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Journal of Virology, May 2008, p. 4612-4619, Vol. 82, No. 9
0022-538X/08/$08.00+0     doi:10.1128/JVI.02233-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Hydrophobic Inactivation of Influenza Viruses Confers Preservation of Viral Structure with Enhanced Immunogenicity

Yossef Raviv,^1,2* Robert Blumenthal,¹ S. Mark Tompkins,³ Jennifer Humberd,³ Robert J. Hogan,⁴ and Mathias Viard^1,2

Nanobiology Program, Center of Cancer Research, National Cancer Institute, National Institutes of Health,¹ Basic Research Program, SAIC-Frederick, Inc., National Cancer Institute, Frederick, Maryland,² Department of Infectious Diseases, University of Georgia, Athens, Georgia,³ Department of Anatomy and Radiology, University of Georgia, Athens, Georgia⁴

Received 15 October 2007/ Accepted 15 February 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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.

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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.

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Viruses, reagents, and cell lines. 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. 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.

Infectivity of influenza virus. 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. 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.

Measurement of viral fusion by photosensitized labeling. 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).

Measurement of lipid mixing by fluorescence dequenching. 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 x [(F – F₀)/(F_t – F₀)], where F₀ and F are the fluorescence intensities at time zero and at a given time point, respectively, and F_t 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).

FRAP acquisition and analysis. 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 40x, 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.

Murine immunizations and challenges. 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.

ELISAs. 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 x 10⁵ PFU/ml) or A/PR/8/34 (4 x 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.

Serological analysis by HI assay. 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.

Enzyme-linked immunospot (ELISPOT) assays. 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-) 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- 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- spot counts were obtained using an AID ELISpot plate reader (Autoimmun Diagnostika GmbH, Strassberg, Germany).

Statistical analysis. 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).

Electron microscopy. 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.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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. 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. 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. 2 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 3 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.

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FIG. 1. Effect of INA on influenza virus. Band-purified H3N2 virus was treated with INA at the indicated concentrations. Infectivity and neuraminidase activity were measured as described in Materials and Methods and are presented as a percentage of the value observed with untreated virus. Squares, infectivity; diamonds, neuraminidase activity. Data points represent means and standard deviations from triplicate experiments.

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FIG. 2. INA has no effect on HA-induced lipid mixing. R-18-labeled influenza virus was bound to erythrocytes, and fusion was triggered by lowering the pH of the buffer as described in Materials and Methods. Results represent the redistribution of viral envelope lipids into the target red blood cell membranes as measured by R-18 dequenching. Dequenching was measured by the increase in fluorescence of R-18 observed with time after lowering the pH to 5.0. The data are presented as the percentage of the maximal fluorescence observed upon lysis of the viral membrane in 1% Triton X-100. A. Control, untreated virus. B. Virus treated with 100 µM INA. Blue, preexposure at pH 7.4; green, dequenching induced by a virus that was preexposed to pH. 5.0 at 4°C and incubated with cells at pH 7.4; red, as for the green line, except that preexposure to low pH was done at 37°C.

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FIG. 3. INA blocks the fusion-driven insertion and redistribution of viral envelope proteins into the target cell membrane. Insertion of viral envelope proteins was measured by photosensitized labeling to follow the extent of 125INA incorporation into influenza virus HA after the triggering of fusion at pH 5.0. The experiment was repeated for every indicated concentration of INA. Values represent radioactivity of HA as measured by phosphorimager-enhanced autoradiography. The insert shows the actual labeling of HA as observed with the phosphorimager. The pH 7.4 lane shows the measurement obtained 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. 4 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. 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.

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FIG. 4. INA treatment blocks the mobility of proteins but not of lipids in cell membranes. Translational diffusion of lipids was measured by FRAP as described in Materials and Methods after treatment with the indicated INA concentrations. The values presented are the mobile fraction. The fluorescent lipid probe was DiO, and the protein probe was GFP-CD4. White bars, protein; black bars, lipid; UV, control cells irradiated with UV in the absence of INA.

