a pcr based method for the identification of equine influenza virus from clinical samples

A PCR based method for the identification of equine

influenza virus from clinical samples

L. Oxburgha,*, AÊ . HagstroÈmb

aDepartment of Veterinary Microbiology, Virology Section, Swedish University of Agricultural Sciences,

Biomedical Centre, Box 585, S-751 23, Uppsala, SwedenbDepartment of Virology, National Veterinary Institute, Biomedical Centre, Box 585, S-751 23, Uppsala, Sweden

Received 22 October 1998; accepted 16 March 1999

Abstract

In this paper we describe the development of a nested RT-PCR assay for the rapid diagnosis and

characterisation of influenza virus directly from clinical specimens. Viral RNA is extracted from

nasal swabs by the guanidine thiocyanate extraction method, and subsequently reverse transcribed.

The complementary DNA is then used as template in a nested PCR reaction. Primers designed for

use in this assay are specific for three templates; (1) the nucleoprotein (NP) gene, (2) the

haemagglutinin gene of the H7N7 equine influenza virus (A1), and (3) the haemagglutinin gene of

the H3N8 equine influenza virus (A2). We show that the assays are specific for the target genes

chosen, and display sensitivity similar to virus isolation. The NP assay detects a variety of different

influenza subtypes, whereas A1 and A2 assays are specific for influenza subtypes H7N7 and H3N8,

respectively. Sequencing of amplicons obtained in the A2 assay yields information on antigenic

regions of the haemagglutinin molecule, and use of this procedure in the routine surveillance of

equine influenza will enable tentative characterisation of circulating viruses despite difficulties in

isolating field strains of the H3N8 subtype. The A1 assay will be useful in ascertaining whether

viruses of the H7N7 subtype still circulate amongst horses, or whether these are extinct. # 1999

Elsevier Science B.V. All rights reserved.

Keywords: Equine in¯uenza; Virus; Horse; Nucleic acids diagnosis-viruses; PCR

Veterinary Microbiology 67 (1999) 161±174

* Corresponding author. Tel.: +46-18-4714030; fax: +46-18-4714572; e-mail: [email protected]

0378-1135/99/$ ± see front matter # 1999 Elsevier Science B.V. All rights reserved.

PII: S 0 3 7 8 - 1 1 3 5 ( 9 9 ) 0 0 0 4 1 - 3

1. Introduction

A number of outbreaks of severe disease caused by influenza viruses have been

reported during recent years, highlighting the importance of rapid and efficient

identification of viral strains and their origins. Since the circulating flora of influenza

virus in a population has the potential to change through the introduction of viruses or

individual viral genes from other species (Guo et al., 1992; Subbarao et al., 1998; Brown

et al., 1998), most notably birds, methods used in surveillance must have a broad

specificity allowing detection of antigenically and genetically diverse members of the

genus influenza A. Less dramatic changes are seen both in human and equine influenza

viruses. Minor changes in the surface glycoproteins allow the circulation of significantly

antigenically altered viruses and necessitate the exchange of virus strains used in vaccine

production on a regular basis. Additionally, in the case of both the H1N1 and H3N2

influenza viruses of humans, strains with differing receptor-specificity have been shown

to co-circulate (Morishita et al., 1996; Ciappi et al., 1997).

Although large numbers of cases of H3N8 influenza virus infection have been

diagnosed serologically in horses during the past years, attempts to isolate the virus by a

number of laboratories have met with limited success (Leif Oxburgh, unpublished data

and Berndt Klingeborn, personal communication). This finding suggests that currently

circulating strains of the H3N8 subtype do not replicate readily in culture systems in

routine use, and indicates that use of an alternative technique to virus isolation (VI) in the

surveillance of equine influenza virus is necessary.

In contrast to the H3N8 subtype, the H7N7 subtype has not been isolated from the

horse for over 20 years. Serological evidence of the virus has been presented (Mumford

and Wood, 1993), but it is unclear whether these responses represent post-infection titres,

post-vaccination titres or cross-reactivity with another pathogen. A technique for rapid

identification of the virus in clinical samples would facilitate screening in order to

ascertain whether this subtype still is in circulation, or whether it can be removed from

existing vaccines.

