An immunisation strategy for the protection of cattle against alcelaphine herpesvirus-1-induced malignant catarrhal fever

Vaccine 26 (2008) 4461–4468

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An immunisation strategy for the protection of cattle against alcelaphine herpesvirus-1-induced malignant catarrhal fever David M. Haig ∗ , Dawn Grant, David Deane, Iris Campbell, Jackie Thomson, Catherine Jepson 1 , David Buxton, George C. Russell Moredun Research Institute, Pentlands Science Park, Bush Loan, Penicuik EH26 0PZ, Scotland, UK

a r t i c l e

i n f o

Article history: Received 2 April 2008 Received in revised form 2 June 2008 Accepted 13 June 2008 Available online 2 July 2008 Keywords: Malignant catarrhal Fever Herpesvirus Vaccine

a b s t r a c t The aim of this study was to stimulate immunity in the oro-nasal-pharyngeal region of cattle to protect them from alcelaphine herpesvirus-1 (AlHV-1)-induced malignant catarrhal fever. Attenuated C500 strain AlHV-1 was used along with Freund’s adjuvant intramuscularly (IM) in the upper neck region to immunise cattle. Virulent C500 strain AlHV-1 was used for intranasal challenge. Nine of ten cattle were protected. Protection was associated with high levels of neutralising antibody in nasal secretions. Some protected animals showed transient low levels of viral DNA in blood samples and in one lymph node sample after challenge whereas viral DNA was detected in the blood and in lymph node samples of all animals with MCF. This is the most promising immunisation strategy to date for the control of malignant catarrhal fever. © 2008 Elsevier Ltd. All rights reserved.

1. Introduction Malignant catarrhal fever (MCF) is a fatal lymphoproliferative disease of cattle and other ungulates caused mainly by the ruminant gamma-herpesviruses alcelaphine herpesvirus 1 (AlHV-1) and ovine herpesvirus 2 (OvHV-2) [1]. These viruses have distinct geographical distributions and infect their reservoir hosts asymptomatically (wildebeest for AlHV-1 and sheep for OvHV-2) but often cause fatal lymphoproliferative disease when they infect susceptible hosts that include cattle, deer, bison, water buffalo and pigs. MCF is an important disease wherever reservoir and susceptible species mix, and is currently a particular problem in Bali cattle in Indonesia, bison in the USA (both OvHV-2 MCF) and in pastoralist cattle herds in eastern and southern Africa (AlHV-1 MCF) [1]. The genomes of both AlHV-1 and OvHV-2 have been sequenced revealing that they are highly related but distinct viruses [2,3]. They both induce MCF with a broadly similar pathology in susceptible animals and there are rabbit and hamster animal models of MCF caused by either virus [4–6]. Clinical signs of MCF include fever, inappetance, a generalised lymphadenopathy, a nasal/ocular discharge and erosions in the

∗ Corresponding author. Current address: School of Veterinary Medicine and Science, Nottingham University, Sutton Bonington LE12 5RD, UK. Tel.: +44 115 951 6464; fax: +44 115 951 6415. E-mail address: [email protected] (D.M. Haig). 1 Current address: Big DNA Ltd., Wallace Building, Roslin Biocentre, Roslin, Midlothian EH25 9PP, UK. 0264-410X/$ – see front matter © 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.vaccine.2008.06.056

mouth. Death can occur within a few days or up to several weeks after the onset of signs. The pathological changes also include vasculitis and interstitial accumulations of lymphoid cells in several tissues, which may be associated with tissue cytolysis, particularly at epithelial surfaces [7–9]. Despite the severe pathology of MCF, virus-infected cells in the tissues appear to be of low frequency [10]. However, recent results in OvHV-2-infected cattle, bison and rabbits suggest that virus-infected cells in MCF are more frequent [11] (D. Haig, unpublished data). These may be the source of large granular lymphocytes cultured from the tissues of MCF-affected animals that are >90% infected and indiscriminately cytotoxic for a variety of target cells [12–15]. Early attempts to immunise cattle using live or inactivated formulations of attenuated strains of AlHV-1 were either unsuccessful or gave equivocal results, providing no clear protection against either parenteral/systemic experimental challenge with virus or natural challenge with virus, despite the development of virus-neutralising antibodies in the serum [16,17]. Later work in rabbits showed that inactivated cell-free virulent AlHV-1 C500 strain could induce partial protection against a cell-free virus challenge systemically [18] but in another study, immunisation with live cell-associated virus protected rabbits from cell-associated virus given systemically whereas inactivated cell-associated virus immunisation did not [19]. Observations on the small numbers of immunised cattle that survived an initial experimental virus challenge suggested that induced immunity was short-lived while cattle surviving a natural infection remained immune despite having lower titres of serum neutralising antibody than immunised