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FIG. 5. Visualization of influenza viruses by electron microscopy. INA-treated and control viruses were fixed and subjected to negative staining. Three different imaging fields are presented for each experimental group.

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 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. 6A and Table 1).

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TABLE 1. Influenza virus (X-31)-specific antibody titers

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FIG. 6. Immunization with INA-inactivated X-31 induces potent mucosal antibodies and cellular immune responses. (A) On day 28 after a single administration of PBS or influenza B/AA, live influenza X-31, or INA-treated X-31 virus, lung washes were performed using 1 ml of PBS per mouse. BAL fluid was collected individually, and pooled samples from each group were then tested by standard ELISA using plates coated with UV-inactivated influenza X-31 virus. Influenza virus-specific IgA levels in the lungs were determined using anti-mouse IgA-specific secondary antibodies. The optical density (OD) represents the average optical density for each group at 405 nM. (B) Spleens were aseptically removed from the mice represented in panel A, and single-cell suspensions of splenocytes were prepared from pooled spleens and used in standard ELISPOT assays with a twofold dilution scheme. After 48 h in culture with 1 µg/ml of either NP147-155 (NP) or HA533-541 (HA), cells were removed by washing and the plates were developed. Values from control wells containing no peptide (background) were subtracted, and numbers were adjusted to reflect cytokine spots per 1 x 106 cells. Spots were enumerated using an AID ELISPOT plate reader.

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--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. 6B).

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 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.

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TABLE 2. Influenza virus-specific HI titers

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. 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. 7B).

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FIG. 7. Immunization with INA-inactivated X-31 induces protection against heterosubtypic challenge, similar to live X-31 infection. (A) Mice immunized as for Fig. 6 were challenged 28 days later with 10 LD50 of PR/8. Mice were monitored daily for weight loss (A) and survival (B) until all animals had succumbed to infection or recovered most of their original weight. Survival after heterosubtypic virus challenge in INA X-31- and live X-31-immunized animals was significantly different from B/AA- and PBS-immunized controls (P < 0.001, log rank).

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
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. 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. 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. 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. 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. 2 and 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. 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. 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 1 and 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.

 
* Corresponding author. Mailing address: Building 469, Room 215, NCI-Frederick, Frederick, MD 21702. Phone: (301) 846-6522. Fax: (301) 846-6210. E-mail: yraviv{at}ncifcrf.gov

