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Immunoblotting analysis of the reaction of wildebeest, sheep and cattle sera with the structural antigens of alcelaphine herpesvirus-1 (malignant catarrhal fever virus)
Veterinary Microbiology, 19 (1989) 205-215 Elsevier Science Publishers B.V., Amsterdam - - Printed in The Netherlands
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Immunoblotting Analysis of the Reaction of Wildebeest, Sheep and Cattle Sera with the Structural Antigens of Alcelaphine Herpesvirus-1 (Malignant Catarrhal Fever Virus) ALAN HERRING, HUGH REID, NEIL INGLIS and IRENE POW
Moredun Research Institute, 408 Gilmerton Road, Edinburgh EH17 7JH (Gt. Britain) (Accepted for publication 3 October 1988)
ABSTRACT Herring, A., Reid, H., Inglis, N. and Pow, I., 1989. Immunoblotting analysis of the reaction of wildebeest, sheep and cattle sera with the structural antigens of alcelaphine herpesvirus- 1 (malignant catarrhal fever virus). Vet. Microbiol., 19: 205-215. Malignant catarrhal fever (MCF) is a disease of cattle and some other ruminants caused by alcelaphine herpesvirus-1 (AHV-1), a virus of wildebeest. The disease also occurs in the absence of wildebeest and is then thought to be caused by a viral agent harboured by the sheep. The structural proteins of AHV-1 have been used as antigens for the immunoblottinganalysis of sera from wildebeest, sheep and cattle infected by either AHV-1 or the "sheep-associated" form of the disease. Wildebeest sera showed a uniform response reacting strongly with six polypeptides. Sheep sera also gave positive results but individual sera reacted with varying subsets of the antigens recognized by wildebeest. These results support the earlier suggestion that sheep harbour a herpesvirus related to AHV-1. A bovine serum from a case of MCF caused by AHV-1 also reacted only with a subset of the six wildebeest-reactive polypeptides. Sera from cattle affected with the "sheep-associated" form of the disease gave reactions in only two of the eight cases tested; both positive sera reacted to a few polypeptides only.
INTRODUCTION
Malignant catarrhal fever (MCF), a fatal lymphoproliferative disease of cattle and other ruminants, occurs in two clinically and pathologically similar forms. One cause of the disease is alcelaphine herpesvirus-1 (AHV-1) which is carried by the wildebeest (Connochaetes taurinus) as an inapparent infection (Plowright et al., 1960; Plowright, 1965, 1968). Epidemiological observations of the other form of the disease strongly suggest that the source of infection is the domestic sheep and it is proposed that this species carries a virus biologically similar to AHV-1 (Rossiter, 1981; Reid and Buxton, 1984; Reid et 0378-1135/89/$03.50
© 1989 Elsevier Science Publishers B.V.
206 al., 1986). This form of the disease is termed "sheep-associated" MCF (SA MCF). Several other antelope of the subfamilies Hippotraginae and Alcelaphinae consistently have neutralizing antibody to AHV-1, implying that related viruses are prevalent in these two groups of African antelope (Reid et al., 1975; Mushi and Karstad, 1981; Hamblin and Hedger, 1984). No such neutralizing antibody can be detected consistently in sheep sera, although Rossiter (1981) found a reaction with all adult sheep sera examined using cells infected with AHV-1 in an indirect immunofluorescent (IIF) antibody test. However, the broadly cross-reactive nature of the IIF test, and the known property of some herpesvirus proteins to bind immunoglobulins (Westmoreland and Watkins, 1974), make interpretation of these results difficult. In order to assess the significance of the above observations, a qualitative comparison of the reaction of wildebeest and sheep sera to proteins prepared from purified virions of AHV1 was made employing the immunoblotting technique. In addition, sera from cattle with both forms of the disease were examined. MATERIALSAND METHODS
Sera Eighteen sheep sera were used. Seven of these came from sheep which had been housed with a group of red deer that developed MCF (Reid et al., 1979). The remainder were from normal adult sheep. The 11 wildebeest sera were kindly supplied by R. Kock from animals that had previously been kept at Whipsnade Park, England. The bovine serum from a calf with MCF induced by AHV-1 was kindly supplied by Dr. N. Edington, and the sera from cattle with SA MCF were from clinical cases referred to the Institute and confirmed histologically as MCF (Reid et al., 1986). The serum against bovine herpesvirus-1 (BHV-1) was derived from a cow which had been vaccinated with a live, temperature-sensitive vaccine (Zygraich et al., 1974) and subsequently challenged with virulent virus. The iodinated detection reagent used throughout was derived from a rabbit hyperimmune serum raised against ovine IgG and subsequently purified on an affinity column bearing ovine IgG Fab2 fragments. This rabbit IgG also reacted with immunoglobulins from bovine, cervine and alcelaphine sources. Class and subclass reactions have not been investigated but IgG is the major species recognized by this reagent in sheep serum.
Antigen The WC 11, cell-culture-adapted, cell-free strain of AHV-1 (Plowright et al., 1965) was propagated in monolayer cultures of bovine kidney cells grown in
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roller vessels. Supernatant fluids, which contained approximately 5 X 106 50% tissue culture-infective doses m l - 1, were harvested when 50% of the cells showed cytopathic effects. After clarification for 20 min at 9000 Xg and 4 ° C, virus was pelleted by centrifugation at 40 000 x g at 5 ° C. Pellets were resuspended in 8 ml of 1 x T N E buffer (10 mM Tris-HC1, 100 mM NaC1, 1 mM EDTA (diNa) pH 7.5 ), loaded onto a pre-formed 7.5 m115-40% w/w Metrizamide (Nyegaard U.K. ) gradient in 1 X TNE pH 7.5 and centrifuged at 114 000 X g for 60 min at + 4 ° C in a Beckman SW40Ti rotor. The visible band of virus which formed at the centre of the gradient was collected diluted to 12 ml in 1 X TNE pH 7.5 and repelleted at 114 000Xg for 40 min at 4°C. The resulting pellet was resuspended in 100 zl of TNE pH 7.5 and assessed for purity by inspection in the electron microscope at a magnification of 25 000 after negative staining with 1% (w/v) phosphotungstic acid. Protein content was estimated using the "BioRad" dye-binding assay (Bradford, 1976). BHV-1 virus was purified in essentially the same fashion except that a 10-40% (w/v) potassium tartrate gradient was employed.