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animals [16,17]. These results suggested that serum neutralising antibody was not a critical component of a protective immune response in cattle and raised the question of what the protective response might be. Work on cellular immunity to MCF virus has been hampered by the lack of a good experimental system in which animals can be immunised and challenged, and by the severe T cell hyperplasia induced by MCF virus which is a central part of disease pathology [20,21]. Current control measures for MCF rely on preventing susceptible livestock contacting carrier animals. As this is not a robust control measure for MCF, there is a need for a vaccine. In this study we wished to test the hypothesis that a protective immune response can be induced in cattle immunised with attenuated AlHV-1 and that this can prevent MCF in cattle subsequently challenged intranasally (IN) with virus, representing a natural route of infection. AlHV-1 was used as, unlike OvHV-2, free virulent and attenuated virus can be obtained from tissue culture and used to challenge cattle and immunise them, respectively [16,18]. 2. Materials and methods 2.1. Animals Disease-free and OvHV-2 seronegative male Friesian-Holstein cross-calves between 3 and 5 months of age were used in the experiments. The animal experiments were carried out with the approval of the Moredun Research Institute’s experiments and ethics committee and complied fully with the Home Office of Great Britain and Northern Ireland “Animals (Scientific Procedures) Act 1986”. Animals exhibiting a rise in rectal temperature >40 ◦ C for 2–4 days along with inappetance and ocular or nasal discharge were euthanized with an overdose of intravenous sodium pentobarbitone. 2.2. Immunisation study design In preliminary experiments 1 and 2, an attempt was made to stimulate a mucosal antibody response in the oro-nasal-pharyngeal region (detected as virus-neutralising antibody in nasal secretions) by IN immunisation. Over the two experiments, performed at different times, five groups of two cattle per group were immunised/treated as follows: group 1, IN prime immunisation with attenuated AlHV-1 [107.3 to 107.5 tcid50 /ml virus, where tcid50 /ml is the tissue culture infective dose of virus giving a cytopathic effect (CPE) in 50% of wells containing cultured fibroblasts] adsorbed to the mucosal adjuvant chitosan (Sigma, Poole, UK, 0.2% (w/v) in sodium acetate buffer pH 7.5). After 2 weeks, an IN boost immunisation was given, as for priming. Group 2 received chitosan IN prime and boost on the same schedule without virus (control group). Group 3 received IN prime and boost with attenuated AlHV1 (107.2 tcid50 virus) in cholera toxin adjuvant (from vibrio cholerae Inaba 569B strain, List Biological Labs, Campbell, CA, USA, 10 ␮g/ml, based on previous dose–response studies in cattle with BoHV-1 [22]). Group 4 received IN prime & boost application of cholera toxin without virus (control group). Group 5 received medium only (no adjuvant, no virus). Experiments 3 and 4 involved groups of calves immunised and challenged as shown in Table 1. Intramuscular (IM, upper neck) immunisation was with attenuated AlHV-1 in complete Freund’s adjuvant (CFA, Sigma, Poole, UK) and for IM boost immunisation, virus in incomplete Freund’s adjuvant (IFA, Sigma). In all experiments, blood and nasal secretions were collected from the animals just prior to primary immunisation/primary treatment and every 2 weeks thereafter. Blood plasma was obtained from heparinised blood and heated at 56 ◦ C for 10 min to inactivate

