- Email: [email protected]
Cytokine responses to EHV-1 infection in immune and non-immune ponies
Veterinary Immunology and Immunopathology 111 (2006) 109–116 www.elsevier.com/locate/vetimm
Short communication
Cytokine responses to EHV-1 infection in immune and non-immune ponies Dane K. Coombs a, T. Patton a, Andrea K. Kohler b, G. Soboll c, Cormac Breathnach d, Hugh G.G. Townsend e, D.P. Lunn b,* a
Department of Medical Sciences, School of Veterinary Medicine, University of Wisconsin, 2015 Linden Drive West, Madison, WI 53706, USA Department of Clinical Sciences, College of Veterinary Medicine, Colorado State University, 300 W. Drake Rd, Fort Collins, CO 80523, USA c Dartmouth Medical Hitchcock Center, Department of Physiology, HB 7700, Room 708 W., 1 Medical Center Dr, Lebanon, NH 03756, USA d Maxwell H. Gluck Equine Research Center, Department of Veterinary Science, University of Kentucky, Lexington, KY 40546-0099, USA e Department of Veterinary Internal Medicine, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, Sask., Canada S7N 5B4 b
Abstract Protecting equids against equine herpesvirus-1 (EHV-1) infection remains an elusive goal. Repeated infection with EHV-1 leads to protective immunity against clinical respiratory disease, and a study was conducted to measure the regulatory cytokine response (IFN-g and IL-4) in repeatedly infected immune ponies compared to non-immune ponies. Two groups of four ponies were established. Group 1 ponies had previously been infected on two occasions, and most recently 7 months before this study. Group 2 ponies had no history no vaccination or challenge infection prior to this study. Both groups were subjected to an intranasal challenge infection with EHV-1, and blood samples were collected pre-infection, and at 7 and 21 days post-infection for preparation of PBMCs. At each time point, the in vitro responses of PBMCs to stimulation with EHV-1 were measured, including IFN-g and IL-4 mRNA production, and lymphoproliferation. Group 1 ponies showed no signs of clinical disease or viral shedding after challenge infection. Group 2 ponies experienced a biphasic pyrexia, mucopurulent nasal discharge, and nasal shedding of virus after infection. Group 1 ponies had an immune response characterized both before and subsequent to challenge infection by an IFN-g response to EHV-1 in the absence of an IL-4 response, and demonstrated increased EHV-1-specific lymphoproliferation post-infection. Group 2 ponies had limited cytokine or lymphoproliferative responses to EHV-1 pre-challenge, and demonstrated increases in both IFN-g and IL-4 responses post-challenge, but without any lymphoproliferative response. Protective immunity to EHV-1 infection was therefore characterized by a polarized IFN-g dependent immunoregulatory cytokine response. # 2006 Elsevier B.V. All rights reserved. Keywords: Horses; Equine herpesvirus-1; IFN-g; IL-4
* Corresponding author. Tel.: +1 970 297 1274; fax: +1 970 297 1275. E-mail address: [email protected] (D.P. Lunn). 0165-2427/$ – see front matter # 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.vetimm.2006.01.013
110
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
1. Introduction Equine herpesvirus-1 (EHV-1) infection causes contagious respiratory disease, epidemic abortion and sporadic neurological disease (Allen et al., 1999; Kydd et al., 2006). Respiratory infection by EHV-1 is common in young horses, with many horses becoming infected in the first weeks or months of life (Foote et al., 2003). Clinical respiratory disease is most common in younger horses, and repeated infections are increasingly associated with subclinical disease as horses grow older (Allen et al., 1999). The duration of acquired immunity is shortlived and re-infection can occur within 3–6 months (Kydd et al., 2006). Life-long latent infection is a common sequelae to primary infection and latently infected horses may develop a recrudescent infection and shed virus with or without demonstrating clinical disease (Allen et al., 1999). Although modified live (MLV) and killed vaccines have been used in the equine industry in an attempt to control EHV-1 infection, success has been limited (Kydd et al., 2006). The basis of protective immunity to EHV-1 infection is not fully understood, although there is substantial evidence that a cellular response involving major histocompatability class I (MHC I) restricted CD8+ cytotoxic T-lymphocytes (CTLs) is protective (Kydd et al., 2006). Recently, the number of EHV-1 specific CTL precursors (CTLp) has been demonstrated to be the most accurate, measurable correlate of immunity to EHV-1 infection. A low frequency of CTLp is associated with an increase in severity of disease from EHV-1 infection, while a high frequency of CTLp reduces the duration and level of viremia post-EHV-1 infection (Kydd et al., 2003; O’Neill et al., 1999). Although CTLp numbers may be a reliable predictor of a protective immune response against EHV-1 infection, the limiting dilution assay used to measure this parameter is laborious and complex. An alternative marker of a cell-mediated immune response is the production of interferon-gamma (IFN-g) (Murali-Krishna et al., 1998), and we have recently demonstrated that the post-exposure immune response to EHV-1 infection is associated with IFN-g production and an increasing CTL response (Breathnach et al., 2005). It is possible
that the IFN-g response to EHV-1 infection is part of a T-helper 1 (Th-1) immunoregulatory response, as is the case for the protective immune response to equine influenza virus infection (Soboll et al., 2003a). Such Th-1 responses may be associated with protective immunity to viral infection in many species, and this type of polarization of the equine immune response has been reported (Horohov, 2000). The cytokines IFN-g and IL-4 are important regulators of the balance between a Th-1 and Th-2 immune response, and there have been no studies documenting their relative production in EHV-1 immune and non-immune horses. This study was designed to characterize the IFN-g/ IL-4 response to EHV-1 infection in ponies with a history that would predict that they would be either immune or non-immune to clinical disease consequent to infection. The goal was to determine the regulatory cytokine profile associated with these different immune responses to EHV-1 infection. To accomplish this, we studied older ponies that had experienced a number of EHV-1 infections (putative immune group), and compared them to younger ponies with no recent history of EHV-1 infection. The purpose of this study was to further characterize protective immunity to EHV-1.
2. Materials and methods 2.1. Animals All animal experiments were reviewed and conducted in accordance with the requirements of the Research and Animal Resources Committee, University of Wisconsin, Madison, WI. Eight clinically normal, intact male ponies were studied. Two groups of ponies were used for challenge experiments: Group 1 ponies (ponies 82, 84, 85 and 86) ranged from 26 to 29 months of age and had previously been intranasally infected twice with EHV-1/A183, with the most recent infection being 7 months prior to this study; Group 2 ponies (ponies 10, 17, 95 and 99) ranged from 14 to 17 months of age and had no prior history of vaccination or experimental challenge with EHV-1. Ponies were housed in separate rooms in a semi-isolation unit for the duration of the experiment.
