Institute for Animal Health, Compton, Newbury, Berkshire RG20 7NN, UK1
Author for correspondence: Chris Howard. Fax +44 1635 577263. e-mail chris.howard{at}bbsrc.ac.uk
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Infection of animals with ncp BVDV generally causes a transient infection, with the animal exhibiting subclinical or mild symptoms of infection (Baker, 1995 ). However, if an animal becomes infected with ncp virus within the first trimester of pregnancy, virus can cross the placenta and infect the foetus (Brownlie et al., 1984
; Fredriksen et al., 1999
; Moennig & Liess, 1995
). This may result in the birth of a calf that is persistently infected (PI) and specifically immunotolerant to the infecting virus. Very little or no antibody to BVDV is evident in sera, and viral antigen has been reported to be widely distributed in tissues, most notably in cells and organs of the immune system (Bielefeldt Ohmann, 1988
; Bielefeldt Ohmann et al., 1987
; Sopp et al., 1994
). Superinfection of PI calves with a cp homologous strain of BVDV can cause death due to the onset of MD (Brownlie et al., 1984
). Calves PI with BVDV may suffer from reduced weight gain, growth retardation and higher rates of neonatal mortality, usually through secondary infection with enteric or respiratory pathogens. Additionally, PI animals are reported to suffer from a degree of immunosuppression, although the molecular basis for this has not yet been defined (Johnson & Muscoplat, 1973
; Muscoplat et al., 1973
; Potgieter, 1995
).
Many viruses are reported to subvert the normal host immune response during infection. This can aid the establishment and maintenance of viable infection within the host and limit the level of immune-mediated damage to host tissues. Several mechanisms by which viruses alter the immune response of the host have been described. These include virus latency, infection of immunoprivileged sites, synthesis of cytokine homologues and receptors, mutation of the viral genome, which subsequently prevents binding of viral peptides to host MHC class I and II molecules, inhibition of antigen processing and presentation pathways and interference with the host cellular machinery (reviewed by Spriggs, 1996 ; Tortorella et al., 2000
). A number of these effects have direct consequences on the ability of the antigen-presenting cells (APC) to stimulate an immune response.
BVDV has been reported to modulate functions of immune cells after infection in vitro, with increased production of nitric oxide from infected macrophages (Adler et al., 1994 ), decreased production of TNF
from lipopolysaccharide (LPS)-stimulated macrophages (Adler et al., 1996
) and reduction of Fc and C3 receptor expression on, and phagocytic activity of, alveolar macrophages (Welsh et al., 1995
). Additionally, several authors have reported that cells, isolated from PI animals, that are pivotal in control of the immune response are infected in vivo. These include the antigen-presenting myeloid cells, CD4+ and CD8+ T lymphocytes and B cells (Bielefeldt Ohmann et al., 1987
; Bielefeldt Ohmann, 1988
; Bruschke et al., 1998
; Sopp et al., 1994
). However, it has not been established whether APC from PI, specifically immunotolerant, cattle are compromised in their ability to induce immune responses to the virus. Any effect might play a role in the pathogenesis of MD or in the generalized immunosuppression noted in PI cattle.
A breeding programme at the Institute for Animal Health has produced cattle that are major histocompatibility complex (MHC) identical. A series of experiments was performed using T cells from cattle that were immune to BVDV, or that had not come into contact with the virus, and APC from an MHC-identical, PI animal, to determine whether being persistently infected compromised the ability of APC to present BVDV antigen to T cells.
![]() |
Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cell separation, culture and storage.
PBMC were separated by density gradient centrifugation (1·083 g/ml Histopaque; Sigma) from blood taken into heparin (10 units per ml blood; Leo). Cells were resuspended in tissue culture medium (TCM) consisting of RPMI-1640 medium with Glutamax I (Life Technologies), supplemented with 10% heat-inactivated FCS, 5x10-5 M 2-mercaptoethanol and 50 µg/ml gentamycin. Monocytes were isolated from PBMC, taken from the PI calf, after staining with an anti-CD14 monoclonal antibody (MAb), CC-G33 (Table 1), and incubation with anti-mouse IgG1 super-paramagnetic particles (Miltenyi Biotech). Labelled cells were isolated using a MiniMacs column (Miltenyi Biotech), following the manufacturers instructions. The purity of the cells was evaluated by flow cytometry and shown to be >96%. CD4+ and CD8+ T lymphocytes were obtained in a similar manner from non-PI calves that were seropositive (immune) or seronegative (naïve) using anti-CD4 or anti-CD8 MAbs, respectively (Table 1
). For convenience, these T cell populations are referred to as immune or naïve.
|
Proliferation assays.
