from febrile patients in Panama (14, 15). Virus stocks were generated from infected Vero cells. The Vero E6 cell line and human hepatocellular carcinoma HepG2 cell line were obtained from the American Type Culture Collection (Rockville, MD). HepG2 cells and Vero E6 cells were cultured at 37°C in 5% CO2 atmosphere in Dulbecco's modified Eagle's medium (Gibco/BRL, Carlsbad, CA) supplemented with 10% fetal bovine serum (FBS), and 100 μg/ml penicillin and 10 μg/ml streptomycin (Gibco/BRL).
Cloning of N and NSs
Total cellular RNA was extracted using an RNA isolation kit (Invitrogen, Carlsbad, CA) and was reverse transcribed into cDNA, using the superscript III reverse transcription system (Invitrogen). Primers were designed from previously published sequences (16, 17) and used for both PTV strains (Adames and Balliet). A negative control was made by inserting an additional base to the forward primers after the initiator codon ATG, which leads to expression of an irrelevant protein. The primers were: N forward, 5′ CACCATGTCATACGAAGAGATTGCC 3′; N reverse, 5′ AAAGAGGGATCTGAAGACTTTTGC 3′; NSs forward, 5′ CACCATGTCCAACAT AAACTATTAT 3′; NSs reverse, 5′ AAATATG TCTTGATTTAGCATTGT 3′; N control forward, 5′ CACCATGATCATACGAAGAGA TTGC 3′; NSs control forward, 5′ CACCATGATCCAACATAAACTATTAT 3′. The first methionine codon (ATG) and four base pair sequences (CACC) were added to all forward primers for TOPO cloning. The genes encoding NA and NB were amplified by PCR from cDNA of PTVA and PTVB, respectively, with the conditions of pre-denaturation at 94°C for 5 min, followed by 35 cycles of denaturation at 94°C for 45 sec, annealing at 60°C for 45 sec, extension at 72°C for 45 sec, and finally extension at 72°C for 10 min. The genes encoding NSsA and NSsB were obtained with the same conditions described above except for annealing at 52°C for 45 sec. The coding sequences for the control proteins, NC and NSsC, were amplified from the cDNA of the Adames virus, using the primers described above. PCR products were purified using the QIAquick PCR purification kit (Qiagen). TOPO cloning was carried out according to Invitrogen's TOPO cloning manual. Briefly, 1 μl purified PCR product, 1 μl salt solution, 1 μl pcDAN3.1/V5/His-TOPO vector (Invitrogen) and 3 μl sterile water were mixed and incubated for 5 min at room temperature. The mixture was transformed into TOP10 competent cells and plated on a Luria Bertani (LB) plate containing 50 μg/ml ampicillin.
Colonies of the six recombinants (pcDNA3.1/NA, NB, NC, NSsA, NSsB and NSsC) were randomly picked from the overnight plates and transferred into 1 ml LB medium containing 50 μg/ml ampicillin and grown at 37°C overnight at 220 r.p.m. Diagnostic PCR was carried out to check for positive clones. Plasmid DNA from positive colonies were sequenced using the ABI 3700 at the Core Laboratory of the UTMB, and digested with restriction enzymes Xho I and BamH I (Promega) followed by electrophoresis, to confirm that all target genes were correctly cloned into the pcDNA3.1/V5/His-TOPO vector.
Expression of the fusion proteins in Vero E6 cells
Vero E6 cells were transfected with pcDNA3.1/NA, NB, NC, NSsA, NSsB and NSsC, respectively, using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions. Briefly, sterilized round coverslips were placed into six-well plates and Vero E6 cells (5 × 105 cells/well) were seeded into the plates and cultured for 20 hr. The medium was removed and cells exposed to the transfection complexes (3 μg DNA and 9 μl liposome mixed in 500 μl of serum-free Opti-MEM I medium) were incubated overnight at 37°C. Cells were then cultured in growth medium for 24, 36, 42 hr post-transfection and fixed with acetone at 4°C for 15 min, respectively. For evaluation of expression of fusion proteins, the coverslips were carefully removed and examined by IFA assay. A PTV mouse immune ascitic fluid, diluted 1: 20 in PBS, or 50 μl anti-His antibody (Invitrogen) diluted 1: 200 in PBS, was added to each slide and incubated at 37°C for 1 hr. These were washed with PBST and incubated at 37°C for 50 min with 50 μl fluorescein isothiocyanate (FITC)-conjugated goat antimouse IgG (Sigma, St Louis, MO), diluted 1: 60 in PBS containing 1% Evan's blue dye. Evidence of specific fluorescence was monitored by fluorescence microscopy using an Olympus BX51 microscope (Leeds Intruments, Irving, TX). Each experiment was carried out three times, independently.