complement, then stored at −20 ◦ C prior to antibody analysis. Nasal secretions were collected by placing a standard tampon into a nostril of each animal for 10 min. The tampons were then compressed in a 10-ml syringe to expel secretions. The secretions were clarified by centrifugation at 1700 × g for 10 min at 4 ◦ C, the supernates collected and stored at −20 ◦ C. At autopsy, the following tissues were collected: brain, buccal mucosa, rumen, reticulum, liver, kidney, urinary bladder, mesenteric lymph node (MLN) and blood. Standard blocks of each tissue (except blood) were fixed in 10% formal saline before processing and embedding in paraffin wax. Sections, 4-␮m thick, were then cut and stained with haematoxylin and eosin. In addition, blood buffy coat cells and freshly isolated pieces of MLN were analysed for the presence of viral DNA (cell extract) or were cultured for virus isolation as described below. 2.3. Viruses The C500 strain of AlHV-1 was originally obtained from an AlHV1-infected ox in Kenya and passaged in New Zealand White rabbits [16,23]. Virus was cloned from the spleen of an infected rabbit by limiting dilution, purified by sucrose density ultracentrifugation and maintained in rabbits [23]. Virulent C500 AlHV-1 (i.e. that causes MCF when inoculated into rabbits or cattle) used for virus challenge in this study was obtained by collection of virus from cultures of bovine turbinate (BT) cells (seeded with infected rabbit lymphoid cells) below pass 5 (low pass virulent virus). At this stage the preparation is a mixture of cell-free and cell-associated virus [16,18]. Culture supernates containing virus were collected and additional virus obtained after a freeze/thaw cycle of the cells. Virus was purified and tested for contamination by bovine viral diaorrhea virus by the Moredun Virus Surveillance Unit. Propagation of C500 virus in BT cells for >30 serial passages resulted in a virus preparation where virus was largely cell-free and non-pathogenic for animals [23]. This cell-free virus was obtained from BT culture supernates, purified and checked for contaminating virus as described above, then used as the source of attenuated virus in the immunisation schedule (see below and Table 1). Highpass attenuated virus and low-pass virulent virus were prepared to coincide with immunisation dates and challenge dates, respectively to avoid freeze–thaw of the virus and loss of infective virus titre. Tcid50 /ml virus titres were obtained retrospectively with respect to the immunisation or challenge dates, using an aliquot of the virus used for immunisation or challenge. For group 3 animals (Table 1), attenuated virus was inactivated by treatment with 1% formalin at room temperature for 18 h. The treated virus did not form a CPE in BT cells in culture, and was therefore nonviable. 2.4. Virus detection: viral DNA and the virus-neutralising antibody test Viral DNA was assayed in pure genomic DNA samples extracted from fresh tissues (blood and MLN) using both quantitative (q)PCR and the more sensitive nested PCR as described previously [24,25]. These tests detected both attenuated and wild-type challenge viruses. For virus isolation, single cell suspensions were seeded on monolayers of BT cells and cultures given fresh medium twice weekly and analysed for virus-induced CPE. The virus neutralisation test was based upon inhibition of AlHV1-induced CPE in BT cells by dilutions of blood plasma or nasal secretion fluid as described [26]. BT cells were maintained in Iscove’s Modified Dulbecco’s Medium (IMDM) supplemented with 10% FBS, 1 mM l-glutamine and 100 Units/ml penicillin and strep-

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Table 1 Immunisation study groups and treatmentsa Group (n) c

1 (10) 2 (6) 3 (5) 4 (6) 5 (5) 6 (6) 7 (7) 8 (4) 9 (4)

Primary immunisationb (week 0) 6.2/6.5

c

10 virus IM in CFA 106.8 virus IM in CFA 6.5 10 Inact. virus IM in CFA 106.2 virus IM in CFA 106.8 virus IM in CFA CFA IM CFA IM PBS IM PBS IM

Immunisation boostb (week 4) 7.3/6.4

c

10 virus IM in IFA 106.5 virus IN in chitosan 6.4 10 Inact. virus IN in chitosan 107.3 virus IM in IFA 106.5 virus IN in chitosan IFA IM IFA IM PBS + chitosan IM PBS + chitosan IM

Virus challenge (week 10) 104/4.5 tcid50 virus INc 104 tcid50 virus IN 104.5 tcid50 virus IN 104.5 tcid50 virus IV 104.5 tcid50 virus IV 104 tcid50 virus IN 104.5 tcid50 virus IV 104 tcid50 virus IN 104.5 tcid50 virus IV

a The virus used for the priming and boost immunisations was attenuated AlHV-1 (from high pass in culture) unless otherwise stated (Inact = inactivated). The challenge virus was virulent AlHV-1 obtained from low pass tissue culture. b Immunisation doses of virus shown as tcid50 titres. c Group 1 consisted of two separate experiments (n = 4; n = 6) where animals were treated in the same way (group 1 tcid50 virus titres for n = 6 and n = 4 animals, respectively are shown) except that the group of four animals were euthanized at 12 weeks after challenge to look for any evidence of virus infection post-mortem. The other animals (n = 6), were kept and monitored for a further 42 weeks prior to euthanasia. CFA = complete Freund’s adjuvant, IFA = incomplete Freund’s adjuvant, IM = intramuscular injection (upper neck region), IN = intranasal challenge, IV = intravenous challenge.

