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Onset and duration of immunity to equine influenza virus resulting from canarypox-vectored (ALVAC®) vaccination
Veterinary Immunology and Immunopathology 135 (2010) 100–107
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Veterinary Immunology and Immunopathology journal homepage: www.elsevier.com/locate/vetimm
Research paper
Onset and duration of immunity to equine influenza virus resulting from canarypox-vectored (ALVAC1) vaccination Gisela Soboll a, Stephen B. Hussey a, Jules M. Minke b, Gabriele A. Landolt a, James S. Hunter c, Shyla Jagannatha d, David P. Lunn a,* a
Department of Clinical Sciences, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, 300 West Drake Road, Fort Collins, CO 80523, USA Merial S.A.S., 254 rue Marcel Merieux, 69007 Lyon, France c Merial Limited, 6498 Jade Road, Fulton, MO 65251, USA d School of Veterinary Medicine, Iowa State University, Ames, IA 50010, USA b
A R T I C L E I N F O
A B S T R A C T
Article history: Received 3 September 2009 Received in revised form 10 November 2009 Accepted 14 November 2009
Equine influenza virus remains an important problem in horses despite extensive use of vaccination. Efficacy of equine influenza vaccination depends on the onset and duration of protective immunity, and appropriate strain specificity of the immune response. This study was designed to test the protective immunity resulting from vaccination with the North American commercial ALVAC1 equine influenza vaccine (RECOMBITEK1 Influenza, Merial, USA)1 against challenge with American lineage influenza viruses. In experiment 1, 12 ponies were vaccinated twice, at a 35 day interval, using the ALVAC1-influenza vaccine expressing the HA genes of influenza A/eq/Newmarket/2/93 and A/eq/Kentucky/94 (H3N8), and 11 ponies served as unvaccinated controls. Six months after the second vaccination, all ponies were challenged with A/eq/Kentucky/91. In experiment 2, 10 ponies received one dose of the ALVAC1-influenza vaccine, 10 ponies served as unvaccinated controls, and all ponies were challenge infected with A/equine/Ohio/03, 14 days after vaccination. Parameters studied included serological responses, and clinical disease and nasal viral shedding following challenge infection. In experiment 1, following the twodose regimen, vaccinated ponies generated high titered anti-influenza virus IgGa and IgGb antibody responses to vaccination and demonstrated statistically significant clinical and virological protection to challenge infection compared to controls. Infection with A/eq/ Kentucky/91 produced unusually severe signs in ponies in the control group, requiring therapy with NSAID’s and antibiotics, and leading to the euthanasia of one pony. In experiment 2 following the one-dose regimen, vaccinates generated IgGa responses prechallenge, and anamnestic IgGa and IgGb responses after challenge. Vaccinates demonstrated statistically significant clinical and virological protection to challenge infection compared to controls. The results of this study clearly demonstrate the early onset, and 6-month duration of protective immunity resulting from ALVAC1-influenza vaccination against challenge with American lineage equine influenza viruses. ß 2009 Elsevier B.V. All rights reserved.
Keywords: ALVAC1-influenza vaccination Horse Influenza virus
1 RECOMBITEK and PROTEQFLU are registered trademarks of Merial in the United States of America and elsewhere; ALVAC is a registered trademark of Connaught Technologies Corporation in the United States of America and elsewhere; SAS is a registered trademark of SAS Institute, Inc. in the United States of America and elsewhere; QIAAMP is a registered trademark of QIAGEN GmbH in the United States of America and elsewhere; MAXEFFICIENCY is a registered trademark of Life Technologies, Inc. in the United States of America; STATXACT is a registered trademark of Cytel, Inc. in the United States of America; STBLA is a trademark of Life Technologies, Inc. * Corresponding author. Tel.: +1 970 297 1274; fax: +1 970 297 1275. E-mail address: [email protected] (D.P. Lunn).
0165-2427/$ – see front matter ß 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.vetimm.2009.11.007
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1. Introduction Influenza virus is one of the primary causes of equine infectious respiratory disease in the horse throughout the world (Landolt, 2007) and is a common reason for medical treatment by equine practitioners in North America (Traub-Dargatz et al., 1991). The severity of the disease is critically dependent on the viral strain and the immune status of the affected animal. In young influenza-naı¨ve foals, infection can result in severe bronchopneumonia and death, as was recently reported in New South Wales, Australia, which was free of equine influenza virus prior to August 2007 (Patterson-Kane et al., 2008). In horses, the current worldwide circulating strain is equine A/2 virus, which is an H3N8 virus (Webster, 1993). This subtype has split into an American and a Eurasian lineage (Daly et al., 1996) which has prompted the World Organization for Animal Health (OIE) to recommend including representative viruses from both lineages in equine influenza virus vaccines (OIE, 2008). Despite their wide use, conventional inactivated vaccines can fail to induce a full spectrum of immune responses, and studies evaluating inactivated influenza virus vaccines have demonstrated the failure of these vaccines to induce long-term protection of horses from infection in the past (Morley et al., 1999; Nelson et al., 1998; Newton and Mumford, 1995; van Maanen et al., 2003). An intranasal modified-live vaccine for equine influenza virus shows more promising results (Lunn et al., 2001; Townsend et al., 2001), however, the use of modified-live vaccines can be problematic in immunocompromised animals (Oehen et al., 1991) and intranasal administration remains unpopular with some veterinarians and owners. DNA vaccination with the HA gene induces immune responses more similar to natural infection (Lunn et al., 1999; Soboll et al., 2003a, 2003b), and is considered safe in pregnant animals, but current techniques are not practical in the field and no vaccine has been licensed at this point. Recently the first vectored vaccine (RECOMBITEK1 Influenza, Merial, USA) based on the ALVAC1 platform (Poulet et al., 2007) has been licensed in North America for use in horses. The ALVAC1-influenza vaccine contains two canarypox constructs expressing the HA gene of influenza A/eq/Newmarket/2/93 (H3N8) and A/eq/Kentucky/94 (H3N8). The vaccine belongs to a group of secondgeneration influenza vaccines, which are thought likely to stimulate both humoral and cell-mediated immunity and have greater efficacy than conventional killed vaccines. It has previously been reported that ALVAC1 equine influenza vaccination using the European commercial product (PROTEQFLU1, Merial, France) can protect ponies from challenge infection with an Eurasian lineage influenza virus for up to 5 months after an initial two-dose primary vaccination and 12 months after a third booster vaccination (Minke et al., 2007). In addition, this vaccine showed early onset of clinical and virological protection to challenge infection with A/Eq/Newmarket/5/03 (an American lineage virus) after only one dose of the vaccine (Edlund Toulemonde et al., 2005). Lastly, there is evidence that this vaccine is able to prime foals in the presence of maternally derived immunity against influenza virus
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(Minke et al., 2007). The studies described in this paper advance these preceding references by addressing two questions about using RECOMBITEK1 Influenza, the ALVAC1 equine influenza virus vaccine formulation licensed and marketed in North America. The first goal of this study was to evaluate the duration of immunity 6months after a primary two-dose vaccination with ALVAC1-influenza to challenge protection using an American lineage virus (A/eq/Kentucky/91) in a severe challenge model. In a second experiment, the onset of immunity to challenge infection with a contemporary American lineage isolate (A/equine/Ohio/03) was tested. In both experiments we also sought to evaluate the utility of a novel real-time reverse transcriptase one-step quantitative PCR test for equine influenza virus. 2. Materials and methods 2.1. Experimental animals A total of 43 male 6-month-old influenza-naı¨ve ponies were used for both experiments. Ponies in each of two separate experiments were commingled and housed in an open-air facility, with access to shelter and feeding areas, but kept separate from all other horses. All ponies were fed twice a day with a diet of hay and pelleted concentrate. The maintenance and experimental protocols followed the animal care guidelines of the Animal Care and Use Committee, Colorado State University. 2.2. Equine influenza virus preparation and isolation For experiment 1 the influenza virus A/equine/Ky/1/91 was used. For experiment 2 A/equine/Ohio/03 was used. Both viruses were propagated and purified as described previously (Olsen et al., 1997). For virus isolation, viral titers were measured as EID50/ml of viral transport media (egg infectious dose 50%) (Palmer et al., 1975). 2.3. Experimental design and vaccine The RECOMBITEK1 Influenza vaccine (Merial, USA) contains 106.5 50% fluorescent antibody infectious dose (FAID50) of two live recombinant canarypox viruses (ALVAC1), one expressing the HA of A/eq/KY/94 (American lineage) and a second expressing the HA of A/eq/NM/2/93 (Eurasian lineage). The RECOMBITEK1-influenza vaccine was reconstituted with 2 ml of diluent containing Carbomer 974P immediately before administration and injected intramuscularly in the neck. Both of the following studies were designed as blinded, formally randomized, controlled challenge studies. Individuals performing vaccination were not involved in any further component of studies. Personnel performing challenge and post-challenge clinical observations and laboratory staff were blinded to group assignments. 2.3.1. Experiment 1 Ponies were randomly assigned to a vaccination group (n = 12) or a control group (n = 11) using SAS1 V8.2 software. During the vaccination phase, which occurred