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
2.2. Virus All experiments described in this paper used viral strain EHV-1 Army 183. The virus was propagated in Kentucky equine dermal (KyED) cells (generously provided by Dr. George Allen, The Gluck Equine Research Center, Lexington, KY). Virus-laden, cellfree supernatant was used for challenge infections, and the viral dose (pfu/ml) and virus isolation was determined by titration on KyED cells. 2.3. Experimental design All challenges were performed by intranasal instillation of 2 107 pfu/ml EHV-1/A183 through a 14-gauge catheter placed in the ventral nasal meatus. Two intranasal challenge experiments were conducted. In a first experiment, Group 1 ponies were challenged with EHV-1, while Group 2 ponies were unchallenged and used as a control group. Ponies were monitored daily for pulse and respiratory rate, rectal temperature and nasal discharge for 10 days following challenge. In the second experiment (conducted 6 weeks later), the Group 2 ponies were challenged with EHV-1 and monitored for 3 weeks post-challenge. Blood was collected by jugular venipuncture for serum sampling. Blood was collected into heparin for PBMC isolation for use in lymphoproliferation and cytokine mRNA assays, and for detection of viremia. Nasal swabs were also collected for detection of nasal virus shedding. 2.4. Virus isolation Detection of nasal virus shedding was performed using dacron nasal swabs (Baxter Healthcare Corporation, McGaw Park, IL). Nasal swabs were collected daily on day 0 (pre-challenge) and on days 1–10 and then on alternate days until day 21 postinfection. Viral shedding in nasal swab samples was measured in PFU/ml of sample. For this purpose, KyED cells were added to six well tissue culture plates (Corning Inc., Corning, NY) and allowed to adhere. When cell layers were 90% confluent, nasal swab samples were thawed and filter sterilized using a 0.45 mM syringe top filter (Pall Corp., Ann Arbor, MI). Filtered samples were added to cell
111
layers at 10-fold dilutions in duplicate and incubated at 37 8C, 4% CO2 for 1.5 h. Inoculums were then removed and plates were rinsed with minimal essential media (MEM: GibcoBRL, Life Technologies, Grand Island, NY) plus 2% fetal calf serum (FCS). Subsequently 3 ml of MEM containing 0.75% methyl cellulose (Sigma), 100 U/ml penicillin, 100 mg/ml streptomycin, and 2% FCS were added to each well. Plates were incubated for 3 days at 37 8C, 4% CO2, before removal of the methylcellulose media and staining with 1 ml of crystal violet (Fisher Scientific, Fair Lawn, NJ) per well for 30 min. After rinsing with water and drying, plaques were counted and the PFU/ml of sample was calculated. Viremia was detected from heparinized blood samples during experiment I on days 0, 2, 4, 7, 8 and 10 days post-infection for Group 1 ponies and on days 0 and 7 for Group 2 ponies. During experiment II, viremia was measured daily on days 0–10, 12, 14 and 21 for the infected Group 2 ponies. For detection of viremia, 15 ml of blood was collected into heparin and red blood cells were allowed to settle. The plasma, containing PBMCs, was collected, white blood cells were pelleted and washed three times in PBS and subsequently added to 25 cm2 tissue culture flasks (Corning Inc.) containing a 90% confluent monolayer of KyED cells. Flasks were incubated for 7 days at 37 8C, 4% CO2 and checked daily for appearance of cytopathic effect (CPE). After 7 days, the virus containing supernatants were recovered by freeze–thawing and 500 ml aliquots were added to a fresh 25 cm2 tissue culture flasks containing a 90% confluent monolayer of KyED cells. These second pass flasks were incubated for further 7 days at 37 8C, 4% CO2 and appearance of cytopathic effect was recorded as a positive result. 2.5. Virus neutralization assay Serum antibody titers to EHV-1 were determined during days 0, 7, 14 and 21 post-infection for Groups 1 and 2 ponies. The amount of neutralizing antibody titer in the sera was determined by a virus neutralization assay at the Wisconsin State Veterinary Diagnostic Laboratory (Madison, WI) as previously described (Soboll et al., 2003b).
112
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
2.6. Proliferation assay Heparinized blood was collected for PBMC isolation on days 0, 7 and 21 post-infection for both groups of ponies. PBMCs were prepared by overlaying leukocyte-rich plasma on Histopaque-1077 (Sigma Chemical, St. Louis, MO) as previously described (Kydd et al., 1994). Isolated PBMCs were re-suspended at 2 106 cells/ml in RPMI 1640 (Gibco/Invitrogen) containing 2 mM L-glutamine, 10 mg streptomycin/ml, 10,000 units penicillin/ml, 250 mg amphotericin B/ml, 2 mM sodium pyruvate, 2 mM B-mercaptoethanol, 10 mM HEPES and 10% FBS. PBMCs were assayed in round-bottom 96-well microplates (Fisher Scientific). A total of 2.5 105 PBMCs/well were incubated with heat-inactivated EHV-1 (2 1010 pfu/ml), or a media control for 5 days at 37 8C in 5% CO2. At that time, 1 mCi [3H] thymidine was added to each well, and the cells were incubated for an additional 12 h before harvesting with a Packard Filter Mate (Packard, Meriden, CT) cell harvester. The uptake of [3H] thymidine was measured by liquid scintillation counting with a Top CountTM cell counter (Hewlett Packard, Palo Alto, CA). Stimulation indices (SI) were calculated based on the following equation: [cpm (virus-stimulated)]/[cpm (medium-stimulated)]. 2.7. Quantification of IFN-g and IL-4 mRNA Aliquots of 1.6 106 medium or EHV-1-stimulated equine PBMC, prepared as described above, were placed in duplicate wells of a 24-well plate, and incubated for 48 h at 37 8C, 5% CO2. Assays for IFNg mRNA expression were performed as described previously (Breathnach et al., 2004, #2825). Briefly, the contents of duplicate wells were pooled and resuspended in 300 ml RNA Stat 60 (Tel-Test, Friendswood, TX) and frozen at 70 8C until RNA extraction according to the manufacturer’s instructions. The RNA concentration was estimated by spectrophotometry, and cDNA was generated using with Moloney’s murine leukaemia virus reverse transcriptase (Promega, Madison, WI) and oligo dT primers (Promega). Expression of equine IFN-g, IL-4 and b-actin in the sample cDNA was determined by performing realtime PCR as previously described (Breathnach et al., 2004). The primer and probe sequences used were as follows: IFN-g forward primer: 50 -CGCAAAGCAA-
TAAGTGAACTCATC-30 ; IFN-g reverse: 50 -CGAAATGGATTCTGACTCCTCTTC-30 ; IFN-g probe: 50 56-FAM-TCTGTCGCCCAAAGCTAACCTGAGGAA-3BHQ-1-30 ; IL-4 forward primer: 50 -TCGTGCATGGAGCTGACTGTA-30 ; IL-4 reverse: 50 -GCCCTGCAGATTTCCTTT-CC-30 ; IL-4 probe: 50 -56-FAMCCTTTGCTGGCCCGAAGAACACAGA-3BHQ-130 ; b-actin forward primer: 50 -AGCGAAATCGTGCGTGACA-30 ; b-actin reverse: 50 -GCCATCTCCTGCTCGAAGT-C-30 ; b-actin probe: 50 -HEX-CAAGGAGAAGCTCTGCTATGTCGCCCT-BHQ2-30 . Fluorescent probes were used to measure amplified product in a thermocycler (iCycler; Bio-Rad Laboratories, Hercules, CA). Assays were run in triplicate in capped 96-well optical plates (Bio-Rad) containing 100 ng cDNA, 0.5 ml forward and reverse primers (10 mM), 5 ml probe (2 mM; IDT, Caralvile, IA), and 12.5 ml master mix (Gibco/Invitrogen); volume of reaction mixtures, 25 ml. Plasmids encoding equine IL-4 (pUC 2.1), equine IFN-g (pWRG 1647) and b-actin (pCR 2.1) were used to establish a standard curve on each 96well plate. Standard curves were generated by using the plasmid DNA at dilutions ranging from 1015 to 1022 M/ml. Each cytokine was analyzed on a separate experimental plate and amplification was performed by using the following procedure: 3 min at 95 8C followed by 45 cycles of 95 8C for 30 s and 60 8C for 60 s. Separate standard curves for IFN-g, IL-4 and bactin were established using the log dilutions of plasmid standards. On each plate, reaction mixtures containing no cDNA were included to control for contaminating DNA within the reagents. The transcript levels of IFN-g and IL-4 were normalized by using the transcript level value for bactin from the same sample (IFN-g/b-actin or IL-4/bactin). The herpes virus-specific cytokine response was expressed as stimulation index (SI) using the following equation: cytokine copy numberðvirus-stimulatedÞ= b-actin copy numberðvirus-stimulatedÞ cytokine copy numberðmedium-stimulatedÞ= b-actin copy numberðmedium-stimulatedÞ Additionally, the tendency towards an IFN-g or IL4 response was determined by the following equation: IFN-g SI/IL-4 SI.