In conclusion, there is a need for a diagnostic procedure which has both the redundancy

required to detect diverse subtypes of influenza virus, and the specificity to discriminate

between these different subtypes. It has previously been shown that the reverse

transcription-polymerase chain reaction procedure (RT-PCR) can be used to detect

influenza virus in nasal swab material from horses (Donofrio et al., 1994). We have

therefore chosen to modify this technique for use in the routine surveillance of equine

influenza virus. As shown by Ilobi and colleagues (Ilobi et al., 1998), the product of the

PCR assay can be sequenced in order to obtain information regarding antigenicity and

receptor-binding affinity of viruses without having to isolate and passage field strains for

subsequent purification of RNA and sequencing. Information of this type is important in

the routine surveillance of equine influenza virus, since significant changes at antigenic

sites on the viral haemagglutinin leading to antigenic drift necessitate the updating of

vaccine strains.

In this paper we report the development of a RT-PCR protocol for use in the laboratory

diagnosis of influenza viruses of equine origin. The procedure consists of three steps; (1)

a general PCR detecting the nucleoprotein gene, capable of detecting both avian and

162 L. Oxburgh, AÊ . HagstroÈm / Veterinary Microbiology 67 (1999) 161±174

equine virus strains, (2) a PCR specific for the H3 (A2) equine virus, and (3) a PCR

specific for the H7 (A1) equine virus. This protocol enables laboratory diagnosis of

infection within one day and distinguishes between the two different subtypes of equine

influenza virus. This system can be further modified to discriminate between different

avian and mammalian strains by addition of primer-pairs specific for evolutionarily

divergent genes.

2. Materials and methods

2.1. Abbreviations

Abbreviations used to denote virus strains are shown in Table 1.

2.2. Virus stocks

Stocks of viruses SKA/88, PRA/56, LON/73, MIN/84, SHA/89, H5 and AIC/68 were

obtained from the repository at the National Veterinary Institute. All viruses were

cultured in embryonated hen's eggs, and were used in the experiments without further

purification from allantoic fluid. Haemagglutination titres of the viruses ranged from

1 : 32 to 1 : 256 assayed with 1% chick erythrocytes.

2.3. Clinical samples

Clinical samples consisting of nasal swabs collected from horses were obtained from

the routine diagnostic laboratory at the National Veterinary Institute. Ten samples which

had previously been tested positive by immunofluorescent staining (IF, see below) were

chosen as positive controls (SW1±10), five samples from Iceland where equine influenza

Table 1Influenza viruses used in this study

Virus Strain Abbreviation Subtype Genbank accession number

Haemagglutinin Nucleoprotein

A/equi 2/Skara/88 SKA/88 H3N8 Y14053 ±

A/equi 2/Miami/63 MIA/63 H3N8 M24719 ±

A/equi 2/Solvalla/79 SOL/79 H3N8 Y14054 ±

A/equi 2/SoÈderala/94 SOD/94 H3N8 Y14058 ±

A/equi 2/BollnaÈs/96 BOL/96 H3N8 Y14060 ±

A/equi 1/Prague/56 PRA/56 H7N7 X62552 M63748

A/equi 1/London/73 LON/63 H7N7 X62560 M30750

A/swine/Bavaria/76 BAV/76 H1N1 ± ±

A/human/Aichi/2/68 AIC/68 H3N2 V01085 ±

A/human/Shanghai/16/89 SHA/89 H3N2 AF008668 L07372

A/mink/Sweden/84 MIN/84 H10N4 M21646 M24454

A/chicken/H5N4 H5 H5N4 ± ±

L. Oxburgh, AÊ . HagstroÈm / Veterinary Microbiology 67 (1999) 161±174 163

has never been shown to circulate were chosen as negative controls (ISW1±5), and 12

unknown samples from horses displaying symptoms of respiratory infection were chosen

(RSW1±12).