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at Merial, Fulton, MO, vaccination group ponies were vaccinated on two occasions 35 days apart. Approximately 4 weeks prior to challenge, the vaccinates and control ponies were transported to Colorado State University, Ft. Collins, CO. Six months after the second vaccination all ponies were challenge infected with 108 EID50 of A/eq/ Kentucky/91 using a nebulizer and a face mask. 2.3.2. Experiment 2 Ponies were randomly assigned to a vaccination group (n = 10) or a control group (n = 10) and the entire experiment was conducted at Colorado State University. The vaccination group ponies were vaccinated once with the ALVAC1-influenza vaccine administered as described above, and all vaccinated and control ponies were challenge infected 14 days following vaccination and studied for 21 days. Challenge infection was performed with 1 108 EID50 of A/Equine/Ohio/03 influenza virus using a nebulizer and a face mask. 2.4. Sample and data collection Physical examinations were conducted throughout the experiment at all sample collection times and following each vaccination, and daily for 21 days after challenge infection. For evaluation of clinical disease a clinical score was established, which is described in Table 1. The clinical signs included coughing, nasal discharge, and dyspnoea. Ponies were also monitored for clinical signs consistent with secondary bacterial bronchopneumonia using the following diagnostic criteria: persistent (>5 days) pyrexia, severe mucopurulent nasal discharge, abnormal lung sounds, anorexia, or weight loss. If these criteria were met, affected animals were treated with antibiotics (procaine penicillin; 22,000 iu/kg, bid, im, for 5–8 days) and/or anti-inflammatory medication (flunixin meglumine; 1.0 mg/kg, iv, as indicated). If ponies were unresponsive to therapy and developed persistent and severe respiratory distress they were humanely euthanized. Serum for antibody assays was sampled at the times indicated in the figures. Serum was prepared from blood samples collected by jugular venipuncture. Serum samples were aliquoted and stored at 20 8C until analysis. Nasal swabs were collected for virus
Table 1 Clinical score. Clinical sign
Description
Score
Coughing
No cough Coughing once Coughing twice or more
0 1 2
Nasal discharge
No discharge Serous discharge Mucopurulent discharge Profuse mucopurulent discharge
0 1 2 3
Dyspnoea
No dyspnea (36 breaths/min) 0 Mild dyspnea (>36 breaths/min) 2 Severe dyspnea (>36 breaths/min 4 and additional signsa) a Extension of the head and neck, increased movement of the thoracic and abdominal wall plus dilation of the nostrils and abduction of the elbows.
isolation, daily for 21 days after challenge infection using Dacron swabs (Baxter Healthcare Corporation, McGaw Park, IL). Swabs were stored in 1 ml of virus transport medium at 80 8C until analysis. 2.5. Antibody immunoassays Serum was assayed for equine influenza virus-specific antibody using an ELISA procedure that has been extensively described (Lunn et al., 1999; Nelson et al., 1998). This procedure allows for identification of influenza virus-specific isotype and sub-isotype antibody responses using monoclonal antibodies to equine IgGa, IgGb, and IgG(T) (Lunn et al., 1998). Results are expressed as titers that are determined by comparison of the test sample to a standard curve generated by serial dilution of a sample of known titer. 2.6. Detection of viral shedding Viral RNA in nasal swab samples was isolated using the QIAAMP1 Viral RNA Mini Kit (Qiagen, Inc., Valencia, CA) and protocol. To determine nasal viral shedding, viral RNA load was determined by a one-step real-time reverse transcriptase-PCR assay that used specific primers and a specific probe recognizing glycoprotein M of equine influenza virus with the Eppendorf Mastercycler RealplexTM (Eppendorf) as previously described (Spindel et al., 2007). Briefly, real-time PCR primers and probe were designed to recognize conserved sequences in the influenza A virus M gene of equine influenza viruses, including equine and canine H3N8 viruses that have been isolated from horses and dogs in North America since 1963. Primers and probe were selected to detect a 144 bp product from the M gene between nucleotides 130 and 274 using the Beacon Designer software (Premier Biosoft Inc.). Primer and probe sequences were as follows: Forward primer (50 GAA CAC CGA TCT TGA GGC ACT C 30 ), Reverse primer (50 GGC ATT TTG GAC AAA GCG TCT AC-30 ). 50 reporter dye (6carboxyfluorescein [FAM]) labeled probe (50 -AGT CCT CGC TCA CTG GGC ACG GT-30 ). For calibration of equine influenza virus copy number, a purified full length M gene RNA was used as a standard. The M gene RNA standard was in vitro transcribed from the corresponding full length M gene template of A/Eq/WI/1/03 cloned into a plasmid vector (pGEM1-T Easy vector, Promega, Madison, WI). The directionality of the insert was confirmed by direct cycle sequencing using T7 primers. RNA was transcribed with the RiboMAX1 Large Scale RNA production systems (Promega, Madison, WI) kit from the T7 promoter according to the manufacturer’s instructions. The RNA generated was treated with RNase-free DNase I, tested for purity by both gel electrophoresis and PCR testing prior to use in real-time RT-PCR. The purified RNA was suspended in RNase-free water, quantified by spectrophotometer and stored at 80 8C in 10 mL aliquots at a concentration of 0.12 ng/mL. To determine the minimum detection level for real-time RT-PCR, the in vitro transcribed RNA was serially diluted in RNase-free water to produce dilutions ranging from 107 to 100 genomic equivalents/mL of viral M gene RNA. To evaluate inter-
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assay variation, the Ct values of 10 RNA standard curves, run on different days were determined. The mean, standard error of the mean (SEM) and coefficient of variation were calculated. This plasmid standard was included in each run as an external standard for the realtime reverse transcriptase PCR assay for quantification of equine influenza in nasal swab samples. Because the realtime assay detects viral RNA and not infectious virus, a cutoff 103 log copy numbers was established based on comparisons of results obtained using conventional methods (data not shown). 2.7. Statistical analysis For experiment 1, statistical analyses were performed using SAS V8.2, STATXACT1 V7.0 and MS Excel 2000. Differences were declared significant at p-value of 0.05 or less using a two-sided test. Pony 556 was euthanized on study day 225; data collected prior to euthanasia are included in statistical analyses. A post-challenge clinical disease score is presented in Table 1. The clinical score was calculated by adding the individual scores for coughing, nasal discharge, and dyspnea to produce a total score for each animal on each day (maximal possible score per day was 9) and severity of clinical disease for each day was defined based on the clinical score for that day. The difference between controls and vaccinates for incidence of severe clinical disease (clinical score > 4) were compared using Fisher’s exact test. The differences in the duration of individual clinical signs between controls and vaccinates were compared using Wilcoxon rank-sum test. For experiment 2, data was analyzed with generalized estimating equation approach using PROC GENMOD in SAS v9.2 (SAS Institute Inc., Cary, NC). Differences were considered significant at a p-value of 0.05 or less. Because the clinical data was not normally distributed, clinical scores and temperature data were dichotomized prior to analysis. Nasal viral shedding data and antibody data was log transformed prior to analysis to meet the assumption of normality. 3. Results 3.1. Experiment 1 3.1.1. Clinical signs following influenza challenge infection Median temperatures, clinical scores and body weights are shown in Fig. 1. There is no data shown in the weight difference graph (Fig. 1c) from days 5 to 9 because the scale was broken for that period. Vaccinated ponies exhibited significantly lower temperatures (Fig. 1a, p-value < 0.05), significantly shorter duration of clinical disease (pvalue < 0.0066) and significantly less weight loss (Fig. 1c, p-value < 0.001) following challenge infection when compared to unvaccinated controls. In addition, there was a significant increase in the incidence of severe disease (clinical score > 4; p-value < 0.0001) in controls (Fig. 1b). In the control group 7 of 10 ponies (#556, #568, #554, #562, #567, #571, #555) exhibited bronchopneumonia that required medical intervention; in the form of antibiotics and anti-inflammatory medication in 6 ponies, and anti-
Fig. 1. Experiment 1 (two-dose vaccination, 6-month duration of immunity): clinical disease post-challenge infection (CH) in each experimental group of ponies. Controls (n = 11): &, vaccinates (n = 12): ^. (a) Median temperature, (b) median clinical score, and (c) median weight difference (kg) from day 1.
