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The use of inactivated influenza virus for the development of vaccines with broad heterosubtypic protection requires selective inactivation techniques that eliminate viral infectivity while preserving structural integrity. Here we tested if a hydrophobic inactivation approach reported for retroviruses could be applied to the influenza virus. By this approach, the transmembrane domains of viral envelope proteins are selectively targeted by the hydrophobic photoactivatable compound 1,5-iodonaphthyl-azide (INA). This probe partitions into the lipid bilayer of the viral envelope and upon far UV irradiation reacts selectively with membrane-embedded domains of proteins and lipids while the protein domains that localize outside the bilayer remain unaffected. INA treatment of influenza virus blocked infection in a dose-dependent manner without disrupting the virion or affecting neuraminidase activity. Moreover, the virus maintained the full activity in inducing pH-dependent lipid mixing, but pH-dependent redistribution of viral envelope proteins into the target cell membrane was completely blocked. These results indicate that INA selectively blocks fusion of the virus with the target cell membrane at the pore formation and expansion step. Using a murine model of influenza virus infection, INA-inactivated influenza virus induced potent anti-influenza virus serum antibody and T-cell responses, similar to live virus immunization, and protected against heterosubtypic challenge. INA treatment of influenza A virus produced a virus that is noninfectious, intact, and fully maintains the functional activity associated with the ectodomains of its two major envelope proteins, neuraminidase and hemagglutinin. When used as a vaccine given intranasally (i.n.), INA-inactivated influenza virus induced immune responses similar to live virus infection.
Influenza virus is a negative-strand RNA virus belonging to the Orthomyxoviridae family of pH-dependent viruses. In spite of the availability of a variety of influenza virus vaccines, yearly epidemics occur affecting 10 to 20% of the general population and as much as 30% of school age children (36). Gradual changes in the coding sequences of the surface proteins hemagglutinin and neuraminidase (HA and NA, respectively) cause changes in these antigens that accumulate over time and are positively selected by immune responses in vaccinated or infected individuals. This process, called antigenic drift, gives rise to variants that can infect individuals immune to the parental strain and gives rise to periodic epidemics every 2 to 5 years (43). Additionally, influenza A virus has a segmented genome, and therefore different influenza A virus subtypes can undergo reassortment in the infected host to give rise to new viruses not present in the vaccine formulation. This process, called antigenic shift, gave rise to pandemics in the past and could result in an influenza pandemic in the near future. For that reason approaches for the rapid preparation of new and more immunogenic vaccines, ideally with enhanced heterosubtypic protection, are of utmost importance (39, 42).
Inactivation of viruses for vaccine application requires methods that kill the virus while preserving the integrity of the immunogenic epitopes on the viral envelope. For influenza virus, several inactivation techniques have been reported, including UV irradiation (5, 46), ionizing radiation (25), heat treatment (9, 10), and chemical inactivation (6, 35). The prevailing inactivation method used today for the preparation of influenza vaccines is treatment with chemicals like formalin or β-propiolactone followed by a detergent disruption process called splitting (2, 20). This procedure results in a degree of structural preservation of the virus sufficient to elicit a protective immune response (20, 36). Whole inactivated viruses that are not split as well as live attenuated viruses have consistently proved to be better immunogens and confer a more efficient protective response (19, 40).
We have recently reported for retroviruses an approach for the preparation of inactivated viruses with preservation of structural integrity for vaccine application (34). In this approach the photoactivatable alkylating membrane probe 1,5-iodonaphthylazide (INA) is used to target selectively the transmembrane segments of proteins in the viral envelope while preserving the epitopes on the surface of the virus. In this work it was of interest to examine if this approach could be generally applied to other enveloped viruses by testing it on influenza A virus and whether improved integrity of the viral antigens would enhance vaccine immunogenicity and induce heterosubtypic immunity.