Western blot analysis
Transfected cells were washed three times in PBS and incubated in RIPA lysis buffer (Sigma) on ice for 10 min. The lysate was centrifuged at 10 000 g for 15 min at 4°C. An equal amount of protein (∼100 μg) from each lysate was subjected to 10% SDS-PAGE (Bio-Rad, Hercules, CA) and transferred onto a Sequi-Blot PVDF membrane (Bio-Rad). After blocking for 1 hr in 5% non-fat dried milk containing 0.1% (v/v) Tween 20, the membrane was incubated at 4°C overnight in the presence of PTV mouse ascitic fluid diluted 1: 20 in PBS, or anti-His antibody diluted 1: 200 in PBS. The membrane was washed and further incubated for 1 hr at room temperature with horseradish peroxidase-conjugated goat antimouse IgG (Abcam, Cambridge, MA), diluted 1: 2000 in PBS. After washing, immune reactive bands were detected using the Western Lightning Chemiluminescence Reagent (Amersham Bioscience, Piscataway, NJ).
Expression of fusion proteins in HepG2 cells
HepG2 cells were transfected with pcDNA3.1/NA, NB, NC, NSsA, NSsB and NSsC, respectively, using Lipofectamine 2000 (Invitrogen). Thirty-six hours later, expression was monitored using the methods described above for Vero E6 cells.
Detection of apoptosis
For each plasmid, three eight-well chamber slides (Nunc, New York, NY) were used. One was used for Annexin V-Fluorescein (BioVision Corporation, Mountain View, CA) staining, the second was used for TUNEL assay, and the third was used for observation of tranfection efficiency by IFA assay. For Annexin V-Fluorescein staining, the cells were grown in eight-well chamber slides (1 × 105 cells/well). Each plasmid (1 μg) was diluted in 125 μl Opti-Mem I medium (Invitrogen Corporation, Carlsbad, CA) and incubated for 5 min at room temperature. Lipofectamine (3 μl) in 125 μl Opti-Mem I were added to the DNA and allowed to form complexes for 20 min at room temperature. The DNA mixture was added to cells in one well containing 0.5 ml Opti-Mem I and incubated overnight. The medium was then exchanged for growth medium. Thirty-six hours post-transfection, the supernatant was discarded, and cells were washed with PBS. After removing the plastic divider, 50 μl staining solution (Roche Diagnostic, Indianapolis, IN) containing Annexin-V-Fluorescein and propidium iodide were added to each well with a coverslip and incubated for 15 min at room temperature in the dark. Analysis was carried out using fluorescence microscopy. Apoptotic cells were identified by green fluorescence and were counted in 10 randomly selected microscope fields (using 20× objective). The latter value was compared with the number of positive cells detected by IFA assay. For TUNEL assay, the cells were fixed in 1% paraformaldehyde in PBS (pH 7.4) in a Coplin jar for 10 min at room temperature, washed twice with PBS, and postfixed in precooled ethanol: acetic acid (2: 1 v/v) for 5 min. After washing with PBS twice, excess liquid was carefully removed. Equilibration buffer (75 μl) was added to each well for 1 min. Then, the TUNEL reaction mixture (Serological Corp., Norcross, GA) containing fluorescein isothiocyanate-labeled dUTP and terminal deoxynucleotidyl transferase was added immediately, and incubated at 37°C for 1 hr in a moist dark chamber. The slide was transferred to a Coplin jar containing working strength stop/wash buffer and incubated for 10 min at room temperature after it was agitated for 15 sec. Excess liquid was carefully removed and mounting medium containing 1 μg/ml propidium iodide was added. The slide was covered with a glass coverslip, and the stained cells were observed under fluorescent microscopy. The number of labeled cells was counted in 10 random microscope fields (using a 20× objective) and compared with the number of positive cells detected by IFA assay; the result was expressed as a percentage of the total transfected cell number.