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
2.8. Statistical analyses Comparisons were made for post-challenge responses between Group 1 ponies and Group 2 ponies, to determine the different clinical and immunological consequences of challenge infection. Clinical data were analyzed using two-way t-tests for comparison of data. The cytokine and proliferation data were not normally distributed, therefore, median values were plotted against time for all graphical representations of data. Data were analyzed for significant differences using a non-parametric Wilcoxon Rank Sum test, and two tailed p-values were calculated. Significant differences were reported when p < 0.05.
3. Results and discussion 3.1. Clinical, virological and serological response to infection Challenge infection resulted in a biphasic pyrexia in Group 2 ponies, but had no effect on temperature of Group 1 ponies (Fig. 1). Group 2 ponies had significantly higher rectal temperatures than Group
113
1 ponies on days 1, 2 and 7 post-infection. Other signs of clinical disease in Group 2 ponies post-infection were limited to muco-purulent nasal discharge, which was evident on days 1–7 inclusive, post-infection. These clinical outcomes were consistent with resistance to challenge infection in the older, previously challenged, Group 1 ponies, as has been reported (Kydd et al., 2006). In contrast, Group 2 ponies showed clinical signs typical of EHV-1 respiratory infection. Results of virological testing demonstrated that post-challenge infection the nasal swabs from Group 1 ponies were negative. In contrast, three out of four Group 2 ponies post-challenge infection shed virus to variable levels: pony #10 shed virus on days 2–5 (13.2, 8.8, 83.6 and 4.4 pfu/ml, respectively), pony #17 shed virus on day 1 (22 pfu/ml) and pony #99 shed virus on days 1–3 (616, 66 and 13.2 pfu/ml, respectively). A single Group 2 pony was EHV-1 viremic postchallenge infection. These results further confirmed the resistance to infection of Group 1 ponies compared to Group 2 ponies, and were consistent with the Group 1 ponies having an immunoprotective memory immune response prior to the EHV-1 challenge. The immune response of both groups of ponies before and after challenge was characterized by measuring the virus-neutralizing antibody titer (Table 1). Prior to challenge infection, Group 1 ponies all exhibited low levels of virus neutralizing antibodies, and showed no sign of seroconversion in the 21 days post-challenge. In contrast, the Group 2 ponies had no detectable antibody in three out of four ponies, and seroconverted within 7–14 days of infection. Table 1 Virus neutralizing serum antibody titer to EHV-1 infection Pony#
Fig. 1. Rectal temperatures of Group 1 (&; n = 4) and Group 2 (*; n = 4) ponies after intranasal challenge infection with EHV-1 on day 0. Means S.E.M.
Group 1 82 84 85 86 Group 2 10 17 95 99
Days post-challenge infection 0
7
14
21
16 16 16 16
16 32 16 8
32 16 32 8
32 16 8 32
0 4 0 0
32 16 16 8
32 64 64 64
32 64 64 64
114
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
The rapidity of seroconversion in Group 2 ponies suggests that these animals had had previous antigenic encounter with EHV-1. The lack of seroconversion in Group 1 ponies suggests a lack of stimulation of a humoral immune response consequent to challenge infection, perhaps consistent with a lack of viral proliferation due to pre-existing immunity. 3.2. Cytokine and proliferative cellular responses The IFN-g and IL-4 mRNA response to in vitro EHV-1 stimulation are shown as stimulation indexes in Fig. 2a and b, respectively. The IFN-g response was consistently high in Group 1 ponies, but appeared to be lower at the time of infection in Group 2 ponies, subsequently increasing by day 21 post-challenge. However, for IFN-g, there were no significant
differences between Groups 1 and 2 on any day. For IL-4, Group 1 responses were low throughout the experiment, while Group 2 ponies demonstrated an increasing response post-infection, which was significantly different from Group 1 ponies on day 21 ( p = 0.03; two-tailed test). When the IFN-g/IL-4 SI ratio is examined (Fig. 2c), Group 1 ponies show consistently high values, while Group 2 ponies have consistently low values. These values were significantly different on day 7 ( p = 0.03), and the difference approached significance on day 21 ( p = 0.06). Overall, it was notable that the IL-4 SI values indicated no change in IL-4 transcription compared to the housekeeping gene in response to EHV-1 stimulation in immune Group 1 ponies (i.e. SI 1), while there were large increases in IFN-g transcription, which were most evident in the immune Group 1 ponies. Similarly,
Fig. 2. Median cytokine mRNA and proliferative cellular responses of PBMCs to in vitro stimulation with EHV-1 of PBMCs from Group 1 (&; n = 4) and Group 2 (*; n = 4) ponies after intranasal challenge infection with EHV-1 on day 0. (a) IFN-g stimulation index; (b) IL-4 stimulation index; (c) IFN-g SI/IL-4 SI; (d) lymphoproliferation stimulation index.