Electrophoresis Most experiments were performed using block gels in which a single large well was loaded with viral protein and protein molecular weight standards were loaded in a small side well. A volume of viral suspension containing 200 ]lg of protein was mixed with an equal volume of 2 X concentration denaturing buffer to yield final concentrations of 63 mM Tris, 2% (v/v) fl-mercaptoethanol, 2% (w/v) sodium dodecyl sulphate (SDS), 10% (w/v) sucrose and 0.001% bromophenol blue. After heating in a 100°C waterbath for 90 s, the viral polypeptides were electrophoretically separated using a 10% discontinuously buffered polyacrylamide gel with a 3% stacking gel (Laemmli, 1970). Electrophoresis was performed under conditions of constant current (current density 1.9 mA cm-2) until the bromophenol blue tracker dye had migrated a distance of 9 cm in the resolving gel.
Immunoblotting Electrophoretic transfer of resolved viral polypeptides to nitrocellulose and the reaction of blotted strips with antibody was performed essentially as described by Burnette (1981) using the modifications of Herring and Sharp (1984). After transfer, the part of the blot containing the edges of the gel and molecular weight standards was cut off and stained with Coomassie blue to reveal the polypeptide profile (Herring and Sharp, 1984). The centre part of the blot was cut into strips 5 mm wide. Non-specific binding sites on the nitrocellulose strips were then blocked using 50% (v/v) rabbit serum for 60 min at 37 oC prior to incubating single strips for a further 60 min at room temperature
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in a 1/40 dilution of test serum. Thorough washing of the strips to remove excess test antibody was followed by reaction of the nitrocellulose-bound immune complexes with 12~I-labelled rabbit anti-sheep Fab2 IgG. After further washing to remove excess label, strips were dried and exposed to photographic film (Kodak X omat-S ) at - 70 ° C with the use of an intensifying screen (Dupont, Cronex, Lighting-Plus). Molecular weights were estimated by using a sonic digitizer to input the band positions into a BBC B microcomputer running a " M A P - G E L 2" program (kindly communicated by J o h n Coadwell). The accuracy of the estimates is, however, limited by band distortion in the edge region of the gel in which the molecular weight standards are run. RESULTS
The antigen used thoughout this study consisted of gradient-purified virions of AHV-1. All the virion preparations gave similar polypeptide profiles, with a pair of major bands at molecular weight of approximately 140 kDa and another single major component at 43 kDa as shown in Fig. 1, Track 1. The number of individual polypeptide components observed was 25-30. Eleven wildebeest sera were tested for their ability to react with AHV-1 polypeptides and nine gave clearly positive reactions while two were only weakly reactive. The patterns of reactivity were very similar with all sera and were especially strong with polypeptides of 140, 77, 50, 35, 34 and 32 kDa as seen in Fig. 1, Track 2 and Fig. 2, Tracks 7-10. Figure 1, Track 2 shows a typical reaction aligned with the stained viral polypeptides and molecular weight standards. The reaction given by serum obtained from a terminal case of M C F caused by AHV-1 is shown in Fig. 2, Track 6. Only three polypeptides reacted strongly (50, 35 and 34 kDa) and there was a weaker reaction with a band of 140 kDa. All these reactions were common with the polypeptides detected by wildebeest sera. An additional weak reaction with a polypeptide of 26 kDa has been observed when another antigen preparation of AHV-1 was reacted with the terminal M C F serum (Fig. 3, Track 4). The reaction of AHV-1 antigens with a convalescent serum against BHV-1 is shown in Fig. 3 (Track 2 and 3) and the reaction of this serum with purified BHV-1 polypeptides is seen in Track 1. The homologous reaction of the antiBHV-1 serum is mainly with polypeptides in the size range 140-50kDa and is consistent with results reported by other workers (Trudel et al., 1987). However, it is notable that, while BHV-1 serum did show some cross reaction with AHV-1 polypeptides, it reacted mainly with bands of 65, 57, 44 and 25 kDa which are not those recognized by wildebeest sera. There was a weak reaction with a band of 140 kDa which is common to sera from all species tested. Sixteen sheep sera tested showed positive reactions and typical profiles are
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2
3
4
a
b C
d
~4
g Fig. 1. Immunoblots of wildebeest and sheep sera aligned with the AHV-1 polypeptide profile. Track 1: blot strip stained to show AHV-1 polypeptides. Track 2: strip reacted with wildebeest serum. Track 3: strip reacted with sheep serum. Track 4: blot strip stained to show molecular weight standards: (a) B-galactosidase, 130 000; (b) phosphorylase-A, 94 000; (c) ovotransferrin, 77 000; (d) bovine serum albumin, 67 000; (e) ovalbumin, 45 000; (f) chymotrypsinogen A, 26 000; {g) (at buffer front) myoglobulin, 17 000.
shown in Fig. 2, Tracks 1-5 and in Fig 1, Track 3. The reactions of a further two sera were masked by very high background reactions. The reactions were not as strong as those shown by wildebeest sera and most were somewhat obscured by high non-specific background which we find is a feature of immunoblots with adult sheep sera in spite of the use of blocking reagents. An important feature was that the majority of the reactions involved polypeptides also recognized by the wildebeest sera. However, the sheep sera were more variable in their recognition patterns with individual sera reacting only to a subset of the polypeptides positive with wildebeest sera. Alignment of the pep-
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~i~ 13
.....
i ili!i i~ -,-a
--b "--C
.-d ---e
i ¸~!!~i~
'i~iiiii!iii
_
f
Fig. 2. I m m u n o b l o t s of AHV-1 polypeptides. Tracks 1-5: sheep sera. Track 6: bovine serum from a terminal case of AHV-1 MCF. Tracks 7-10: wildebeest sera. Tracks 11-12: bovine sera from sheep-associated cases of MCF. T r a c k 13: serum-free control. T h e mobilites of the molecular weight standards a - f are shown (see Fig. 1 ). T h e arrowheads indicate faint bands.