Purified influenza virus strain X-31 (H3N2) was prepared by and obtained from Charles River Laboratories (N. Franklin, CT). INA was synthesized and supplied by Combinix Inc. (San Mateo, CA). 3,3′-Dioctadecyloxacarbocyanine perchlorate (DiO), chloromethylfluorescein diacetate (CMFDA), and the Amplex red neuraminidase assay kit were from Invitrogen-Molecular Probes (Carlsbad, CA). PKH-67 and PKH-26 were from Sigma. The 3-1 KB carcinoma cell line was generously supplied by Suresh Ambudkar from the Laboratory of Cell Biology, National Cancer Institute, National Institutes of Health. The construct with CD4 conjugated to green fluorescent protein (CD4-GFP) was a generous gift from W. Popik from the Oncology Center, The Johns Hopkins University School of Medicine, Baltimore, MD. Mouse-adapted A/Puerto Rico/8/34 (PR/8; H1N1; kindly provided by Suzanne Epstein, Center for Biologics Evaluation and Research, Food and Drug Administration, Bethesda, MD) was propagated in the allantoic cavity of embryonated hen eggs at 35°C for 72 h.
Inactivation of influenza virus by INA was carried out essentially as previously described for retroviruses (34). In short, band-purified X-31 influenza virus was suspended in phosphate-buffered saline (PBS) at a concentration of 1 mg/ml protein. INA from a 40 mM stock solution in dimethyl sulfoxide was added to the virus suspension in several installments and thoroughly mixed. The virus was incubated for 15 min in the dark and irradiated with UV light using a 100-W mercury lamp source for 2 min. The INA-treated virus was divided into two experimental groups. One group was subjected to infectivity assays, and the second group was tested for neuraminidase activity.
To monitor infectivity of inactivated viruses, we assessed binding of red blood cells to influenza virus-infected cells as described previously (31). Briefly, 25 μg of the virus in Dulbecco's modified Eagle's medium plus 2% bovine serum was added in duplicates to KB carcinoma cells grown to confluence in six-well plates. The virus was incubated with the cells for 2 to 3 h at 37°C, and then the cells were washed and further incubated with fresh Dulbecco's modified Eagle's medium plus 10% bovine serum overnight at the same temperature. The cells were washed again and labeled with the cytosolic fluorescent probe CMFDA (excitation at 485 nm, emission at 525 nm) following the manufacturer's protocol (Molecular Probes). The cells were then washed with PBS and incubated with a suspension of 0.5% human erythrocytes prelabeled with the fluorescent membrane probe PKH-26 (excitation at 530 nm, emission at 580 nm; Sigma). After 15 min of incubation at room temperature the cells were washed with PBS five times and lysed with a solution of 1% Triton X-100. The fluorescence values of PKH26/CMFDA at 580/525 nm were measured using a Cytofluor fluorescence plate reader.
Neuraminidase activity was determined by measuring the hydrogen peroxide-induced fluorescence of a probe using the Amplex red neuraminidase assay kit and following the manufacturer's protocol.
Fusion by photosensitized labeling was measured essentially as previously described for enveloped viruses (29, 33, 34). In short, human erythrocyte resealed ghosts (1 ml) were labeled with the fluorescent membrane probe PKH-67 (Sigma) and incubated with 20 μCi 125INA for 15 min on ice. The ghosts were washed and divided into five experimental groups. Influenza virus at 1 mg/ml protein was treated with different concentrations of INA and irradiated as described above (34). The virus was mixed with the resealed ghosts for 30 min at room temperature and washed, and then the appropriate amount of citric acid was added to the mixture to lower the pH to 5.0. The tubes were further incubated at 37°C for 15 min and then irradiated at 488 nm for 30 seconds using an argon laser at an intensity of 0.5 W. The ghosts were lysed, and the HA was isolated by immunoprecipitation and run on an sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel. The radioactivity incorporated in the HA molecules was measured by autoradiography using a phosphorimager. Controls were the following: (i) untreated virus (100%); (ii) untreated virus that was bound to the ghosts and kept at neutral pH 7.4 (signal of 0).
Labeling of virus with octadecyl rhodamine (R-18) and fusion with red blood cell (RBC) ghosts were performed as previously described (8, 29, 31). Influenza virus was inactivated with 100 μM INA as described above and labeled with R-18. The R-18-labeled virus was prebound to RBC resealed ghosts at room temperature for 20 min and washed. The sample was transferred to a cuvette in the fluorimeter at 37°C, and the pH was lowered to 5.0 by adding citric acid. Fluorescence was monitored before and after the addition of the citric acid. Percent fluorescence dequenching was determined as follows: 100 × [(F − F0)/(Ft − F0)], where F0 and F are the fluorescence intensities at time zero and at a given time point, respectively, and Ft is the total fluorescence obtained after disruption of cells by Triton X-100. As a control for nonspecific dequenching, the virus was inactivated by preexposure to pH 5.0 followed by incubation with cells at pH 7.4 (22).