Published ahead of print on 27 February 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Armstrong, R. T., A. S. Kushnir, and J. M. White. 2000. The transmembrane domain of influenza hemagglutinin exhibits a stringent length requirement to support the hemifusion to fusion transition. J. Cell Biol. 151:425-437.[Abstract/Free Full Text]
2
2. Bardiya, N., and J. H. Bae. 2005. Influenza vaccines: recent advances in production technologies. Appl. Microbiol. Biotechnol. 67:299-305.[CrossRef][Medline]
3
3. Blumenthal, R., M. J. Clague, S. R. Durell, and R. M. Epand. 2003. Membrane fusion. Chem. Rev. 103:53-69.[CrossRef][Medline]
4
4. Bravo, A., I. Gomez, J. Conde, C. Munoz-Garay, J. Sanchez, R. Miranda, M. Zhuang, S. S. Gill, and M. Soberon. 2004. Oligomerization triggers binding of a Bacillus thuringiensis Cry1Ab pore-forming toxin to aminopeptidase N receptor leading to insertion into membrane microdomains. Biochim. Biophys. Acta 1667:38-46.[Medline]
5
5. Budowsky, E. I., S. E. Bresler, E. A. Friedman, and N. V. Zheleznova. 1981. Principles of selective inactivation of viral genome. I. UV-induced inactivation of influenza virus. Arch. Virol. 68:239-247.[CrossRef][Medline]
6
6. Budowsky, E. I., A. Smirnov Yu, and S. F. Shenderovich. 1993. Principles of selective inactivation of viral genome. VIII. The influence of beta-propiolactone on immunogenic and protective activities of influenza virus. Vaccine 11:343-348.[CrossRef][Medline]
7
7. Bullough, P. A., F. M. Hughson, J. J. Skehel, and D. C. Wiley. 1994. Structure of influenza haemagglutinin at the pH of membrane fusion. Nature 371:37-43.[CrossRef][Medline]
8
8. Clague, M. J., J. R. Knutson, R. Blumenthal, and A. Herrmann. 1991. Interaction of influenza hemagglutinin amino-terminal peptide with phospholipid vesicles: a fluorescence study. Biochemistry 30:5491-5497.[CrossRef][Medline]
9
9. De Flora, S., and G. Badolati. 1973. Inactivation of A2-Hong Kong influenza virus by heat and by freeze-thawing. Comparison of untreated and gamma-irradiated preparations. Boll. Ist. Sieroter. Milan. 52:293-305.[Medline]
10
10. De Flora, S., and G. Badolati. 1973. Thermal inactivation of untreated and gamma-irradiated A2-Aichi-2-68 influenza virus. J. Gen. Virol. 20:261-265.[Abstract/Free Full Text]
11
11. Earp, L. J., S. E. Delos, H. E. Park, and J. M. White. 2005. The many mechanisms of viral membrane fusion proteins. Curr. Top. Microbiol. Immunol. 285:25-66.[Medline]
12
12. Epand, R. F., C. M. Yip, L. V. Chernomordik, D. L. LeDuc, Y. K. Shin, and R. M. Epand. 2001. Self-assembly of influenza hemagglutinin: studies of ectodomain aggregation by in situ atomic force microscopy. Biochim. Biophys. Acta 1513:167-175.[Medline]
13
13. Epstein, S. L. 2003. Control of influenza virus infection by immunity to conserved viral features. Expert Rev. Anti Infect. Ther. 1:627-638.[CrossRef][Medline]
14
14. Epstein, S. L., W. P. Kong, J. A. Misplon, C. Y. Lo, T. M. Tumpey, L. Xu, and G. J. Nabel. 2005. Protection against multiple influenza A subtypes by vaccination with highly conserved nucleoprotein. Vaccine 23:5404-5410.[CrossRef][Medline]
15
15. Epstein, S. L., T. M. Tumpey, J. A. Misplon, C. Y. Lo, L. A. Cooper, K. Subbarao, M. Renshaw, S. Sambhara, and J. M. Katz. 2002. DNA vaccine expressing conserved influenza virus proteins protective against H5N1 challenge infection in mice. Emerg. Infect. Dis. 8:796-801.[Medline]
16
16. Finnegan, M. C., S. S. Satinder, E. H. Cho, D. L. Guiffre, S. Lockett, A. H. Merill, Jr., and R. Blumenthal. 2007. Sphingomyelinase restricts the lateral diffusion of CD4 and inhibits human immunodeficiency virus fusion. J. Biol. Chem. 81:5294-5304.