inflammatory medication alone in an additional 2 ponies. Therapeutic interventions were needed between days 5 and 14 following infection. One of the control ponies did not respond to the treatment and was humanely euthanized on day 9 post-infection (#556) because of worsening respiratory disease and dyspnoea. On necropsy this pony exhibited severe, diffuse, subacute, fibrinonecrotic bronchointerstitial pneumonia. In contrast, none of the vaccinated animals required medical treatment post-challenge infection. 3.1.2. Virus shedding Nasal shedding was determined by real-time PCR measurement of influenza virus M-gene copies and mean log copy numbers are shown in Fig. 2. All animals were negative for viral shedding prior to challenge infection. All animals tested positive for 1 or more days post-challenge. The duration of viral shedding calculated as the first
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ponies showed significant increases in IgGa and IgGb levels following the second vaccination, while unvaccinated controls did not show IgGa or IgGb responses prior to challenge infection. IgGa levels declined to baseline 2 months after the second vaccination and IgGb levels declined to base line levels 4–5 months after the second vaccination. Following challenge infection IgGa and IgGb increased in both groups but levels were significantly higher in the vaccinates when compared to the control animals (p < 0.05). Fig. 2. Experiment 1 (two-dose vaccination, 6-month duration of immunity): median log M gene copy numbers in nasal swabs as determined by real-time PCR. Controls are shown by black bars, vaccinates are shown in white bars.
positive post-challenge day until the last positive postchallenge day was not significantly different from each other in the two treatment groups (p-value = 0.18) using a t-test. However, the amount of daily post-challenge viral shedding was significantly higher in the controls as compared to vaccinates on days 2–7 and 9 post-challenge infection (all p-values < 0.05).
3.2. Experiment 2 3.2.1. Clinical signs following influenza challenge infection Median temperatures, clinical scores and body weights are shown in Fig. 4. The number of days on which ponies
3.1.3. Antibody responses Median IgGa responses are shown in Fig. 3a, and median IgGb responses are shown in Fig. 3b. Vaccinated
Fig. 3. Experiment 1 (two-dose vaccination, 6-month duration of immunity): median IgGa and IgGb antibody titers for each experimental group. Controls (n = 11): &, vaccinates (n = 12): ^. Vaccinations (V) occurred on days 0 and 35 and all ponies were challenge infected (CH) on day 217. (a) IgGa and (b) IgGb.
Fig. 4. Experiment 2 (one-dose vaccination, onset of immunity): clinical disease post-challenge infection (CH) 14 days after a one-dose vaccination regimen. Controls (n = 10): &, vaccinates (n = 10): ^. (a) Median temperature, (b) median clinical score, and (c) median weight difference (kg) from day 1.
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exhibited increased temperatures was significantly lower (Fig. 1a, p-value < 0.0004) in vaccinates than in control ponies. In addition, vaccinates showed significantly less disease (Fig. 1b, p-value < 0.0001) and lost less weight (Fig. 1c) following challenge infection when compared to unvaccinated controls, although these weight loss differences between treatment groups were not statistically significant (p-value = 0.1). Starting at day 11 ponies in both groups gained weight, which is to be expected in young growing animals after recovery from the infection. No therapeutic interventions were needed in either experimental group, however one of the control ponies was humanely euthanized on day 11 post-infection due to a traumatic accident unrelated to the challenge. Upon necropsy this pony exhibited a severe, diffuse, subacute, suppurative pneumonia probably caused by bacteria secondary to viral infection. 3.2.2. Virus shedding Nasal shedding was determined by real-time PCR measurement of influenza virus M-gene copies and is shown as log copy numbers in Fig. 5. All animals were negative for viral shedding prior to challenge infection. All animals tested positive for one or more days postchallenge infection. The amount of daily post-challenge viral shedding of control ponies was significantly higher when compared to that of vaccinated ponies from days 2 to 9 post-challenge infection (p-value < 0.0001). 3.2.3. Antibody responses Median IgGa and IgGb responses are shown in Fig. 6. Vaccinated ponies showed a significant increase in IgGa titer following vaccination on day 13 post-vaccination – the day prior to challenge (p-value < 0.0001), while no increases in IgGb levels were seen in the vaccinates until after challenge infection. Antibody titers following challenge infection for both IgGa and IgGb were significantly higher in the vaccinates then in the unvaccinated controls indicating that these animals were primed by the vaccine (IgGa p-values 1 week (day 21) and 2 weeks (day 28) after challenge infection: <0.0001, IgGb p-values: day 21 = 0.0014, day 28 <0.0001). Unvaccinated controls did not show IgG responses prior to challenge infection and IgGb levels did not rise in the unvaccinated controls until 14 days following challenge infection (day 28), while IgGa
Fig. 5. Experiment 2 (one-dose vaccination, onset of immunity): median log M gene copy numbers in nasal swabs as determined by real-time PCR. Swabs were collected for 14 days post-challenge infection after a onedose vaccination regimen. Controls are shown by black bars, vaccinates are shown in white bars.
Fig. 6. Experiment 2 (one-dose vaccination, onset of immunity): median IgG antibody titers for each experimental group. Controls (n = 11): &, vaccinates (n = 12): ^. The vaccination (V) occurred on day 0 and all ponies were challenge infected (CH) on day 14. (a) IgGa and (b) IgGb.