2.4. Analysis of the influenza virus content of the samples

In order to determine the virus content of nasal swabs, the samples were subjected to

VI and IF. In cases where both of these assays were negative, paired acute and

convalescent sera from sampled horses were tested by haemagglutination inhibition (HI)

assay in order to assess whether the animals had been infected with equine influenza

virus.

2.4.1. Virus isolation

Nasal swabs were resuspended in 0.5 ml PBS supplemented with 0.5% penicillin,

0.01% streptomycin and 0.00025% fungizone. 100 ml of this fluid was inoculated into the

allantoic cavity of two embryonated hen's eggs (9±10 days old). After 48 h incubation at

378C, allantoic fluid was harvested, and inoculated undiluted into the allantoic cavities of

embryonated hen's eggs. After three passages, allantoic fluid was assayed for the

presence of virus by haemagglutination assay with 1% chick erythrocytes.

2.4.2. Immunofluorescent staining

Cells in the fluid extracted from nasal swabs were sedimented by low-speed

centrifugation, the supernatant was removed, and the pellet was smeared onto a glass

slide. Fixation with methanol and subsequent immunostaining were performed according

to the protocol described by Harlow and Lane (Harlow and Lane, 1988). Polyclonal rabbit

antiserum raised against strain VIS/90 was used as the primary antibody, and FITC-

labelled anti-rabbit immunoglobin was used as the secondary antibody. Stained slides

were examined using a UV microscope, and the presence of fluorescing cells and cell-

associated material was determined.

2.4.3. Haemagglutination inhibition

In the HI test, sera were assayed for antibodies against the most recently isolated strain

of H3N8 equine influenza virus (BOL/96) using the method described by Klingeborn and

colleagues (Klingeborn et al., 1980). Serum samples were collected on two occasions

from the same animals; (i) simultaneously with the nasal swab, and (ii) 10±14 days after

the initial sampling. A fourfold increase in HI titre between the first and the second

sample was interpreted as indicating that the horse had been infected with equine

influenza virus during or immediately prior to the sampling period.

2.5. RNA purification from nasal swabs and virus stocks

Viral RNA was purified from nasal swab material and allantoic fluid using Promega's

SV Total RNA Purification Kit (Promega Corporation), essentially according to the

manufacturer's instructions. The kit is based on disruption of cellular and viral proteins


by treatment with guanidine thiocyanate, binding of the liberated nucleic acid to a silica

matrix and treatment with DN:ase. Neutralisation of DN:ase activity and several washing

steps ensure that no DN:ase is carried over to the purified RNA preparation. Briefly, 30 ml

of material was suspended in lysis buffer containing guanidine thiocyanate, heated to

708C for 3 min, centrifuged, and the supernatant bound to the silica matrix packed in a

minicolumn. After washing, nucleic acid bound to the column was treated with DN:ase

for 15 min at room temperature, DN:ase activity was neutralised by addition of guanidine

thiocyanate, and the bound RNA was washed twice. Finally, RNA was eluted in 100 ml

DEPC-treated dH2O, and the RNA was immediately frozen in liquid nitrogen in 20 ml

aliquots.

2.6. Primer design

Nucleotide sequences of viral genes shown in Fig. 1 were retrieved from the NCBI

database, and were aligned using the Megalign program of the DNASTAR software package

(DNAstar Inc.). After consensus and non-consensus regions were identified as putative

stretches for primer synthesis, the individual genes sequences were processed using the

OLIGO 4.05 software package (National Biosciences) to identify suitable sequences for use

as oligonucleotides in RT-PCR. The oligonucleotide primers shown in Fig. 1 were

subsequently commercially synthesised (DNA Technology AS, Aarhus, Denmark).

