VIRAL IMMUNOLOGYVolume 15, Number 2, 2002© Mary Ann Liebert, Inc.Pp. 385–393

BHV-1–Specific CD41, CD81, and gd T Cells in Calves Vaccinated with One Dose of a

Modified Live BHV-1 Vaccine

JANICE J. ENDSLEY,1 MARK J. QUADE,1 BRETT TERHAAR,2

and JAMES A. ROTH1

ABSTRACT

Expression of the high-affinity interleukin 2 receptor a chain (CD25) was used to monitorantigen-specific activation of T lymphocyte subsets (CD41, CD81, and gd T cells) from cat-tle immunized with modified live bovine herpesvirus–1 (BHV-1). Calves seronegative forBHV-1 were either vaccinated with one dose of modified live vaccine containing BHV-1 ornot vaccinated to serve as negative controls. Two animals vaccinated 7 and 5 weeks beforethe start of the experiment with two doses of modified live vaccine containing BHV-1 servedas positive controls. Blood samples were taken from the nonvaccinate group, the positivecontrol group, and the vaccinate group at 0, 21, 35, 60, and 90 days postinoculation (PI).Isolated peripheral blood mononuclear cells from immunized and control animals were in-cubated for 5 days with and without live BHV-1 ISU99. Compared to the nonvaccinates, asignificant (p , 0.05) increase in expression of CD25 by CD41, CD81, and gd T lympho-cytes from the vaccinate group was detected following in vitro exposure to live BHV-1 aftervaccination. This is apparently the first report using live BHV-1 to stimulate lymphocytesin vitro and showing CD81 T cell activation. Peripheral blood from the positive control an-imals was depleted of CD41, CD81, or gd T lymphocytes prior to incubation with BHV-1to assess bystander activation in the CD25 expression assay. When incubated with live BHV-1, depletion of CD41 T cells depressed the expression of CD25 by CD81 T cells, but not gdT cells. Depleting CD81 or gd T cells prior to in vitro culture with BHV-1 did not affect theexpression of CD25 by the remaining T lymphocyte subsets. Vaccinates were protected fromchallenge with virulent BHV-1 at 110 days postvaccination compared to nonvaccinates. Ex-pression of CD25 appears to be a useful marker for evaluating induction of antigen-specificT lymphocyte subset responses following vaccination.

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1Department of Veterinary Microbiology and Preventative Medicine, College of Veterinary Medicine, Iowa StateUniversity, Ames, Iowa.

2Agri Laboratories Ltd., St. Joseph, Missouri.

INTRODUCTION

BOVINE HERPESVIRUS 1 (BHV-1) is a DNA virus that causes significant respiratory and reproductive dis-ease in cattle (1). BHV-1–induced disease continues despite the availability of numerous attenuated

and inactivated BHV-1 vaccines. Determining the relative importance of the lymphocyte subsets involvedin a protective memory response to BHV-1 would provide important direction for the development of moreeffective vaccines. The humoral immune response mediated by B lymphocytes following vaccination ornatural exposure to BHV-1 is farily well characterized (1,17). Protective antibodies produced in responseto infection or vaccination with BHV-1 act by inhibiting attachment and penetration of host cells by virus(1). Antibodies against BHV-1 have been detected as long as 30 months following vaccination (16). Whetherthese antibody responses correlate with long-term protection from BHV-1 infection has not been demon-strated, as challenge studies are usually done within a few weeks of vaccination.

