• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of cjvetresCVMACanadian Journal of Veterinary ResearchSee also Canadian Journal of Comparative MedicineJournal Web siteHow to Submit
Can J Vet Res. Oct 2010; 74(4): 271–278.
PMCID: PMC2949340

Language: English | French

Surveillance of equine respiratory viruses in Ontario


The objective of this project was to develop and implement an active surveillance program for the early and rapid detection of equine influenza viruses in Ontario. For this purpose, from October 2003 to October 2005, nasopharyngeal swabs and acute and convalescent serum samples were collected from 115 client-owned horses in 23 outbreaks of respiratory disease in Ontario. Sera were paired and tested for antibody to equine influenza 1 (AE1-H7N7), equine influenza 2 (AE2-H3N8), equine herpesvirus 1 and 4 (EHV1 and EHV4), and equine rhinitis A and B (ERAV and ERBV). Overall, the cause-specific morbidity rate of equine influenza virus in the respiratory outbreaks was 56.5% as determined by the single radial hemolysis (SRH) test. The AE2-H3N8 was isolated from 15 horses in 5 outbreaks. A 4-fold increase in antibody levels or the presence of a high titer against ERAV or ERBV was observed in 10 out of 13 outbreaks in which AE2-H3N8 was diagnosed as the primary cause of disease. In conclusion, AE2-H3N8 was found to be an important contributor to equine respiratory viral disease. Equine rhinitis A and B (ERAV and ERBV) represented an important component in the equine respiratory disease of performing horses.


L’objectif du présent projet était de développer et mettre en place un programme de surveillance active pour la détection hâtive et rapide des virus de l’influenza équin en Ontario. À cette fin, durant la période allant de octobre 2003 à octobre 2005, des écouvillons naso-pharyngés et des échantillons de sérum prélevés en phase aiguë et de convalescence ont été pris chez 115 chevaux de clients lors de 23 épisodes de maladies respiratoires en Ontario. Les sérums ont été pairés et testés pour la présence d’anticorps contre l’influenza équin de type 1 (AE1-H7N7), l’influenza équin de type 2 (AE2-H3N8), les herpès virus équins de type 1 et 4 (EHV1 et EHV4), et les virus de la rhinite équine A et B (ERAV et ERBV). De manière globale, le taux de morbidité spécifique associé au virus de l’influenza équin dans les poussées de cas de maladies respiratoires était de 56,5 % tel que déterminé par l’épreuve d’hémolyse radiale simple (SRH). Le virus AE2-H3N8 a été isolé de 15 chevaux dans 5 épisodes. Une augmentation d’un facteur de 4 des titres d’anticorps ou la présence d’un titre d’anticorps élevé envers ERAV ou ERBV a été observée dans 10 des 13 épisodes lors desquels le virus AE2-H3N8 a été identifié comme la cause première de la maladie. En conclusion, le virus AE2-H3N8 a été identifié comme étant un contributeur important des maladies respiratoires équines. Les virus de la rhinite équine A et B (ERAV et ERBV) pourrait également représenté une composante importante des maladies respiratoires équines chez les chevaux de performance.

(Traduit par Docteur Serge Messier)


Equine influenza A virus is considered one of the most common viruses affecting the respiratory tract of young horses worldwide (1,2). The equine influenza A virus was first isolated in 1956 during an equine respiratory outbreak in Eastern Europe and was characterized as AE1-H7N7 (3). The AE1-H7N7 subtype has not been isolated since 1979. In 1963, the AE2-H3N8 subtype was identified during an outbreak of equine influenza in Miami, Florida, USA (4). The AE2-H3N8 virus soon spread throughout America and Europe and has been the cause of many respiratory outbreaks in the last 25 y (57). In 1986, the AE2-H3N8 virus was introduced into a naïve equine population in South Africa and subsequently had a negative effect on the racing industry (8). More recently, this virus has been introduced into Australia (9) with major consequences in a susceptible population. Since 1990, variations in the viral ribonucleic acid (RNA) and antigenic epitopes have been identified (10). At that time, 2 diverse lineages were recognized: the European and the American lineages (11).