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
the relative levels of IFN-g to IL-4 transcription were consistently higher in the immune, older and repeatedly infected Group 1 ponies. In sharp contrast, EHV-1 infection in non-immune younger Group 2 ponies led to increases not only in IFN-g transcription, but also in IL-4. The pattern of cytokine production in Group 1 animals was consistent with a response in immune animals that was strongly polarized towards IFN-g, possibly consistent with a Th-1 response, while non-immune Group 2 ponies with limited preinfection immune responses, generated a non-polarized cytokine response on challenge infection. Results of lymphoproliferation assays give an interesting perspective on the cytokine data. Postchallenge infection, lymphoproliferative responses increased in Group 1 ponies, but showed no change in Group 2 ponies (Fig. 2d); on day 21, the Group 1 lymphoproliferative response was significantly greater than that of the Group 2 ponies ( p = 0.03). The increase in lymphoproliferation in Group 1 ponies was evident on day 7, although it was not significantly different from Group 2 ponies at that time. This response in Group 1 ponies is consistent with a recall response consequent to the challenge exposure, and in the absence of evidence of seroconversion in these ponies or signs of viral infection, it is the one piece of data confirming exposure of the immune system of Group 1 ponies to EHV-1. The lack of lymphoproliferative responses in Group 2 ponies (there was actually a small decline) occurred despite increases in both IFN-g and IL-4 production, and may be consistent with previous reports of decreased lymphoproliferative responses to either mitogens, inactivated or live EHV-1, in the weeks post-EHV-1 infection, that have been interpreted as evidence of immunosuppression (Hannant et al., 1991; Kydd et al., 2006). These phenomena have been attributed to both soluble (TGF-b) and cellular factors (Charan et al., 1997; Hannant et al., 1999). It is also possible that the decreased or suppressed lymphoproliferative response may result from a failure of antigen presentation, which could result from the down-regulation of MHC I expression that follows EHV-1 infection (Ambagala et al., 2004; Rappocciolo et al., 2003). Another possibility, raised by the results of this study, is that the immunoregulatory cytokine response of non-immune ponies, characterized by both IFN-g and IL-4 production, was not capable of supporting lympho-
115
proliferation, which is a hallmark of immunity. In contrast, the mature and polarized IFN-g response of the older immune ponies, did support lymphoproliferation. In conclusion, this study demonstrated that ponies immune to EHV-1 infection have an immune response characterized by an IFN-g response to EHV-1, while non-immune ponies have a limited cytokine response prior to challenge infection, and develop an unpolarized IFN-g and IL-4 responses post-challenge. These characteristics of the immunoregulatory responses of immune and non-immune ponies are consistent with other protective responses to viral infection in horses, and should be considered in vaccine design for EHV-1.
Acknowledgements This study was supported by grants from the USDA and the Grayson–Jockey Club Research Foundation.
References Allen, G.P., Kydd, J.H., Slater, J.D., Smith, K.C., 1999. Advances in understanding of the pathogenesis, epidemiology, and immunological control of equid herpesvirus) abortion. In: Wernery, U., Wade, J.F., Mumford, J.A., Kaaden, O.-R. (Eds.), Equine infectious diseases. VIII. Proceedings of the Eighth International Conference, Dubai, 23–26 March 1998, R&W Publications, Newmarket, pp. 129–146. Ambagala, A.P., Gopinath, R.S., Srikumaran, S., 2004. Peptide transport activity of the transporter associated with antigen processing (TAP) is inhibited by an early protein of equine herpesvirus-1. J. Gen. Virol. 85, 349–353. Breathnach, C.C., Rudersdorf, R., Lunn, D.P., 2004. Use of recombinant modified vaccinia Ankara viral vectors for equine influenza vaccination. Vet. Immunol. Immunopathol. 98, 127–136. Breathnach, C.C., Soboll, G., Suresh, M., Lunn, D.P., 2005. Equine herpesvirus-1 infection induces IFN-g production by equine T lymphocyte subsets. Vet. Immunol. Immunopathol. 103, 207–215. Charan, S., Palmer, K., Chester, P., Mire-Sluis, A.R., Meager, A., Edington, N., 1997. Transforming growth factor-beta induced by live or ultraviolet-inactivated equid herpes virus type-1 mediates immunosuppression in the horse. Immunology 90, 586–591. Foote, C.E., Gilkerson, J.R., Whalley, J.M., Love, D.N., 2003. Seroprevalence of equine herpesvirus 1 in mares and foals on a large Hunter Valley stud farm in years pre- and post-vaccination. Aust. Vet. J. 81, 283–288. Hannant, D., O’Neill, T., Jessett, D.M., Mumford, J.A., 1991. Evidence for non-specific immunosuppression during the development of immune responses to equid herpesvirus-1. Equine Vet. J. Suppl. 1, 41–45.
116
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
Hannant, D., O’Neill, T., Ostlund, E.N., Kydd, J.H., Hopkin, P.J., Mumford, J.A., 1999. Equid herpesvirus-induced immunosuppression is associated with lymphoid cells and not soluble circulating factors. Viral Immunol. 12, 313–321. Horohov, D.W., 2000. Equine T-cell cytokines. Protection and pathology. Vet. Clin. North Am. Equine Pract. 16, 1–14. Kydd, J.H., Antczak, D.F., Allen, W.R., Barbis, D., Butcher, G., Davis, W., Duffus, W.P.H., Edington, N., Grunig, G., Holmes, M.A., Lunn, D.P., McCullock, J., O’Brien, M.A., Perryman, L.E., Tavernor, A.S., Williamson, S., Zhang, C., 1994. Report of the First International Workshop on Equine Leucocyte Antigens, Cambridge, UK, July 1991. Vet. Immunol. Immunopathol. 42, 1–60. Kydd, J.H., Townsend, H.G.G., Hannant, D., 2006. The equine immune response to equine herpesvirus-1: the virus and its vaccines. Vet. Immunol. Immunopathol. 111, 15–30. Kydd, J.H., Wattrang, E., Hannant, D., 2003. Pre-infection frequencies of equine herpesvirus-1 specific, cytotoxic T lymphocytes correlate with protection against abortion following experimental infection of pregnant mares. Vet. Immunol. Immunopathol. 96, 207–217.
Murali-Krishna, K., Altman, J.D., Suresh, M., Sourdive, D., Zajac, A., Ahmed, R., 1998. In vivo dynamics of anti-viral CD8 T cell responses to different epitopes. An evaluation of bystander activation in primary and secondary responses to viral infection. Adv. Exp. Med. Biol. 452, 123–142. O’Neill, T., Kydd, J.H., Allen, G.P., Wattrang, E., Mumford, J.A., Hannant, D., 1999. Determination of equid herpesvirus 1-specific, CD8+, cytotoxic T lymphocyte precursor frequencies in ponies. Vet. Immunol. Immunopathol. 70, 43–54. Rappocciolo, G., Birch, J., Ellis, S.A., 2003. Down-regulation of MHC class I expression by equine herpesvirus-1. J. Gen. Virol. 84, 293–300. Soboll, G., Horohov, D.W., Aldridge, B.M., Olsen, C.W., McGregor, M.W., Drape, R.J., Macklin, M.D., Swain, W.F., Lunn, D.P., 2003a. Regional antibody and cellular immune responses to equine influenza virus infection, and particle mediated DNA vaccination. Vet. Immunol. Immunopathol. 94, 47–62. Soboll, G., Whalley, J.M., Koen, M.T., Allen, G.P., Fraser, D.G., Macklin, M.D., Swain, W.F., Lunn, D.P., 2003b. Identification of equine herpesvirus-1 antigens recognized by cytotoxic T lymphocytes. J. Gen. Virol. 84, 2625–2634.