The importance of immunity mediated by T lymphocytes in recovery or protection from BHV-1 is notwell characterized. Peripheral blood mononuclear cells (PBMC) from animals exposed to BHV-1 prolifer-ate (6,15) and secrete IFNg and IL-2 in response to inactivated BHV-1 in vitro (15). The in vitro prolifer-ative response of PBMC to inactivated BHV-1 is ameliorated following depletion of the CD41 T cell sub-set. Cytotoxic activity by CD41 and CD81 T cells against BHV-1–infected targets has also been described(7,18). Proliferation and cytotoxicity are important diagnostic indicators of memory responses by T lym-phocytes to vaccine antigens. Common methods for measuring antigen-specific lymphocyte proliferationand cytotoxicity use radioactive material and require additional steps to identify the subset of lymphocytesresponding. The use of an activation marker allows for the simultaneous detection of antigen-specific re-sponses and T cell phenotype via multicolor flow cytometry. The interleukin 2 receptor a chain (CD25) isa very sensitive activation marker that is expressed on the surface of lymphocytes following recognition ofspecific antigen. When compared as activation markers for mitogen-stimulated lymphocytes, CD25 andMHC II yield very similar results (12). CD25 expression has also been used to demonstrate antigen-spe-cific T cell responses to inactivated BHV-1 (12), bovine respiratory syncytial virus (11), and Mycobac-terium avium subspecies paratuberculosis (19). The bovine CD41 and WC11 gdT cell subsets, known tohave predominant roles in immunity to bovine tuberculosis, proliferate and express high levels of CD25 af-ter in vitro exposure to Mycobacterium bovis (14). Selection of cells that express CD25 following in vitroculture with Epstein Barr virus (EBV) greatly increases the concentration of specific cytotoxic T lympho-cytes that can be generated to treat EBV lymphoproliferative disorders (13). The CD41, CD81, and gd Tcells each recognize antigen differently. It is important to be able to determine if a vaccine activates eachof these T cell subsets. CD25 expression is an excellent indicator of antigen specific activation, but doesnot provide information about the functional role of the T cell subset responding to in vitro antigen. Iden-tifying the T cell subsets activated by in vitro BHV-1 exposure, however, is an important diagnostic indi-cator of previous antigen exposure and the first step towards focusing on T cell subsets whose function canbe exploited for vaccine design. The objectives of this study were (1) to use expression of CD25 to deter-mine if cattle vaccinated with modified live BHV-1 according to label directions (one dose) have CD41,CD81, and/or gd T cells that become activated when exposed to live BHV-1 in vitro, (2) to determine theeffect of depleting individual T lymphocyte subsets from the peripheral blood mononuclear cell pool on ex-pression of CD25 by remaining T lymphocytes in response to in vitro BHV-1, and (3) to determine if amodified live BHV-1 vaccine provides protective immunity at 110 days after vaccination.

MATERIALS AND METHODS

Animals. Twenty-one cross-bred beef steers, age 12–16 months, were used in the study. Animals werenegative for BHV-1 based on virus antibody titer and in vitro lymphocyte responsiveness to antigen. Ani-mals were randomly assigned to treatment groups; six animals served as negative controls, while 15 weregiven a 2-mL intramuscular dose of Titanium 5 L5 (Agri Labs, St. Joseph, MO) containing modified liveBHV-1, bovine viral diarrhea virus genotype 1 (BVDV1), BVDV genotype 2 (BVDV2) , bovine respira-tory syncytial virus (BRSV), parainfluenza 3 (PI3), and killed Leptospira bacterin (canicola, grippotyphosa,

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hardjo, icterohaemorragiae, pomona). Two previously vaccinated animals with known positive T cell re-sponses to BHV-1 were used as positive controls to validate the assay at each time point. These animalshad received 2 doses of BRSV Vac-4 (Miles Inc., Shawnee Mission, KS) containing modified live BVDV1,BHV-1, PI3, and BRSV approximately 7 and 5 weeks before onset of the trial. Animals were monitoredimmediately after injection and for the next 7 days for adverse reactions or problems associated with vac-cination.

Challenge. At 110 days after vaccination animals in all treatment groups were challenged intranasallywith 108.2 TCID50 of live Cooper strain BHV-1 (obtained from USDA-APHIS National Veterinary Ser-vices Laboratories, Ames, IA). Clinical signs and rectal temperatures were recorded for 14 days followingchallenge. Scores for clinical signs of BHV-1 infection were assigned based on the relative level of lethargy,nasal discharge, ocular discharge, nasal mucosa lesions, dyspnea, diarrhea, coughing, and anorexia.