Caspase activation and inhibition assay
To examine transfected cells for caspase-3, -8 and -9 activation, caspase-3, -8 and -9 Colorimetric Activity Assay Kits (Chemicon Millipore, Billerica, MA) were used. HepG2 cells (5 × 105 cells/well) were seeded into 24-well plates. Thirty-six hours after transfection, cell pellets were resuspended in 50 μl cell lysis buffer and incubated on ice for 10 min and then centrifuged at 10 000 g, 4°C for 10 min. The supernatant was transferred to a clean Eppendorf tube and mixed with 5× Assay Buffer (with 10 mM dithiothreitol) and 10 μl of 3 mg/ml caspase substrate (Ac-DEVD-pNA for caspase-3; Ac-IETD-pNA for caspase-8; Ac-LEHD-pNA for caspase-9) at 37°C for 2 hr. The mixture was then transferred to a 96-well plate and the absorbance value at 405 nm wavelength (A405) was recorded as the relative activity of caspase-3, -8 and -9.
For inhibition of caspases, cells were transfected and examined as for Annexin V-Fluorescein staining. The cell-permeable caspase-8 inhibitor Z-IETD-FMK, caspase-9 inhibitor Z-LEHD-FMK (R&D systems, Minneapolis, MN) was added to a final concentration of 200 μM 1 hr before plasmids were transfected. Dimethy sulfoxide (DMSO, 4%) was used as vehicle control.
To clarify whether TNF-α or Fas was also playing a role in N- and NSs-induced apoptosis, transfected cells were treated with antibodies (BD Biosciences, Sparks, MD) that inhibit signaling by either TNF-α or Fas ligand. Antibodies were added at a final concentration of 10 μg/ml in medium 1 hr prior to transfection with plasmids. Apoptotic cells were monitored by Annexin V-Fluorescein staining. The experiments described above were performed in triplicate.
The results are expressed as mean ± SD of at least four wells or ten 20× microscope fields. The images shown below are representatives for a set of experiments. The different numbers of apoptotic cells between groups were calculated for significance by ANOVA Student's t-test with StatView software. P < 0.05 was considered statistically significant.
Recombinant eukaryotic expression vectors for N and NSs proteins
Figure 1a shows the target genes amplified by RT-PCR. These were successfully cloned to make the constructed eukaryotic expression vectors: pcDNA3.1-V5-His-Topo-Adames-NSs (NSsA), pcDNA3.1-V5-His-Topo-Balliet-NSs (NSsB), pcDNA3.1-V5-His-Topo-NSs control (NSsC), pcDNA3.1-V5-His-Topo-Adames-N (NA), pcDNA3.1-V5-His-Topo-Balliet-N (NB), pcDNA3.1-V5-His-Topo-N control (NC). They expressed proteins of approximately 31 kDa (Table 1). Digestion with XhoI and BamHI confirmed the expected sizes of the cloned fragments (Fig. 1b). These were sequenced and confirmed by comparing to published sequences of Adames (ACCESSION # EF201835) and Balliet (ACCESSION # EF201834).
Table 1. Nucleocapsid and non-structural proteins of PTV and their target genes
No. base pairs
No. amino acids of fusion protein
Size of fusion protein (kDa)
N of Adames (NA)
N of Balliet (NB)
Mutant of N (NC)
Non-structural of Adames (NSsA)
Non-structural of Balliet (NSsB)
Mutant of NSs (NSsC)
Expression of recombinant vectors in Vero E6 cells
As both PTV Adames and Balliet proliferate well in Vero E6 cells (18), we chose this cell line for initial transfection experiments to examine for efficient expression of each target protein. The expressed proteins of NA, NB, NSsA, NSsB, NC and NSsC were detected by IFA at three time points: 24, 36 and 42 hr after transfection. Maximum expression of each protein was detected at 36 hr after transfection (Fig. 2a,b), with cytoplasmic localization.
For further confirmation, the cells were collected at 36 hr post-transfection, and 100 μg of each protein from the cell lysate was subjected to western blot analysis (Fig. 2c) using PTV mouse immune ascitic fluid. NC and NSsC were non-reactive by PTV antibody, but were detected by anti-6His (C-terminal) antibody contained in the fusion protein. These fusion proteins were approximately 31 kDa, corresponding to the predicted sizes of the PTV N and NSs proteins.
Expression of recombinant vectors in HepG2 cells
To determine the roles of individual proteins encoded by the S segment of PTV on virus-induced hepatocytic apoptosis, we transfected a widely used human hepatoma cell line HepG2 with the constructed plasmids. Thirty-six hours after transfection, IFA detected the intranuclear localization of the expressed proteins (Fig. 3a) in HepG2 cells. Interestingly, NC and NSsC showed cytoplasmic localization in HepG2 cells, the same as in Vero E6 cells (Fig. 3b).