Short communication
Cytokine responses to EHV-1 infection in immune and non-immune ponies Dane K. Coombs a, T. Patton a, Andrea K. Kohler b, G. Soboll c, Cormac Breathnach d, Hugh G.G. Townsend e, D.P. Lunn b,* a
Department of Medical Sciences, School of Veterinary Medicine, University of Wisconsin, 2015 Linden Drive West, Madison, WI 53706, USA Department of Clinical Sciences, College of Veterinary Medicine, Colorado State University, 300 W. Drake Rd, Fort Collins, CO 80523, USA c Dartmouth Medical Hitchcock Center, Department of Physiology, HB 7700, Room 708 W., 1 Medical Center Dr, Lebanon, NH 03756, USA d Maxwell H. Gluck Equine Research Center, Department of Veterinary Science, University of Kentucky, Lexington, KY 40546-0099, USA e Department of Veterinary Internal Medicine, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, Sask., Canada S7N 5B4 b
Abstract Protecting equids against equine herpesvirus-1 (EHV-1) infection remains an elusive goal. Repeated infection with EHV-1 leads to protective immunity against clinical respiratory disease, and a study was conducted to measure the regulatory cytokine response (IFN-g and IL-4) in repeatedly infected immune ponies compared to non-immune ponies. Two groups of four ponies were established. Group 1 ponies had previously been infected on two occasions, and most recently 7 months before this study. Group 2 ponies had no history no vaccination or challenge infection prior to this study. Both groups were subjected to an intranasal challenge infection with EHV-1, and blood samples were collected pre-infection, and at 7 and 21 days post-infection for preparation of PBMCs. At each time point, the in vitro responses of PBMCs to stimulation with EHV-1 were measured, including IFN-g and IL-4 mRNA production, and lymphoproliferation. Group 1 ponies showed no signs of clinical disease or viral shedding after challenge infection. Group 2 ponies experienced a biphasic pyrexia, mucopurulent nasal discharge, and nasal shedding of virus after infection. Group 1 ponies had an immune response characterized both before and subsequent to challenge infection by an IFN-g response to EHV-1 in the absence of an IL-4 response, and demonstrated increased EHV-1-specific lymphoproliferation post-infection. Group 2 ponies had limited cytokine or lymphoproliferative responses to EHV-1 pre-challenge, and demonstrated increases in both IFN-g and IL-4 responses post-challenge, but without any lymphoproliferative response. Protective immunity to EHV-1 infection was therefore characterized by a polarized IFN-g dependent immunoregulatory cytokine response. # 2006 Elsevier B.V. All rights reserved. Keywords: Horses; Equine herpesvirus-1; IFN-g; IL-4
* Corresponding author. Tel.: +1 970 297 1274; fax: +1 970 297 1275. E-mail address: [email protected] (D.P. Lunn). 0165-2427/$ – see front matter # 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.vetimm.2006.01.013
110
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
1. Introduction Equine herpesvirus-1 (EHV-1) infection causes contagious respiratory disease, epidemic abortion and sporadic neurological disease (Allen et al., 1999; Kydd et al., 2006). Respiratory infection by EHV-1 is common in young horses, with many horses becoming infected in the first weeks or months of life (Foote et al., 2003). Clinical respiratory disease is most common in younger horses, and repeated infections are increasingly associated with subclinical disease as horses grow older (Allen et al., 1999). The duration of acquired immunity is shortlived and re-infection can occur within 3–6 months (Kydd et al., 2006). Life-long latent infection is a common sequelae to primary infection and latently infected horses may develop a recrudescent infection and shed virus with or without demonstrating clinical disease (Allen et al., 1999). Although modified live (MLV) and killed vaccines have been used in the equine industry in an attempt to control EHV-1 infection, success has been limited (Kydd et al., 2006). The basis of protective immunity to EHV-1 infection is not fully understood, although there is substantial evidence that a cellular response involving major histocompatability class I (MHC I) restricted CD8+ cytotoxic T-lymphocytes (CTLs) is protective (Kydd et al., 2006). Recently, the number of EHV-1 specific CTL precursors (CTLp) has been demonstrated to be the most accurate, measurable correlate of immunity to EHV-1 infection. A low frequency of CTLp is associated with an increase in severity of disease from EHV-1 infection, while a high frequency of CTLp reduces the duration and level of viremia post-EHV-1 infection (Kydd et al., 2003; O’Neill et al., 1999). Although CTLp numbers may be a reliable predictor of a protective immune response against EHV-1 infection, the limiting dilution assay used to measure this parameter is laborious and complex. An alternative marker of a cell-mediated immune response is the production of interferon-gamma (IFN-g) (Murali-Krishna et al., 1998), and we have recently demonstrated that the post-exposure immune response to EHV-1 infection is associated with IFN-g production and an increasing CTL response (Breathnach et al., 2005). It is possible
that the IFN-g response to EHV-1 infection is part of a T-helper 1 (Th-1) immunoregulatory response, as is the case for the protective immune response to equine influenza virus infection (Soboll et al., 2003a). Such Th-1 responses may be associated with protective immunity to viral infection in many species, and this type of polarization of the equine immune response has been reported (Horohov, 2000). The cytokines IFN-g and IL-4 are important regulators of the balance between a Th-1 and Th-2 immune response, and there have been no studies documenting their relative production in EHV-1 immune and non-immune horses. This study was designed to characterize the IFN-g/ IL-4 response to EHV-1 infection in ponies with a history that would predict that they would be either immune or non-immune to clinical disease consequent to infection. The goal was to determine the regulatory cytokine profile associated with these different immune responses to EHV-1 infection. To accomplish this, we studied older ponies that had experienced a number of EHV-1 infections (putative immune group), and compared them to younger ponies with no recent history of EHV-1 infection. The purpose of this study was to further characterize protective immunity to EHV-1.
2. Materials and methods 2.1. Animals All animal experiments were reviewed and conducted in accordance with the requirements of the Research and Animal Resources Committee, University of Wisconsin, Madison, WI. Eight clinically normal, intact male ponies were studied. Two groups of ponies were used for challenge experiments: Group 1 ponies (ponies 82, 84, 85 and 86) ranged from 26 to 29 months of age and had previously been intranasally infected twice with EHV-1/A183, with the most recent infection being 7 months prior to this study; Group 2 ponies (ponies 10, 17, 95 and 99) ranged from 14 to 17 months of age and had no prior history of vaccination or experimental challenge with EHV-1. Ponies were housed in separate rooms in a semi-isolation unit for the duration of the experiment.