1
2
3
4
-a
"b mC
.d • e
~"
.f
.g Fig. 3. I m m u n o b l o t s of BHV-1 a n d AHV-1 polypeptides. Track 1 : B H V - 1 polypeptides reacted with a bovine convalescent serum to BHV-1. Tracks 2 a n d 3: two separate preparations of AHVI polypeptides reacted with the a n t i - B H V - 1 convalescent serum. Track 4: the same AHV-1 antigen as Track 2 reacted with a bovine serum from a terminal case of AHV-1 M C F (cf. Fig. 2, Track 6). T h e mobilities of the molecular weight standards a - g are shown (see Fig. 1 ). T h e arrowheads indicate faint bands.
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tides recognized by a typical ovine serum with the viral polypeptide pattern is shown in Fig. 1, Track 3. Only two of the six sera from terminal bovine cases of SA MCF showed positive reactions, both sera recognizing peptides which were among those bound by antibodies in wildebeest and sheep sera. One serum gave a strong reaction to the 50 kDa peptide and a weak reaction to three other polypeptides (Fig. 2, Track 12) whilst the other showed only weak reactions, the strongest of which was with a peptide of 43 kDa (Fig. 2, Track 11). DISCUSSION
Gel electrophoresis of the polypeptides from purified virions of AHV-1 yielded 25-30 components, a similar number to that observed with other herpesviruses using one-dimensional gel systems (Spear et al., 1978; Misra et al., 1981 ). Additionally, a major component was detected with a molecular weight of 140 kDa, several other herpesviruses have been shown to have major nucleocapsid proteins of this size (Honess and Watson, 1977). Previous attempts to detect antibody to AHV-1 have relied primarily on virus neutralization (Plowright, 1965; Reid et al., 1975; Mushi and Karstad, 1981) and IIF (Rossiter et al., 1977; Rossiter, 1981; Heuschele, 1982; Heuschele et al., 1984) although complement fixation (Rossiter and Jessett, 1980), indirect immunoperoxidase and agar gel precipitation tests (Rossiter et al., 1980) have been described also. Collectively, these studies have shown that, within the Bovidae subfamilies Alcelaphinae and Hippotraginae, neutralizing antibody to AHV-1 is prevalent, indicating that these species are infected w " h closely related herpesviruses (reviewed by Plowright, 1986). Several groups have been able to demonstrate by IIF that antibody to AHV-1 was prevalent in sera from the family Caprinae suggesting the presence of a related virus in this group also (Rossiter, 1981; Heuschele et al., 1984; Harkness, 1985). Neutralizing antibody was not consistently observed although Harkness (1985) reports finding low titres in 17% of U.K. sheep sera. The majority of cattle experimentally infected with virulent strains of AHV-1 succumb to acute lethal MCF. Studies of virus neutralization by serum from the terminal stages of the disease show that antibody is absent or present only at a low titres (Reid and Rowe, 1973; Rossiter et al., 1980). However, IIF antibody can be detected regularly in the final stages of the disease (Rossiter et al., 1977, 1978, 1980). Antibody to AHV-1, detectable by IIF, has also been described in some bovine cases with the sheep-associated form of the disease (Rossiter, 1983; Harkness, 1985). However, as AHV-1 shows some antigenic cross-reaction with bovid herpesviruses-1, -2 and bovine cytomegalovirus (formerly termed bovid herpesvirus-4, now named bovine herpesvirus-3; Roizman et al., 1981), these observations are not easily interpreted (Rossiter et al., 1977, 1988; Heuschele et al., 1984). BHV-1 is known to infect wildebeest (Karstad et al., 1973), but the
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results presented above show that a convalescent serum raised by sequential infection with BHV-1 reacted weakly to AHV-1 antigens by immunoblotting. Moreover, the BHV-1 serum recognized a set of antigens largely different from those which reacted with wildebeest and sheep sera. Thus it seems unlikely that cross reactions with BHV-1 were contributing to the reactions which were observed with wildebeest sera. Sera from wildebeest, the natural host of AHV-1, contained antibodies that reacted strongly with six distinct components resolved by electrophoresis and the relative intensity of the reaction of individual sera with these bands was consistent. This finding contrasts with the results obtained with sheep sera all of which reacted with at least some of the AHV-1 polypeptides recognized by the wildebeest. Individual sera varied considerably in the number and intensity of their reactions. This result provides further evidence that sheep are infected with a virus that shares at least some antigenic determinants with AHV-1. The more extensive studies of sheep sera utilizing IIF (Rossiter, 1981 ) can now be interpreted with greater confidence as indicating the widespread presence of a related herpesvirus. Serum from the bovine case with AHV-l-induced MCF reacted strongly to only three polypeptides, all of which were amongst those recognized by wildebeest sera. Following AHV-1 infection, the humoral immune response of cattle and rabbits affected with acute MCF is ineffective in preventing the lethal course of infection despite the high titres of IIF antibody which are present terminally (Rossiter et al., 1977, 1978, 1980 ). Cattle may react to only a limited number of viral determinants either due to a very low level of viral replication or to incomplete expression of the full genetic repertoire of AHV-1 in this host. Although it is clearly desirable to extend the investigation to a larger number of animals, incomplete viral gene expression in cattle is an attractive explanation in view of the absence of detectable specific virus-induced cytopathology or of virus particles in affected tissues (Rossiter, 1980; Edington et al., 1979). Only two of the six bovine sera obtained from cattle infected with SA MCF were positive and they reacted with only a few polypeptides. This result is in accord with previous reports of IIF antibody to AHV-1 antigens in a proportion of affected animals (Rossiter, 1983; Harkness, 1985). Transmission of sheepassociated MCF, although difficult, was achieved many years ago (Gotze and Leiss, 1930) and has been demonstrated several times since (Pierson et al., 1974; Selman et al., 1978; Hoffmann et al., 1984; Reid et al., 1986). However, no aetiological agent, capable of reproducing the disease, has ever been recovered from these experiments. These observations are consistent with incomplete expression of the sheep-associated agent genome, as has been suggested for AHV-1 infections of cattle. Our present efforts are directed towards extending these results to rodent models of MCF and exploring immune reactions with non-structural proteins of AHV-1 using immunoblotting and immune-precipitation techniques.