Measurement of T cell activation. Antigen-specific T cell subset activation was detected using a mod-ification of a CD25 expression procedure described previously (12). In the modified procedure, lympho-cytes were cultured and later stained for flow cytometry in one 96-well plate. Due to time constraints, itwas not feasible to perform the CD25 expression assay on peripheral blood from all animals in the trial.Instead, 10 animals from the vaccinate group and three animals from the nonvaccinate group were randomlychosen to monitor in vitro T cell activation by BHV-1 following vaccination. The same animals were usedduring all sampling periods. Two animals previously vaccinated with two doses of modified live vaccinewere also sampled for CD25 expression at the same time points as the vaccinate and nonvaccinate group,to validate the assay at each time point. Peripheral blood mononuclear cells (PBMC) were isolated from 50mL of citrated blood, washed twice in Hanks balanced salt solution (HBSS) without calcium and magne-sium, and resuspended at 4 3 105 in 200 mL of RPMI media containing 2 mM L-glutamine, penicillin (60mg/mL), 20 mM HEPES, and 10% fetal bovine serum. Triplicate wells of PBMCs were cultured alone andwith a laboratory strain of BHV-1 (strain ISU99), or Concanavalin A, for 5 days at 39°C. The ISU99 strainwas found to be useful as a live antigen for stimulating lymphocytes in vitro. We therefore did not have touse a killed BHV-1 virus as we had in a previous trial (12). The ISU99 isolate caused mild cytopathic ef-fects in cultured MDBK cells and was identified as BHV-1 using fluorescently labeled antibody providedby the Iowa State University Veterinary Diagnostic Laboratory (Ames, IA). The titer of the BHV-1 isolatewas 108 TCID50 and was given the designation of BHV-1 ISU99 for reference purposes. The amount ofviral antigen to use in the CD25 expression assay was optimized using positive control animals.

Following 5 days of culture, cells were washed twice with HBSS and twice with a PBS solution con-taining 0.5% BSA and 0.1% NaN3 (PBS11). The washed cells were then stained with murine monoclonalantibodies specific for bovine CD4 (CACT138A; VMRD, Pullman, WA), CD8 (CACT80C; VMRD), andgd (GB21A; VMRD) T cells and a murine monoclonal antibody specific for bovine CD25 (CACT108A;VMRD). The antibodies specific for CD4, CD8, and a matched isotype control were an IgG1 isotype, whileantibodies specific for gd TCR and a matched isotype control were an IgG2b isotype. The antibody spe-cific for CD25 was an IgG2a isotype. Following a 30-min incubation at 4°C, cells were washed four timeswith PBS11 and then stained with phycoerythrin-labeled anti-IgG2a and either fluorescein isothiocyanate(FITC)–labeled anti-IgG2b (gd and isotype control) or FITC-labeled anti-IgG1 (CD4, CD8, and isotypecontrol; Caltag, Burlingame, CA). After 30 min at 4°C, cells were washed five times in HBSS (withoutCa21, Mg21, or phenol red) and fixed in 1% paraformaldehyde (Fisher, Fair Lawn, N.J.). To lower auto-fluorescence, a 1% formaldehyde solution that was made from dilution of a 10% ultrapure formaldehyde(PolySciences, Warrington, PA) was used as a fixative from the 21-day sampling point to the end of thetrial. Triplicate wells were pooled prior to flow cytometric analysis.

Depletion of T lymphocyte subsets. Following isolation, PBMC were resuspended at a concentrationof 2 3 107 cells/mL of coating buffer (PBS containing 10% FCS and 10 mM HEPES). Separate cell sus-pensions were incubated for 30 min at room temperature with murine monoclonal antibodies specific forbovine CD4 (CACT138A; VMRD), CD8 (CACT80C; VMRD), or gd TCR (GB21A; VMRD) at a con-centration of 1 mg/106 PBMC. Following incubation with primary antibody, cells were centrifuged at 1,000rpm and washed twice with coating buffer. After the second wash, cells were resuspended in 2 mL coat-ing buffer containing 4 3 107 dynabeads conjugated to a goat antibody specific for mouse IgG (DynalBiotech, Oslo, Norway). Cells were incubated with the dynabead conjugated secondary antibody for 30 min

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at room temperature on a slow rocking platform. Tubes were then applied to magnetic particle concentra-tor model MPC-L (Dynal Biotech, Oslo, Norway) on a rocking platform for 5 min. Supernatant was care-fully removed to clean tubes and then reapplied to the MPC-L for 5 min. Supernatant was again collected,and an aliquot was taken for counting cell concentration. The depleted population was resuspended at 4 3

105 cells in 200 mL of RPMI media containing 2 mM L-glutamine, 60 mg/mL penicillin, 20 mM HEPES,and 10% fetal bovine serum. Activation of the depleted lymphocyte population by in vitro BHV-1 was de-termined as described above.

Data analysis. Antigen-specific T cell responsiveness to in vitro antigen was determined by flow cyto-metric analysis. A total of 5,000 gated cells per sample were counted on a Becton Dickinson FACScan (SanJose, CA). Data are reported as an expression index, which is a ratio of the lymphocyte responsiveness fromeach animal when stimulated with BHV-1 versus when not stimulated with antigen (12).