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
2.2. Virus All experiments described in this paper used viral strain EHV-1 Army 183. The virus was propagated in Kentucky equine dermal (KyED) cells (generously provided by Dr. George Allen, The Gluck Equine Research Center, Lexington, KY). Virus-laden, cellfree supernatant was used for challenge infections, and the viral dose (pfu/ml) and virus isolation was determined by titration on KyED cells. 2.3. Experimental design All challenges were performed by intranasal instillation of 2 107 pfu/ml EHV-1/A183 through a 14-gauge catheter placed in the ventral nasal meatus. Two intranasal challenge experiments were conducted. In a first experiment, Group 1 ponies were challenged with EHV-1, while Group 2 ponies were unchallenged and used as a control group. Ponies were monitored daily for pulse and respiratory rate, rectal temperature and nasal discharge for 10 days following challenge. In the second experiment (conducted 6 weeks later), the Group 2 ponies were challenged with EHV-1 and monitored for 3 weeks post-challenge. Blood was collected by jugular venipuncture for serum sampling. Blood was collected into heparin for PBMC isolation for use in lymphoproliferation and cytokine mRNA assays, and for detection of viremia. Nasal swabs were also collected for detection of nasal virus shedding. 2.4. Virus isolation Detection of nasal virus shedding was performed using dacron nasal swabs (Baxter Healthcare Corporation, McGaw Park, IL). Nasal swabs were collected daily on day 0 (pre-challenge) and on days 1–10 and then on alternate days until day 21 postinfection. Viral shedding in nasal swab samples was measured in PFU/ml of sample. For this purpose, KyED cells were added to six well tissue culture plates (Corning Inc., Corning, NY) and allowed to adhere. When cell layers were 90% confluent, nasal swab samples were thawed and filter sterilized using a 0.45 mM syringe top filter (Pall Corp., Ann Arbor, MI). Filtered samples were added to cell
111
layers at 10-fold dilutions in duplicate and incubated at 37 8C, 4% CO2 for 1.5 h. Inoculums were then removed and plates were rinsed with minimal essential media (MEM: GibcoBRL, Life Technologies, Grand Island, NY) plus 2% fetal calf serum (FCS). Subsequently 3 ml of MEM containing 0.75% methyl cellulose (Sigma), 100 U/ml penicillin, 100 mg/ml streptomycin, and 2% FCS were added to each well. Plates were incubated for 3 days at 37 8C, 4% CO2, before removal of the methylcellulose media and staining with 1 ml of crystal violet (Fisher Scientific, Fair Lawn, NJ) per well for 30 min. After rinsing with water and drying, plaques were counted and the PFU/ml of sample was calculated. Viremia was detected from heparinized blood samples during experiment I on days 0, 2, 4, 7, 8 and 10 days post-infection for Group 1 ponies and on days 0 and 7 for Group 2 ponies. During experiment II, viremia was measured daily on days 0–10, 12, 14 and 21 for the infected Group 2 ponies. For detection of viremia, 15 ml of blood was collected into heparin and red blood cells were allowed to settle. The plasma, containing PBMCs, was collected, white blood cells were pelleted and washed three times in PBS and subsequently added to 25 cm2 tissue culture flasks (Corning Inc.) containing a 90% confluent monolayer of KyED cells. Flasks were incubated for 7 days at 37 8C, 4% CO2 and checked daily for appearance of cytopathic effect (CPE). After 7 days, the virus containing supernatants were recovered by freeze–thawing and 500 ml aliquots were added to a fresh 25 cm2 tissue culture flasks containing a 90% confluent monolayer of KyED cells. These second pass flasks were incubated for further 7 days at 37 8C, 4% CO2 and appearance of cytopathic effect was recorded as a positive result. 2.5. Virus neutralization assay Serum antibody titers to EHV-1 were determined during days 0, 7, 14 and 21 post-infection for Groups 1 and 2 ponies. The amount of neutralizing antibody titer in the sera was determined by a virus neutralization assay at the Wisconsin State Veterinary Diagnostic Laboratory (Madison, WI) as previously described (Soboll et al., 2003b).
112
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
2.6. Proliferation assay Heparinized blood was collected for PBMC isolation on days 0, 7 and 21 post-infection for both groups of ponies. PBMCs were prepared by overlaying leukocyte-rich plasma on Histopaque-1077 (Sigma Chemical, St. Louis, MO) as previously described (Kydd et al., 1994). Isolated PBMCs were re-suspended at 2 106 cells/ml in RPMI 1640 (Gibco/Invitrogen) containing 2 mM L-glutamine, 10 mg streptomycin/ml, 10,000 units penicillin/ml, 250 mg amphotericin B/ml, 2 mM sodium pyruvate, 2 mM B-mercaptoethanol, 10 mM HEPES and 10% FBS. PBMCs were assayed in round-bottom 96-well microplates (Fisher Scientific). A total of 2.5 105 PBMCs/well were incubated with heat-inactivated EHV-1 (2 1010 pfu/ml), or a media control for 5 days at 37 8C in 5% CO2. At that time, 1 mCi [3H] thymidine was added to each well, and the cells were incubated for an additional 12 h before harvesting with a Packard Filter Mate (Packard, Meriden, CT) cell harvester. The uptake of [3H] thymidine was measured by liquid scintillation counting with a Top CountTM cell counter (Hewlett Packard, Palo Alto, CA). Stimulation indices (SI) were calculated based on the following equation: [cpm (virus-stimulated)]/[cpm (medium-stimulated)]. 2.7. Quantification of IFN-g and IL-4 mRNA Aliquots of 1.6 106 medium or EHV-1-stimulated equine PBMC, prepared as described above, were placed in duplicate wells of a 24-well plate, and incubated for 48 h at 37 8C, 5% CO2. Assays for IFNg mRNA expression were performed as described previously (Breathnach et al., 2004, #2825). Briefly, the contents of duplicate wells were pooled and resuspended in 300 ml RNA Stat 60 (Tel-Test, Friendswood, TX) and frozen at 70 8C until RNA extraction according to the manufacturer’s instructions. The RNA concentration was estimated by spectrophotometry, and cDNA was generated using with Moloney’s murine leukaemia virus reverse transcriptase (Promega, Madison, WI) and oligo dT primers (Promega). Expression of equine IFN-g, IL-4 and b-actin in the sample cDNA was determined by performing realtime PCR as previously described (Breathnach et al., 2004). The primer and probe sequences used were as follows: IFN-g forward primer: 50 -CGCAAAGCAA-
TAAGTGAACTCATC-30 ; IFN-g reverse: 50 -CGAAATGGATTCTGACTCCTCTTC-30 ; IFN-g probe: 50 56-FAM-TCTGTCGCCCAAAGCTAACCTGAGGAA-3BHQ-1-30 ; IL-4 forward primer: 50 -TCGTGCATGGAGCTGACTGTA-30 ; IL-4 reverse: 50 -GCCCTGCAGATTTCCTTT-CC-30 ; IL-4 probe: 50 -56-FAMCCTTTGCTGGCCCGAAGAACACAGA-3BHQ-130 ; b-actin forward primer: 50 -AGCGAAATCGTGCGTGACA-30 ; b-actin reverse: 50 -GCCATCTCCTGCTCGAAGT-C-30 ; b-actin probe: 50 -HEX-CAAGGAGAAGCTCTGCTATGTCGCCCT-BHQ2-30 . Fluorescent probes were used to measure amplified product in a thermocycler (iCycler; Bio-Rad Laboratories, Hercules, CA). Assays were run in triplicate in capped 96-well optical plates (Bio-Rad) containing 100 ng cDNA, 0.5 ml forward and reverse primers (10 mM), 5 ml probe (2 mM; IDT, Caralvile, IA), and 12.5 ml master mix (Gibco/Invitrogen); volume of reaction mixtures, 25 ml. Plasmids encoding equine IL-4 (pUC 2.1), equine IFN-g (pWRG 1647) and b-actin (pCR 2.1) were used to establish a standard curve on each 96well plate. Standard curves were generated by using the plasmid DNA at dilutions ranging from 1015 to 1022 M/ml. Each cytokine was analyzed on a separate experimental plate and amplification was performed by using the following procedure: 3 min at 95 8C followed by 45 cycles of 95 8C for 30 s and 60 8C for 60 s. Separate standard curves for IFN-g, IL-4 and bactin were established using the log dilutions of plasmid standards. On each plate, reaction mixtures containing no cDNA were included to control for contaminating DNA within the reagents. The transcript levels of IFN-g and IL-4 were normalized by using the transcript level value for bactin from the same sample (IFN-g/b-actin or IL-4/bactin). The herpes virus-specific cytokine response was expressed as stimulation index (SI) using the following equation: cytokine copy numberðvirus-stimulatedÞ= b-actin copy numberðvirus-stimulatedÞ cytokine copy numberðmedium-stimulatedÞ= b-actin copy numberðmedium-stimulatedÞ Additionally, the tendency towards an IFN-g or IL4 response was determined by the following equation: IFN-g SI/IL-4 SI.