213 ACKNOWLEDGEMENTS
We thank A. Dawson of the Moredun Research Institute for preparation of the rabbit anti-sheep immunoglobulin, R. Kock of the Zoological Society of London, Whipsnade Park, for the wildebeest sera, Dr. N. Edington of the Royal Veterinary College, London University, for the serum from a bovine terminal case of AHV-1 MCF and Dr. John Coadwell of the Institute for Animal Physiology and Genetics Research, Babraham for the "Mapgel 2" programme. REFERENCES Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem., 72: 248-254. Burnette, W.N., 1981. 'Western Blotting', electrophoretic transfer of proteins from SDS-polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A. Anal. Biochem., 112: 195-203. Edington, N., Patel, J., Russell, P.H. and Plowright, W., 1979. The nature of the acute lymphoid proliferation in rabbits infected with herpesvirus of bovine malignant catarrhal fever. Eur. J. Cancer, 5: 1515-1522. Gotze, R. and Leiss, J., 1930. Untersuchungen uber das bosartigen Katarrhalfieber des Rindes. 11. Schafe als Ubertrager. Dtsch. Tieraerztl. Wochenschr., 38: 194-200. Hamblin, C. and Hedger, R.S., 1984. Neutralising antibodies to wildebeest derived malignant catarrhal fever virus in African wildlife. Comp. Immunol. Microbiol. Infect. Dis., 7: 195-199. Harkness, J.W., 1985. Bovine malignant catarrhal fever in the United Kingdom. State Vet. J., 38: 60-64. Herring, A.J. and Sharp, J.M., 1984. Protein blotting: the basic method and its role in viral diagnosis. In: M.S. McNulty and J.B. McFerran (Editors), Recent Advances in Virus Diagnosis. Martinus Nijhoff, Boston, The Hague, Dordrecht, Lancaster, for the Commission of the European Communities, pp. 115-124. Heuschele, W.P., 1982. Malignant catarrhal fever in wild ruminants - - a review and current status report. Proc. 86th Annu. Meet. U.S. Anim. Health Assoc., pp. 552-570. Heuschele, W.P., Fletcher, H.R., Oosterhuis, J., Janssen, D. and Robinson, P.T., 1984. Epidemiologic aspects of malignant catarrhal fever in the U.S.A. Proc. 88th Annu. Meet. U.S. Anim. Health Assoc., pp. 640-651. Honess, R.W. and Watson, D.H., 1977. Unity and diversity in the herpesviruses. J. Gen. Virol., 37: 15-37. Hoffmann, D., Sobironingsih, S., Clarke, B.C., Young, P.J. and Sendow, I., 1984. Transmission and virological studies of a malignant catarrhal fever syndrome in the Indonesian swamp buffalo (Bubalus bubalis). Aust. Vet. J., 61: 113-116. Laemmli, U.K., 1970. Cleavage of stuctural proteins during the assembly of the head of bacteriophage T4. Nature (London), 227: 680-685. Karstad, L.S., Drevemo, S., Otema, J.C. and Jessett, D.M., 1973. Vulvovaginitis in wildebeest caused by the virus of infectious bovine rhinotracheitis. J. Wildl. Dis, 10: 392-396. Misra, V., Blumenthal, R.M. and Babiuk, L.A., 1981. Proteins specified by Bovine Herpesvirus 1 (Infectious Bovine Rhinotracheitis Virus ). J. Virol., 40: 367-378. Mushi, E.Z. and Karstad, L., 1981. Prevalence of virus neutralising antibodies to Malignant Catarrhal Fever virus in Oryx {Oryx beisa callotis). J. Wildl. Dis., 17: 467-470. Pierson, R.E., Storz, J., McChesney, A.E. and Thake, D., 1974. Experimental transmission of Malignant catarrhal fever. Am. J. Vet. Res., 35: 532-525.
214 Plowright, W., 1965. Malignant catarrhal fever in East Africa. 1. Behaviour of the virus in free living populations of blue wildebeest (Gorgon, taurinus taurinus Burchell). Res. Vet. Sci., 6: 56-68. Plowright, W., 1968. Malignant catarrhal fever. J. Am. Vet. Med. Assoc., 152: 795-804. Plowright, W., 1986. Malignant catarrhal fever. Rev. Sci. Tech. Off. Int. Epiz., 5: 897-918. Plowright, W., Ferris, R.D. and Scott, G.R., 1960. Blue wildebeest and the aetiological agent of bovine malignant catarrhal fever. Nature (London), 188:1167 - 1169. Plowright, W., Macadam, R.F. and Armstrong, J.A., 1965. Growth and characterisation of the virus of bovine malignant catarrhal fever in East Africa. J. Gen. Microbiol., 39: 253-266. Reid, H.W. and Buxton, D., 1984. Malignant catarrhal fever of deer. Proc. R. Soc. Edinburgh, 82B: 261-283. Reid, H.W. and Rowe, L., 1973. The attenuation of a herpesvirus (malignant catarrhal fever virus ) isolated from hartebeest (Alcelaphus buselaphus cokei Gunther). Res. Vet. Sci., 15: 144-146. Reid, H.W., Plowright, W. and Rowe, L.W., 1975. Neutralising antibody to herpesviruses derived from wildebeest and hartebeest in wild animals in East Africa. Res. Vet. Sci., 18: 269-273. Reid, H.W., Buxton, D., Corrigall, W., Hunter, A.R., McMartin, D.A. and Rushton, R., 1979. An outbreak of malignant catarrhal fever in red deer (Cervus elaphus). Vet. Rec., 104: 120-123. Reid, H.W., Buxton, D., Pow, I. and Finlayson, J., 1986. Malignant catarrhal fever: experimental transmission of the 'sheep-associated' form of the disease from cattle and deer to cattle, deer, rabbits and hamsters. Res. Vet. Sci., 41: 76-81. Roizman, B., Carmichael, L.E., Dienhard, T.F., De The, G., Nahmias, A.N., Plowright, W., Rapp, F., Sheldrick, P., Takahashi, M. and