Fluorescence recovery after photobleaching (FRAP) was performed as previously described (16) using a Zeiss LSM 510 confocal laser scanning microscope (Carl Zeiss, Jena, Germany). HeLa cells were plated on 35-mm glass-bottom dishes (MatTek, Ashland, MA) and either transfected with CD4-GFP 24 h prior to confocal analysis as described previously (30) or labeled with DiO 1 hour prior to the FRAP analysis. INA was added to the cells from a stock of 30 mM in dimethyl sulfoxide to a final concentration of 20 or 100 μM. After 10 min of incubation at room temperature, the sample was irradiated with UV light at 10 mW/cm2 for 2 minutes. The cells were then submitted to FRAP while being kept under physiological conditions of 37°C and 5% CO2 in a stage incubation system (Incubator S; PeCon GmbH, Erbach, Germany). A 488-nm Ar+ laser line was used for excitation, and emission light was collected with a 500-550 band-pass filter. A 40×, 1.3 numerical aperture oil immersion objective lens was used with a zoom factor of 4. The detector pinhole was opened slightly to acquire an optical section of 2-μm thickness. This allowed more light to be collected, for better quantification. Three prebleach images were acquired to determine the rate of nonpurposeful photobleaching. Photobleaching was performed by increasing the transmission of the laser to 100% for 20 to 50 iterations to optimize the extent of bleaching. Following photobleaching, 8 to 10 images were acquired at 1-second intervals and then the acquisition rate was changed to 10-second intervals to follow the recovery to completion. A total of 20 to 40 images were acquired. FRAP analysis was performed using the MIPAV software package (CIT, National Institutes of Health, Bethesda, MD) with a one-dimensional diffusion FRAP model to retrieve the mobile fraction (24). Data were automatically corrected with background subtraction as well as normalization for the nonpurposeful photobleaching rate calculated from the whole cell membranes. All in vitro experiments were repeated three times except for the photosensitized labeling, which was performed twice.
BALB/c mice (5 to 7 weeks old, female; Harlan, Indianapolis, IN) were anesthetized with 2,2,2-tribromoethanol via intraperitoneal injection. For immunizations, each mouse received 30 μl of PBS, 30 μl of live X-31 (15 μg total protein), 30 μl of INA-treated X-31 (15 μg total protein), or 30 μl of influenza B/AA/1/86 virus (100 50% tissue culture infective dose) i.n. For lethal challenge, 28 days postimmunization, mice were infected i.n. with 10 50% lethal doses (LD50) of PR/8 in 50 μl of PBS, under anesthesia with an intraperitoneal injection of 0.2 ml of 2,2,2-tribromoethanol in tert-amyl alcohol (Avertin; Aldrich Chemical Co., Milwaukee, WI). Some animals were euthanized on day 5 postchallenge for analysis of lung virus titers. The remaining animals were monitored for body weight and mortality until all animals had succumbed to infection or were recovering, based on body weight.
Influenza virus-specific antibody titers were measured in serum and lung lavage (broncheoalveolar [BAL]) fluid using an isotype-specific enzyme-linked immunosorbent assay (ELISA). Enzyme immunoassay plates (Corning, Lowell, MA) were coated with 50 μl/well of X-31 (3 × 105 PFU/ml) or A/PR/8/34 (4 × 105 PFU/ml) overnight at 4°C. Viruses coated on plates were UV inactivated, and the plates were washed. The wells were blocked with 200 μl/well starting block buffer (Pierce, Rockford, IL) twice for 1 min. After washing, dilutions of sera or BAL fluid samples were added (50 μl/well) and incubated at room temperature for 2 hours. Plates were washed, and 100 μl/well of a 1:1,000 dilution of alkaline phosphate-labeled rat anti-mouse immunoglobulin M (IgM), IgG2a, IgG2b, and/or IgA (KPL, Gaithersburg, MD, or Southern Biotechnology, Birmingham, AL) was added. After incubation for 1 hour at room temperature, the plates were washed, 100 μl/well pNPP phosphatase substrate (KPL) was added, and the enzymatic reaction was allowed to develop at room temperature. The optical density was measured at 405 nm on a BioTek Powerwave plate reader.