The influenza virus affects specifically the respiratory tract, diminishing the performance potential of the animal and raising the risk of secondary bacterial complications (7,12). As a result, this viral respiratory infection is a threat to the equine population due to the loss of training days and the high risk of severe epidemics (7). Influenza virus infection has traditionally been diagnosed through virus isolation and/or serology [hemagglutinin inhibition (HI)]. However, the single radial hemolysis (SRH) test has been introduced as a technique with more accurate quantitative titers for determining protection in vaccinated horses (13). The SRH test is based on the passive hemolysis of virus-sensitized sheep erythrocytes by the anti-hemagglutinin antibodies in the test serum. The hemolysis observed has been shown to be directly proportional to the amount of strain-specific antibody in the serum being tested (11). An increase of 25 mm2 or a doubling of the hemolysis area is considered to be a significant increase. An international surveillance program for equine influenza has been established in an attempt to increase the identification of outbreaks and recognize new strains affecting the world horse population. Canada has not actively been part of this program. This project was therefore established to develop and implement an active surveillance program for the early and rapid detection of equine influenza viruses in Ontario.

Materials and methods

Study design

The study was designed for sample collection over 2 consecutive years, from September 2003 to November 2005. Equine practitioners in the province of Ontario were asked to identify young performing horses (preferably not older than 6 y) with clinical signs consistent with equine influenza virus infection based on the following criteria: acute signs of viral respiratory disease less than 24 h from the onset of signs; temperature of ≥ 39°C; cough, serous nasal discharge, clear ocular discharge, depression, anorexia, no history of recent influenza vaccination (horses were not vaccinated at least 2 wk prior to the outbreak), and rapid spread of clinical signs within the stable. On the first visit, the main investigator recorded the history and physical examination with emphasis on the respiratory tract on a questionnaire that had been previously prepared. At the time, 2 nasopharyngeal swabs per horse were collected for antigen-detection enzyme-linked immunosorbent assay (ELISA), reverse transcriptase-polymerase chain reaction (RT-PCR), and viral isolation and acute blood samples were collected for subsequent antibody analysis. On a second visit 21 d later, convalescent blood samples were collected for paired serology for antibody titers against the following common equine respiratory viruses: equine influenza 1 (AE1-H7N7), equine influenza 2 (AE2-H3N8), equine herpesvirus 1 and 4 (EHV1 and EHV4), and equine rhinitis A and B (ERAV and ERBV).


The subjects were included in the study based on the criteria described above. The health status of the horses was evaluated and 5 horses from the stable were included. Horses that presented clinical signs of respiratory viral infection on the morning of the first visit were the primary candidates for the study. Three symptomatic horses and 2 non-diseased stablemates in neighboring stalls were sampled. The Animal Care Committee at the University of Guelph approved an animal utilization protocol (AUP) in September 2003.

Clinical examination

Physical examination records included rectal temperature, respiratory rate, presence or absence of cough, characterization of submandibular lymph nodes (enlargement), and the presence or absence of nasal and ocular discharge and its characteristics (clear, mucoid, or mucopurulent).

Sampling techniques

Whenever possible, samples for virus isolation were collected within 24 h of the onset of clinical signs. Two nasopharyngeal swabs (Kalayjian Industries; Signal Hill, California, USA) were taken from each horse, the tips were cut off and placed in 2 different tubes containing virus transport medium (VTM), the tubes were shaken and kept in a cooler with ice, and were submitted to the Animal Health Laboratory (AHL) at the University of Guelph (Guelph, Ontario). At the onset of the respiratory outbreak, blood samples were obtained for serology. Serum samples were stored at −80°C for subsequent analysis of antibody titers. Approximately 21 d later, a convalescent sample was also taken and stored.

Laboratory analysis

Virus isolation

Virus isolation was carried out in cell culture (MDCK and RK-13 cells) and in embryonated eggs as previously described (6). The influenza A virus isolates recovered were initially identified using hemagglutination and the Directigen Flu A Antigen ELISA (Becton, Dickinson; Franklin Lakes, New Jersey, USA). They were confirmed by RT-PCR using nucleoprotein primers (14) and the hemagglutinin was subtyped using gel-based RT-PCR (14,15). The equine rhinitis viruses recovered in RK-13 cells were identified using type-specific monospecific antisera in virus neutralization assays as previously described (6).


Microtiter hemagglutination-inhibition tests for AE1-H7N7 and AE2-H3N8 and microtiter virus neutralization assays for ERAV, ERBV, EHV1, and EHV4 were performed as previously described (6). A significant change in antibody levels was defined as a change from negative to positive or a 4-fold increase in the titer between acute and convalescent samples. The single radial hemolysis (SRH) test was validated in our laboratory to assess the level of antibodies to equine influenza virus on the paired serum samples collected. This validation was based on standards established by the World Health Organization (WHO) (11,13).