Purified monocytes from the PI calf (PI monocytes), used directly or cultured for 3 days, were irradiated (20 Gy from a 137Cs source) and dilutions were incubated with 105 CD4+ or CD8+ T lymphocytes from naïve or immune calves. Total volumes were made up to 200 µl with TCM in 96-well U-bottomed microtitre plates (Becton Dickinson). Triplicate cultures were incubated for 5 days and 37 Bq [3H]thymidine (3H-TdR; DuPont) was added for 16 h (overnight) before harvesting. Incorporated radioactivity was determined by liquid scintillation counting. In some experiments, monocytes were purified from immune calves and infected by adding BVDV strain Pec515 at an m.o.i. of 2 per monocyte (Brownlie et al., 1984 ) for 3 days. These cells and uninfected control cells were used as APC with autologous CD4+ or CD8+ T lymphocytes.
Flow-cytometric analysis.
Two-colour staining of PBMC for leukocyte differentiation antigens and intracellular BVDV NS3 (p80) protein was performed by using a slight modification of the procedure described by Sopp et al. (1994) . All samples were diluted and washed in PBSa (PBS containing 1% BSA) and 0·1% sodium azide. Mouse MAbs to bovine CD antigens (Table 1
), optimally diluted, were added to 106 PBMC for 10 min at room temperature, washed three times and incubated for 10 min with optimally diluted goat anti-mouse secondary antibodies, conjugated to either FITC or PE (Southern Biotechnology Associates). After three washes, PBMC were fixed in 1% paraformaldehyde (Sigma) in PBS for 10 min. All subsequent washing used PBS/0·1% sodium azide/0·1% saponin (Sigma). After fixation, cells were washed three times and optimally diluted mouse anti-BVDV MAb or an isotype-matched control MAb was added to the samples. Cells were incubated for 10 min, washed and an FITC- or PE-conjugated secondary goat anti-mouse MAb to a different mouse Ig isotype was added. Immunofluorescent staining was analysed by using a FACScan (Becton Dickinson) and data were analysed by using WinMDI (obtained from Joseph Trotter, Scripps Research Institute, San Diego, CA, USA) and FCS Express (De novo Software).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Monocytes isolated from a calf PI with BVDV (PI monocytes) stimulate the proliferation of resting CD4+ T memory cells
PI monocytes, freshly isolated (Fig. 2a) or cultured for 3 days (Fig. 2b
), stimulated a proliferative response in immune CD4+ T lymphocytes. The proliferative response of the CD4+ T cells to PI monocytes that had been cultured for 3 days was consistently higher (approximately 10-fold) than the proliferative response induced by freshly isolated PI monocytes. CD4+ cells or CD14+ cells cultured alone did not incorporate [3H]thymidine (<500 c.p.m.). The data shown in Fig. 2
are representative of 14 separate experiments, with CD4+ T cells isolated from two BVDV-immune animals.
|
|
|
Flow-cytometric analysis of the CD8+ cells after 5 days cultured with the PI monocytes (Fig. 4h) indicated that the
TCR+ population comprised less than 10% of the total live cells in the wells, indicated by gated region 1 (R1) in Fig. 4(d)
.
TCR+ cells also accounted for 11% of the large blast-like cells indicated, by size, in region 2 (R2) (Fig. 4i
, m
). At time zero, analysis of the PBMC indicated that approximately 4% of the total PBMC were CD8+
TCR+(Fig. 4c
; upper-right quadrant). This was approximately 28% of the total cells that stained for CD8+ antigen. Therefore, the percentage of
TCR+ cells within the CD8+ population decreased with incubation from 28% to <10%. This implied that the proliferative response of CD8+ T cells to PI monocytes was due to an expansion of the population of
TCR+ T cells, indicating a BVDV antigen-specific response. The data presented in Fig. 4
are representative of three separate experiments.
|
CD4+ and CD8+ T lymphocytes from immune calves proliferate in response to in vitro-infected monocytes
Monocytes from calves immune to BVDV, cultured with BVDV Pec515 for 3 days, were shown to be infected with BVDV, with over 90% of the monocyte population being NS3+ (p80+) when assessed by flow cytometry (data not shown). Fig. 5 shows the proliferative response of autologous, purified CD4+ and CD8+ T lymphocytes to monocytes infected with BVDV or non-infected. The response is similar to that seen with 3-day-cultured PI monocytes and immune CD4+ (Fig. 2b
) and CD8+ (Fig. 3b
) T lymphocytes. The data are representative of three separate experiments.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In vitro studies with BVDV have shown that infection of monocytes or macrophages causes the synthesis of cytokines that may be responsible for the reduced ability to stimulate T cell responses to specific antigens and mitogens. Adler et al. (1996) reported a decrease in the secretion of TNF
from LPS-or Salmonella-stimulated bone marrow-derived macrophages that were infected with either ncp or cp BVDV. Differential priming of monocytes, by ncp or cp BVDV, for nitric oxide production (Adler et al., 1994
), the reduction of monocyte responses to chemotactic stimuli (Ketelsen et al., 1979
) and production of an inhibitor of IL-1 activity by infected monocytes (Jensen & Schultz, 1991
) have all been reported. Thus, it was possible that the specific immunotolerance to BVDV that is evident in PI cattle and is a central component of the pathogenesis of MD is a consequence of infection of APC in vivo.