levels were elevated by 7 days post-challenge infection (day 21). 4. Discussion This study confirms the observations of previous studies of ALVAC1-influenza vaccine technology (Edlund Toulemonde et al., 2005; Minke et al., 2007; Paillot et al., 2006), and advances them in a number of important ways. The work described here is the first study conducted with the commercial brand marketed in North America, and the first to demonstrate long-term (6-month) protective immunity resulting from this type of vaccine against an American lineage challenge strain. Furthermore, the onset of immunity study is the first report of a challengeprotection study using the contemporary North American isolate (A/equine/Ohio/03) which is one of the strains currently recommended by the OIE for inclusion in equine influenza vaccines (OIE, 2008). Finally this is the first report of influenza virus-specific equine IgG sub-class responses to an ALVAC1-influenza vaccine, and demonstrates that they are characterized by IgGa and IgGb responses, which are typically associated with protective immunity (Nelson et al., 1998). The principal finding of this report is the protection against a severe American lineage influenza strain challenge infection 6 months after completion of a primary series of two vaccinations. This observation provides significant evidence for the duration of protection of
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ALVAC1-influenza from the principle equine influenza virus strain in circulation worldwide today. The challenge model used was severe and was chosen because of its extensive past characterization (Lunn et al., 2001; Townsend et al., 2001). In this instance, the use of a challenge dose of 108 EID50, perhaps combined with conducting the challenge in Colorado at an altitude of 1520 m above sea level in the heat of summer, led to particularly severe disease in the control group. The severity of disease in controls required extensive medical intervention and the euthanasia of one animal. In the pony that had to be euthanized on day 9 post-infection, there were findings of severe, diffuse, subacute, fibrinonecrotic bronchointerstitial pneumonia on necropsy. These pathological findings are similar to reports from the first Australian equine influenza outbreak in 2007, where a number of foals born to influenza-naı¨ve mothers died of bronchointerstitial pneumonia, a disease that is usually rare in this age group (Patterson-Kane et al., 2008). In contrast, the minimal signs of disease in the vaccinated ponies further support the value of the vaccination regime. A significant reduction in viral shedding on 8 of the 10 days after challenge provided evidence for virological protection, however, there was no difference in the duration of shedding. This may indicate that 6 months after vaccination, equids may be able to propagate an infection, even if its clinical effects are limited. These findings are highly consistent with those reported previously by Minke et al. (2007) after a Eurasian viral challenge with the European brand of this vaccine. We chose to investigate the onset of immunity to a single ALVAC1-influenza vaccination of naı¨ve animals by challenge infection with a contemporary American lineage equine influenza virus (A/equine/Ohio/03). The results demonstrated good evidence of clinical and virological protection, similar to a previously reported but much smaller study of onset of immunity to the European brand (Edlund Toulemonde et al., 2005). However, in the study reported here, all vaccinated ponies shed virus on 2 or more days, whereas in the previous study vaccinates did not shed virus. This may be the result of differences in the challenge strain, route of infection or the detection methodology, although the former seems more likely. Both onset and duration of immunity after ALVAC1influenza vaccination were associated with induction of IgGa and IgGb antibody responses. These IgG sub-classes have previously been reported to be associated with protection from challenge infection in the absence of a nasal IgA response, while IgG(T) has not shown to provide protection (Nelson et al., 1998; Soboll et al., 2003a, 2003b). These responses are very likely responsible for the single radial hemolysis reactions previously described after use of ALVAC1 equine influenza virus vaccination (Edlund Toulemonde et al., 2005; Minke et al., 2007; Paillot et al., 2006). Interferon-g responses were not evaluated in this study, but a recent study by Paillot et al. (2006) has shown that two doses of the ALVAC1-influenza vaccine did prime horses to show significantly increased IFN-g responses to in vitro re-stimulation in PBMCs collected 7 and 14 days post-challenge infection. Together these results are consistent with induction of a type 1 immune response. These immunological outcomes are similar to vaccine responses
following DNA vaccination of horses with the hemagglutinin gene (Soboll et al., 2003a), but offer a more practical alternative to particle-mediated vaccine delivery. In addition to reduction of clinical disease, vaccination in our study significantly reduced the amount of viral shedding. For detection of viral nasal shedding, we employed a novel one-step real-time PCR technique that quantified viral shedding using probes and primers recognizing conserved sequences in the influenza A virus M gene of equine influenza viruses (Spindel et al., 2007). For quantification purposes this assay uses a plasmid containing the full length M gene and purified full length M gene RNA as standards. The assay system proved to be economical, fast and sensitive, with significant advantages over determination of viral titers in eggs or tissue culture systems. One disadvantage of this system is that it does not directly measure the infectious load in a sample. Nevertheless, the system has many advantages, and will be easier to establish in laboratories, particularly when egg incubators are not available. This is the first challenge infection study for equine influenza virus to report the use of a real-time PCR assay. Using this assay system we were able to detect differences in the amount of virus shed between treatment groups, which are an important factor for determination of vaccine efficacy. In summary, our study showed that the ALVAC1 equine influenza vaccine significantly reduced the clinical and virological sequelae of equine influenza virus infection in both onset and duration of immunity studies. This is the first study documenting the efficacy of this vaccine in a 6month duration of immunity study using an American lineage challenge strain. Acknowledgements We would like to thank Dr. Sangeta Rao and Dr. Ashley Hill for assistance with the statistical analysis of the second experiment in this study. In addition, Merial and sanofi aventis R&D are gratefully acknowledged for their valuable contribution. References Daly, J.M., Lai, A.C., Binns, M.M., Chambers, T.M., Barrandeguy, M., Mumford, J.A., 1996. Antigenic and genetic evolution of equine H3N8 Influenza A viruses. Journal of General Virology 77, 661–671. Edlund Toulemonde, C., Daly, J., Sindle, T., Guigal, P.M., Audonnet, J.C., Minke, J.M., 2005. Efficacy of a recombinant equine influenza vaccine against challenge with an American lineage H3N8 influenza virus responsible for the 2003 outbreak in the United Kingdom. Veterinary Record 156, 367–371. Landolt, G., 2007. Equine influenza infection. In: Sellon, D.C., Long, M.T. (Eds.), Equine Infectious Diseases. pp. 124–134. Lunn, D.P., Holmes, M.A., Antczak, D.F., Agerwal, N., Baker, J., BendaliAhcene, S., Blanchard-Channell, M., Byrne, K.M., Cannizzo, K., Davis, W., Hamilton, M.J., Hannant, D., Kondo, T., Kydd, J.H., Monier, M.C., Moore, P.F., O’Neil, T., Schram, B.R., Sheoran, A., Stott, J.L., Sugiura, T., Vagnoni, K.E., 1998. Report of the Second Equine Leucocyte Antigen Workshop, Squaw valley, California, July 1995. Veterinary Immunology & Immunopathology 62, 101–143. Lunn, D.P., Hussey, S., Sebring, R., Rushlow, K.E., Radecki, S.V., WhitakerDowling, P., Youngner, J.S., Chambers, T.M., Holland Jr., R.E., Horohov, D.W., 2001. Safety, efficacy, and immunogenicity of a modified-live equine influenza virus vaccine in ponies after induction of exerciseinduced immunosuppression. Journal of the American Veterinary Medical Association 218, 900–906.