Pooled serum samples collected from mice before infection and 3 weeks postinfection were treated with Vibrio cholera receptor-destroying enzyme (Denka-Seiken, Tokyo, Japan), heat inactivated at 56°C for 30 min, and absorbed with packed chicken red blood cells (cRBCs). Pooled serum samples were tested for HA inhibition (HI) antibodies to the virus with which the mice were infected using an HI assay with 0.5% cRBCs, as described elsewhere (40). Viruses were diluted to contain four agglutinating units in sterile PBS solution.
Plates (ELISPOT IP; Millipore, Billerica, MA) were coated with 50 μl of Hanks' balanced salt solution containing 5 μg/ml of anti-gamma interferon (anti-IFN-γ) monoclonal antibody AN18 overnight at 4°C. After washing, the membrane was blocked with medium containing 10% fetal bovine serum for 60 to 90 min. Spleens were aseptically removed from euthanized mice, a single-cell suspension was prepared, and red blood cells were lysed. Dilutions (twofold) of splenocytes were added to wells starting at 250,000 cells/well in a volume of 50 μl. Peptides NP147-155 (TYQRTRALV; NP) and HA533-541 (IYSTVASSL; HA) were added at a final concentration of 1 μg/ml. After incubation for 48 h at 37°C, bound IFN-γ was detected by the addition of 50 μl per well of biotinylated monoclonal antibody R4-6A2 at 1 μg/ml. Spots were developed using alkaline phosphatase-labeled streptavidin (BD Biosciences, San Jose, CA) and 5-bromo-4-chloro-3-indolylphosphate-nitroblue tetrazolium substrate (KPL). IFN-γ spot counts were obtained using an AID ELISpot plate reader (Autoimmun Diagnostika GmbH, Strassberg, Germany).
Weight loss after challenge was compared for survivors on each day using one-way analysis of variance (ANOVA) statistical analysis, followed by pairwise multiple comparison using the Holm-Sidak method. This overestimates body weight in groups with deaths, as generally the animals with the lowest body weights succumb to infection. Comparison of survival used the log rank test, followed by pairwise multiple comparison using the Holm-Sidak method. Overall significance level for post hoc tests was at a P level of 0.05. All statistical analysis was done with SigmaStat software v3.11 (Systat Software, Point Richmond, CA).
Influenza virus pellet was fixed in 2% glutaraldehyde in 0.1 M sodium cacodylate buffer at pH 7.0 for 2 h. Two μl of sample was placed directly on a Formvar-coated transmission electron microscopy grid, dried, and then stained with 1% phosphotungstic acid. After drying, the samples were examined in an electron microscope (Hitachi, Tokyo, Japan) operated at 75 kV, and digital images were obtained with a charge-coupled-device camera.
INA reduced the infectivity of the virus to 0 in a dose-dependent manner, while the catalytic activity of the viral envelope enzyme neuraminidase was not affected (Fig. (Fig.1).1). Infectivity is mediated by the fusion protein of influenza virus HA, which is an integral membrane protein of the viral envelope whose transmembrane segment's integrity is essential for full fusion and infectivity of the virus (11). Neuraminidase is also an integral membrane protein of the viral envelope, but its catalytic site is located on the hydrophilic segment that protrudes outside the membrane. In order to test if INA has an effect on the ability of the virus to fuse with the target cell membrane, fusion was measured directly by two different methods: dequenching of R-18 and photosensitized labeling. Dequenching measures the pH-dependent mixing of lipids from the viral envelope with lipids of the target cell, and this function was not affected by INA relative to nontreated viruses (Fig. (Fig.2).2). Similar results were obtained when the dequenching experiments were repeated using the nonexchangeable fluorescent lipid analogue PKH-26 instead of R-18. When X-31 influenza virus is exposed to low pH at 37°C prior to the mixing with the target cells, its fusion activity is inactivated. The ability of HA to mediate lipid mixing is regarded as a manifestation of the conformational change that the HA molecules undergo in response to low pH (3). The data shown in Fig. Fig.22 indicate that the conformational transitions of HA are not affected by INA, as both INA-treated and nontreated viruses were equally inactivated by preexposure to low pH at 37°C but not at 4°C. Photosensitized labeling, on the other hand, monitors fusion by measuring the pH-dependent redistribution of proteins from the viral envelope into the target cell membrane. Figure Figure33 shows that treatment of virus with INA blocked the redistribution of viral envelope proteins into the cell membrane in a dose-dependent manner and was dramatically reduced at the highest concentration of INA tested (75 μM). At this concentration the viral envelope proteins did not incorporate into the target cell membrane upon lowering the pH and remained in a position similar to where they were before the onset of fusion at neutral pH.