tomycin. Assays were carried out in 96-well tissue culture plates with BT cells at greater than 80% confluence. The samples were added to wells of 96-well plates (in duplicate for each dilution) prior to adding attenuated AlHV-1 virus (100 tcid50 /(0.1 ml well)). A high titre bovine anti-AlHV-1 serum was used as a standard. After 4 h of culture at 37 ◦ C in a moist box, 105 BT cells/well were added and plates incubated at 37 ◦ C for 5–7 days. Titres were the reciprocal of the sample dilution that neutralised virus in 50% of the wells. Non-specific toxicity-control wells contained sample and cells without virus. 2.5. Immunoglobulin isotype of the anti-AlHV-1 neutralising antibody Nasal secretions (NSs) samples were first analysed for virusneutralising activity as described above. To define the isotype of virus-neutralising antibodies in NS, anti-isotype antibodies were used to specifically inhibit the immunoglobulin isotypes IgM and IgA and the IgG subtypes IgG1 and IgG2 that were present in the NS. Aliquots of anti-bovine IgM, IgA, IgG1 and IgG2 Mabs (AbD Serotec, Oxford, UK; all Mabs IgG1 subtype) supplied as tissue culture supernatants were diluted 1:1 in PBS and dialysed overnight against PBS to remove sodium azide before filter sterilisation. A dilution of NS corresponding to its virusneutralising titre was incubated with an equal volume of a 1:10 dilution of anti-isotype Mab for 2 h at 37 ◦ C. This dilution of antibody was determined from a dose–response analysis. AlHV-1 was then added and the mixture incubated for a further 2 h before adding 100 ␮l to BT cells in triplicate wells of 96-well plates. The assay plates were incubated for 7 days and analysed as described above for the virus-neutralising antibody assay. Wells contained virus at a final concentration of 100 tcid50 /well. In control wells, to measure any effect of the Mabs on the cell growth and virus infectivity, NS was replaced by medium. An isotype-matched (IgG1) anti-border disease Mab (VPM21) [27] was also included as a non-specific antibody control.

and readily palpated. Post-mortem examination confirmed these observations and in addition in the kidney there were small white foci in the renal cortex. The histopathological changes consisted of three types of related change. There was expansion of lymph nodes, vasculitis and interstitial accumulations of mononuclear cells, predominantly lymphoid in appearance. The latter in non-lymphoid tissues were often associated with tissue cytolysis, particularly at epithelial surfaces. Thus in the buccal papillae there were focal erosions of the mucosa with associated haemorrhage and inflammation, and changes in the rumen and reticulum were generally more severe. In the kidney there were multiple well-defined foci of lymphoid cell accumulations, with or without associated vasculitis. In the renal pelvis, lymphoid inflammation associated with the transitional epithelium was sometimes seen, and similar lesions were also found in the urinary bladder in a number of cases. In the liver, prominent periportal lymphoid inflammation was consistently observed and lymph nodes were invariably enlarged due to expansion of the cortex and paracortex, and the follicles appeared activated. In the brain there was often a mild non-suppurative meningo-encephalitis in which instances of vasculitis were confined to the meninges. 2.7. Statistical analysis Data for post-challenge survival between groups were analysed by Fisher’s exact test [28] (www.matforsk.no/ola/fisher.htm) to compare experimental groups. The effect of challenge route on mean survival time within control groups was analysed by twosample t-test within Genstat (version 7; VSN International, Hemel Hempstead, UK). Virus neutralisation data were analysed by twosample t-test within Genstat. 3. Results 3.1. Preliminary experiments: challenge virus dose–response and intranasal immunisation with virus along with mucosal adjuvants

2.6. Pathology and histopathology In this study MCF was diagnosed by clinical and pathological criteria. The clinical symptoms included fever, nasal and ocular discharge, initially serous but progressing to catarrhal. There was conjunctivitis and development of peripheral opacity of the cornea. Close examination of the buccal papillae usually revealed small ecchymotic haemorrhages. The surface lymph nodes, such as the prefemoral, prescapular and submandibular nodes were enlarged

A preliminary dose–response experiment was performed using groups of three cattle to determine the optimal dose of virulent AlHV-1 for intranasal (IN) challenge of cattle. Doses of 104 tcid50 /ml and 103 tcid50 ml induced MCF in 100% of cattle within 2.5–6 weeks of challenge. 102 tcid50 /ml virus IN induced MCF in two animals at 4 and 5 weeks, respectively after challenge. A dose of ∼104 tcid50 /ml was therefore selected for use in the subsequent experiments involving virus challenge.