G. Soboll et al. / Veterinary Immunology and Immunopathology 135 (2010) 100–107 Lunn, D.P, Soboll, G., Schram, B.R., Quass, J., McGregor, M.W., Drape, R.J., Macklin, M.D., McCabe, D.E., Swain, W.F., Olsen, C.W., 1999. Antibody responses to DNA vaccination of horses using the influenza virus hemagglutinin gene. Vaccine 17, 2245–2258. Minke, J.M., Toulemonde, C.E., Coupier, H., Guigal, P.-M., Dinic, S., Sindle, T., Jessett, D., Black, L., Bublot, M., Pardo, M.C., Audonnet, J.-C., 2007. Efficacy of a canarypox-vectored recombinant vaccine expressing the hemagglutinin gene of equine influenza H3N8 virus in the protection of ponies from viral challenge. American Journal of Veterinary Research 68, 213–219. Morley, P.S., Townsend, H.G., Bogdan, J.R., Haines, D.M., 1999. Efficacy of a commercial vaccine for preventing disease caused by influenza virus infection in horses. Journal of the American Veterinary Medical Association 215, 61–66. Nelson, K.M., Schram, B.R., McGregor, M.W., Olsen, C.W., Lunn, D.P., 1998. Local and systemic isotype-specific antibody responses to equine influenza virus infection versus conventional vaccination. Vaccine 16, 1306–1313. Newton, J.R., Mumford, J.A., 1995. Equine influenza in vaccinated horses. Veterinary Record 137 (19), 495–496. Oehen, S., Hengartner, H., Zinkernagel, R.M., 1991. Vaccination for disease. Science 251, 195–198. OIE Bulletin, 2008, Expert Surveillance Panel on Equine Influenza Vaccines – Conclusions and Recommendations. Olsen, C.W., McGregor, M.W., Dybdahl-Sissoko, N., Schram, B.R., Nelson, K.M., Lunn, D.P., Macklin, M.D., Swain, W.F., Hinshaw, V.S., 1997. Immunogenicity and efficacy of baculovirus-expressed and DNAbased equine influenza virus hemagglutinin vaccines in mice. Vaccine 15, 1149–1156. Paillot, R., Kydd, J.H., Sindle, T., Hannant, D., Edlund Toulemonde, C., Audonnet, J.C., Minke, J.M., Daly, J.M., 2006. Antibody and IFN-gamma responses induced by a recombinant canarypox vaccine and challenge infection with equine influenza virus. Veterinary Immunology & Immunopathology 112, 225–233. Palmer, D.F., Dowdle, W.R., Coleman, M.T., 1975. Advanced Laboratory Techniques for Influenza Diagnosis, 6th edition. United States
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Department of Health, Education and Welfare Immunology Series. Patterson-Kane, J.C., Carrick, J.B., Axon, J.E., Wilkie, I., Begg, A.P., 2008. The pathology of bronchointerstitial pneumonia in young foals associated with the first outbreak of equine influenza in Australia. Equine Veterinary Journal 40, 199–203. Poulet, H., Minke, J., Pardo, M.C., Juillard, V., Nordgren, B., Audonnet, J.C., 2007. Development and registration of recombinant veterinary vaccines. The example of the canarypox vector platform. Vaccine 25, 5606–5612. Soboll, G., Horohov, D.W., Aldridge, B.M., Olsen, C.W., McGregor, M.W., Drape, R.J., Macklin, M.D., Swain, W.F., Lunn, D.P., 2003a. Regional antibody and cellular immune responses to equine influenza virus infection, and particle mediated DNA vaccination. Veterinary Immunology & Immunopathology 94, 47–62. Soboll, G., Nelson, K.M., Leuthner, E.S., Clark, R.J., Drape, R., Macklin, M.D., Swain, W.F., Olsen, C.W., Lunn, D.P., 2003b. Mucosal co-administration of cholera toxin and influenza virus hemagglutinin-DNA in ponies generates a local IgA response. Vaccine 21, 3081–3092. Spindel, M., Dillion, S., Lunn, K., Landolt, G., 2007. Detection and quantification of canine influenza virus by one-step real-time reverse transcription PCR. Journal of Veterinary Internal Medicine 21, 576. Townsend, H.G., Penner, S.J., Watts, T.C., Cook, A., Bogdan, C., Haines, D.M., Griffin, G.E., Chambers, C.A., Holland, R.E., Whitaker-Dowling, P., Younger, J.S., Sebring, R.W., 2001. Efficacy of a cold adapted, intranasal, equine influenza vaccine: challenge trials. Equine Veterinary Journal 33, 637–643. Traub-Dargatz, J.L., Salman, M.D., Voss, J.L., 1991. Medical problems of adult horses, as ranked by equine practitioners. Journal of the American Veterinary Medical Association 198, 1745–1747. van Maanen, C, van Essen, G.J., Minke, J., Daly, J.M., Yates, P.J., 2003. Diagnostic methods applied to analysis of an outbreak of equine influenza in a riding school in which vaccine failure occurred. Veterinary Microbiology 93, 291–306. Webster, R.G., 1993. Are equine 1 influenza viruses still present in horses? Equine Veterinary Journal 25, 537–538.
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Research paper
Onset and duration of immunity to equine influenza virus resulting from canarypox-vectored (ALVAC1) vaccination Gisela Soboll a, Stephen B. Hussey a, Jules M. Minke b, Gabriele A. Landolt a, James S. Hunter c, Shyla Jagannatha d, David P. Lunn a,* a
Department of Clinical Sciences, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, 300 West Drake Road, Fort Collins, CO 80523, USA Merial S.A.S., 254 rue Marcel Merieux, 69007 Lyon, France c Merial Limited, 6498 Jade Road, Fulton, MO 65251, USA d School of Veterinary Medicine, Iowa State University, Ames, IA 50010, USA b
A R T I C L E I N F O
A B S T R A C T
Article history: Received 3 September 2009 Received in revised form 10 November 2009 Accepted 14 November 2009
Equine influenza virus remains an important problem in horses despite extensive use of vaccination. Efficacy of equine influenza vaccination depends on the onset and duration of protective immunity, and appropriate strain specificity of the immune response. This study was designed to test the protective immunity resulting from vaccination with the North American commercial ALVAC1 equine influenza vaccine (RECOMBITEK1 Influenza, Merial, USA)1 against challenge with American lineage influenza viruses. In experiment 1, 12 ponies were vaccinated twice, at a 35 day interval, using the ALVAC1-influenza vaccine expressing the HA genes of influenza A/eq/Newmarket/2/93 and A/eq/Kentucky/94 (H3N8), and 11 ponies served as unvaccinated controls. Six months after the second vaccination, all ponies were challenged with A/eq/Kentucky/91. In experiment 2, 10 ponies received one dose of the ALVAC1-influenza vaccine, 10 ponies served as unvaccinated controls, and all ponies were challenge infected with A/equine/Ohio/03, 14 days after vaccination. Parameters studied included serological responses, and clinical disease and nasal viral shedding following challenge infection. In experiment 1, following the twodose regimen, vaccinated ponies generated high titered anti-influenza virus IgGa and IgGb antibody responses to vaccination and demonstrated statistically significant clinical and virological protection to challenge infection compared to controls. Infection with A/eq/ Kentucky/91 produced unusually severe signs in ponies in the control group, requiring therapy with NSAID’s and antibiotics, and leading to the euthanasia of one pony. In experiment 2 following the one-dose regimen, vaccinates generated IgGa responses prechallenge, and anamnestic IgGa and IgGb responses after challenge. Vaccinates demonstrated statistically significant clinical and virological protection to challenge infection compared to controls. The results of this study clearly demonstrate the early onset, and 6-month duration of protective immunity resulting from ALVAC1-influenza vaccination against challenge with American lineage equine influenza viruses. ß 2009 Elsevier B.V. All rights reserved.
Keywords: ALVAC1-influenza vaccination Horse Influenza virus
1 RECOMBITEK and PROTEQFLU are registered trademarks of Merial in the United States of America and elsewhere; ALVAC is a registered trademark of Connaught Technologies Corporation in the United States of America and elsewhere; SAS is a registered trademark of SAS Institute, Inc. in the United States of America and elsewhere; QIAAMP is a registered trademark of QIAGEN GmbH in the United States of America and elsewhere; MAXEFFICIENCY is a registered trademark of Life Technologies, Inc. in the United States of America; STATXACT is a registered trademark of Cytel, Inc. in the United States of America; STBLA is a trademark of Life Technologies, Inc. * Corresponding author. Tel.: +1 970 297 1274; fax: +1 970 297 1275. E-mail address: [email protected] (D.P. Lunn).