These results suggest that INA may have a general effect on the translational mobility of proteins in the membrane. To test this hypothesis we measured the diffusion of proteins and lipids in the HeLa cell membrane after INA treatment by FRAP. For these experiments we used CD4-GFP as the fluorescently labeled transmembrane protein and DiO as the lipid fluorescent probe. The results presented in Fig. Fig.44 show that the protein mobile fraction was reduced to the background level after treatment with INA, whereas the mobile fraction of the lipid was not affected. In order to further analyze the structural preservation of the inactivated virus, the viruses were visualized with an electron microscope by negative staining (Fig. (Fig.5).5). The electron microscopy images show clearly that the structure of the INA-treated viruses is preserved and that both groups are similar in appearance of the virions and their spikes. An interesting observation is the fact that the INA-treated viruses seem less permeable to phosphotungstate, which results in a lighter appearance of the lumen of the virus. This result may indicate that INA treatment may have a stabilizing effect on the membrane that causes the sealing of pores through which the negative stain can penetrate into the virus.
A murine model of influenza virus immunization was used to compare the immunogenicity of INA-inactivated X-31 to infection with live X-31 influenza virus. Female BALB/c mice were immunized one time either i.n. or subcutaneously (s.c.), and serum and in some cases bronchial alveolar lavage samples were collected for testing of specific antibody responses. Both INA-inactivated X-31 and live X-31 intranasally immunized mice exhibited robust serum antibody responses (Table (Table1),1), although with the exception of IgG2a, live X-31 infection induced about a threefold-greater serum antibody response with other IgG isotypes. While s.c. immunization with INA-inactivated X-31 induced serum IgG responses, they were 10- to 100-fold lower than in intranasally immunized animals. Intranasal immunization induced potent respiratory antibody responses as well, and in this case INA-inactivated X-31 induced threefold-greater IgA titers compared to live X-31 infection (Fig. (Fig.6A6A and Table Table11).
To test whether INA-inactivated X-31 induced strong cellular immune responses in addition to humoral responses, BALB/c mice were immunized intranasally as before and 28 days postimmunization, and splenic T-cell responses were measured by ELISPOT analysis. Both INA-inactivated X-31 and live X-31 immunization induced a high frequency of influenza virus-specific IFN-γ-producing cells in the spleen. Peptide-specific responses were measured for both an external influenza virus antigen, HA, and an internal, conserved antigen, NP, and in both cases were two- to threefold higher in INA-inactivated X-31-immunized mice (Fig. (Fig.6B6B).
Immunization with live or inactivated virus can induce potent neutralizing antibody responses and protect against homologous challenge. So, it was likely that mice immunized with live or INA-inactivated X-31 would be protected from homologous (X-31) challenge. To test for neutralizing activity against homologous virus, sera were tested for HI titer against X-31, PR8, and B/AA (H3N2, H1N1, and influenza B virus, respectively). Prebleed sera were tested for HI activity and were all negative (data not shown). Similarly, serum mock (PBS)-immunized animals had no measurable HI activity (Table (Table2).2). Both INA-inactivated X-31- and live X-31-immunized groups had robust HI titers against X-31 virus, suggesting they were immune to X-31 challenge, as HI titers of >40 are indicative of immunity to matched challenge. None of the groups had HI titers against PR8, and only B/AA-immunized mice had HI titers against the influenza B virus.