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Preliminary experiments 1 and 2 were an attempt to stimulate a mucosal antibody response detectable in nasal secretions after IN prime and IN boost with virus and the mucosal adjuvants cholera toxin or chitosan. After priming immunisation, virus-neutralising antibody was not detected in nasal secretions from any of the immunised animals. However, after IN boost (2 weeks after priming) with virus adsorbed to chitosan in the viruschitosan primed animals, low titres of virus-neutralising antibody were detected (titres of between 2 and 8) 2–4 weeks after boost. Virus-neutralising antibody was not detected in any virus-cholera toxin-boosted animal. This provided the background against which the immunisation and challenge experiments 3 and 4 were performed. 3.2. Prime and boost immunisation with attenuated virus in Freund’s adjuvant IM in the upper neck protects cattle from MCF after intranasal challenge with virulent virus Table 1 shows details of the immunisation and control groups in this study. Fig. 1 shows that nine of ten animals in group 1 (IM prime with attenuated virus in CFA and IM boost with attenuated virus in IFA) and three of five animals in group 2 (IM prime with attenuated virus in CFA and IN boost with attenuated virus adsorbed to chitosan adjuvant) survived a lethal challenge with AlHV-1 given by the intranasal route. Both immunisation regimes produced significant protective effects (P < 0.0005, group 1 vs. groups 6 + 8 and P = 0.02, group 2 vs. groups 6 + 8). Five of the group 1 animals were euthanized 12 weeks after challenge to look for signs of infection or disease. Four of the remaining five group 1 animals survived for a year after challenge, when they were euthanized to terminate the experiment. Control animals given adjuvant or PBS without immunising virus and challenged intranasally with virulent virus succumbed with MCF between 3 and 7 weeks after challenge, with the exception of one group 6 animal that succumbed to MCF 16 weeks after challenge. Group 4 animals, given the same immunisation regime that protected group 1 cattle against intranasal virus challenge, all succumbed with MCF when the virus was given systemically. These animals developed MCF over a similar time period as unimmunised control group animals given virus systemically and compared with groups 7 and 9, no statistical difference was found (Fig. 1). Immunisation with formalin-inactivated virus did not induce protection against MCF. Adjuvant alone had no MCF-protective effect. Finally, of the unprotected animals, those challenged intravenously with virus developed MCF earlier than those that were intranasally challenged (P = 0.03; groups 7 and 9 vs. groups 6 and 8; Fig. 1).

3.3. Protection and neutralising antibody titres in nasal secretions and blood plasma Fig. 2 shows the neutralising antibody titres in nasal secretions and blood plasma for groups 1–5. Titres were higher in plasma samples compared to nasal secretions for each group. The highest titres in both nasal secretions and blood plasma were obtained in animals that were primed and boosted intramuscularly with attenuated virus in Freunds’ adjuvant compared to those boosted intranasally (Fig. 2A and C. P < 0.001 for nasal secretion neutralising antibody titres and P = 0.002 for serum neutralising antibody titres immediately pre-challenge: groups 1 and 4 vs. groups 2 and 5). This was associated with protection in group 1 animals challenged intranasally but not in group 4 animals that were challenged with virus systemically and had all succumbed to MCF by 3 weeks after challenge (week 13 of the experiment, Fig. 2A and C). In group 1 animals, neutralising antibody titres in nasal secretions and blood plasma increased after boost immunisation. In addition, titres in nasal secretions rose after intranasal virus challenge (reaching a group highest titre 6–8 weeks after challenge (week 16–18 of the experiment, Fig. 2A)). However, a similar rise in titre was not seen in blood plasma samples from the same animals at this time. Group 2 animals developed the next highest neutralising antibody titres (group mean) in nasal secretions and blood plasma, followed by group 5 that received the same immunisation procedure as group 2. This was associated with the protection of three of five cattle from MCF after intranasal virus challenge (group 2), but not after systemic inoculation of virus (group 5). A more complete time course of antibody titres was not obtained in group 2 animals as survivors were euthanized 12 weeks after challenge to look for evidence of MCF. Immunisation with inactivated virus stimulated low levels of virus-neutralising antibody and no protection against disease. Significantly, neutralising antibody was not detected in either serum or nasal secretions from any of the control-infected animals in groups 6–9. This includes animal 632 that survived 129 days after challenge before the onset of MCF. 3.4. Neutralising antibody is contained predominantly within the IgA class and IgG1 subclass of bovine immunoglobulins Nasal secretion material was analysed for IgM, IgA and IgG virus-neutralising antibody from five animals in group 1 that were protected against MCF (Fig. 3). For each of the individual NS samples, there was a significant difference (P < 0.01) between the effects of anti-IgA and anti-IgG1 treatment on virus neutralisation at all time points compared to the control antibody or no antibody

Fig. 1. Post-challenge outcome of the immunisation procedures. The relative timing of the onset of MCF in the different treatment groups is shown in the left panel while animals that survived for autopsy at the end of the experiment are indicated on the right (*). The symbols used for the treatment groups are as follows: group 1 (); group 2 (䊉); group 3 (); group 4 (); group 5 (); group 6 (); group 7 (); group 8 (♦); group 9 (). Abbreviations: Att, attenuated; Inact, inactivated; IM, intramuscular; IN, intranasal; IV, intravenous; chall, challenge.