0165-2427/$ – see front matter ß 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.vetimm.2009.11.007
G. Soboll et al. / Veterinary Immunology and Immunopathology 135 (2010) 100–107
1. Introduction Influenza virus is one of the primary causes of equine infectious respiratory disease in the horse throughout the world (Landolt, 2007) and is a common reason for medical treatment by equine practitioners in North America (Traub-Dargatz et al., 1991). The severity of the disease is critically dependent on the viral strain and the immune status of the affected animal. In young influenza-naı¨ve foals, infection can result in severe bronchopneumonia and death, as was recently reported in New South Wales, Australia, which was free of equine influenza virus prior to August 2007 (Patterson-Kane et al., 2008). In horses, the current worldwide circulating strain is equine A/2 virus, which is an H3N8 virus (Webster, 1993). This subtype has split into an American and a Eurasian lineage (Daly et al., 1996) which has prompted the World Organization for Animal Health (OIE) to recommend including representative viruses from both lineages in equine influenza virus vaccines (OIE, 2008). Despite their wide use, conventional inactivated vaccines can fail to induce a full spectrum of immune responses, and studies evaluating inactivated influenza virus vaccines have demonstrated the failure of these vaccines to induce long-term protection of horses from infection in the past (Morley et al., 1999; Nelson et al., 1998; Newton and Mumford, 1995; van Maanen et al., 2003). An intranasal modified-live vaccine for equine influenza virus shows more promising results (Lunn et al., 2001; Townsend et al., 2001), however, the use of modified-live vaccines can be problematic in immunocompromised animals (Oehen et al., 1991) and intranasal administration remains unpopular with some veterinarians and owners. DNA vaccination with the HA gene induces immune responses more similar to natural infection (Lunn et al., 1999; Soboll et al., 2003a, 2003b), and is considered safe in pregnant animals, but current techniques are not practical in the field and no vaccine has been licensed at this point. Recently the first vectored vaccine (RECOMBITEK1 Influenza, Merial, USA) based on the ALVAC1 platform (Poulet et al., 2007) has been licensed in North America for use in horses. The ALVAC1-influenza vaccine contains two canarypox constructs expressing the HA gene of influenza A/eq/Newmarket/2/93 (H3N8) and A/eq/Kentucky/94 (H3N8). The vaccine belongs to a group of secondgeneration influenza vaccines, which are thought likely to stimulate both humoral and cell-mediated immunity and have greater efficacy than conventional killed vaccines. It has previously been reported that ALVAC1 equine influenza vaccination using the European commercial product (PROTEQFLU1, Merial, France) can protect ponies from challenge infection with an Eurasian lineage influenza virus for up to 5 months after an initial two-dose primary vaccination and 12 months after a third booster vaccination (Minke et al., 2007). In addition, this vaccine showed early onset of clinical and virological protection to challenge infection with A/Eq/Newmarket/5/03 (an American lineage virus) after only one dose of the vaccine (Edlund Toulemonde et al., 2005). Lastly, there is evidence that this vaccine is able to prime foals in the presence of maternally derived immunity against influenza virus
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(Minke et al., 2007). The studies described in this paper advance these preceding references by addressing two questions about using RECOMBITEK1 Influenza, the ALVAC1 equine influenza virus vaccine formulation licensed and marketed in North America. The first goal of this study was to evaluate the duration of immunity 6months after a primary two-dose vaccination with ALVAC1-influenza to challenge protection using an American lineage virus (A/eq/Kentucky/91) in a severe challenge model. In a second experiment, the onset of immunity to challenge infection with a contemporary American lineage isolate (A/equine/Ohio/03) was tested. In both experiments we also sought to evaluate the utility of a novel real-time reverse transcriptase one-step quantitative PCR test for equine influenza virus. 2. Materials and methods 2.1. Experimental animals A total of 43 male 6-month-old influenza-naı¨ve ponies were used for both experiments. Ponies in each of two separate experiments were commingled and housed in an open-air facility, with access to shelter and feeding areas, but kept separate from all other horses. All ponies were fed twice a day with a diet of hay and pelleted concentrate. The maintenance and experimental protocols followed the animal care guidelines of the Animal Care and Use Committee, Colorado State University. 2.2. Equine influenza virus preparation and isolation For experiment 1 the influenza virus A/equine/Ky/1/91 was used. For experiment 2 A/equine/Ohio/03 was used. Both viruses were propagated and purified as described previously (Olsen et al., 1997). For virus isolation, viral titers were measured as EID50/ml of viral transport media (egg infectious dose 50%) (Palmer et al., 1975). 2.3. Experimental design and vaccine The RECOMBITEK1 Influenza vaccine (Merial, USA) contains 106.5 50% fluorescent antibody infectious dose (FAID50) of two live recombinant canarypox viruses (ALVAC1), one expressing the HA of A/eq/KY/94 (American lineage) and a second expressing the HA of A/eq/NM/2/93 (Eurasian lineage). The RECOMBITEK1-influenza vaccine was reconstituted with 2 ml of diluent containing Carbomer 974P immediately before administration and injected intramuscularly in the neck. Both of the following studies were designed as blinded, formally randomized, controlled challenge studies. Individuals performing vaccination were not involved in any further component of studies. Personnel performing challenge and post-challenge clinical observations and laboratory staff were blinded to group assignments. 2.3.1. Experiment 1 Ponies were randomly assigned to a vaccination group (n = 12) or a control group (n = 11) using SAS1 V8.2 software. During the vaccination phase, which occurred
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at Merial, Fulton, MO, vaccination group ponies were vaccinated on two occasions 35 days apart. Approximately 4 weeks prior to challenge, the vaccinates and control ponies were transported to Colorado State University, Ft. Collins, CO. Six months after the second vaccination all ponies were challenge infected with 108 EID50 of A/eq/ Kentucky/91 using a nebulizer and a face mask. 2.3.2. Experiment 2 Ponies were randomly assigned to a vaccination group (n = 10) or a control group (n = 10) and the entire experiment was conducted at Colorado State University. The vaccination group ponies were vaccinated once with the ALVAC1-influenza vaccine administered as described above, and all vaccinated and control ponies were challenge infected 14 days following vaccination and studied for 21 days. Challenge infection was performed with 1 108 EID50 of A/Equine/Ohio/03 influenza virus using a nebulizer and a face mask. 2.4. Sample and data collection Physical examinations were conducted throughout the experiment at all sample collection times and following each vaccination, and daily for 21 days after challenge infection. For evaluation of clinical disease a clinical score was established, which is described in Table 1. The clinical signs included coughing, nasal discharge, and dyspnoea. Ponies were also monitored for clinical signs consistent with secondary bacterial bronchopneumonia using the following diagnostic criteria: persistent (>5 days) pyrexia, severe mucopurulent nasal discharge, abnormal lung sounds, anorexia, or weight loss. If these criteria were met, affected animals were treated with antibiotics (procaine penicillin; 22,000 iu/kg, bid, im, for 5–8 days) and/or anti-inflammatory medication (flunixin meglumine; 1.0 mg/kg, iv, as indicated). If ponies were unresponsive to therapy and developed persistent and severe respiratory distress they were humanely euthanized. Serum for antibody assays was sampled at the times indicated in the figures. Serum was prepared from blood samples collected by jugular venipuncture. Serum samples were aliquoted and stored at 20 8C until analysis. Nasal swabs were collected for virus
Table 1 Clinical score. Clinical sign
Description
Score
Coughing
No cough Coughing once Coughing twice or more
0 1 2
Nasal discharge
No discharge Serous discharge Mucopurulent discharge Profuse mucopurulent discharge
0 1 2 3
Dyspnoea
No dyspnea (36 breaths/min) 0 Mild dyspnea (>36 breaths/min) 2 Severe dyspnea (>36 breaths/min 4 and additional signsa) a Extension of the head and neck, increased movement of the thoracic and abdominal wall plus dilation of the nostrils and abduction of the elbows.