2.7. Nested RT-PCR

A nested RT-PCR protocol was designed to optimise sensitivity in order to be able to

amplify small quantities of genetic material present in swabs. A modification of

Promega's Access RT-PCR was used for first-strand synthesis and the first PCR step. The

second (nested) PCR was carried out using a standard protocol. Briefly, a mix containing

3 ml 5x Access reaction buffer, 1 ml reverse primer (to a final concentration of 1 pmol/ml),

1 ml (5 u) Tfl DNA polymerase and 15 ml dH20 was sealed into the bottom of a Hot-Start

tube (Molecular BIO-products) under a layer of wax by melting the wax bead at 808C for

3 min, then rapidly cooling on ice. A mixture containing 7 ml 5x Access reaction buffer,

1 ml dNTP (to a final concentration of 0.2 mM each of dATP, dCTP, dGTP and dTTP),

4 ml MgSO4 (to a final concentration of 1.92 mM), 1 ml forward primer (to a final

concentration of 1 pmol/ml) and 1 ml (5 u) AMV reverse transcriptase was layered on top

of the solidified wax. Two drops of mineral oil were added to prevent evaporation of the

sample during the reverse transcription step. An aliquot of purified RNA was rapidly

thawed and denatured for 15 min at 658C. After snap-cooling on ice, 16 ml of the RNA

solution was added through the mineral oil to the reverse transcription mix. The tube was

subsequently transferred to a thermal cycler (Minicycler, MJ Research) and the following

program was run; 488C for 45 min, 948C for 3 min, then 20 cycles of 948C for 30 s, 528Cfor 1 min and 688C for 2 min. For the NP and A2 reactions, 1 ml of the product of this first

RT-PCR step was used as template in the second PCR. For the A1 reaction, the first RT-

PCR mix was diluted 10 times in dH2O before transfer of a 1 ml aliquot to the second

PCR. This dilution step is necessary in order to minimise appearance of bands caused by

carry-over of primers from the first RT-PCR.


Fig. 1. Primer design for the three nested PCR assays described in this paper. An alignment of influenza virus genes spanning the region that was used for

oligonucleotide synthesis is shown. Sequences corresponding to primers used in the assays are underlined. The DNA sequence is in mRNA sense, with arrows

pointing downstream denoting forward primers, and arrows pointing upstream representing reverse primers. The sequences of reverse primers used in the assay

are the reverse complement of those underlined in the figure. (I) Primer set for the NP RT-PCR, (II) primer set for the A1 RT-PCR, (III) primer set for the A2

RT-PCR; this assay is semi-nested, with the same reverse primer being used in both the first PCR and the nested PCR.

16

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For the nested PCR reaction three different MgCl2 concentrations were used. For the

NP assay, 5 ml Promega Mg-free reaction buffer, 3 ml MgCl2 (to a final concentration of

1.5 mM), 1 ml dNTP (to a final concentration of 0.05 mM) and 1 ml each primer (to a final

concentration of 1 pmol/ml) were mixed with 1 ml of template. For the A1 assay, 5 ml

Promega Mg-free reaction buffer, 4 ml MgCl2 (to a final concentration of 2 mM), 1 ml

dNTP (to a final concentration of 0.05 mM) and 1 ml each primer (to a final concentration

of 1 pmol/ml) were mixed with 1 ml of template. For the A2 assay, 5 ml Promega Mg-free

reaction buffer, 2 ml MgCl2 (to a final concentration of 1 mM), 1 ml dNTP (to a final

concentration of 0.05 mM) and 1 ml each primer (to a final concentration of 1 pmol/ml)

were mixed with 1 ml of template. The tubes were transferred to a thermal cycler, heated

to 948C and put on hold at this temperature. 5 ml diluted Promega Taq polymerase

containing 2.5 u enzyme was added to each tube at this denaturing temperature. The

thermal cycle was subsequently initiated. Thermal cycle programs used for each of the

assays consisted of 3 min of 948C denaturation followed by 32 cycles of 948C for 1 min,

annealing for 1 min and 728C for 1 min. The cycle was terminated with an elongation

step of 728C for 10 min. Annealing temperatures for the three assays were; 508C for the

NP and A1 assays and 608C for the A2 assay.

2.8. Analysis of the nested PCR product

5 ml of each PCR product was run on a 2% agarose gel in a TAE buffer system and

subsequently stained with ethidium bromide before viewing on a UV lamp. A 100 base

pair ladder (Boehringer Mannheim Scandinavia AB) was used as a molecular weight

marker. The marker contains highlighted bands, with increased intensity at 500, 1000 and

1500 base pairs.