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Fig. 2. Virus-neutralising antibody in the nasal secretions (A and B) and blood plasma (C and D) of animals in the treatment groups. Symbols used are as in Fig. 1. P = priming immunisation (week 0), B = boost immunisation (week 4), C = virus challenge (week 10). Titres are expressed as means for each group ± S.E.M. For group descriptions see Table 1.

control groups. There was no evidence of IgM virus-neutralising antibody and samples from only two animals (one on week 10 and another on week 14) showed some IgG2 virus-neutralising antibody activity (P < 0.02 for a comparison with the control groups). Furthermore, IgA virus-neutralising antibody predominated over

IgG1 virus-neutralising antibody in the early stages after immunisation (at 4 weeks, just prior to boost immunisation), whereas both contributed in roughly equal proportions on weeks 8, 10 and 12 after immunisation. By 14 weeks after immunisation (and 4 weeks after virus challenge), IgG1 virus-neutralising antibody predominated. Note that Fig. 3 shows the group means for all five animals and not individual animal sample results.

3.5. Protection was associated with a reduced establishment of virus in the blood

Fig. 3. Immunoglobulin isotype analysis of the virus-neutralising antibody in nasal secretions (NSs) of five group 1 animals protected against MCF. Mean cytopathic effect (CPE) and S.E.M. of triplicate samples tested from animals 119, 631, 830, 833 and 843. Cattle were immunisation boosted at 4 weeks after priming immunisation (week 0) and virus-challenged intranasally 6 weeks after the boost immunisation. Each of the NS samples was used at a dilution corresponding to its virus neutralisation assay titre (previously determined), in order to provide a near zero or low CPE background against which anti-immunoglobulin-isotype antibody inhibition of virus-neutralising antibody could be measured (as an increase in CPE compared to controls). Note that day 0 samples contained no virus-neutralising activity, therefore CPE was approximately 100%. The analysis is not quantitative but shows relative values for the sample group mean at each of the different time points. Control wells with virus were all 90–100% CPE on day 7 of the assay, and control wells with cells and reagents but no virus exhibited 0% CPE (i.e. no non-specific cytotoxicity).

To determine whether the immunisation strategy in group 1 had prevented infection and the establishment of a persistent (latent) state, nasal secretions and blood were analysed for AlHV-1 DNA during the course of the experiments and MLN (a sentinel tissue for infection) was analysed at autopsy. Although the PCR did not distinguish between attenuated virus used for immunisation and virulent virus used for challenge, viral DNA was not detected in any nasal secretion or blood sample collected at any time point after immunisation and prior to virus challenge. After virus challenge, Table 2 shows the result of the tissue survey for virus DNA. In unprotected groups, where all animals succumbed to MCF following either intranasal or intravenous challenge (groups 3–9), AlHV-1 was generally detected in blood prior to the onset of clinical signs (27 of 37 animals; Table 2). Blood samples were taken at 2-weekly intervals and this may have contributed to the lack of virus detection in animals that succumbed to MCF in the few days preceding blood sampling. Where animals survived long enough, all subsequent DNA samples were virus-positive and all animals in groups 3–9 had detectable virus DNA in lymph node tissue samples taken at post-mortem. In contrast, animals in groups 1 and 2 that were protected from MCF did not reliably show the presence of viral DNA in blood samples. Two of nine protected animals in group 1 had single blood samples that were positive for AlHV-1 DNA and one animal showed viral DNA in two consecutive blood samples.