Expression of PTV NSs or N protein induces apoptosis of HepG2 cells
As described above, detection of apoptosis was done on two eight-well chamber slides after transfection with the plasmid DNA using both Annexin-V-Fluorescein and TUNEL staining; another eight-well chamber slide was used to observe transfection efficiency by immunofluorescence assay to detect fusion tag 6His. Annexin V is a Ca2+-dependent phospholipid-binding protein with high affinity for phosphatidylserine, which is translocated from the inner part of the plasma membrane to the outer layer in the early stages of apoptosis (19). TUNEL assay detects DNA strand breakage by using the terminal deoxynucleotidyl transferase (TdT), which is a hallmark in the last phase of apoptosis (20, 21).
Apoptotic cells were counted at 36 hr after transfection. With annexin V staining (Fig. 4a,b), apoptotic cells induced by NA and NB were 2.6-fold and 3.3-fold greater than that by NC (P < 0.01), respectively; and apoptotic cells induced by NSsA and NSsB were 5.1-fold and 7.2-fold greater than that produced by NSsC (P < 0.01), respectively. By TUNEL assay, apoptotic cells induced by NA and NB were 5.1-fold and 5.3-fold higher than that induced by NC (P < 0.01), respectively; apoptotic cells induced by NSsA and NSsB were 8.4-fold and 8.1-fold higher than that produced by NSsC (P < 0.01), respectively (Fig. 5a,b).
Using IFA antigen-positive cells as the base (total number of HepG2 cells expressing the target protein), overall, NSs or N led to apoptosis of 45% and 28% of transfected cells, respectively, on average, four- to eightfold higher than that of control proteins.
Activation of caspases in NSs- and N protein-expressing cells
Multiple apoptotic pathways may lead to cell death, involving several caspases. In our previous studies, it was shown that PTV induces apoptosis through both caspase-8- and caspase-9-dependent pathways (18, 22).
To determine whether or not these pathways are involved in NSs- and N-induced apoptosis, caspase-3, -8- and -9 activities were assayed using Ac-DEVD-pNA, Ac-IETD-pNA and Ac-LEHD-pNA, respectively. These are caspase substrates allowing direct detection of caspase activation. Comparisons were made of caspase-3, -8 and -9 activities in NSs- or N-expressing cells to NSsC- and NC-expressing cells. As shown in Figure 6, caspase-3, -8 and -9 activities were significantly higher in NSs- and N-expressing cells. Caspase-3 activity significantly increased in NA (P= 0.0142) and NB (P= 0.0044) compared to NC in NSsA (P= 0.0025) and NSsB (P= 0.017) as compared with NSc. Caspase-8 activity significantly increased in NA, NB, NSsA and NSsB compared with NC or NSsC (P= 0.0064, 0.0243, 0.0001 and 0.0063, respectively). Caspase-9 activation was also seen in NA, NB, NSsA and NSsB compared with NC or NSsC (P= 0.0127, 0.0045, 0.0121 and 0.0262, respectively).
To further examine the roles of caspases-3, -8 and -9 activation, inhibition of caspase activity was tested. HepG2 cells were incubated with specific cell-permeable caspase inhibitors: Z-IETD-FMK for caspase-8 and Z-LEHD-FMK caspase-9. NA-, NB-, NssA- and NSsB-transfected HepG2 cells treated with caspase-8 inhibitor, caspase-9 inhibitor or both showed a significant decrease in apoptosis (Fig. 7). These findings indicate that N or NSs induces apoptosis of HepG2 cells through caspase-8- and caspase-9-dependent pathways, suggesting that both the intrinsic and extrinsic apoptosis pathways are involved in N- or NSs-induced HepG2 cell apoptosis.
In our previous study using intact PTV, it was shown that TNF-α and its receptor also play a role in HepG2 cells apoptosis (death-receptor activation-induced apoptosis) (22). To investigate the role of these molecules in N- and NSs-induced apoptosis of liver cells, transfected cells were treated with antibodies that inhibit the signaling pathway by either TNF-α or Fas ligand. There was no change in the level of apoptosis in the transfected cells with treatment of either antibody as compared with untreated controls (Fig. 8). These findings suggest that TNF-α or Fas is unlikely play a significant role in N- or NSs-induced apoptosis in liver cells.