2.9. Precautions to minimise contamination risks

Sample purification and RT-PCR reaction preparation, second PCR step reaction

preparation and agarose gel analysis were all performed in separate laboratories. All

solutions were aliquoted to volumes suitable for one set of 10 reactions, and each aliquot

was discarded after one use. Negative (dH2O) controls were run with each assay,

comprising every fifth sample in each series. No contamination was detected at any

time.

2.10. Dilution series of virus template to assess the sensitivity of the assays

Titration of egg infectious doses (EID) for SKA/88 and PRA/56 was performed by

inoculation of 10-fold dilutions of these virus stocks into 10-day embryonated hen's

eggs. Allantoic fluid was harvested after 2 days incubation, and the end-point of the

titration was determined by haemagglutination assay with 1% chick erythrocytes. The

lowest concentration of inoculum yielding growth in eggs was designated 1 EID. Titrated

virus was diluted in negative nasal swab fluid so that 30 ml would contain 1 EID, 10 EID

or 100 EID. These samples were subsequently assayed in the same way as the swab

samples.


2.11. Sequence analysis

PCR products generated by each of the assays were analysed by automated sequencing.

SKA/88 PCR products from the NP and A2 assays and the PRA/56 product from the A1

assay were purified using Promega's Wizard PCR preps, and sequenced using the ABI

sequencing kit and the second forward PCR primer. Sequence data was aligned with the

corresponding sequence from the NCBI database to assess homology.

3. Results

3.1. All nested primer sets amplify targets specifically

Oligonucleotide primers were designed to amplify motifs in the influenza virus

genome with differing evolutionary conservation (Fig. 1). The forward primer of each

first PCR step primer pair was used to reverse transcribe viral RNA prior to the first PCR

reaction. The NP primer set amplifies a 241 base pair fragment from each of the viruses

which have been assayed (Fig. 2). These viruses are evolutionarily distantly related and

are isolated from different animal species. The A1 primer set specifically amplifies a 327

base pair fragment from the H7N7 equine influenza viruses. No PCR product can be seen

with any other template. The A2 primer set amplifies a 522 base pair fragment from the

H3N8 equine influenza virus, and does not show any positive result with the other

templates. It can be concluded that the three RT-PCR assays can be used in combination

to identify diverse subtypes of influenza virus, and subsequently to discriminate between

the two subtypes of equine virus.

3.2. The amplified fragments are specific for influenza virus

Sequencing of amplicons from each of the assays reveals a 99±100% homology with

the sequence predicted to be amplified by that primer set (data not shown). SKA/88

Fig. 2. RT-PCR assay of a selection of different viruses to demonstrate the specificities of the three assays.

Assays were performed as described in Materials and methods. Virus used was unpurified allantoic fluid.


amplicons were sequenced in order to verify the NP assay and A2 assay, and the PRA/56

amplicon was sequenced to verify the A1 assay. One single, non-coding substitution

could be seen in the SKA/88 A2 amplicon. From the sequencing data we conclude that

the assays amplify influenza genes specifically and that there is no background generated

by priming of sequences other than those from influenza virus in the template.

3.3. The assay system displays sensitivity similar to VI

Owing to the difficulties in isolation of field strains of the H3N8 subtype (see above

and Table 2), and the lack of clinical material from horses infected with the H7N7

subtype, we chose to compare the detection limit of the RT/PCR assays with VI

indirectly. Egg-adapted strains of both subtypes were used for this purpose. Purification

and amplification of serial dilutions of allantoic fluid containing defined titres of virus

mixed with negative nasal swab material were performed in order to assess sensitivity

(Fig. 3). The NP assay was found to amplify as little as 1 EID of virus, whereas the A1

and A2 assays showed a lower sensitivity, amplifying 10 EID of virus. This demonstrates

the sensitivity of the method, suggesting that it is similar to VI.