Expression index 5 [(% stimulated T cells positive for CD25) * (mean fluorescent intensity of stimu-lated T cells positive for CD25)]/[(% nonstimulated T cells positive for CD25) * (mean fluorescent in-tensity of nonstimulated T cells positive for CD25)]

Prior to calculation of the expression index, isotype values are subtracted from sample values to correctfor background fluorescence. A Wilcoxon rank sum test was used to determine the level of significance ofthe differences in expression indices between non-vaccinate and vaccinate responses to BHV-1. Tempera-ture data and clinical score data were analyzed by the Mixed Procedure of SAS (SAS Institute, Cary, NC).The error associated with the repeated measures analysis of the temperature data was modeled using a firstorder autoregressive structure. In this analysis, a significant day * treatment group interaction was detected;thus, means within a day were further assessed by a least significant difference test, in which all pairs ofmeans within a day were compared. Clinical scores were assessed by the Kruskal-Wallis nonparametrictest. When this test revealed significant differences (p , 0.05), the groups were compared by Dunn’s pro-cedure in an all pair-wise fashion.

RESULTS

Adverse reactions to vaccination were not observed following inoculation. One animal in the vaccinategroup died at day 32 postvaccination. Necropsy determined the cause of death to be bloat. The deceasedanimal was from the larger vaccinate group, but was not part of the smaller vaccinate group randomly se-lected to assay BHV-1–specific T lymphocytes. Peripheral blood CD41, CD81, and gd T cells from ani-mals in both treatment groups responded to in vitro stimulation with Concanavalin A by substantially in-creasing expression of CD25, indicating that all T cell subsets were capable of responding. The averageCD25 expression during the trial by following Concanavalin A exposure CD41, CD81, and gd T cells fromthe vaccinate group was 31.6, 55.6, and 44.5, respectively, CD25 expression by CD41, CD81, and gd Tcells from the nonvaccinates in response to Concanavalin A during the trial averaged 32.2, 34.4, and 31.1,respectively. All three T cell subsets from the positive control animals were also activated by in vitro ex-posure to live BHV-1 (Table 1), indicating that the antigen used in the assay was capable of stimulating Tcells from positive animals. In response to in vitro BHV-1 antigen, CD41 T cells from the vaccinate groupsignificantly (p , 0.05) increased CD25 expression compared to the nonvaccinates at 21, 60, and 90 dayspostvaccination (Fig. 1). Compared to the nonvaccinate group, the vaccinate group had significantly (p ,

0.05) increased expression of CD25 by CD81 T cells in response to BHV-1 at 35–90 days PI. Expressionof CD25 by gd T cells from the vaccinate group in response to BHV-1 exposure was significantly greater(p , 0.05) than the nonvaccinates at 21, 35, and 90 days PI.

Peripheral blood mononuclear cells from the positive control animals were depleted of CD41, CD81, orgd T cells to determine the effect of each subset on in vitro BHV-1 specific activation by the remainingsubsets. Using magnetic bead separation, 92% , 88%, and 94% of the CD41, CD81, and gd T cells, re-spectively, were removed from isolated peripheral blood mononuclear cells. Depletion of CD41 T cellsprior to in vitro BHV-1 exposure decreased the percentage of CD81 T cells that expressed CD25 and the

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mean fluorescent intensity of the CD81/CD25 double positive cells (Table 1). While the activation responseof CD81 T cells to in vitro BHV-1 was decreased by CD41 T cell depletion, the response was not com-pletely prevented compared to background levels. The gd T cell activation response to in vitro BHV-1 wasnot prevented by depletion of the CD41 T cell population. Depleting CD81 T cells prior to in vitro BHV-1exposure did not prevent the activation response of the CD41 or gd T cell subsets. Depleting the PBMCpool of gd T cells prior to in vitro BHV-1 exposure did not decrease the activation response of the CD41

or CD81 T cell subsets.Animals in all treatment groups were seronegative for BHV-1 before vaccination and were clinically nor-

mal before challenge. Before BHV-1 challenge, cattle in the nonvaccinate treatment group were verified tobe seronegative for this virus and all vaccinated cattle had developed serum neutralizing antibody. Fol-lowing challenge with virulent BHV-1 at 110 days postvaccination, animals were observed for clinical signsand rectal temperatures were taken daily for 14 days. Nonvaccinated animals had significantly higher (p ,

0.05) rectal temperatures from 2 to 7 days postchallenge compared to either vaccinate group (Fig. 2). Clin-ical score data followed a pattern similar to the temperature data, with nonvaccinated animals having sig-nificantly higher (p , 0.05) clinical scores than vaccinates at 5, 6, 7, and 9 days after challenge (Fig. 2).