Reverse transcriptase-polymerase chain reaction (RT-PCR) and RT-PCR typing

Ribonucleic acid (RNA) was extracted from nasal swabs and allantoic fluids infected with influenza A virus using QIAamp Viral RNA Mini Kit (Qiagen Sciences; Germantown, Maryland, USA) as described by the manufacturer. Primers to the nucleoprotein (14) were used in gel-based RT-PCR to identify virus from nasopharyngeal swabs and allantoic fluids. The hemagglutinin was subtyped using gel-based RT-PCR (14,15).

Hemagglutinin genome sequencing

A total of 15 AE2-H3N8 isolates from Ontario horses sampled from 2003 to 2005 were grouped into 4 groups according to period of outbreak (time of the year). A representative isolate from each group was subjected to hemagglutinin genome sequencing. Viral RNA was extracted using the QIAamp Viral RNA Mini Kit (Qiagen Sciences). Complementary DNA (cDNA) was synthesized by reverse transcription of viral RNA (2 μL) with Stratascript cDNA synthesis kit (Stratagene; La Jolla, California, USA) using the universal primer (Uni-12) (16). Full-length hemagglutinin (HA) gene amplicon was generated by PCR amplification of 2 μL of cDNA with Taq DNA polymerase (Invitrogen, Carlsbad, California, USA). Amplified PCR products were purified with the QIAquick Gel extraction kit (Qiagen Sciences). Using the set of equine influenza HA gene-specific primers (primer sequences available on request), sequencing reaction (10 μL) was set up with BigDye Terminator v1.1 (Applied Biosystems, Carlsbad, California, USA) following the manufacturer’s instructions. Sequencing reaction was precipitated by ethanol/EDTA as outlined in the user’s manual. Purified sequencing reaction was dissolved in 12 μL of HiDi Formamide (Applied Biosystems) and electrophoresis was performed using an ABI 310 Genetic Analyzer (Applied Biosystems).

Sequence fragments were assembled and edited on Lasergene v7.2 (DNASTAR, Madison, Wisconsin, USA). Hemagglutinin (HA) gene sequences of 4 AE2-H3N8 isolates determined in this study were aligned with relevant HA gene sequences available in the influenza virus database at the National Center for Biotechnology Information (NCBI), (CLUSTALX version 1.83) (17). The phylogenetic tree was constructed from the aligned sequences by the neighbor-joining Kimura-2 method implemented in the PHYLIP (Phylogeny Inference Package) 3.67 package (18). The reliability of the branching orders was estimated by bootstrapping (100 replicates). The tree topology was produced in MEGA 4 (19) and was rooted to A/equine/ Miami/1963.

Data analysis

An outbreak was considered positive for equine influenza if at least 1 horse from the outbreak seroconverted, as determined by the SRH assay. To calculate the case morbidity rate of equine influenza, the number of positive outbreaks diagnosed by SRH titers (an increase in the hemolysis area of 25 mm2 or doubling of the hemolysis area) was divided by the total number of outbreaks sampled. Descriptive statistics were performed on the prevalence of clinical signs, results from serology tests, antigen-detection ELISA, RT-PCR, and virus isolation. Results obtained by SRH and hemagglutination inhibition (HI) tests for equine influenza viruses were categorized as positive or negative for seroconversion or significant increase in antibody levels (dichotomous variables) and were evaluated for agreement. Agreement between the 2 serological tests (SRH and HI) was analyzed at the outbreak level and also assessed at the individual level. Agreement among serology results for AE1-H7N7, AE2-H3N8, EHV1 and EHV4, and ERAV and ERBV was also determined. All variables were compared using the simple kappa coefficient test (k).

The chi-squared (X2) test was used to compare the proportions of horses that had a significant increase in antibody levels (positive SRH) in vaccinated and unvaccinated groups and to compare the proportion of horses that developed clinical signs of equine influenza in vaccinated and unvaccinated groups. Horses with unknown vaccination history (n = 23) were not included in this analysis. The chi-squared (X2) test was also used to compare the proportion of horses in both clinically ill and healthy groups that had a positive test for equine influenza virus (ELISA, RT-PCR, virus isolation, SRH, and HI). Statistical significance was set at P < 0.05.