3.4. Assay of clinical samples demonstrates the utility of the system for diagnostic use

3.4.1. Assay of positive and negative control samples

Ten nasal swab samples which had previously been tested positive by IF (SW1±10),

and five samples from horses tested negative for antibodies by IF and HI (ISW1±5) were

used as positive and negative controls, respectively. Virus culture in embryonated hen's

eggs was attempted from each of the positive swab samples without result. VI and IF

were performed directly on arrival of the samples at the laboratory. The samples were

Fig. 3. RT-PCR demonstrating the sensitivity of the assay. Egg-grown virus was titrated as described in

Materials and methods. Defined amounts of virus corresponding to egg infectious doses (EID) of 1, 10 and

100 were assayed in the RT-PCR.


subsequently stored at ÿ708C for 2±6 months prior to RT/PCR assay. Since there has not

been any circulation of equine H7N7 equine influenza virus in Sweden for over 20 years

(Berndt Klingeborn, personal communication), it was not possible to obtain clinical

material for assessment of the A1 assay. Instead, 100 EID of viruses PRA/56 and LON/73

in allantoic fluid were mixed with negative nasal swab material in order to simulate

clinical material.

As shown in Table 2 and Fig. 4, all samples from infected horses tested positive in the

NP assay, without amplification of template from non-infected horses. As expected, none

Table 2Comparison of results of three different assays for the detection of influenza virus in nasal swab samples

Sample NPa A1b A2c VId IFe HIf

SW1g � ÿ � ÿ � NDh

SW2 � ÿ � ÿ � ND

SW3 � ÿ � ÿ � ND

SW4 � ÿ � ÿ � ND

SW5 � ÿ � ÿ � ND

SW6 � ÿ � ÿ � ND

SW7 � ÿ � ÿ � ND

SW8 � ÿ � ÿ � ND

SW9 � ÿ � ÿ � ND

SW10 � ÿ � ÿ � ND

ISW1i ÿ ÿ ÿ ÿ ÿ ÿISW2 ÿ ÿ ÿ ÿ ÿ ÿISW3 ÿ ÿ ÿ ÿ ÿ ÿISW4 ÿ ÿ ÿ ÿ ÿ ÿISW5 ÿ ÿ ÿ ÿ ÿ ÿRSW1j � ÿ � ÿ ÿ �RSW2 � ÿ � ÿ � ND

RSW3 � ÿ � ÿ � ND

RSW4 � ÿ � ÿ ÿ �RSW5 ÿ ÿ ÿ ÿ ÿ ÿRSW6 � ÿ � ÿ � ND




RSW10 � ÿ � ÿ ÿ �RSW11 ÿ ÿ ÿ ÿ ÿ ÿRSW12 � ÿ � ÿ � ND

a RT/PCR assay for nucleoprotein.b RT/PCR assay for H7 haemagglutinin.c RT/PCR assay for H3 haemagglutinin.d Virus isolation in the allantoic cavity of 9±10-day embryonated hen's eggs.e Immunostaining of virus antigens in respiratory secretions.f Haemagglutination inhibition assay. A four-fold titre increase in paired sera taken with 10±14 days interval isscored as positive.g Positive control samples are designated SW.h Not Done.i Negative control samples are designated ISW.j Unknown samples are designated RSW.


of the swabs tested were positive in the A1 assay. The simulated samples containing

PRA/56 and LON/73, however, showed a positive result, demonstrating the ability of the

assay to identify the H7N7 viruses in the background of RNA present in swab material.

The A2 assay amplified sequences in each of the positive samples, but none of the

negative samples, showed a 100% correlation with IF results. There is a slight variability

in the intensity of bands in each of the three assays, presumably reflecting the abundance

of virus present in the swab. The high molecular weight band seen in some samples

corresponds to the amplicon from the first PCR reaction.