Wolf, K., 1981. Herpesviridae-definition, provisional nomenclature and taxonomy. Intervirology, 16: 201-217. Rossiter, P.B., 1980. A lack of demonstrable virus antigens in the tissues of rabbits and cattle infected with malignant catarrhal fever virus. Br. Vet. J., 136: 478-483. Rossiter, P.B., 1981. Antibodies to malignant catarrhal fever virus in sheep sera. J. Comp. Pathol., 91: 303-311. Rossiter, P.B., 1983. Antibodies to malignant catarrhal fever virus in cattle and non-wildebeestassociated malignant catarrhal fever. J. Comp. Pathol., 93: 93-97. Rossiter, P.B. and Jessett, D.M., 1980. A complement fixation test for antigens and antibodies to malignant catarrhal fever virus. Res. Vet. Sci., 28: 228-233. Rossiter, P.B., Mushi, E.Z. and Plowright, W., 1977. The development of antibodies in rabbits and cattle infected experimentally with an african strain of malignant catarrhal fever virus. Vet. Microbiol., 2: 57-66. Rossiter, P.B., Mushi, E.Z. and Plowright, W., 1978. Antibody response in cattle and rabbits to early antigens of malignant catarrhal fever virus in cultured cells. Res. Vet. Sci., 25: 207-210. Rossiter, P.B., Jessett, D.M. and Mushi, E.Z., 1980. Antibodies to malignant catarrhal fever virus antigens in the sera of normal and naturally infected cattle in Kenya. Res. Vet. Sci., 29: 235239. Rossiter, P.B., Gumm, I.D. and Mirangi, P.K., 1988. Immunological relationships between Malignant Catarrhal Fever virus (alcelaphine herpesvirus-1 ) and Bovine cytomegalovirus (Bovine herpesvirus-3 ). Vet. Microbiol., 16: 211-218. Selman, I.E., Wiseman, A., Wright, N.G. and Murray, M., 1978. Transmission studies with bovine malignant catarrhal fever. Vet. Rec., 102: 252-257. Spear, P.G., Sarmiento, M. and Manservigi, R., 1978. The structural proteins and glycoproteins of herpesviruses: a review. In: G. de The, W. Henle and F. Rapp (Editors), Oncogenesis and Herpesviruses III Part 1. IARC Sci. Publ. No. 24, Lyon. Trudel, M., Seguin, C., Nadon, F., Boulay, G., Trepanier, P. and Lussier, G., 1987. Haemagglutination inhibition and virus neutralising response of rabbits inoculated with bovine herpesvirus subunit vaccine. Vet. Microbiol., 13: 107-119.
215 Westmoreland, D. and Watkins, J.F., 1974. studies using radioiodinated IgG. J. Gen. Zygraich, N., Lobmann, M., Vascobionic, E. erties of a temperature sensitive mutant Sci., 16: 328-335.
The IgG receptor induced by herpes simplex virus: Virol., 24: 167-178. and Huygelen, C., 1974. 'In vivo' and 'in vitro' propof infectious bovine rhinotracheitis virus. Res. Vet.
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Immunoblotting Analysis of the Reaction of Wildebeest, Sheep and Cattle Sera with the Structural Antigens of Alcelaphine Herpesvirus-1 (Malignant Catarrhal Fever Virus) ALAN HERRING, HUGH REID, NEIL INGLIS and IRENE POW
Moredun Research Institute, 408 Gilmerton Road, Edinburgh EH17 7JH (Gt. Britain) (Accepted for publication 3 October 1988)
ABSTRACT Herring, A., Reid, H., Inglis, N. and Pow, I., 1989. Immunoblotting analysis of the reaction of wildebeest, sheep and cattle sera with the structural antigens of alcelaphine herpesvirus- 1 (malignant catarrhal fever virus). Vet. Microbiol., 19: 205-215. Malignant catarrhal fever (MCF) is a disease of cattle and some other ruminants caused by alcelaphine herpesvirus-1 (AHV-1), a virus of wildebeest. The disease also occurs in the absence of wildebeest and is then thought to be caused by a viral agent harboured by the sheep. The structural proteins of AHV-1 have been used as antigens for the immunoblottinganalysis of sera from wildebeest, sheep and cattle infected by either AHV-1 or the "sheep-associated" form of the disease. Wildebeest sera showed a uniform response reacting strongly with six polypeptides. Sheep sera also gave positive results but individual sera reacted with varying subsets of the antigens recognized by wildebeest. These results support the earlier suggestion that sheep harbour a herpesvirus related to AHV-1. A bovine serum from a case of MCF caused by AHV-1 also reacted only with a subset of the six wildebeest-reactive polypeptides. Sera from cattle affected with the "sheep-associated" form of the disease gave reactions in only two of the eight cases tested; both positive sera reacted to a few polypeptides only.
INTRODUCTION
Malignant catarrhal fever (MCF), a fatal lymphoproliferative disease of cattle and other ruminants, occurs in two clinically and pathologically similar forms. One cause of the disease is alcelaphine herpesvirus-1 (AHV-1) which is carried by the wildebeest (Connochaetes taurinus) as an inapparent infection (Plowright et al., 1960; Plowright, 1965, 1968). Epidemiological observations of the other form of the disease strongly suggest that the source of infection is the domestic sheep and it is proposed that this species carries a virus biologically similar to AHV-1 (Rossiter, 1981; Reid and Buxton, 1984; Reid et 0378-1135/89/$03.50
© 1989 Elsevier Science Publishers B.V.