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
2.8. Statistical analyses Comparisons were made for post-challenge responses between Group 1 ponies and Group 2 ponies, to determine the different clinical and immunological consequences of challenge infection. Clinical data were analyzed using two-way t-tests for comparison of data. The cytokine and proliferation data were not normally distributed, therefore, median values were plotted against time for all graphical representations of data. Data were analyzed for significant differences using a non-parametric Wilcoxon Rank Sum test, and two tailed p-values were calculated. Significant differences were reported when p < 0.05.
3. Results and discussion 3.1. Clinical, virological and serological response to infection Challenge infection resulted in a biphasic pyrexia in Group 2 ponies, but had no effect on temperature of Group 1 ponies (Fig. 1). Group 2 ponies had significantly higher rectal temperatures than Group
113
1 ponies on days 1, 2 and 7 post-infection. Other signs of clinical disease in Group 2 ponies post-infection were limited to muco-purulent nasal discharge, which was evident on days 1–7 inclusive, post-infection. These clinical outcomes were consistent with resistance to challenge infection in the older, previously challenged, Group 1 ponies, as has been reported (Kydd et al., 2006). In contrast, Group 2 ponies showed clinical signs typical of EHV-1 respiratory infection. Results of virological testing demonstrated that post-challenge infection the nasal swabs from Group 1 ponies were negative. In contrast, three out of four Group 2 ponies post-challenge infection shed virus to variable levels: pony #10 shed virus on days 2–5 (13.2, 8.8, 83.6 and 4.4 pfu/ml, respectively), pony #17 shed virus on day 1 (22 pfu/ml) and pony #99 shed virus on days 1–3 (616, 66 and 13.2 pfu/ml, respectively). A single Group 2 pony was EHV-1 viremic postchallenge infection. These results further confirmed the resistance to infection of Group 1 ponies compared to Group 2 ponies, and were consistent with the Group 1 ponies having an immunoprotective memory immune response prior to the EHV-1 challenge. The immune response of both groups of ponies before and after challenge was characterized by measuring the virus-neutralizing antibody titer (Table 1). Prior to challenge infection, Group 1 ponies all exhibited low levels of virus neutralizing antibodies, and showed no sign of seroconversion in the 21 days post-challenge. In contrast, the Group 2 ponies had no detectable antibody in three out of four ponies, and seroconverted within 7–14 days of infection. Table 1 Virus neutralizing serum antibody titer to EHV-1 infection Pony#
Fig. 1. Rectal temperatures of Group 1 (&; n = 4) and Group 2 (*; n = 4) ponies after intranasal challenge infection with EHV-1 on day 0. Means S.E.M.
Group 1 82 84 85 86 Group 2 10 17 95 99
Days post-challenge infection 0
7
14
21
16 16 16 16
16 32 16 8
32 16 32 8
32 16 8 32
0 4 0 0
32 16 16 8
32 64 64 64
32 64 64 64
114
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
The rapidity of seroconversion in Group 2 ponies suggests that these animals had had previous antigenic encounter with EHV-1. The lack of seroconversion in Group 1 ponies suggests a lack of stimulation of a humoral immune response consequent to challenge infection, perhaps consistent with a lack of viral proliferation due to pre-existing immunity. 3.2. Cytokine and proliferative cellular responses The IFN-g and IL-4 mRNA response to in vitro EHV-1 stimulation are shown as stimulation indexes in Fig. 2a and b, respectively. The IFN-g response was consistently high in Group 1 ponies, but appeared to be lower at the time of infection in Group 2 ponies, subsequently increasing by day 21 post-challenge. However, for IFN-g, there were no significant
differences between Groups 1 and 2 on any day. For IL-4, Group 1 responses were low throughout the experiment, while Group 2 ponies demonstrated an increasing response post-infection, which was significantly different from Group 1 ponies on day 21 ( p = 0.03; two-tailed test). When the IFN-g/IL-4 SI ratio is examined (Fig. 2c), Group 1 ponies show consistently high values, while Group 2 ponies have consistently low values. These values were significantly different on day 7 ( p = 0.03), and the difference approached significance on day 21 ( p = 0.06). Overall, it was notable that the IL-4 SI values indicated no change in IL-4 transcription compared to the housekeeping gene in response to EHV-1 stimulation in immune Group 1 ponies (i.e. SI 1), while there were large increases in IFN-g transcription, which were most evident in the immune Group 1 ponies. Similarly,
Fig. 2. Median cytokine mRNA and proliferative cellular responses of PBMCs to in vitro stimulation with EHV-1 of PBMCs from Group 1 (&; n = 4) and Group 2 (*; n = 4) ponies after intranasal challenge infection with EHV-1 on day 0. (a) IFN-g stimulation index; (b) IL-4 stimulation index; (c) IFN-g SI/IL-4 SI; (d) lymphoproliferation stimulation index.