3.4.2. Assay of unknown samples

Twelve nasal swab samples sent in to the National Veterinary Institute for diagnosis

were assayed using the RT/PCR technique, in addition to IF and VI. In cases where

neither IF or VI were positive, HI assay was performed on paired serum samples to

ascertain whether the sampled horses had been infected with H3N8 influenza virus. The

results shown in Table 2 demonstrate a complete correlation of RT/PCR with IF in cases

where the IF result was positive. In total, four samples were negative by IF. Viral nucleic

acid could be detected in two of these using RT/PCR. These two samples were positive by

Fig. 4. RT-PCR of clinical samples. Ten positive samples (SW) were assayed as described in Materials and

methods. Cultured SKA/88 (H3N8 sybtype) virus was included as a control. A1 influenza virus strains

were not available as clinical samples and 100 EID virus in allantoic fluid diluted in a negative swab

sample were used instead. Negative samples (ISW) consisted of a series of nasal swabs collected from

horses in Iceland where equine influenza does not circulate. These horses were serologically tested in order

to confirm that they were free from infection. The results of the assay shown in the figure are tabulated

together with results of virus isolation, immunoflurescent staining and haemagglutination inhibition in

Table 2.


HI, indicating that infection had taken place in both horses, and that concentrations of

viral antigen in the samples were below the detection limit for IF. The other two IF-

negative samples were negative by all techniques tested, indicating that symptoms

displayed by these horses were not caused by equine influenza virus infection. VI was not

successful for any sample.

4. Discussion

The data presented in this paper demonstrate that the PCR can be a valuable tool in the

surveillance of equine influenza virus. Several aspects of our method make it suitable for

this purpose. First, the method is versatile, allowing amplification either of evolutionarily

conserved regions, giving an assay recognising many different subtypes of virus, or non-

conserved regions, giving a discriminating assay, identifying individual virus subtypes.

Since there are many influenza virus gene sequences in the genome database at this time,

it should be possible to tailor the method to discriminate between most influenza virus

subtypes. Secondly, the method is rapid. Identification of the infecting virus subtypes can

be performed within 1 day of receipt of clinical material, and more detailed information

can be obtained by sequencing of the amplicon (see below). Thirdly, the method does not

rely on the presence of viable virus in the swab sample, allowing diagnosis and

identification to be made on material which has been inactivated during transport, stored

for a long time and which contains virus which will not grow in the culture system used

for isolation at the laboratory. The fourth, which may be the most important aspect is that

the RT-PCR generates an amplicon which can be used for nucleotide sequencing of genes

coding for antigenic determinants of circulating influenza viruses.

Since equine influenza viruses undergo antigenic drift,(Burrows and Denyer, 1982;

Berg et al., 1990; Oxburgh et al., 1993, 1998; Daly et al., 1996), it is important to monitor

the variation in antigenicity of circulating viruses. The best method to this end is

serotyping using the HI assay with mono- and polyclonal antibodies. A major drawback

of this method, however, is that it necessitates isolation and culture of virus. As

demonstrated in this paper, VI is often unsuccessful. A tentative evaluation of the

antigenicity of a new strain could be made from sequencing of the haemagglutinin gene

segment amplified in this RT/PCR assay. Sufficient sequence data has been generated to

enable comparisons of antigenic regions of strains to be made, and major antigenic

changes could be identified.

The protocol described in this paper is, however, labour-intensive and costly, and

cannot be used to replace standard routine methods such as immunoassay of swab

material and serological testing using the HI test. It should, however, be incorporated as a

complement to these methods to facilitate the identification of viruses circulating in new

outbreaks, and as a rapid method for typing virus from outbreaks with unusual

characteristics. The finding by Donofrio and colleagues (Donofrio et al., 1994) that viral

RNA is present in the upper airways during a limited time, from day 3 to approximately

day 8 post infection suggests that pooled samples of nasal swabs from symptomatic

horses in the same stable could be assayed in order to make a certain laboratory dia-

gnosis of an outbreak. Any detailed studies of circulating virus genotypes such as


nucleotide sequencing would, however, have to be performed on samples from individual

horses.

This technique will be extremely useful in elucidating the epidemiology of the H7N7

viruses. The lack of isolations of this virus during the past years raises the question of

whether it could be extinct, and a detailed survey of samples using RT-PCR should give a

picture of whether this is actually a fact.