206 al., 1986). This form of the disease is termed "sheep-associated" MCF (SA MCF). Several other antelope of the subfamilies Hippotraginae and Alcelaphinae consistently have neutralizing antibody to AHV-1, implying that related viruses are prevalent in these two groups of African antelope (Reid et al., 1975; Mushi and Karstad, 1981; Hamblin and Hedger, 1984). No such neutralizing antibody can be detected consistently in sheep sera, although Rossiter (1981) found a reaction with all adult sheep sera examined using cells infected with AHV-1 in an indirect immunofluorescent (IIF) antibody test. However, the broadly cross-reactive nature of the IIF test, and the known property of some herpesvirus proteins to bind immunoglobulins (Westmoreland and Watkins, 1974), make interpretation of these results difficult. In order to assess the significance of the above observations, a qualitative comparison of the reaction of wildebeest and sheep sera to proteins prepared from purified virions of AHV1 was made employing the immunoblotting technique. In addition, sera from cattle with both forms of the disease were examined. MATERIALSAND METHODS
Sera Eighteen sheep sera were used. Seven of these came from sheep which had been housed with a group of red deer that developed MCF (Reid et al., 1979). The remainder were from normal adult sheep. The 11 wildebeest sera were kindly supplied by R. Kock from animals that had previously been kept at Whipsnade Park, England. The bovine serum from a calf with MCF induced by AHV-1 was kindly supplied by Dr. N. Edington, and the sera from cattle with SA MCF were from clinical cases referred to the Institute and confirmed histologically as MCF (Reid et al., 1986). The serum against bovine herpesvirus-1 (BHV-1) was derived from a cow which had been vaccinated with a live, temperature-sensitive vaccine (Zygraich et al., 1974) and subsequently challenged with virulent virus. The iodinated detection reagent used throughout was derived from a rabbit hyperimmune serum raised against ovine IgG and subsequently purified on an affinity column bearing ovine IgG Fab2 fragments. This rabbit IgG also reacted with immunoglobulins from bovine, cervine and alcelaphine sources. Class and subclass reactions have not been investigated but IgG is the major species recognized by this reagent in sheep serum.
Antigen The WC 11, cell-culture-adapted, cell-free strain of AHV-1 (Plowright et al., 1965) was propagated in monolayer cultures of bovine kidney cells grown in
207
roller vessels. Supernatant fluids, which contained approximately 5 X 106 50% tissue culture-infective doses m l - 1, were harvested when 50% of the cells showed cytopathic effects. After clarification for 20 min at 9000 Xg and 4 ° C, virus was pelleted by centrifugation at 40 000 x g at 5 ° C. Pellets were resuspended in 8 ml of 1 x T N E buffer (10 mM Tris-HC1, 100 mM NaC1, 1 mM EDTA (diNa) pH 7.5 ), loaded onto a pre-formed 7.5 m115-40% w/w Metrizamide (Nyegaard U.K. ) gradient in 1 X TNE pH 7.5 and centrifuged at 114 000 X g for 60 min at + 4 ° C in a Beckman SW40Ti rotor. The visible band of virus which formed at the centre of the gradient was collected diluted to 12 ml in 1 X TNE pH 7.5 and repelleted at 114 000Xg for 40 min at 4°C. The resulting pellet was resuspended in 100 zl of TNE pH 7.5 and assessed for purity by inspection in the electron microscope at a magnification of 25 000 after negative staining with 1% (w/v) phosphotungstic acid. Protein content was estimated using the "BioRad" dye-binding assay (Bradford, 1976). BHV-1 virus was purified in essentially the same fashion except that a 10-40% (w/v) potassium tartrate gradient was employed.
Electrophoresis Most experiments were performed using block gels in which a single large well was loaded with viral protein and protein molecular weight standards were loaded in a small side well. A volume of viral suspension containing 200 ]lg of protein was mixed with an equal volume of 2 X concentration denaturing buffer to yield final concentrations of 63 mM Tris, 2% (v/v) fl-mercaptoethanol, 2% (w/v) sodium dodecyl sulphate (SDS), 10% (w/v) sucrose and 0.001% bromophenol blue. After heating in a 100°C waterbath for 90 s, the viral polypeptides were electrophoretically separated using a 10% discontinuously buffered polyacrylamide gel with a 3% stacking gel (Laemmli, 1970). Electrophoresis was performed under conditions of constant current (current density 1.9 mA cm-2) until the bromophenol blue tracker dye had migrated a distance of 9 cm in the resolving gel.
Immunoblotting Electrophoretic transfer of resolved viral polypeptides to nitrocellulose and the reaction of blotted strips with antibody was performed essentially as described by Burnette (1981) using the modifications of Herring and Sharp (1984). After transfer, the part of the blot containing the edges of the gel and molecular weight standards was cut off and stained with Coomassie blue to reveal the polypeptide profile (Herring and Sharp, 1984). The centre part of the blot was cut into strips 5 mm wide. Non-specific binding sites on the nitrocellulose strips were then blocked using 50% (v/v) rabbit serum for 60 min at 37 oC prior to incubating single strips for a further 60 min at room temperature
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in a 1/40 dilution of test serum. Thorough washing of the strips to remove excess test antibody was followed by reaction of the nitrocellulose-bound immune complexes with 12~I-labelled rabbit anti-sheep Fab2 IgG. After further washing to remove excess label, strips were dried and exposed to photographic film (Kodak X omat-S ) at - 70 ° C with the use of an intensifying screen (Dupont, Cronex, Lighting-Plus). Molecular weights were estimated by using a sonic digitizer to input the band positions into a BBC B microcomputer running a " M A P - G E L 2" program (kindly communicated by J o h n Coadwell). The accuracy of the estimates is, however, limited by band distortion in the edge region of the gel in which the molecular weight standards are run. RESULTS
The antigen used thoughout this study consisted of gradient-purified virions of AHV-1. All the virion preparations gave similar polypeptide profiles, with a pair of major bands at molecular weight of approximately 140 kDa and another single major component at 43 kDa as shown in Fig. 1, Track 1. The number of individual polypeptide components observed was 25-30. Eleven wildebeest sera were tested for their ability to react with AHV-1 polypeptides and nine gave clearly positive reactions while two were only weakly reactive. The patterns of reactivity were very similar with all sera and were especially strong with polypeptides of 140, 77, 50, 35, 34 and 32 kDa as seen in Fig. 1, Track 2 and Fig. 2, Tracks 7-10. Figure 1, Track 2 shows a typical reaction aligned with the stained viral polypeptides and molecular weight standards. The reaction given by serum obtained from a terminal case of M C F caused by AHV-1 is shown in Fig. 2, Track 6. Only three polypeptides reacted strongly (50, 35 and 34 kDa) and there was a weaker reaction with a band of 140 kDa. All these reactions were common with the polypeptides detected by wildebeest sera. An additional weak reaction with a polypeptide of 26 kDa has been observed when another antigen preparation of AHV-1 was reacted with the terminal M C F serum (Fig. 3, Track 4). The reaction of AHV-1 antigens with a convalescent serum against BHV-1 is shown in Fig. 3 (Track 2 and 3) and the reaction of this serum with purified BHV-1 polypeptides is seen in Track 1. The homologous reaction of the antiBHV-1 serum is mainly with polypeptides in the size range 140-50kDa and is consistent with results reported by other workers (Trudel et al., 1987). However, it is notable that, while BHV-1 serum did show some cross reaction with AHV-1 polypeptides, it reacted mainly with bands of 65, 57, 44 and 25 kDa which are not those recognized by wildebeest sera. There was a weak reaction with a band of 140 kDa which is common to sera from all species tested. Sixteen sheep sera tested showed positive reactions and typical profiles are
209
2
3
4
a
b C
d
~4
g Fig. 1. Immunoblots of wildebeest and sheep sera aligned with the AHV-1 polypeptide profile. Track 1: blot strip stained to show AHV-1 polypeptides. Track 2: strip reacted with wildebeest serum. Track 3: strip reacted with sheep serum. Track 4: blot strip stained to show molecular weight standards: (a) B-galactosidase, 130 000; (b) phosphorylase-A, 94 000; (c) ovotransferrin, 77 000; (d) bovine serum albumin, 67 000; (e) ovalbumin, 45 000; (f) chymotrypsinogen A, 26 000; {g) (at buffer front) myoglobulin, 17 000.