DISCUSSION

Vaccination with one dose of modified live BHV-1 generated CD41, CD81, and gd T cells in the pe-ripheral blood of cattle that became activated in response to live BHV-1 in culture. Activation of all threeT cell subsets by in vitro BHV-1 was evident until at least 90 days after vaccination. The vaccinated ani-mals were also shown to be protected from virulent BHV-1 challenge at 110 days after vaccination. Thisis the first report, that the authors are aware of, to successfully use live BHV-1 in vitro to stimulate anti-gen-specific T cell responses. Live BHV-1 is usually not used as an antigen in vitro due to cytopathic ef-fects on cultured cells. BHV-1 preferentially infects CD41 cells compared to CD81 and gd T cells (8) andnormally induces apoptosis in CD41 T cells (8,18). Previous work indicated that cattle vaccinated with

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TABLE 1. CD25 EXPRESSION INDEX , MEAN FLUORESCENT INTENSITY OF CD25, AND

THE PERCENTAGE OF T CELLS EXPRESSING CD25 BEFORE AND AFTER DEPLETION

OF CD41, CD81, OR gd T CELLS PRIOR TO IN VITRO EXPOSURE TO BHV-1

CD41 T cells CD81 T cells gd T cells

Treatment Exp Ina MFIb %CD251c Exp In MFI %CD251 Exp In MFI %CD251

In vitro BHV-1PBMCd 472.0 68.7 23.2 96.0 24.1 9.7 20.9 25.3 19.4CD41 depleted NA NA NA 36.7 14.8 5.6 50.9 20.5 20.6CD81 depleted 245.1 60.1 31.3 NA NA NA 15.6 19.9 15.4gd depleted 307.9 51.9 23.7 75.8 18.8 11.4 NA NA NA

Nonstimulated PBMC NA 10.6 2.6 NA 10.9 3.0 NA 11.4 4.6

aExpression index 5 [(% stimulated T cells positive for CD25) * (mean fluorescent intensity of stimulatd T cells posi-tive for CD25)]/[(% nonstimulated T cells positive for CD25) * (mean fluorescent intensity of nonstimulated T cells pos-itive for CD25)]. Isotype control values from each animal were subtracted from expression index to adjust for nonspecificinterference.

bGeometric mean fluorescent intensity (MFI) from phycoerythrin-labeled, CD25 double positive, cells from each sub-set.

cPercentage of the T lymphocyte subset population that is also positive for CD25.dPeripheral blood mononuclear cells (PBMC) and CD41, CD81, and gd T cell–depleted populations were obtained

from the two positive control animals vaccinated with two doses of modified live BHV-1 vaccine. The isolated PBMCpool was depleted of CD41, CD81, and gd T cells by 92%, 88%, and 94%, respectively.

NA, not applicable.

MLV BHV-1 have CD41 and gd T cells, but not CD81 T cells, which can be activated by killed BHV-1in vitro from 2 to 7 weeks after vaccination (12). The kinetic responses of T lymphocyte subsets to BHV-1 in the current trial are in general agreement with the previous report (12), with the exception thatpositive CD81 T cell responses were detected in the current trial. The BHV-1 used in the current trial wasa laboratory strain that had been passaged in vitro 58 times, and may have had reduced virulence and abil-ity to induce apoptosis. This may be why the BHV-1 isolate used here successfully stimulated lymphocytesin vitro, whereas other isolates tried previously did not (12). The BHV-1 ISU99 isolate has not yet beencharacterized at the molecular level to identify genetic alterations that may affect cytopathic capabilities.