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Table 2 Detection of AlHV-1 DNA in the tissues of immunised and control cattle by PCR after intranasal or intravenous virus challenge Groupa

Animalb

Virus DNA detectionc

Euthanizedd e

Blood

Nasal secretions

MLN (VI)

608 612 714 897 947

− − − − −

+ −

−(+) +(−) −(+) − −

1b

119 631 830 833 843

+ d28,35 + d81 − + d252 + d28

2

735 932 939 815 251

− − − + d21,28 + d21,35

− − −

713 736 905 933 944

+ d28 + d14 + d28 + d14 + d28

+ +

4

109 117 453 635 838 845

+ d7,14 + d7 − + d14 − −

5

640 707 906 941 946

+ d28 + d14 + d14,28 + d14,28 + d14

6

632 646 648 835 849 860

7

1a

With MCF d45

d86 d86 d85

− + − − − −(+) +(+) −

d378 d378 d378 d378 d378 d85 d29 d85 d82 d43

+(+) +(+) +(+) +(+) +(+)

d30 d29 d34 d27 d35

+ + + + + +

d24 d15 d14 d24 d17 d15

+(−) +(+) +(+) +(+) +(+)

d45 d24 d34 d29 d20

+ d7,35 + d7,21,35 + d21,35 − − + d21

+ + + + + +

d129 d37 d42 d35 d35 d35

121 636 641 831 832 840 852

− + d7 − + d7 + d7 + d21 + d7

+ + + + + +

d14 d14 d15 d15 d14 d24 d15

8

614 710 712 927

− + d28 + d14 + d28

+ + + +

d43 d31 d24 d30

9

709 926 929 943

+ d14 + d14 + d14 + d14

+ + + +

d16 d17 d17 d21

3

No MCF d86

+

− + − −

− − + +

Blanks indicate samples not available. a Experimental groups as follows: 1 (IM, IM; INchall); 2 (IM, IN; INchall); 3 (inactIM, IN; INchall); 4 (IM, IM; IVchall); 5 (IM, IN; IVchall); 6 (CFA, IFA; INchall); 7 (CFA, IFA; IVchall); 8 (med con x2; INchall); 9 (med con x2; IVchall). b Animals (identified by abbreviated ear-tag numbers) were sampled at approximately 2-weekly intervals in an alternating pattern with groups 1b, 4, 6, 7 beginning on week 1 and groups 1a, 2, 3, 5, 8, 9 beginning on week 2 post-immunisation. c Detection of AlHV-1 DNA in genomic DNA samples prepared from blood (positive on the post-challenge days indicated), nasal secretions (tested on day 15 post-challenge where available) or mesenteric lymph node samples taken at autopsy. d Animals were euthanized on the day post-challenge indicated, following the onset of clinical MCF or for autopsy in the absence of clinical signs of MCF. e Available results of virus isolation from MLN samples are given in brackets, where + indicates the development of cytopathic effect in BT cells overlaid with MLN single cell suspension.

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In each case no viral DNA was detected in subsequent blood samples (Table 2). One of these animals also had viral DNA detectable in lymph node tissue at post-mortem but no signs of MCF were detected by either gross or histological examination. Virus DNA was also detected in some nasal secretion samples (Table 2) but the variable yield of these samples and the primary requirement for analysis of the antibody responses meant that material for DNA purification was available only from a small number of animals in the trial. These samples appear to demonstrate, however, that AlHV-1 virus DNA could be found in nasal secretions 15 days after intranasal challenge in both protected and susceptible cattle. It was not possible to define the source of this DNA. 4. Discussion In this study, we have demonstrated that prime and boost immunisation of cattle intramuscularly in the upper neck with attenuated AlHV-1 emulsified in Freund’s adjuvant induced an immune response that protected nine of ten animals from MCF. Protection was associated with high titres of neutralising antibody in nasal secretions and blood plasma. To achieve this, we exploited the availability of low-pass cultured AlHV-1 as virulent, cell-free virus to challenge cattle by the intranasal route. This is presumed to be a natural route of virus infection. Furthermore, cell-free high-pass (in tissue culture) attenuated C500 AlHV-1 was readily harvested and used as a vaccine. These factors allowed for a robust virus immunisation and challenge procedure. In this study, we wished to test the hypothesis that stimulation of a neutralising antibody response in the oro-nasal-pharyngeal region of cattle would induce a mucosal immune barrier to infection. Although this study does not identify directly the nature of protective immunity, the strategy was effective in preventing disease. However, it did not prevent infection in all cases, although number of animals in the protected groups (1 and 2) with one or more virus-positive tissue samples (blood, or MLN at autopsy) were significantly lower than in other groups where animals succumbed to MCF (P = 0.02 for IN challenge groups 1 and 2 vs. groups 3, 6 and 8). This suggests, as one possibility, that the quantity of virus entering the body of groups 1 and 2 immunised animals was reduced compared with the other groups and that a fatal infection was consequently avoided, possibly through the action of a protective immune response that had time to develop in these animals. Virus was more readily detected in the blood and MLN of animals that succumbed to MCF. Indeed, all MCF-affected animals had detectable AlHV-1 DNA in the mesenteric lymph node at autopsy. It remains unclear whether MCF viruses go through the classical herpesvirus lytic and latent modes of replication in their susceptible hosts. AlHV-1 viral DNA in the blood appears to be cell associated and virus cannot be cultured from these cells. Infected cell lines can be established from lymphoid tissues of animals affected by MCF and, in the case of AlHV-1, infectious pathogenic virus may be obtained following subculture onto BT cells. However the status of the virus in the infected cells in vivo is unknown. Studies of OvHV-2-infected cell lines from cattle and rabbits suggest that the pattern of gene expression in MCF-susceptible species has aspects of both latent and lytic replication and, for OvHV-2 at least, no infectious virus can be produced [29]. Furthermore, neutralising antibody does not develop in MCF-affected animals [26], and control-infected animals in this study. This may be due to lack of availability of viral antigens and/or the severe, normally acute MCF pathology inhibiting an immune response. The fact that virus delivered by the IM route with Freund’s adjuvant was better at stimulating a neutralising antibody response in both blood and nasal secretions of cattle than the IN route with