shown in Fig. 2, Tracks 1-5 and in Fig 1, Track 3. The reactions of a further two sera were masked by very high background reactions. The reactions were not as strong as those shown by wildebeest sera and most were somewhat obscured by high non-specific background which we find is a feature of immunoblots with adult sheep sera in spite of the use of blocking reagents. An important feature was that the majority of the reactions involved polypeptides also recognized by the wildebeest sera. However, the sheep sera were more variable in their recognition patterns with individual sera reacting only to a subset of the polypeptides positive with wildebeest sera. Alignment of the pep-
210
~i~ 13
.....
i ili!i i~ -,-a
--b "--C
.-d ---e
i ¸~!!~i~
'i~iiiii!iii
_
f
Fig. 2. I m m u n o b l o t s of AHV-1 polypeptides. Tracks 1-5: sheep sera. Track 6: bovine serum from a terminal case of AHV-1 MCF. Tracks 7-10: wildebeest sera. Tracks 11-12: bovine sera from sheep-associated cases of MCF. T r a c k 13: serum-free control. T h e mobilites of the molecular weight standards a - f are shown (see Fig. 1 ). T h e arrowheads indicate faint bands.
1
2
3
4
-a
"b mC
.d • e
~"
.f
.g Fig. 3. I m m u n o b l o t s of BHV-1 a n d AHV-1 polypeptides. Track 1 : B H V - 1 polypeptides reacted with a bovine convalescent serum to BHV-1. Tracks 2 a n d 3: two separate preparations of AHVI polypeptides reacted with the a n t i - B H V - 1 convalescent serum. Track 4: the same AHV-1 antigen as Track 2 reacted with a bovine serum from a terminal case of AHV-1 M C F (cf. Fig. 2, Track 6). T h e mobilities of the molecular weight standards a - g are shown (see Fig. 1 ). T h e arrowheads indicate faint bands.
211
tides recognized by a typical ovine serum with the viral polypeptide pattern is shown in Fig. 1, Track 3. Only two of the six sera from terminal bovine cases of SA MCF showed positive reactions, both sera recognizing peptides which were among those bound by antibodies in wildebeest and sheep sera. One serum gave a strong reaction to the 50 kDa peptide and a weak reaction to three other polypeptides (Fig. 2, Track 12) whilst the other showed only weak reactions, the strongest of which was with a peptide of 43 kDa (Fig. 2, Track 11). DISCUSSION
Gel electrophoresis of the polypeptides from purified virions of AHV-1 yielded 25-30 components, a similar number to that observed with other herpesviruses using one-dimensional gel systems (Spear et al., 1978; Misra et al., 1981 ). Additionally, a major component was detected with a molecular weight of 140 kDa, several other herpesviruses have been shown to have major nucleocapsid proteins of this size (Honess and Watson, 1977). Previous attempts to detect antibody to AHV-1 have relied primarily on virus neutralization (Plowright, 1965; Reid et al., 1975; Mushi and Karstad, 1981) and IIF (Rossiter et al., 1977; Rossiter, 1981; Heuschele, 1982; Heuschele et al., 1984) although complement fixation (Rossiter and Jessett, 1980), indirect immunoperoxidase and agar gel precipitation tests (Rossiter et al., 1980) have been described also. Collectively, these studies have shown that, within the Bovidae subfamilies Alcelaphinae and Hippotraginae, neutralizing antibody to AHV-1 is prevalent, indicating that these species are infected w " h closely related herpesviruses (reviewed by Plowright, 1986). Several groups have been able to demonstrate by IIF that antibody to AHV-1 was prevalent in sera from the family Caprinae suggesting the presence of a related virus in this group also (Rossiter, 1981; Heuschele et al., 1984; Harkness, 1985). Neutralizing antibody was not consistently observed although Harkness (1985) reports finding low titres in 17% of U.K. sheep sera. The majority of cattle experimentally infected with virulent strains of AHV-1 succumb to acute lethal MCF. Studies of virus neutralization by serum from the terminal stages of the disease show that antibody is absent or present only at a low titres (Reid and Rowe, 1973; Rossiter et al., 1980). However, IIF antibody can be detected regularly in the final stages of the disease (Rossiter et al., 1977, 1978, 1980). Antibody to AHV-1, detectable by IIF, has also been described in some bovine cases with the sheep-associated form of the disease (Rossiter, 1983; Harkness, 1985). However, as AHV-1 shows some antigenic cross-reaction with bovid herpesviruses-1, -2 and bovine cytomegalovirus (formerly termed bovid herpesvirus-4, now named bovine herpesvirus-3; Roizman et al., 1981), these observations are not easily interpreted (Rossiter et al., 1977, 1988; Heuschele et al., 1984). BHV-1 is known to infect wildebeest (Karstad et al., 1973), but the
212
results presented above show that a convalescent serum raised by sequential infection with BHV-1 reacted weakly to AHV-1 antigens by immunoblotting. Moreover, the BHV-1 serum recognized a set of antigens largely different from those which reacted with wildebeest and sheep sera. Thus it seems unlikely that cross reactions with BHV-1 were contributing to the reactions which were observed with wildebeest sera. Sera from wildebeest, the natural host of AHV-1, contained antibodies that reacted strongly with six distinct components resolved by electrophoresis and the relative intensity of the reaction of individual sera with these bands was consistent. This finding contrasts with the results obtained with sheep sera all of which reacted with at least some of the AHV-1 polypeptides recognized by the wildebeest. Individual sera varied considerably in the number and intensity of