17
17. Fiore, A. E., D. K. Shay, P. Haber, J. K. Iskander, T. M. Uyeki, G. Mootrey, J. S. Bresee, and N. J. Cox. 2007. Prevention and control of influenza. Recommendations of the Advisory Committee on Immunization Practices (ACIP), 2007. MMWR Recommend. Rep. 56:1-54.
18
18. Haan, L., W. R. Verweij, M. Holtrop, R. Brands, G. J. van Scharrenburg, A. M. Palache, E. Agsteribbe, and J. Wilschut. 2001. Nasal or intramuscular immunization of mice with influenza subunit antigen and the B subunit of Escherichia coli heat-labile toxin induces IgA- or IgG-mediated protective mucosal immunity. Vaccine 19:2898-2907.[CrossRef][Medline]
19
19. Hovden, A. O., R. J. Cox, and L. R. Haaheim. 2005. Whole influenza virus vaccine is more immunogenic than split influenza virus vaccine and induces primarily an IgG2a response in BALB/c mice. Scand. J. Immunol. 62:36-44.[Medline]
20
20. Hovden, A. O., R. J. Cox, A. Madhun, and L. R. Haaheim. 2005. Two doses of parenterally administered split influenza virus vaccine elicited high serum IgG concentrations which effectively limited viral shedding upon challenge in mice. Scand. J. Immunol. 62:342-352.[CrossRef][Medline]
21
21. Kemble, G. W., T. Danieli, and J. M. White. 1994. Lipid-anchored influenza hemagglutinin promotes hemifusion, not complete fusion. Cell 76:383-391.[CrossRef][Medline]
22
22. Korte, T., K. Ludwig, F. P. Booy, R. Blumenthal, and A. Herrmann. 1999. Conformational intermediates and fusion activity of influenza virus hemagglutinin. J. Virol. 73:4567-4574.[Abstract/Free Full Text]
23
23. Laver, W. G. 1978. Crystallization and peptide maps of neuraminidase "heads" from H2N2 and H3N2 influenza virus strains. Virology 86:78-87.[CrossRef][Medline]
24
24. Lippincott-Schwartz, J., E. Snapp, and A. Kenworthy. 2001. Studying protein dynamics in living cells. Nat. Rev. Mol. Cell Biol. 2:444-456.[CrossRef][Medline]
25
25. Lowy, R. J., G. A. Vavrina, and D. D. LaBarre. 2001. Comparison of gamma and neutron radiation inactivation of influenza A virus. Antivir. Res. 52:261-273.[CrossRef][Medline]
26
26. Melikyan, G. B., S. A. Brener, D. C. Ok, and F. S. Cohen. 1997. Inner but not outer membrane leaflets control the transition from glycosylphosphatidylinositol-anchored influenza hemagglutinin-induced hemifusion to full fusion. J. Cell Biol. 136:995-1005.[Abstract/Free Full Text]
27
27. Nir, S., F. Nicol, and F. C. Szoka, Jr. 1999. Surface aggregation and membrane penetration by peptides: relation to pore formation and fusion. Mol. Membr. Biol. 16:95-101.[CrossRef][Medline]
28
28. Noad, R., and P. Roy. 2003. Virus-like particles as immunogens. Trends Microbiol. 11:438-444.[CrossRef][Medline]
29
29. Pak, C. C., M. Krumbiegel, R. Blumenthal, and Y. Raviv. 1994. Detection of influenza hemagglutinin interaction with biological membranes by photosensitized activation of [¹²⁵I]iodonaphthylazide. J. Biol. Chem. 269:14614-14619.[Abstract/Free Full Text]
30
30. Popik, W., and T. M. Alce. 2004. CD4 receptor localized to non-raft membrane microdomains supports HIV-1 entry. Identification of a novel raft localization marker in CD4. J. Biol. Chem. 279:704-712.[Abstract/Free Full Text]
31
31. Puri, A., F. P. Booy, R. W. Doms, J. M. White, and R. Blumenthal. 1990. Conformational changes and fusion activity of influenza virus hemagglutinin of the H2 and H3 subtypes: effects of acid pretreatment. J. Virol. 64:3824-3832.[Abstract/Free Full Text]
32
32. Quan, F.-S., C. Huang, R. W. Compans, and S.-M. Kang. 2007. Virus-like particle vaccine induces protective immunity against homologous and heterologous strains of influenza virus. J. Virol. 81:3514-3524.[Abstract/Free Full Text]