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mucosal adjuvants was not expected, although the relative lack of efficacy of cholera toxin in cattle compared to some other species had been recorded [22]. We had anticipated that it would be necessary to stimulate mucosal-epithelial tissue in the nares directly at least once in an immunisation regime but the results presented here suggest this is not the case. The use of chitosan in an intranasal boost led to protection of four animals out of six (group 2; Fig. 1), suggesting that this approach may be worth optimising further. However, the lower success of this method may be partly attributed to difficulties in delivering a protective dose of vaccine to the correct site in the nasopharynx. Thus intramuscular immunisation may also have practical advantages. The neutralising antibody in nasal secretions was of the IgA class and IgG1 subclass predominantly, with a small contribution of IgG2 antibody in some animals. Dominance of IgA compared to IgG1 was noted early after immunisation, but this disappeared with time to be replaced by an IgG1 virus-neutralising antibody dominance in the later stage of the analysis. The reason for this is not known. Also not known is whether virus challenge had any effect on the IgA to IgG1 virus-neutralising antibody ratio. The study also demonstrated that formalin-inactivated virus was ineffective, compared to untreated virus given by the IM route in delivering protection against MCF. It is possible that the treatment had inactivated key antigens, as the neutralising antibody titres (in blood and nasal secretions) from animals in this group were lower than those of animals given attenuated virus (Fig. 2). In addition, there was no evidence (by PCR for viral DNA in the blood) that the attenuated virus used for immunisation had established as an infection in cattle. Indeed the possibility that a proportion of the virus was no longer live, having been inactivated by emulsification in Freund’s adjuvant, cannot be ruled out at this stage. There is a need to further investigate virus inactivation and the ability to stimulate immunity to MCF. Importantly, the immunisation strategy that successfully protected cattle from IN virus challenge did not protect cattle challenged with virulent virus given intravenously. This supports the suggestion that a mucosal barrier is an important aspect of host immunity as, in spite of high serum neutralising antibody in the IV challenge group, virus was able to infect these cattle and induce MCF. Neutralising antibody therefore appears to be ineffective against virulent AlHV-1 given systemically. This is in accord with a previous study [16], which showed that neutralising antibody was ineffective against cell-free AlHV-1 given systemically. Interestingly the pathology, including the distribution of tissues affected by MCF, was the same in all cattle with MCF, whether the virus was introduced by the IV or IN route. The main difference observed was in the time to onset of disease, which was significantly faster in IV-challenged animals. This indicated that the cell association of the virus given systemically does not affect the natural tropism and pathogenesis of MCF. In conclusion, this study has demonstrated that immunity to malignant catarrhal fever can be induced by the use of an attenuated virus vaccine administered by intramuscular injection. While the adjuvants used successfully here, CFA and IFA, are not licensed for use as veterinary vaccine components in the UK, their use in this study provides a baseline against which other adjuvants may be tested. Future experiments will be aimed at developing a vaccine formulation that uses licensed adjuvants but retains the significant efficacy reported here. It will also be important to investigate the mechanism of protection in more detail, by studying the local mucosal as well as systemic serological and cellular responses to identify protective immunity. Nevertheless, by identifying AlHV1 protective antigens recognised by nasal secretion antibody from immune animals, we should be able to identify equivalent antigens in OvHV-2 as potential vaccine candidates for OvHV-2 MCF.

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