The current trial provides preliminary evidence that detection of CD25 on T lymphocytes, following invitro exposure to BHV-1, may be a useful diagnostic indicator of a successful immune response to vacci-nation with live BHV-1. The technique is limited, however, to detecting T cell activation and does not pro-vide functional information about the responding T cell subset. The vaccinated cattle had solid protection

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FIG. 1. Expression of IL2ra chain (CD25) on T lymphocyte subsets after 5 days of in vitro exposure to live BHV-1. *p , 0.05, **p , 0.01; level of statistical significance between values for vaccinate and nonvaccinate treatmentgroups (mean 1 SEM).

from challenge with virulent BHV1 at 110 days after vaccination. The relative contribution of antibody andthe various T cell subsets to protection is not known. The roles of CD41 and CD81 cells in control of in-fectious disease of cattle have been more thoroughly examined than the roles of gd T cells. These subsetswould be expected to function as helper (CD41) and cytotoxic (CD81) T cells in controlling BHV-1 in-fection in cattle, as described for other viral infections. The decrease in CD81 T cell activation by BHV-1observed in the current trial, following in vitro depletion of CD41 T cells, supports previous research in-dicating an important helper role for CD41 T cells in protection from BHV-1 infection (6). The activatedCD81 T cells measured following vaccination in the current trial may represent bystander activation by cy-tokines produced from BHV-1–specific CD41 T cells. However, bystander activation of CD81 T cells didnot occur in a previous trial where killed BHV-1 antigen was used to stimulate lymphocytes from animalsvaccinated with a modified live BHV-1 vaccine (12). BHV-1–specific CD81 T cells may require cytokine

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FIG. 2. Mean daily rectal temperature and mean clinical score of cattle following challenge with virulent BHV-1at 110 days postvaccination. *p , 0.05; level of statistical significance between values for vaccinate and nonvaccinatetreatment groups.

help from CD41 T cells in order to proliferate to a level measurable in the CD25 expression assay. gd Tlymphocytes specific for BHV-1 were also detected following vaccination of cattle with a single dose ofmodified live vaccine in the current trial. This subset had a very high level of activation, but could poten-tially include CD81/gd double positive T cells. Cattle have a subset of gd T cells that also express CD81,but this population is very sparse in peripheral blood (10). The role that activated gd T cells specific forBHV-1 have in protection from BHV1 infection is unclear. Gamma delta T cells have been demonstratedto act as antigen-presenting cells and as cytotoxic T lymphocytes in cattle (4,5) and as helper T cells inmice (2,9). The results from the current trial indicate that activation of gd T cells by BHV-1 may not bedependent on cytokines produced by CD41 T cells.

The use of CD25 as an activation marker may represent a simple, nonradioactive alternative to lympho-cyte blastogenesis for detecting T cell–mediated immunity following vaccination. An additional advantgeof the CD25 expression assay is the ability to detect activation of specific subsets of T lymphocytes. De-tecting T cell subset activation will be especially useful for evaluating vaccines that are designed to clon-ally expand those subsets with a predominant role in protective immunity. A low level of background ex-pression can be a concern when using CD25 as an activation marker. The WC11 gd T cell populationconstitutively expresses a low level of CD25 under nonstimulatory conditions (3). The background level ofCD25 expression needs to be normalized for each animal for appropriate determination of the activationstatus, as described in the current trial. Converting the flow cytometric data into an index value normalizedbackground CD25 expression and simplifies the presentation of results from a fairly large number of ani-mals.

ACKNOWLEDGMENTS

We thank Bruce Pesch for his excellent flow cytometry instruction and Dr. Ken Platt for providing thelaboratory strain of BHV-1 that was designated ISU99 following several passages. This work was supportedby the John G. Salisbury Endowed Chair and by grants from Agri Laboratories, Diamond Laboratories, andfrom the Center for Advanced Technology Development Iowa Industrial Incentives program.

REFERENCES

1. Babiuk, L.A., S. Drunen Littel-van den Hurk, and S.K. Tikoo. 1996. Immunology of bovine herpesvirus 1 infec-tion. Vet. Microbiol. 53:31–42.

2. Boismenu, R., L. Feng, Y.Y. Xia, J.C. Chang, and W.L. Havran. 1996. Chemokine expression by intraepithelialgd T cells. Implications for the recruitment of inflammatory cells to damaged epithelia. J. Immunol. 157:985–992.

3. Collins, R.A., P. Sopp, K.I. Gelder, W.I. Morrison, and C.J. Howard. 1996. Bovine gdTcR1 T lymphocytes arestimulated to proliferate by autologous Theileria annulata–infected cells in the presence of interleukin-2. Scand. J.Immunol. 44:444–452.

4. Collins, R.A., D. Werling, S.E. Duggan, A.P. Bland, K.R. Parsons, and C.J. Howard. 1998. gd T cells present anti-gen to CD41 ab T cells. J. Leukoc. Biol. 63:707–714.