isolation, daily for 21 days after challenge infection using Dacron swabs (Baxter Healthcare Corporation, McGaw Park, IL). Swabs were stored in 1 ml of virus transport medium at 80 8C until analysis. 2.5. Antibody immunoassays Serum was assayed for equine influenza virus-specific antibody using an ELISA procedure that has been extensively described (Lunn et al., 1999; Nelson et al., 1998). This procedure allows for identification of influenza virus-specific isotype and sub-isotype antibody responses using monoclonal antibodies to equine IgGa, IgGb, and IgG(T) (Lunn et al., 1998). Results are expressed as titers that are determined by comparison of the test sample to a standard curve generated by serial dilution of a sample of known titer. 2.6. Detection of viral shedding Viral RNA in nasal swab samples was isolated using the QIAAMP1 Viral RNA Mini Kit (Qiagen, Inc., Valencia, CA) and protocol. To determine nasal viral shedding, viral RNA load was determined by a one-step real-time reverse transcriptase-PCR assay that used specific primers and a specific probe recognizing glycoprotein M of equine influenza virus with the Eppendorf Mastercycler RealplexTM (Eppendorf) as previously described (Spindel et al., 2007). Briefly, real-time PCR primers and probe were designed to recognize conserved sequences in the influenza A virus M gene of equine influenza viruses, including equine and canine H3N8 viruses that have been isolated from horses and dogs in North America since 1963. Primers and probe were selected to detect a 144 bp product from the M gene between nucleotides 130 and 274 using the Beacon Designer software (Premier Biosoft Inc.). Primer and probe sequences were as follows: Forward primer (50 GAA CAC CGA TCT TGA GGC ACT C 30 ), Reverse primer (50 GGC ATT TTG GAC AAA GCG TCT AC-30 ). 50 reporter dye (6carboxyfluorescein [FAM]) labeled probe (50 -AGT CCT CGC TCA CTG GGC ACG GT-30 ). For calibration of equine influenza virus copy number, a purified full length M gene RNA was used as a standard. The M gene RNA standard was in vitro transcribed from the corresponding full length M gene template of A/Eq/WI/1/03 cloned into a plasmid vector (pGEM1-T Easy vector, Promega, Madison, WI). The directionality of the insert was confirmed by direct cycle sequencing using T7 primers. RNA was transcribed with the RiboMAX1 Large Scale RNA production systems (Promega, Madison, WI) kit from the T7 promoter according to the manufacturer’s instructions. The RNA generated was treated with RNase-free DNase I, tested for purity by both gel electrophoresis and PCR testing prior to use in real-time RT-PCR. The purified RNA was suspended in RNase-free water, quantified by spectrophotometer and stored at 80 8C in 10 mL aliquots at a concentration of 0.12 ng/mL. To determine the minimum detection level for real-time RT-PCR, the in vitro transcribed RNA was serially diluted in RNase-free water to produce dilutions ranging from 107 to 100 genomic equivalents/mL of viral M gene RNA. To evaluate inter-
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assay variation, the Ct values of 10 RNA standard curves, run on different days were determined. The mean, standard error of the mean (SEM) and coefficient of variation were calculated. This plasmid standard was included in each run as an external standard for the realtime reverse transcriptase PCR assay for quantification of equine influenza in nasal swab samples. Because the realtime assay detects viral RNA and not infectious virus, a cutoff 103 log copy numbers was established based on comparisons of results obtained using conventional methods (data not shown). 2.7. Statistical analysis For experiment 1, statistical analyses were performed using SAS V8.2, STATXACT1 V7.0 and MS Excel 2000. Differences were declared significant at p-value of 0.05 or less using a two-sided test. Pony 556 was euthanized on study day 225; data collected prior to euthanasia are included in statistical analyses. A post-challenge clinical disease score is presented in Table 1. The clinical score was calculated by adding the individual scores for coughing, nasal discharge, and dyspnea to produce a total score for each animal on each day (maximal possible score per day was 9) and severity of clinical disease for each day was defined based on the clinical score for that day. The difference between controls and vaccinates for incidence of severe clinical disease (clinical score > 4) were compared using Fisher’s exact test. The differences in the duration of individual clinical signs between controls and vaccinates were compared using Wilcoxon rank-sum test. For experiment 2, data was analyzed with generalized estimating equation approach using PROC GENMOD in SAS v9.2 (SAS Institute Inc., Cary, NC). Differences were considered significant at a p-value of 0.05 or less. Because the clinical data was not normally distributed, clinical scores and temperature data were dichotomized prior to analysis. Nasal viral shedding data and antibody data was log transformed prior to analysis to meet the assumption of normality. 3. Results 3.1. Experiment 1 3.1.1. Clinical signs following influenza challenge infection Median temperatures, clinical scores and body weights are shown in Fig. 1. There is no data shown in the weight difference graph (Fig. 1c) from days 5 to 9 because the scale was broken for that period. Vaccinated ponies exhibited significantly lower temperatures (Fig. 1a, p-value < 0.05), significantly shorter duration of clinical disease (pvalue < 0.0066) and significantly less weight loss (Fig. 1c, p-value < 0.001) following challenge infection when compared to unvaccinated controls. In addition, there was a significant increase in the incidence of severe disease (clinical score > 4; p-value < 0.0001) in controls (Fig. 1b). In the control group 7 of 10 ponies (#556, #568, #554, #562, #567, #571, #555) exhibited bronchopneumonia that required medical intervention; in the form of antibiotics and anti-inflammatory medication in 6 ponies, and anti-
Fig. 1. Experiment 1 (two-dose vaccination, 6-month duration of immunity): clinical disease post-challenge infection (CH) in each experimental group of ponies. Controls (n = 11): &, vaccinates (n = 12): ^. (a) Median temperature, (b) median clinical score, and (c) median weight difference (kg) from day 1.
inflammatory medication alone in an additional 2 ponies. Therapeutic interventions were needed between days 5 and 14 following infection. One of the control ponies did not respond to the treatment and was humanely euthanized on day 9 post-infection (#556) because of worsening respiratory disease and dyspnoea. On necropsy this pony exhibited severe, diffuse, subacute, fibrinonecrotic bronchointerstitial pneumonia. In contrast, none of the vaccinated animals required medical treatment post-challenge infection. 3.1.2. Virus shedding Nasal shedding was determined by real-time PCR measurement of influenza virus M-gene copies and mean log copy numbers are shown in Fig. 2. All animals were negative for viral shedding prior to challenge infection. All animals tested positive for 1 or more days post-challenge. The duration of viral shedding calculated as the first
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ponies showed significant increases in IgGa and IgGb levels following the second vaccination, while unvaccinated controls did not show IgGa or IgGb responses prior to challenge infection. IgGa levels declined to baseline 2 months after the second vaccination and IgGb levels declined to base line levels 4–5 months after the second vaccination. Following challenge infection IgGa and IgGb increased in both groups but levels were significantly higher in the vaccinates when compared to the control animals (p < 0.05). Fig. 2. Experiment 1 (two-dose vaccination, 6-month duration of immunity): median log M gene copy numbers in nasal swabs as determined by real-time PCR. Controls are shown by black bars, vaccinates are shown in white bars.
positive post-challenge day until the last positive postchallenge day was not significantly different from each other in the two treatment groups (p-value = 0.18) using a t-test. However, the amount of daily post-challenge viral shedding was significantly higher in the controls as compared to vaccinates on days 2–7 and 9 post-challenge infection (all p-values < 0.05).
3.2. Experiment 2 3.2.1. Clinical signs following influenza challenge infection Median temperatures, clinical scores and body weights are shown in Fig. 4. The number of days on which ponies
3.1.3. Antibody responses Median IgGa responses are shown in Fig. 3a, and median IgGb responses are shown in Fig. 3b. Vaccinated
Fig. 3. Experiment 1 (two-dose vaccination, 6-month duration of immunity): median IgGa and IgGb antibody titers for each experimental group. Controls (n = 11): &, vaccinates (n = 12): ^. Vaccinations (V) occurred on days 0 and 35 and all ponies were challenge infected (CH) on day 217. (a) IgGa and (b) IgGb.
Fig. 4. Experiment 2 (one-dose vaccination, onset of immunity): clinical disease post-challenge infection (CH) 14 days after a one-dose vaccination regimen. Controls (n = 10): &, vaccinates (n = 10): ^. (a) Median temperature, (b) median clinical score, and (c) median weight difference (kg) from day 1.
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exhibited increased temperatures was significantly lower (Fig. 1a, p-value < 0.0004) in vaccinates than in control ponies. In addition, vaccinates showed significantly less disease (Fig. 1b, p-value < 0.0001) and lost less weight (Fig. 1c) following challenge infection when compared to unvaccinated controls, although these weight loss differences between treatment groups were not statistically significant (p-value = 0.1). Starting at day 11 ponies in both groups gained weight, which is to be expected in young growing animals after recovery from the infection. No therapeutic interventions were needed in either experimental group, however one of the control ponies was humanely euthanized on day 11 post-infection due to a traumatic accident unrelated to the challenge. Upon necropsy this pony exhibited a severe, diffuse, subacute, suppurative pneumonia probably caused by bacteria secondary to viral infection. 3.2.2. Virus shedding Nasal shedding was determined by real-time PCR measurement of influenza virus M-gene copies and is shown as log copy numbers in Fig. 5. All animals were negative for viral shedding prior to challenge infection. All animals tested positive for one or more days postchallenge infection. The amount of daily post-challenge viral shedding of control ponies was significantly higher when compared to that of vaccinated ponies from days 2 to 9 post-challenge infection (p-value < 0.0001). 3.2.3. Antibody responses Median IgGa and IgGb responses are shown in Fig. 6. Vaccinated ponies showed a significant increase in IgGa titer following vaccination on day 13 post-vaccination – the day prior to challenge (p-value < 0.0001), while no increases in IgGb levels were seen in the vaccinates until after challenge infection. Antibody titers following challenge infection for both IgGa and IgGb were significantly higher in the vaccinates then in the unvaccinated controls indicating that these animals were primed by the vaccine (IgGa p-values 1 week (day 21) and 2 weeks (day 28) after challenge infection: <0.0001, IgGb p-values: day 21 = 0.0014, day 28 <0.0001). Unvaccinated controls did not show IgG responses prior to challenge infection and IgGb levels did not rise in the unvaccinated controls until 14 days following challenge infection (day 28), while IgGa
Fig. 5. Experiment 2 (one-dose vaccination, onset of immunity): median log M gene copy numbers in nasal swabs as determined by real-time PCR. Swabs were collected for 14 days post-challenge infection after a onedose vaccination regimen. Controls are shown by black bars, vaccinates are shown in white bars.