Immunized mice were given a heterosubtypic challenge of 10 LD50 of PR/8, 28 days after the single immunization described above. All of the mock-immunized (PBS) and negative control (B/AA-infected) mice rapidly lost weight and succumbed to the challenge, with only a single B/AA-infected mouse surviving (Fig. 7A and B, respectively). In contrast, INA-inactivated X-31 and live X-31 immunization provided robust protection, with animals losing only 10 to 15% of their body weight on average and none of them succumbing to infection. Average percent body weights of PBS and B/AA groups were significantly different from the live X-31 group on days 3 and 6 (P ≤ 0.004, ANOVA), while PBS and B/AA groups were significantly different from the INA X-31 group on day 6 only (P < 0.001, ANOVA) (Fig. (Fig.7A).7A). The limited numbers in the PBS and B/AA groups restricted weight comparisons of all groups after day 6; however, the average weight loss of INA-inactivated X-31- and live X-31-immunized mice was never significantly different at the measured time points. Survival rates between groups were also significantly different. Survival rates of B/AA- and PBS-immunized mice were significantly less than INA-inactivated X-31- and live X-31-immunized mice (P < 0.001, log rank), but B/AA and PBS groups or INA-inactivated X-31 and live X-31 groups were not significantly different from each other (P > 0.05) (Fig. (Fig.7B7B).
Previously we have shown for retroviruses that hydrophobic alkylating compounds like INA can selectively target the viral envelope and inactivate fusion while preserving the conformational integrity of the viruses (34). In this study we further examined this inactivation approach by application to the influenza virus, whose mechanism of fusion and cell entry is pH dependent and thus different from retroviruses. In influenza virus the pH-triggered redistribution of lipids and proteins during fusion can be measured separately by R-18 dequenching and photosensitized labeling, respectively. This feature provided the opportunity to obtain a better insight into the mechanism of inactivation and on the role of the hydrophobic domain of the viral envelope in viral fusion.
As expected, INA completely blocked the infectivity of influenza virus in a dose-dependent manner but had no effect on neuraminidase activity (Fig. (Fig.1).1). The influenza A viral envelope contains two major transmembrane proteins, HA and NA, with the enzymatic activity of NA residing in the hydrophilic ectodomain of the enzyme (23). Surprisingly, INA had no effect on the pH-dependent lipid mixing between virus and cells as measured by R-18 and PKH-26 dequenching, at concentrations higher than the one at which complete inactivation of infectivity is observed (Fig. (Fig.1).1). Both the rates and extents of dequenching were similar in INA-treated and control experiments, indicating that the total binding of the virus to the cell membrane was also not affected. R-18 or PKH-26 dequenching occurs as a result of pH-induced conformational changes of the HA molecule that trigger fusion (3, 7, 11). Another manifestation of the molecular transitions of HA is the observation that preexposure of the virus to low pH at 37°C in the absence of target cells results in inactivation of fusion (8, 31, 37), and no effect of INA could be detected on this parameter either (Fig. (Fig.2B).2B). In contrast to lipid mixing, treatment with INA blocked the pH-dependent integration and redistribution of viral envelope proteins into the target membrane as measured by photosensitized labeling of HA (Fig. (Fig.3).3). Interestingly, at 75 μM INA, conditions under which the viral envelope lipids seem to all be released into the target cell membrane, viral hemagglutinin mostly remains outside (Fig. (Fig.22 and and3).3). Previous studies in cell-cell fusion systems have shown that mutations or deletions in the transmembrane segment of HA produce a phenotype able to induce lipid mixing without pore formation (1, 21, 26). This phenomenon, called hemifusion, may also be manifested by the INA-inactivated virus. The profound difference between the movement of proteins and lipids during fusion of the INA-inactivated virus correlates with the differences in the translational mobilities of proteins and lipids in cell membranes that were similarly treated with this probe (Fig. (Fig.4).4). It has been previously shown in lipid bilayers that the ectodomain of BHA, which is capable of promoting hemifusion, is driven to aggregate by low pH (12). Likewise, the dynamics of pore formation by bacterial toxin peptides were also shown to require aggregation, oligomerization, and reorientation of the peptides after their insertion into the lipid bilayer. With the structural integrity of the INA-inactivated viral ectodomains and internal antigens maintained (Fig. (Fig.5),5), INA-treated X-31 induced immune responses comparable to