33
33. Raviv, Y., M. Viard, J. Bess, Jr., and R. Blumenthal. 2002. Quantitative measurement of fusion of HIV-1 and SIV with cultured cells using photosensitized labeling. Virology 293:243-251.[CrossRef][Medline]
34
34. Raviv, Y., M. Viard, J. W. Bess, Jr., E. Chertova, and R. Blumenthal. 2005. Inactivation of retroviruses with preservation of structural integrity by targeting the hydrophobic domain of the viral envelope. J. Virol. 79:12394-12400.[Abstract/Free Full Text]
35
35. Redfield, D. C., D. D. Richman, M. N. Oxman, and L. H. Kronenberg. 1981. Psoralen inactivation of influenza and herpes simplex viruses and of virus-infected cells. Infect. Immun. 32:1216-1226.[Abstract/Free Full Text]
36
36. Rose, G. W., and C. L. Cooper. 2006. Fluarix, inactivated split-virus influenza vaccine. Expert Opin. Biol. Ther. 6:301-310.[CrossRef][Medline]
37
37. Stegmann, T., F. P. Booy, and J. Wilschut. 1987. Effects of low pH on influenza virus. Activation and inactivation of the membrane fusion capacity of the hemagglutinin. J. Biol. Chem. 262:17744-17749.[Abstract/Free Full Text]
38
38. Stephenson, I., M. C. Zambon, A. Rudin, A. Colegate, A. Podda, R. Bugarini, G. Del Giudice, A. Minutello, S. Bonnington, J. Holmgren, K. H. Mills, and K. G. Nicholson. 2006. Phase I evaluation of intranasal trivalent inactivated influenza vaccine with nontoxigenic Escherichia coli enterotoxin and novel biovector as mucosal adjuvants, using adult volunteers. J. Virol. 80:4962-4970.[Abstract/Free Full Text]
39
39. Tamura, S., T. Tanimoto, and T. Kurata. 2005. Mechanisms of broad cross-protection provided by influenza virus infection and their application to vaccines. Jpn. J. Infect. Dis. 58:195-207.[Medline]
40
40. Tumpey, T. M., M. Renshaw, J. D. Clements, and J. M. Katz. 2001. Mucosal delivery of inactivated influenza vaccine induces B-cell-dependent heterosubtypic cross-protection against lethal influenza A H5N1 virus infection. J. Virol. 75:5141-5150.[Abstract/Free Full Text]
41
41. Ulmer, J. B., J. J. Donnelly, S. E. Parker, G. H. Rhodes, P. L. Felgner, V. J. Dwarki, S. H. Gromkowski, R. R. Deck, C. M. DeWitt, A. Friedman, et al. 1993. Heterologous protection against influenza by injection of DNA encoding a viral protein. Science 259:1745-1749.[Abstract/Free Full Text]
42
42. Wood, J. M., and J. S. Robertson. 2004. From lethal virus to life-saving vaccine: developing inactivated vaccines for pandemic influenza. Nat. Rev. Microbiol. 2:842-847.[CrossRef][Medline]
43
43. Wright, P. F., G. Neumann, and Y. Kawaoka. 2007. Orthomyxoviruses, p. 1692-1740. In D. M. Knipe, P. M. Howley, D. E. Griffin, M. A. Martin, R. A. Lamb, B. Roizman, and S. E. Straus (ed.), Fields virology, 5th ed., vol. 2. Lippincott Williams & Wilkins, Philadelphia, PA.
44
44. Yao, Q., Z. Bu, A. Vzorov, C. Yang, and R. W. Compans. 2003. Virus-like particle and DNA-based candidate AIDS vaccines. Vaccine 21:638-643.[CrossRef][Medline]
45
45. Zemel, A., A. Ben-Shaul, and S. May. 2005. Perturbation of a lipid membrane by amphipathic peptides and its role in pore formation. Eur. Biophys. J. 34:230-242.[CrossRef][Medline]
46
46. Zheleznova, N. V. 1982. Method of influenza virus UV inactivation applied to the manufacture of an inactivated chromatographic influenza vaccine. Tr. Inst. Im. Pastera 59:81-88. (In Russian.)[Medline]

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Journal of Virology, May 2008, p. 4612-4619, Vol. 82, No. 9
0022-538X/08/$08.00+0     doi:10.1128/JVI.02233-07
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