Fig. 6. Experiment 2 (one-dose vaccination, onset of immunity): median IgG antibody titers for each experimental group. Controls (n = 11): &, vaccinates (n = 12): ^. The vaccination (V) occurred on day 0 and all ponies were challenge infected (CH) on day 14. (a) IgGa and (b) IgGb.
levels were elevated by 7 days post-challenge infection (day 21). 4. Discussion This study confirms the observations of previous studies of ALVAC1-influenza vaccine technology (Edlund Toulemonde et al., 2005; Minke et al., 2007; Paillot et al., 2006), and advances them in a number of important ways. The work described here is the first study conducted with the commercial brand marketed in North America, and the first to demonstrate long-term (6-month) protective immunity resulting from this type of vaccine against an American lineage challenge strain. Furthermore, the onset of immunity study is the first report of a challengeprotection study using the contemporary North American isolate (A/equine/Ohio/03) which is one of the strains currently recommended by the OIE for inclusion in equine influenza vaccines (OIE, 2008). Finally this is the first report of influenza virus-specific equine IgG sub-class responses to an ALVAC1-influenza vaccine, and demonstrates that they are characterized by IgGa and IgGb responses, which are typically associated with protective immunity (Nelson et al., 1998). The principal finding of this report is the protection against a severe American lineage influenza strain challenge infection 6 months after completion of a primary series of two vaccinations. This observation provides significant evidence for the duration of protection of
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ALVAC1-influenza from the principle equine influenza virus strain in circulation worldwide today. The challenge model used was severe and was chosen because of its extensive past characterization (Lunn et al., 2001; Townsend et al., 2001). In this instance, the use of a challenge dose of 108 EID50, perhaps combined with conducting the challenge in Colorado at an altitude of 1520 m above sea level in the heat of summer, led to particularly severe disease in the control group. The severity of disease in controls required extensive medical intervention and the euthanasia of one animal. In the pony that had to be euthanized on day 9 post-infection, there were findings of severe, diffuse, subacute, fibrinonecrotic bronchointerstitial pneumonia on necropsy. These pathological findings are similar to reports from the first Australian equine influenza outbreak in 2007, where a number of foals born to influenza-naı¨ve mothers died of bronchointerstitial pneumonia, a disease that is usually rare in this age group (Patterson-Kane et al., 2008). In contrast, the minimal signs of disease in the vaccinated ponies further support the value of the vaccination regime. A significant reduction in viral shedding on 8 of the 10 days after challenge provided evidence for virological protection, however, there was no difference in the duration of shedding. This may indicate that 6 months after vaccination, equids may be able to propagate an infection, even if its clinical effects are limited. These findings are highly consistent with those reported previously by Minke et al. (2007) after a Eurasian viral challenge with the European brand of this vaccine. We chose to investigate the onset of immunity to a single ALVAC1-influenza vaccination of naı¨ve animals by challenge infection with a contemporary American lineage equine influenza virus (A/equine/Ohio/03). The results demonstrated good evidence of clinical and virological protection, similar to a previously reported but much smaller study of onset of immunity to the European brand (Edlund Toulemonde et al., 2005). However, in the study reported here, all vaccinated ponies shed virus on 2 or more days, whereas in the previous study vaccinates did not shed virus. This may be the result of differences in the challenge strain, route of infection or the detection methodology, although the former seems more likely. Both onset and duration of immunity after ALVAC1influenza vaccination were associated with induction of IgGa and IgGb antibody responses. These IgG sub-classes have previously been reported to be associated with protection from challenge infection in the absence of a nasal IgA response, while IgG(T) has not shown to provide protection (Nelson et al., 1998; Soboll et al., 2003a, 2003b). These responses are very likely responsible for the single radial hemolysis reactions previously described after use of ALVAC1 equine influenza virus vaccination (Edlund Toulemonde et al., 2005; Minke et al., 2007; Paillot et al., 2006). Interferon-g responses were not evaluated in this study, but a recent study by Paillot et al. (2006) has shown that two doses of the ALVAC1-influenza vaccine did prime horses to show significantly increased IFN-g responses to in vitro re-stimulation in PBMCs collected 7 and 14 days post-challenge infection. Together these results are consistent with induction of a type 1 immune response. These immunological outcomes are similar to vaccine responses
following DNA vaccination of horses with the hemagglutinin gene (Soboll et al., 2003a), but offer a more practical alternative to particle-mediated vaccine delivery. In addition to reduction of clinical disease, vaccination in our study significantly reduced the amount of viral shedding. For detection of viral nasal shedding, we employed a novel one-step real-time PCR technique that quantified viral shedding using probes and primers recognizing conserved sequences in the influenza A virus M gene of equine influenza viruses (Spindel et al., 2007). For quantification purposes this assay uses a plasmid containing the full length M gene and purified full length M gene RNA as standards. The assay system proved to be economical, fast and sensitive, with significant advantages over determination of viral titers in eggs or tissue culture systems. One disadvantage of this system is that it does not directly measure the infectious load in a sample. Nevertheless, the system has many advantages, and will be easier to establish in laboratories, particularly when egg incubators are not available. This is the first challenge infection study for equine influenza virus to report the use of a real-time PCR assay. Using this assay system we were able to detect differences in the amount of virus shed between treatment groups, which are an important factor for determination of vaccine efficacy. In summary, our study showed that the ALVAC1 equine influenza vaccine significantly reduced the clinical and virological sequelae of equine influenza virus infection in both onset and duration of immunity studies. This is the first study documenting the efficacy of this vaccine in a 6month duration of immunity study using an American lineage challenge strain. Acknowledgements We would like to thank Dr. Sangeta Rao and Dr. Ashley Hill for assistance with the statistical analysis of the second experiment in this study. In addition, Merial and sanofi aventis R&D are gratefully acknowledged for their valuable contribution. References Daly, J.M., Lai, A.C., Binns, M.M., Chambers, T.M., Barrandeguy, M., Mumford, J.A., 1996. Antigenic and genetic evolution of equine H3N8 Influenza A viruses. Journal of General Virology 77, 661–671. Edlund Toulemonde, C., Daly, J., Sindle, T., Guigal, P.M., Audonnet, J.C., Minke, J.M., 2005. Efficacy of a recombinant equine influenza vaccine against challenge with an American lineage H3N8 influenza virus responsible for the 2003 outbreak in the United Kingdom. Veterinary Record 156, 367–371. Landolt, G., 2007. Equine influenza infection. In: Sellon, D.C., Long, M.T. (Eds.), Equine Infectious Diseases. pp. 124–134. Lunn, D.P., Holmes, M.A., Antczak, D.F., Agerwal, N., Baker, J., BendaliAhcene, S., Blanchard-Channell, M., Byrne, K.M., Cannizzo, K., Davis, W., Hamilton, M.J., Hannant, D., Kondo, T., Kydd, J.H., Monier, M.C., Moore, P.F., O’Neil, T., Schram, B.R., Sheoran, A., Stott, J.L., Sugiura, T., Vagnoni, K.E., 1998. Report of the Second Equine Leucocyte Antigen Workshop, Squaw valley, California, July 1995. Veterinary Immunology & Immunopathology 62, 101–143. Lunn, D.P., Hussey, S., Sebring, R., Rushlow, K.E., Radecki, S.V., WhitakerDowling, P., Youngner, J.S., Chambers, T.M., Holland Jr., R.E., Horohov, D.W., 2001. Safety, efficacy, and immunogenicity of a modified-live equine influenza virus vaccine in ponies after induction of exerciseinduced immunosuppression. Journal of the American Veterinary Medical Association 218, 900–906.
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