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
the relative levels of IFN-g to IL-4 transcription were consistently higher in the immune, older and repeatedly infected Group 1 ponies. In sharp contrast, EHV-1 infection in non-immune younger Group 2 ponies led to increases not only in IFN-g transcription, but also in IL-4. The pattern of cytokine production in Group 1 animals was consistent with a response in immune animals that was strongly polarized towards IFN-g, possibly consistent with a Th-1 response, while non-immune Group 2 ponies with limited preinfection immune responses, generated a non-polarized cytokine response on challenge infection. Results of lymphoproliferation assays give an interesting perspective on the cytokine data. Postchallenge infection, lymphoproliferative responses increased in Group 1 ponies, but showed no change in Group 2 ponies (Fig. 2d); on day 21, the Group 1 lymphoproliferative response was significantly greater than that of the Group 2 ponies ( p = 0.03). The increase in lymphoproliferation in Group 1 ponies was evident on day 7, although it was not significantly different from Group 2 ponies at that time. This response in Group 1 ponies is consistent with a recall response consequent to the challenge exposure, and in the absence of evidence of seroconversion in these ponies or signs of viral infection, it is the one piece of data confirming exposure of the immune system of Group 1 ponies to EHV-1. The lack of lymphoproliferative responses in Group 2 ponies (there was actually a small decline) occurred despite increases in both IFN-g and IL-4 production, and may be consistent with previous reports of decreased lymphoproliferative responses to either mitogens, inactivated or live EHV-1, in the weeks post-EHV-1 infection, that have been interpreted as evidence of immunosuppression (Hannant et al., 1991; Kydd et al., 2006). These phenomena have been attributed to both soluble (TGF-b) and cellular factors (Charan et al., 1997; Hannant et al., 1999). It is also possible that the decreased or suppressed lymphoproliferative response may result from a failure of antigen presentation, which could result from the down-regulation of MHC I expression that follows EHV-1 infection (Ambagala et al., 2004; Rappocciolo et al., 2003). Another possibility, raised by the results of this study, is that the immunoregulatory cytokine response of non-immune ponies, characterized by both IFN-g and IL-4 production, was not capable of supporting lympho-
115
proliferation, which is a hallmark of immunity. In contrast, the mature and polarized IFN-g response of the older immune ponies, did support lymphoproliferation. In conclusion, this study demonstrated that ponies immune to EHV-1 infection have an immune response characterized by an IFN-g response to EHV-1, while non-immune ponies have a limited cytokine response prior to challenge infection, and develop an unpolarized IFN-g and IL-4 responses post-challenge. These characteristics of the immunoregulatory responses of immune and non-immune ponies are consistent with other protective responses to viral infection in horses, and should be considered in vaccine design for EHV-1.
Acknowledgements This study was supported by grants from the USDA and the Grayson–Jockey Club Research Foundation.
References Allen, G.P., Kydd, J.H., Slater, J.D., Smith, K.C., 1999. Advances in understanding of the pathogenesis, epidemiology, and immunological control of equid herpesvirus) abortion. In: Wernery, U., Wade, J.F., Mumford, J.A., Kaaden, O.-R. (Eds.), Equine infectious diseases. VIII. Proceedings of the Eighth International Conference, Dubai, 23–26 March 1998, R&W Publications, Newmarket, pp. 129–146. Ambagala, A.P., Gopinath, R.S., Srikumaran, S., 2004. Peptide transport activity of the transporter associated with antigen processing (TAP) is inhibited by an early protein of equine herpesvirus-1. J. Gen. Virol. 85, 349–353. Breathnach, C.C., Rudersdorf, R., Lunn, D.P., 2004. Use of recombinant modified vaccinia Ankara viral vectors for equine influenza vaccination. Vet. Immunol. Immunopathol. 98, 127–136. Breathnach, C.C., Soboll, G., Suresh, M., Lunn, D.P., 2005. Equine herpesvirus-1 infection induces IFN-g production by equine T lymphocyte subsets. Vet. Immunol. Immunopathol. 103, 207–215. Charan, S., Palmer, K., Chester, P., Mire-Sluis, A.R., Meager, A., Edington, N., 1997. Transforming growth factor-beta induced by live or ultraviolet-inactivated equid herpes virus type-1 mediates immunosuppression in the horse. Immunology 90, 586–591. Foote, C.E., Gilkerson, J.R., Whalley, J.M., Love, D.N., 2003. Seroprevalence of equine herpesvirus 1 in mares and foals on a large Hunter Valley stud farm in years pre- and post-vaccination. Aust. Vet. J. 81, 283–288. Hannant, D., O’Neill, T., Jessett, D.M., Mumford, J.A., 1991. Evidence for non-specific immunosuppression during the development of immune responses to equid herpesvirus-1. Equine Vet. J. Suppl. 1, 41–45.
116
D.K. Coombs et al. / Veterinary Immunology and Immunopathology 111 (2006) 109–116
Hannant, D., O’Neill, T., Ostlund, E.N., Kydd, J.H., Hopkin, P.J., Mumford, J.A., 1999. Equid herpesvirus-induced immunosuppression is associated with lymphoid cells and not soluble circulating factors. Viral Immunol. 12, 313–321. Horohov, D.W., 2000. Equine T-cell cytokines. Protection and pathology. Vet. Clin. North Am. Equine Pract. 16, 1–14. Kydd, J.H., Antczak, D.F., Allen, W.R., Barbis, D., Butcher, G., Davis, W., Duffus, W.P.H., Edington, N., Grunig, G., Holmes, M.A., Lunn, D.P., McCullock, J., O’Brien, M.A., Perryman, L.E., Tavernor, A.S., Williamson, S., Zhang, C., 1994. Report of the First International Workshop on Equine Leucocyte Antigens, Cambridge, UK, July 1991. Vet. Immunol. Immunopathol. 42, 1–60. Kydd, J.H., Townsend, H.G.G., Hannant, D., 2006. The equine immune response to equine herpesvirus-1: the virus and its vaccines. Vet. Immunol. Immunopathol. 111, 15–30. Kydd, J.H., Wattrang, E., Hannant, D., 2003. Pre-infection frequencies of equine herpesvirus-1 specific, cytotoxic T lymphocytes correlate with protection against abortion following experimental infection of pregnant mares. Vet. Immunol. Immunopathol. 96, 207–217.
Murali-Krishna, K., Altman, J.D., Suresh, M., Sourdive, D., Zajac, A., Ahmed, R., 1998. In vivo dynamics of anti-viral CD8 T cell responses to different epitopes. An evaluation of bystander activation in primary and secondary responses to viral infection. Adv. Exp. Med. Biol. 452, 123–142. O’Neill, T., Kydd, J.H., Allen, G.P., Wattrang, E., Mumford, J.A., Hannant, D., 1999. Determination of equid herpesvirus 1-specific, CD8+, cytotoxic T lymphocyte precursor frequencies in ponies. Vet. Immunol. Immunopathol. 70, 43–54. Rappocciolo, G., Birch, J., Ellis, S.A., 2003. Down-regulation of MHC class I expression by equine herpesvirus-1. J. Gen. Virol. 84, 293–300. Soboll, G., Horohov, D.W., Aldridge, B.M., Olsen, C.W., McGregor, M.W., Drape, R.J., Macklin, M.D., Swain, W.F., Lunn, D.P., 2003a. Regional antibody and cellular immune responses to equine influenza virus infection, and particle mediated DNA vaccination. Vet. Immunol. Immunopathol. 94, 47–62. Soboll, G., Whalley, J.M., Koen, M.T., Allen, G.P., Fraser, D.G., Macklin, M.D., Swain, W.F., Lunn, D.P., 2003b. Identification of equine herpesvirus-1 antigens recognized by cytotoxic T lymphocytes. J. Gen. Virol. 84, 2625–2634.