and in some cases superior to live X-31 infection. Serum antibody responses, a measure of immune protection for inactivated influenza virus vaccines, were similar between immunizations, with IgG titers from virus-infected animals being at most threefold greater than INA-inactivated X-31-immunized mice and HI titers being equivalent (Tables (Tables11 and and2).2). At the same time, INA-inactivated X-31-immunized mice had increased mucosal IgA titers, which have been associated with enhanced homosubtypic and hetersubtypic protection (18, 38, 40). IgA antibodies are probably the major humoral contributor for the heterosubtypic protection observed in this study, although the effect of neutralizing IgG antibodies that are potent in vivo cannot be excluded. The INA-inactivated vaccine induced enhanced cytotoxic T-lymphocyte (CTL) responses, particularly to the highly conserved internal antigen NP. NP-specific CTL responses have been shown to provide robust protection to a spectrum of influenza A viruses, including highly pathogenic avian H5N1 influenza viruses (14, 15, 41). Taken together, the immune response analyses suggest INA-inactivated influenza virus could provide robust protection to matched, drift variant, or heterosubtypic virus infections.
Inactivated influenza virus vaccines approved for use in the Unites States consist of subvirion (“split”) or recombinant subunit antigen preparations and do not contain intact virus particles (17). These vaccines provide robust protection to matched influenza virus infection. The INA-inactivated vaccine leaves the virus particle and antigen functions largely intact, presenting potential advantages over traditional inactivated vaccines. A variety of studies have shown improved immunogenicity with particle-based vaccines, including enhanced antibody responses and stimulation of CD4+ helper T-cell and CD8+ CTL immune responses (reviewed in references 28 and 44). Combined, this multifaceted immune response could not only provide improved protection to matched virus challenge but also improve immunity to heterosubtypic infection (13). In a variety of experimental models, subunit and particle-based vaccines with or without immune-stimulating compounds or adjuvants have provided heterosubtypic immunity; however, in most cases, these vaccination regimens utilize a prime-boost strategy with multiple immunizations (18, 32, 40). Here, we show that similar to live virus infection, a single intranasal immunization with INA-inactivated influenza virus provides protection against heterosubtypic challenge.
The superior CTL response observed against NP antigens for the inactivated virus could be a major factor causing the heterosubtypic immunity. A possible explanation for the improved immunity relative to the live virus is that the intact and fusion-impaired inactivated virus is a superior immunogen that is more efficiently processed for antigen presentation.
In general terms this study demonstrates that hydrophobic alkylating compounds like INA completely inactivate the influenza virus by blocking fusion at the protein redistribution step while preserving the structural integrity and the biological activity of the ectodomains of the viral envelope proteins. Animal immunization experiments show that INA-inactivated influenza virus, an intact particle vaccine, induces robust serum and mucosal antibody responses as well as CD8+ T-cell responses. The breadth and potency of these responses provide protection to heterosubtypic infection, similar to a wild-type influenza virus infection, and suggest that INA inactivation may provide an opportunity for an improved influenza vaccine.
We thank Suresh Ambudkar from the Laboratory of Cell Biology, NCI, NIH, for his expert support and providing cell lines and materials. We thank Kunio Nagashima and M. Jason de la Cruz from the Image Analysis Laboratory Advanced Technology Program, SAIC-Frederick, for their expert support and help with electron microscopy. We thank Stephen Lockett from the image analysis laboratory for his expert advice about FRAP. We thank Frank Michel and Kari Kramer for excellent technical assistance and David Steinhauer (Emory University, Atlanta, GA) for scientific discussions. We thank W. Popik from the Oncology Center, The Johns Hopkins University School of Medicine, Baltimore, MD, for his support with the CD4-GFP construct.
This research was supported in part by the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research. This research was supported in part by the National Institute of Allergy and Infectious Diseases. This project has been funded in whole or in part with federal funds from the National Cancer Institute, National Institutes of Health, under contract N01-CO-12400.
The content of this publication does not necessary reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. government.
Published ahead of print on 27 February 2008.