The versatility of this assay system enables it to be used in the identification of a broad

spectrum of influenza viruses. Addition of primer sets specific for other subtypes of the

virus will enable characterisation of an unknown influenza virus to be made from a

clinical sample within a day, and will give important information regarding the subtypes

of virus. This information will help in monitoring the epidemiology of influenza virus in

various animal species.

Acknowledgements

This work was made possible by a grant from the Swedish Horse Racing Board (ATG).

References

Berg, M., Desselberger, U., Abusugra, I.A., Klingeborn, B., Linne, T., 1990. Genetic drift of equine 2 in-

fluenza A virus (H3N8), 1963±1988: analysis by oligonucleotide mapping. Vet. Microbiol. 22(2/3), 225±

236.

Brown, I.H., Harris, P.A., McCauley, J.W., Alexander, D.J., 1998. Multiple genetic reassortment of avian and

human influenza A viruses in European pigs, resulting in the emergence of an H1N2 virus of novel

genotype. J. Gen. Virol. 79(12), 2947±2955.

Burrows, R., Denyer, M., 1982. Antigenic properties of some equine influenza viruses. Arch. Virol. 73(1),

15±24.

Ciappi, S., Azzi, A., Stein, C.A., De Santis, R., Oxford, J.S., 1997. Isolation of influenza A (H3N2) virus with

`̀ O''!`̀ D'' phase variation. Res. Virol. 148(6), 427±431.

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. J. Gen. Virol. 77(4), 661±671.

Donofrio, J.C., Coonrod, J.D., Chambers, T.M., 1994. Diagnosis of equine influenza by the polymerase chain

reaction. J. Vet. Diagn. Invest. 6(1), 39±43.

Guo, Y., Wang, M., Kawaoka, Y., Gorman, O., Ito, T., Saito, T., Webster, R.G., 1992. Characterization of a new

avian-like influenza A virus from horses in China. Virology 188(1), 245±255.

Harlow, E., Lane, D., 1988. Antibodies: a Laboratory Manual, Cold Spring Harbor Laboratory Press.

Ilobi, C.P., Nicolson, C., Taylor, J., Mumford, J.A., Wood, J.M., Robertson, J.S., 1998. Direct sequencing of the

HA gene of clinical equine H3N8 influenza virus and comparison with laboratory derived viruses. Arch.

Virol. 143, 891±901.

Klingeborn, B., Rockborn, G., Dinter, Z., 1980. Significant antigenic drift within the influenza equi 2 subtype in

Sweden. Vet. Rec. 106(16), 363±364.

Morishita, T., Nobusawa, E., Nakajima, K., Nakajima, S., 1996. Studies on the molecular basis for loss of the

ability of recent influenza A (H1N1) virus strains to agglutinate chicken erythrocytes. J. Gen. Virol. 77(10),

2499±2506.

Mumford, J., Wood, J., 1993. In: Proc. WHO/OIE meeting: consultation on newly emerging strains of equine

influenza, 18±19 May 1992, Animal Health Trust, Newmarket, Suffolk, UK, Vaccine 11(11), 1172±

1175.


Oxburgh, L., Akerblom, L., Fridberger, T., Klingeborn, B., Linne, T., 1998. Identification of two antigenically

and genetically distinct lineages of H3N8 equine influenza virus in Sweden. Epidemiol. Infect. 120(1),

61±70.

Oxburgh, L., Berg, M., Klingeborn, B., Emmoth, E., Linne, T., 1993. Equine influenza virus from the 1991

Swedish epizootic shows major genetic and antigenic divergence from the prototype virus. Virus Res.

28(3), 263±272.

Subbarao, K., Klimov, A., Katz, J., Regnery, H., Lim, W., Hall, H., Perdue, M., Swayne, D., Bender, C., Huang,

J., Hemphill, M., Rowe, T., Shaw, M., Xu, X., Fukuda, K., Cox, N., 1998. Characterization of an avian

influenza A (H5N1) virus isolated from a child with a fatal respiratory illness (see comments). Science

279(5349), 393±396.


a pcr based method for the identification of equine influenza virus from clinical samples

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