5. Daubenberger, C.A. , E.L. Taracha, L. Gaidulis, W.C. Davis, and D.J. McKeever. 1999. Bovine gd T-cell responsesto the intracellular protozoan parasite Theileria parva. Infect. Immunol. 67:2241–2249.

7. Denis, M., M.J. Kaashoek, J.T. van Oirschot, P.P. Pastoret, and E. Thiry. 1994. Quantitative assessment of the spe-cific CD41 T lymphocyte proliferative response in bovine herpesvirus 1 immune cattle. Vet. Immunol. Im-munopathol. 42:275–286.

7. Denis, M., M. Slaoui, G. Keil, L.A. Babiuk, E. Ernst, P.P. Pastoret, and E. Thiry. 1993. Identification of differenttarget glycoproteins for bovine herpes virus type 1–specific cytotoxic T lymphocytes depending on the method ofin vitro stimulation. Immunology. 78:7–13.

8. Eskra, L., and G.A. Splitter. 1997. Bovine herpesvirus-1 infects activated CD41 lymphocytes. J. Gen. Virol. 78(Pt9):2159–2166.

ENDSLEY ET AL.

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9. Ferrick, D.A., M.D. Schrenzel, T. Mulvania, B. Hsieh, W.G. Ferlin, and H. Lepper. 1995. Differential productionof interferon-g and interleukin-4 in response to Th1- and Th2-stimulating pathogens by gd T cells in vivo. Nature.373:255–257.

10. MacHugh, N.D., J.K. Mburu, M.J. Carol, C.R. Wyatt, J.A. Orden, and W.C. Davis. 1997. Identification of two dis-tinct subsets of bovine gd T cells with unique cell surface phenotype and tissue distribution. Immunology92:340–345.

11. Mcinnes, E., P. Sopp, C.J. Howard, and G. Taylor. 1999. Phenotypic analysis of local cellular responses in calvesinfected with bovine respiratory syncytial virus. Immunology. 96:396–403.

12. Quade, M.J., and J.A. Roth. 1999. Antigen-specific in vitro activation of T-lymphocyte subsets of cattle immu-nized with a modified live bovine herpesvirus 1 vaccine. Viral Immunol. 12:9–21.

13. Savoldo, B., M.L. Cubbage, A.G. Durett, J. Goss, M.H. Huls, Z. Liu, L. Teresita, A.P. Gee, P.D. Ling, M.K. Bren-ner, H.E. Heslop, and C.M. Rooney. 2002. Generation of EBV-specific CD41 cytotoxic T cells from virus naiveindividuals. J. Immunol. 168:909–918.

14. Smyth, A.J., M.D. Welsh, R.M. Girvin, and J.M. Pollock. 2001. In vitro responsiveness of gd T cells from My-cobacterium bovis–infected cattle to mycobacterial antigens: predominant involvement of WC1(1) cells. Infect.Immun. 69:89–96.

15. Tikoo, S.K., M. Campos, Y.I. Popowych, S. Drunen Littel–van den Hurk, and L.A. Babiuk. 1995. Lymphocyteproliferative responses to recombinant bovine herpes virus type 1 (BHV-1) glycoprotein gD (gIV) in immune cat-tle: identification of a T cell epitope. Viral Immunol. 8:19–25.

16. Van der Poel, W.H., J.A. Kramps, J. Quak, A. Brand, and J.T. van Oirschot. 1995. Persistence of bovine her-pesvirus-1–specific antibodies in cattle after intranasal vaccination with a live virus vaccine. Vet. Rec. 137:347–348.

17. van Oirschot, J.T., C.J. Bruschke, and P.A. van Rijn. 1999. Vaccination of cattle against bovine viral diarrhoea.Vet. Microbiol. 64:169–183.

18. Wang, C., and G.A. Splitter. 1998. CD41 cytotoxic T-lymphocyte activity against macrophages pulsed with bovineherpesvirus 1 polypeptides. J. Virol. 72:7040–7047.

19. Whist, S.K., A.K. Storset, and H.J. Larsen. 2000. The use of interleukin-2 receptor expression as a marker of cell-mediated immunity in goats experimentally infected with Mycobacterium avium ssp. paratuberculosis. Vet. Im-munol. Immunopathol. 73:207–218.

Address reprint requests to:Dr. James A. Roth

2156 College of Veterinary MedicineIowa State University

Ames, IA 50011

E-mail: jaroth@iastate.edu

Received November 20, 2001; accepted March 7, 2002.

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