Twenty-three outbreaks of equine respiratory infection were identified from October 2003 to October 2005, with a total of 115 horses sampled (Table I). These outbreaks were distributed throughout southern Ontario. Acute and convalescent serum samples were available from 113 of 115 horses, as 2 horses were unavailable for collection of convalescent samples.

Table I
Results for serological and virus detection methods in each respiratory outbreak (2003 to 2005)

The average age of the horses was 3.2 y and 98 horses were ≤ 5 y old. Horses included in this study were involved primarily in racing, showing, and breeding. Of the 115 horses sampled, 86 (75%) were clinically ill and 29 (25%) were stablemates in clinically healthy condition. Of the 86 diseased horses (Table II), 65 had a harsh cough and some degree of nasal discharge, of which 40 had serous and clear discharge, 15 had mucoid discharge, and 10 presented mucopurulent discharge. Twenty-nine clinically ill horses were presented with a respiratory rate of ≥ 24 respirations/min and only 18 presented a rectal temperature of ≥ 39°C. Enlarged submandibular lymph nodes were observed in 13 horses and ocular discharge was observed in 12 of the ill horses. Depression, anorexia, and muscle soreness were reported in 26, 21, and 4 diseased horses, respectively. Of those sampled, 41 horses (36%) had been given an equine influenza vaccine at least once during the previous year, 51 horses (44%) had a history of no vaccination, and the vaccine history of 23 horses (20%) was unknown.

Table II
Clinical signs observed in clinically ill horses (n = 86) involved in the study of equine viral respiratory outbreaks in Ontario and results by diagnostic tests (2003 to 2005)

From the 87 nasopharyngeal swabs that were suitable and submitted for virus isolation, 57 were also screened for equine influenza virus with the commercial antigen-detection ELISA, 16 with both the antigen-detection ELISA assay and RT-PCR, and 14 with RT-PCR alone. Ten out of 16 samples tested positive to AE2-H3N8 as detected by RT-PCR, compared to only 6 positive samples with the antigen-detection ELISA from the same horses. Virus isolation in embryonated eggs yielded 15 equine influenza virus isolates from 5 outbreaks that were subsequently typed as H3 using RT-PCR. Equine rhinitis A (ERAV) was isolated from 1 horse and equine rhinitis B (ERBV) from 3 horses using cell culture, each from separate outbreaks.

Further sequencing of the HA1 region of 4 influenza virus isolates revealed a high identity (98%) to Kentucky/2002 and Ohio/2003 at the nucleotide level (GenBank accession numbers: 1106728, 1107170, 1107171, and 1107172). Additionally, construction and analysis of the phylogenetic tree confirmed the closeness of these isolates (Figure 1).

Figure 1
Phylogenetic relationship of Guelph EIV A [H3N8] isolates used in the study with other EIV A [H3N8] viruses. Bootstrap value at each node is 100.

A significant 4-fold increase in antibody levels to AE2-H3N8-HI was identified in 28 horses. Of these 28 horses, only 14 yielded positive results in virus isolation in eggs and had significant changes in antibody levels in the HI test. When influenza virus was isolated, a significant 4-fold increase in antibody levels by HI (AE2-H3N8) was observed in 14 of 15 horses and in 15 of 15 horses when tested by SRH (AE2-H3N8). The SRH test demonstrated a significant increase in antibody levels to AE2-H3N8 in 45 of 113 horses (40%). In addition, the SRH assay detected a significant increase in antibody levels to AE1-H7N7 in 4 horses, while HI detected a significant increase against this subtype in only 1 horse.

Seroconversion or significant increase (4-fold increase) in antibody levels to ERAV by virus neutralization (VN) was observed in only 3 horses. However, high titers to ERAV, for example ≥ 1:1024, were detected in 32 horses (28%). Interestingly, a 4-fold increase in antibody levels or the presence of a high titer against ERAV-VN and AE2-H3N8-SRH was observed concomitantly in 19 horses (17%).

Furthermore, a 4-fold increase in virus-neutralizing antibody levels to ERBV was identified in only 4 horses. Overall, antibody to ERAV and ERBV was detected in 18 of 23 outbreaks (Table I). A 4-fold increase in antibody levels to EHV1 and EHV4 was observed in only 1 horse.