their reactions. This result provides further evidence that sheep are infected with a virus that shares at least some antigenic determinants with AHV-1. The more extensive studies of sheep sera utilizing IIF (Rossiter, 1981 ) can now be interpreted with greater confidence as indicating the widespread presence of a related herpesvirus. Serum from the bovine case with AHV-l-induced MCF reacted strongly to only three polypeptides, all of which were amongst those recognized by wildebeest sera. Following AHV-1 infection, the humoral immune response of cattle and rabbits affected with acute MCF is ineffective in preventing the lethal course of infection despite the high titres of IIF antibody which are present terminally (Rossiter et al., 1977, 1978, 1980 ). Cattle may react to only a limited number of viral determinants either due to a very low level of viral replication or to incomplete expression of the full genetic repertoire of AHV-1 in this host. Although it is clearly desirable to extend the investigation to a larger number of animals, incomplete viral gene expression in cattle is an attractive explanation in view of the absence of detectable specific virus-induced cytopathology or of virus particles in affected tissues (Rossiter, 1980; Edington et al., 1979). Only two of the six bovine sera obtained from cattle infected with SA MCF were positive and they reacted with only a few polypeptides. This result is in accord with previous reports of IIF antibody to AHV-1 antigens in a proportion of affected animals (Rossiter, 1983; Harkness, 1985). Transmission of sheepassociated MCF, although difficult, was achieved many years ago (Gotze and Leiss, 1930) and has been demonstrated several times since (Pierson et al., 1974; Selman et al., 1978; Hoffmann et al., 1984; Reid et al., 1986). However, no aetiological agent, capable of reproducing the disease, has ever been recovered from these experiments. These observations are consistent with incomplete expression of the sheep-associated agent genome, as has been suggested for AHV-1 infections of cattle. Our present efforts are directed towards extending these results to rodent models of MCF and exploring immune reactions with non-structural proteins of AHV-1 using immunoblotting and immune-precipitation techniques.
213 ACKNOWLEDGEMENTS
We thank A. Dawson of the Moredun Research Institute for preparation of the rabbit anti-sheep immunoglobulin, R. Kock of the Zoological Society of London, Whipsnade Park, for the wildebeest sera, Dr. N. Edington of the Royal Veterinary College, London University, for the serum from a bovine terminal case of AHV-1 MCF and Dr. John Coadwell of the Institute for Animal Physiology and Genetics Research, Babraham for the "Mapgel 2" programme. REFERENCES Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem., 72: 248-254. Burnette, W.N., 1981. 'Western Blotting', electrophoretic transfer of proteins from SDS-polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A. Anal. Biochem., 112: 195-203. Edington, N., Patel, J., Russell, P.H. and Plowright, W., 1979. The nature of the acute lymphoid proliferation in rabbits infected with herpesvirus of bovine malignant catarrhal fever. Eur. J. Cancer, 5: 1515-1522. Gotze, R. and Leiss, J., 1930. Untersuchungen uber das bosartigen Katarrhalfieber des Rindes. 11. Schafe als Ubertrager. Dtsch. Tieraerztl. Wochenschr., 38: 194-200. Hamblin, C. and Hedger, R.S., 1984. Neutralising antibodies to wildebeest derived malignant catarrhal fever virus in African wildlife. Comp. Immunol. Microbiol. Infect. Dis., 7: 195-199. Harkness, J.W., 1985. Bovine malignant catarrhal fever in the United Kingdom. State Vet. J., 38: 60-64. Herring, A.J. and Sharp, J.M., 1984. Protein blotting: the basic method and its role in viral diagnosis. In: M.S. McNulty and J.B. McFerran (Editors), Recent Advances in Virus Diagnosis. Martinus Nijhoff, Boston, The Hague, Dordrecht, Lancaster, for the Commission of the European Communities, pp. 115-124. Heuschele, W.P., 1982. Malignant catarrhal fever in wild ruminants - - a review and current status report. Proc. 86th Annu. Meet. U.S. Anim. Health Assoc., pp. 552-570. Heuschele, W.P., Fletcher, H.R., Oosterhuis, J., Janssen, D. and Robinson, P.T., 1984. Epidemiologic aspects of malignant catarrhal fever in the U.S.A. Proc. 88th Annu. Meet. U.S. Anim. Health Assoc., pp. 640-651. Honess, R.W. and Watson, D.H., 1977. Unity and diversity in the herpesviruses. J. Gen. Virol., 37: 15-37. Hoffmann, D., Sobironingsih, S., Clarke, B.C., Young, P.J. and Sendow, I., 1984. Transmission and virological studies of a malignant catarrhal fever syndrome in the Indonesian swamp buffalo (Bubalus bubalis). Aust. Vet. J., 61: 113-116. Laemmli, U.K., 1970. Cleavage of stuctural proteins during the assembly of the head of bacteriophage T4. Nature (London), 227: 680-685. Karstad, L.S., Drevemo, S., Otema, J.C. and Jessett, D.M., 1973. Vulvovaginitis in wildebeest caused by the virus of infectious bovine rhinotracheitis. J. Wildl. Dis, 10: 392-396. Misra, V., Blumenthal, R.M. and Babiuk, L.A., 1981. Proteins specified by Bovine Herpesvirus 1 (Infectious Bovine Rhinotracheitis Virus ). J. Virol., 40: 367-378. Mushi, E.Z. and Karstad, L., 1981. Prevalence of virus neutralising antibodies to Malignant Catarrhal Fever virus in Oryx {Oryx beisa callotis). J. Wildl. Dis., 17: 467-470. Pierson, R.E., Storz, J., McChesney, A.E. and Thake, D., 1974. Experimental transmission of Malignant catarrhal fever. Am. J. Vet. Res., 35: 532-525.
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