Host cell death by viral pathogens depends on both viral and host cell factors. Virus-driven host cell apoptosis is considered to play a critical role in host pathogenesis. So, investigating viral pathogens that induce cell death is important. In the present study, we provide evidence for the induction of host cell death induced by the S segment of PTV, which encodes the NSs protein in the antigenomic (positive-sense) and N protein in the genomic (negative-sense) orientation, and is very important in the determination of viral virulence (6, 23). Here, we found for the first time that expression of either NSs or N in the absence of other PTV genes could directly induce apoptosis in HepG2 cells, which includes the induction of cell membrane blebbing and convolution, and chromatin disintegration. Immunofluorescent studies on the expression of N and NSs revealed a cytoplasmic distribution pattern in Vero E6 cells, but a perinuclear localization pattern in HepG2 cells. Perhaps this is essential for the apoptosis processing function of HepG2 cells.
The mechanisms by which N and NSs induced apoptosis in HepG2 cells were examined, which involve caspases-3, -8 and -9. Expression of NSs and N initiated the activation of caspase-8 and -9, which in turn, trigger the activation of effector caspase-3. The use of cell-permeable caspase inhibitors such as Z-IETD-FMK and Z-LEHD-FMK that are specific to caspase-8 and -9, respectively, confirmed the participation of these caspases in the observed apoptotic process. As we know, two well-studied distinct pathways of apoptosis include the death receptor (extrinsic) pathway and the stress (mitochondria-dependent, intrinsic) pathway. The death receptor pathway requires FADD (Fas Associated protein with Death Domain), which is a key adaptor molecule transmitting the death signal by activating the caspase-8/caspase-3 cascade leading to apoptosis. In contrast, stress factors, such as reactive oxygen species, induce apoptosis through mitochondria, which release cytochrome c that, in turn, activates the caspase cascade through caspase-9 (23, 24). In this study, blocking of the Fas receptor or TNF-α with anti-Fas or anti-TNF-α antibody did not affect the apoptosis. These results demonstrated that N and NSs induced apoptosis of HepG2 cells through activation of caspase-8- and caspase-9-mediated activation of caspase-3, but not through a TNF-α- or Fas-triggered signal. This is understandable, as the viral proteins were expressed within the cells, bypassing the cell surface receptors. However, experimental results from the detection of caspase activity and the use of caspase inhibitors demonstrated that both intrinsic and extrinsic pathways were shown to be involved in the NSs- and N-induced apoptosis.
The use of cell lines to study virus-induced cell death may generate results in which apoptosis occurs in a non-viral protein-specific manner. For example, in the present study, NSs or N protein-initiated HepG2 cell death cannot be accounted for entirely through caspase-8- and -9-dependent pathways. This indicates that additional pathways may also be involved, and may not be viral protein specific. To address this concern of non-specific induction, we included mutation genes to express non-specific proteins as a control in this study, which allows assessment of whether or not the apoptosis of HepG2 cells induced by NSs and N is a specific result.
The biological functions of NSs of other bunyaviruses have been extensively studied, including promoting apoptosis and/or inhibiting cellular translation (22, 25). Analysis of RVFV by the T7 RNA polymerase-driven minigenome system demonstrated that NSs protein enhances the RNA-dependent RNA polymerase activities of L and N proteins to promote efficient virus RNA replication (26). NSs has also been shown to mediate the pan-down regulation of mRNA production by the inhibition of RNA polymerase II activity and, via this mechanism, the NSs protein plays a critical role in mammalian host pathogenesis by indirectly disrupting the host cell antiviral response (11, 27, 28). Being a major structural component, the N protein of bunyaviruses also carries out functions interfering with important regulatory pathways in the infected cells, which plays a critical role in viral pathogenesis (28–30). In the present study, we extended the functions of NSs and N of PTV in the viral pathogenesis.
In conclusion, we have found that both N and NSs proteins induce apoptosis of hepatocytes, which is a main pathogenic component in Phlebovirus-induced liver injury and disease. This is the first study demonstrating the direct roles of N and NSs proteins in this process, which is associated with both the extrinsic and intrinsic signal pathway. Further studies are needed to gain insight into how the cellular proteins interact with N or NSs proteins. Also, our findings have implications regarding targeted therapeutics for Phlebovirus-associated hemorrhagic fevers.
We thank Konstantin Tsetserkin for fine technical assistance in western blot and Dora Salinas for help in preparing the manuscript. This work was supported by contracts NO1-AI25489 from the National Institutes of Health.