Overall, the case-specific morbidity rate of AE2-H3N8 in the respiratory outbreaks in Ontario was 56.5%. This calculation was based on a significant increase in antibody levels as demonstrated in the AE2-H3N8-SRH test at the outbreak level based on having 1 positive horse per herd (Table I). The sensitivity and specificity for the SRH test in our laboratory were calculated as 100% and 80%, respectively. The average age of positive horses in the AE2-H3N8-SRH test was 4 y and the average age of negative horses in the AE2-H3N8-SRH test was 2.7 y. As shown in Table III, both assays (AE2-H3N8-SRH and AE2-H3N8-HI) identified 28 positive horses and 68 negative horses. Additionally, 17 horses were classified as positive by the AE2-H3N8-SRH, but negative with AE2-H3N8-HI. The kappa (k) of 0.66 indicates a moderate level of agreement between the 2 tests (P ≤ 0.0001) (Table III). These 2 serology tests were also assessed for agreement at the outbreak level (significant rise in antibody levels to AE2-H3N8 of at least 1 horse in an outbreak), finding a similar moderate level of agreement with a k value of 0.66 (P = 0.0016).

Table III
Agreement between single radial hemolysis (SRH) and hemagglutination inhibition (HI)

In our study, horses that had respiratory clinical signs were less likely to be positive in the antigen-detection ELISA test (relative risk = 0.11, P = 0.01). No statistically significant association between clinically ill and healthy groups was found when results from other tests (RT-PCR, virus isolation, SRH, and HI) were compared. In this study, horses that were unvaccinated (variety of commercial influenza vaccines) were 2.42 times more likely to seroconvert based on the SRH test during an influenza outbreak than vaccinated horses (P = 0.01). Vaccine status was not associated with the presence or absence of clinical respiratory signs (P = 0.8).


The outbreaks included in this study represent only the proportion of horses reported by veterinarians willing to participate in the study. Nevertheless, the distribution of outbreaks was not linked to a specific region. Despite veterinary recommendations, only a small proportion of the population included in this study had been vaccinated in the previous year. As shown in similar studies, young horses were the population most likely to be positive for viral isolation since age is an important factor in determining susceptibility to viral infections (20). The clinical signs found in horses sampled during this study were consistent with signs previously observed in both natural and induced respiratory viral infections (7,21,22). In some of the outbreaks, however, the severity of the clinical signs may have been due to the virulence of viral strains, a combination of 2 or more viral agents, and possibly concomitant secondary bacterial infections. Furthermore, having early access to the clinical cases made it possible to recognize the more severe clinical signs that are characteristic of viral respiratory infections (22).

A combination of time delay and difficulty in isolating the influenza virus from clinical cases may explain why influenza virus was isolated from only 15 horses in 5 outbreaks, since these viruses are isolated only in the first 24 to 48 h after the onset of illness. These factors may have influenced the results in 2 outbreaks of respiratory disease where neither rise in antibody levels, nor was isolation of any virus achieved. Previous and similar studies conducted in racehorses by Sherman et al (12) and Carman et al (6) demonstrate that the success rate of isolating influenza virus using the same methods was relatively low (13.7% and 5.4%, respectively). It is possible that our results may have differed from Sherman’s study (12) due to improved sample handling, processing, and technical methods or a combination of these factors. In 1977, Sherman found that equine herpes virus 2 (EHV2) was the most common isolate recovered from respiratory outbreaks in Ontario, whereas we found influenza and rhinitis viruses to be more commonly isolated from such outbreaks. On the other hand, although Carman et al (6) recovered AE2-H3N8 from 5 of 92 horses, they found that the most prevalent virus isolated from equine respiratory outbreaks was ERBV, which they isolated from 28 of 92 horses. Equine herpes viruses were not a common cause of respiratory disease in animals included in this study, which could have been associated with the age of the target population.

Diverse diagnostic techniques have been used to rapidly identify and diagnose influenza viral infection in humans. Sensitivity to these tests may be compromised depending on viral strains and other factors (23). In veterinary medicine, Directigen Flu A ELISA has been utilized as a rapid diagnostic test, both on site and in laboratory settings. A high sensitivity and specificity was shown for this commercial ELISA in previous studies, (24,25). In contrast, our study found a low rate of positive detection when direct clinical samples were tested with this ELISA. This lack of sensitivity could have been influenced by either insufficient viral protein in the samples or the presence of antigen-antibody complexes that would prevent a positive reaction from developing on the test. The reaction obtained in this test is based on the detection of the influenza virus nucleoprotein, which is highly conserved among influenza A viruses. In some of our cases, viral isolation in embryonated eggs was achieved from the same material used to perform the antigen-detection ELISA, but the Directigen Flu A ELISA was able to detect a positive case using allantoic fluid from infected eggs only after amplification of the virus by a second or third passage in eggs. A recent study comparing the sensitivity of equine influenza virus isolation, antigen-detection ELISA, and nucleic acid amplification found that Directigen Flu A ELISA was the least sensitive of these 3 methods (26). This evidence demonstrates that this commercial assay may be useful to support or confirm the presence of the virus in heavily virus-loaded nasal swabs or in virus-infected allantoic fluids after egg inoculation where virus is present in large concentrations. A negative test, however, does not rule out influenza virus infection. Others have reported high sensitivity and specificity when analyzing equine secretions with this human immunoassay (24,25).

In our study, the reverse transcriptase-polymerase chain reaction (RT-PCR) test identified the samples with influenza virus better than the ELISA. In 2004, Quinlivan and collaborators demonstrated the high sensitivity of the RT-PCR test for identifying equine respiratory viral infection (26). Our results and findings by others may indicate that RT-PCR is a more reliable tool than the ELISA for diagnosing influenza virus in the horse. In addition, it has been proven that RT-PCR is highly sensitive and specific, requiring very small amounts of genetic material for analysis (27).

In this study, the single radial hemolysis test (SHR) for AE2-H3N8 confirmed its greater ability to detect changes in antibody levels (56.5% of outbreaks) when compared to the hemagglutination inhibition (HI) test for AE2-H3N8 (39% of outbreaks) (Table I). Studies have demonstrated that the SRH is a more efficient technique for detecting small changes in antibody levels between acute and convalescent samples (28,29). Specifically, a collaborative study to establish an equine influenza antiserum as a reference preparation demonstrated poor repeatability and reproducibility in the hemagglutination inhibition (HI) test compared to results obtained in the SRH assay. Consequently, the SRH results were the only parameter considered to establish the antibody titer for the European Pharmacopoeia (30). This evidence and the results obtained in this study are in agreement with their findings.

Antibody levels determined using SRH have been evaluated in experimental challenge-infection studies (28). When used in combination with valuable information collected during an influenza outbreak, SRH antibody levels have demonstrated that 90% of horses with antibody levels above 160 mm2 are likely to be protected (31). When evaluating post-vaccination antibody levels in young animals or post-infection changes in specific equine influenza antibody, the SRH test offers a straightforward value to interpret, with a greater possibility of detecting small variations in the antibody levels. Studies have also demonstrated that horses with an SRH antibody level of >120 mm2 and > 90 mm2 respectively were protected from infection and clinical signs of disease when challenged with homologous viruses (32).

Interestingly, in this study an association was observed between infection with ERAV and AE2-H3N8. It is known that antibody levels against equine rhinitis viruses increase rapidly and peak very fast so that a 4-fold increase in antibody titer cannot be easily demonstrated (acute and convalescent) (33). Other studies have reported that equine ERAV is a primary cause of respiratory disease in horses (34). In our study, a significant increase in antibody levels to ERAV was observed in only 3 horses, although high titers, for example ≥ 1:1024, were seen among horses (n = 47) in some of the outbreaks. This evidence suggests that ERAV might have played an important role in clinical disease observed in those cases. Based on previous studies, ERAV might have been underestimated when evaluating equine respiratory outbreaks (34). A previous study in Ontario (5,6) found that ERBV was the most common cause of respiratory disease in study horses. Our study confirmed that ERBV, which was recovered in 3 of 23 outbreaks, is still a potential cause of respiratory disease in horses in Ontario. Using both virus identification and serological titers, ERAV and ERBV were associated with 18 of 23 outbreaks. Rhinoviruses have been extensively investigated in humans and associated not only with upper respiratory infections, but also with asthma exacerbations and inflammatory airways disease (35). Similarly, ERAV and ERBV may represent a cause for increased severity in clinical viral infections and prolonged periods of respiratory disease in susceptible horses. Further investigations of AE2-H3N8, alone and in association with ERAV and ERBV, could clarify the role of these viruses in the equine respiratory viral complex.

Genetic differences among influenza virus isolates recently recovered in Ontario and other equine influenza viruses in Ontario (36) and North America suggest that strains circulating in southern Ontario have diverged. All of our isolates, however, fell within the American lineage (Clade 1), which is consistent with other North American isolates. Our data confirm that the continuous evolution of the equine influenza virus requires active surveillance in order to provide updated information for infection control and vaccine development.

In summary, we have confirmed a high prevalence of AE2-H3N8 in equine respiratory outbreaks in Ontario. This virus remains an important cause of equine respiratory viral disease affecting young performance horses. However, ERAV and ERBV might also represent a significant and central component in the equine respiratory disease complex, as shown by virus identification and serological methods. For these reasons, ERAV and ERBV should be taken into account when identifying the cause of equine respiratory outbreaks.

Our study suggests that the SRH assay might be considered as an alternative technique for evaluating antibody levels to equine influenza. Active vaccination protocols, quarantine measures, prompt laboratory detection, and active surveillance programs remain the best options for preventing and controlling disease due to equine influenza virus.


The authors thank the equine practitioners of Ontario for their collaboration in identifying the outbreaks, Dr. Janet Daly from The Animal Health Trust for providing guidance for SRH testing and supplying the equine influenza antiserum for technique validation, and Dr. Hugh Townsend (University of Saskatchewan) for providing sera for validation of the SRH test. This study would not have been possible without the generous support of Boehringer Ingelheim (Canada) Ltd., Vetmedica, Burlington, Ontario, the EP Taylor Equine Research Fund, and the Ontario Racing Commission.


1. Powell DG. A study of infectious respiratory disease among horses in Great Britain, 1971–1976. Proc 4th Intern Conf Equine Infectious Diseases. 1978:451–459.
2. Van Maanen C, Cullinane A. Equine influenza virus infections: An update. Vet Q. 2002;24:79–94. [PubMed]
3. Sovinova O, Tumova B, Pouska F, Nemec J. Isolation of a virus causing respiratory disease in horses. Acta Virol. 1958;2:51–61. [PubMed]
4. Waddell GH, Teigland MB, Sigel MM. A new influenza virus associated with equine respiratory disease. J Am Vet Med Assoc. 1963;15:587–90. [PubMed]
5. Willoughby RA, Huber L, Vie1 L. Culture and serological results in acute upper respiratory infections in horses. Proc Am Coll Vet Intern Med (ACVIM) Forum. 1989:604–605.
6. Carman S, Rosendal S, Huber L, et al. Infectious agents in acute respiratory disease in horses in Ontario. J Vet Diagn Invest. 1997;9:17–23. [PubMed]
7. Newton JR, Verheyen K, Wood JLN, Yates PJ, Mumford JA. Equine influenza in the United Kingdom in 1998. Vet Rec. 1999;145:449–52. [PubMed]
8. Guthrie AJ, Stevens KB, Bosman PP. The circumstances surrounding the outbreak and spread of equine influenza in South Africa. Rev Sci Tech. 1999;18:179–85. [PubMed]
9. Cowled B, Ward MP, Hamilton S, Garner G. The equine influenza epidemic in Australia: Spatial and temporal descriptive analyses of a large propagating epidemic. Prev Vet Med. 2009;92:60–70. [PubMed]
10. Daly JM, Lai ACK, Binns MM, Chambers TM, Barrandeguy M, Mumford J. Antigenic and genetic evolution of equine H3N8 influenza A viruses. J Gen Virol. 1996;77:661–671. [PubMed]
11. Oxburgh L, Berg M, Klingeborn B, Emmoth E, Linne T. Evolution of H3N8 equine influenza virus from 1963 to 1991. Virus Res. 1994;34:153–65. [PubMed]
12. Sherman J, Thorsen J, Barnum DA, Mitchell WR, Ingram DG. Infectious causes of equine respiratory disease on Ontario Standardbred racetracks. J Clin Microbiol. 1977;5:285–289. [PMC free article] [PubMed]
13. Newton JN, Townsend HGG, Wood JLN, Sinclair R, Hannant D, Mumford JA. Immunity to equine influenza: Relationship of vaccine-induced antibody in young thoroughbred racehorses to protection against field infection with influenza A/equine-2 viruses (H3N8) Equine Vet J. 2000;32:65–74. [PubMed]
14. Lee MS, Change PC, Shien JH, Cheng MC, Shieh HK. Identification and subtyping of avian influenza viruses by reverse transcription-PCR. J Viro Methods. 2001;97:13–22. [PubMed]
15. Schorr E, Wentworth D, Hinshaw VS. Use of polymerase chain reaction to detect swine influenzavirus in nasal swabs specimens. Am J Vet Res. 1994;55:952–956. [PubMed]
16. Huddleston JA, Brownlee GG. The sequence of the nucleoprotein gene of human influenza A virus strain, A/NT/60/68. Nucleic Acids Res. 1982;10:1029–1037. [PMC free article] [PubMed]
17. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG. The ClustalX windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 1997;25:4876–4882. [PMC free article] [PubMed]
18. Felsenstein J. Distributed by the author. Department of Genome Sciences, University of Washington; Seattle: 2005. PHYLIP (Phylogeny Inference Package) version 3.6.
19. Tamura K, Dudley J, Nei M, Kumar S. MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol Biol Evoln. 2007;24:1596–1599. [PubMed]
20. Wood JLN, Newton JR, Chanter N, Mumford JA. Association between respiratory disease and bacterial and viral infections in British race horses. J Clin Microbiol. 2005;43:120–126. [PMC free article] [PubMed]
21. Willoughby R, Ecker G, McKee S, et al. The effects of equine rhinovirus, influenza virus and herpesvirus on tracheal clearance rate in horses. Can J Vet Res. 1992;56:115–121. [PMC free article] [PubMed]
22. Sutton G. Master of Science thesis. Guelph, Ontario: University of Guelph; 1992. Immunocytochemical identification of equine influenza-2 virus and equine herpesvirus-1 in experimentally infected pony foals.
23. Weinberg A, Mettenbrink CJ, Ye D, Yang C. Sensitivity of diagnostic tests for influenza varies with the circulating strains. J Clin Virol. 2005;35:172–175. [PubMed]
24. Chambers TM, Shortridge KF, Li PH. Rapid diagnosis of equine influenza by the Directigen Flu A enzyme immunoassay. Vet Rec. 1994;135:275–279. [PubMed]
25. Morley PS, Bogdan JR, Townsend HGG, Haines DM. Evaluation of Directigen Flu A assay for detection of influenza antigen in nasal secretions of horses. Equine Vet J. 1995;27:131–134. [PubMed]
26. Quinlivan M, Cullinane A, Nelly M, Van Maanen K, Heldens J, Arkins S. Comparison of sensitivities of virus isolation, antigen detection, and nucleic acid amplification for detection of equine influenza virus. J Clin Microbiol. 2004;42:759–763. [PMC free article] [PubMed]
27. Quinlivan M, Dempsey E, Ryan F, Arkins S, Cullinane A. Real-time reverse transcription PCR for detection and quantitative analysis of equine influenza virus. J Clin Microbiol. 2005;43:5055–5057. [PMC free article] [PubMed]
28. Mumford JA, Hannant D, Jessett DM. Experimental infection of ponies with equine influenza (H3N8) viruses by intranasal inoculation or exposure to aerosols. Equine Vet J. 1990;22:93–98. [PubMed]
29. Morley PS, Bogdan JR, Townsend HGG, Haines DM. The effect of changing single radial haemolysis assay method when quantifying influenza A antibodies in serum. Vet Microbiol. 1995;44:101–110. [PubMed]
30. Mumford JA. Collaborative study for the establishment of three European pharmacopoeia biological reference preparation for equine influenza horse antiserum. Pharmeuropa. 2000:7–21.
31. Mumford JA. Progress in the control of equine influenza. Proc. 6th Inter Conf Equine Infectious Diseases. 1992:207–218.
32. Mumford JA, Jessett DM, Dunleavy U, Wood JLN, Hannant D, Sundquist B. Antigenicity and immunogenicity of experimental equine influenza ISCOM vaccines. Vaccine. 1994;12:857–63. [PubMed]
33. Powell DG. Viral respiratory disease of the horse. Vet Clin North Am Equine Pract. 1991;7:27–52. [PubMed]
34. Li F, Drummer HE, Ficorilli N, Studdert MJ, Crabb BS. Identification of noncytopathic equine rhinovirus 1 as a cause of acute febrile respiratory disease in horses. J Clin Microbiol. 1997;35:937–943. [PMC free article] [PubMed]
35. Gern JE, Busse WW. Association of hinovirus infections with asthma. Clin Microbiol Rev. 1999;12:9–18. [PMC free article] [PubMed]
36. Gagnon CA, Elahi SM, Tremblay D, et al. Genetic relatedness of recent Canadian equine influenza virus isolates with vaccine strains used in the field. Can Vet J. 2007;48:1028–1030. [PMC free article] [PubMed]

Articles from Canadian Journal of Veterinary Research are provided here courtesy of Canadian Veterinary Medical Association
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...