Making use of cattle of the same MHC haplotype, it was established that ex vivo monocytes from a PI animal were able to stimulate resting CD4+ T memory cells. This implies that the APC have taken up exogenous BVDV antigen, processed that antigen via the endosomal pathway and presented the resultant peptides in association with MHC class II molecules. The response of the CD4+ T cells to BVDV antigen was equivalent to that seen in other antigen-specific systems (Knight & Macatonia, 1991 ; Schlesier et al., 1994
) and is similar to that reported by Rhodes et al. (1999)
.
Monocytes from a PI calf, cultured for 3 days, were more effective than fresh monocytes at inducing proliferative responses of memory T cells. This could be for a number of reasons. There was an increase in the number of infected cells after 3 days in culture, as almost 90% of the monocytes stained with MAb to the NS3 (p80) antigen whereas only approximately 43% of the freshly isolated monocytes stained. Additionally, placing monocytes into culture is known to cause a transitory increase in the ability of the cells to stimulate T cells and a corresponding upregulation of co-stimulatory and adhesion molecules, before full differentiation into macrophages, which display poor accessory cell function (Mayernik et al., 1983 ; Najar et al., 1990
). It is likely that the combination of an increase in APC function, due to culture, and an increase in the number of infected cells that present viral antigen is responsible for the increase in T cell responses to cultured PI monocytes when compared with ex vivo PI monocytes. Thus, APC infected in vivo and isolated from the PI animal and APC cultured for 3 days, which were a mixture of in vivo- and in vitro-infected cells, did not appear to be compromised in their ability to present BVDV antigen to CD4+ T cells and to stimulate a MHC class II-restricted T cell response.
PI monocytes were also able to stimulate a CD8+ MHC class I-restricted T cell response in CD8+ T cells isolated from BVDV-immune cattle. As with the CD4+ T cell response, 3-day-cultured monocytes stimulated a 10-fold higher proliferative response in CD8+ T cells than did the freshly isolated monocytes. Cross-presentation is a feature of dendritic cells and is a means by which non-replicating antigen, normally processed by the exogenous pathway and presented in the context of MHC class II molecules, is presented by MHC class I molecules with the subsequent induction of CD8+ T cell responses (Grommé et al., 1999 ). However, this pathway has not been reported for monocytes. Thus, the CD8+ T cell responses demonstrated here would be expected to be directed against viral antigen, processed via the endogenous pathway only and presented in association with MHC class I. The APC used here were infected naturally in vivo for the freshly isolated cells, or were a mixture of in vitro- and in vivo-infected cells when the APC were cultured for 3 days. These results suggest that APC from the PI animal were not compromised in their ability to stimulate an MHC class I-restricted T cell response and that BVDV does not exert a suppressive effect on the endogenous pathway of antigen processing.
Furthermore, the proliferative response of CD8+ as well as CD4+ T cells from immune calves to 3-day-cultured PI monocytes was similar to that noted with monocytes that were obtained from an immune calf that was not PI and which had been infected in vitro by incubation for 3 days with BVDV. This indicates that PI monocytes do not differ from normal monocytes in their capacity to differentiate into potent APC. The lack of a response with T cells from naïve animals confirmed that neither recognition of major or minor histocompatibility antigens by the responding T cells nor a non-specific effect of the virus on the APC was responsible for major component of the response seen.
Previous studies, from experiments that involved depletion of CD4+ and CD8+ T cells in vivo (Howard et al., 1992 ), and the observation that passive antibody can protect against transient, acute BVDV infection (Howard et al., 1989
) have been taken to indicate that CD4+ T cells and antibody, and not cytolytic CD8+ T cells, are the main component in the recovery from and immunity of animals to BVDV. This mechanism has also been demonstrated for mice infected with lymphocytic choriomeningitis virus (Planz et al., 1997
). However, a critical role for CD8+ T cells in the immune response should not be ruled out. The results presented here show that a CD8 memory T cell response is evident in previously infected cattle. Furthermore, shedding of BVDV in nasal secretions has been noted that persisted in the presence of serum neutralizing antibodies (Fray et al., 1998
; Howard et al., 1999
) and different effector cell populations may have varying degrees of importance in different tissues, as suggested for a murine gammaherpesvirus (Ehtisham et al., 1993
).
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Adler, H., Jungi, T. W., Pfister, H., Strasser, M., Sileghem, M. & Peterhans, E. (1996). Cytokine regulation by virus infection: bovine viral diarrhea virus, a flavivirus, downregulates production of tumor necrosis factor alpha in macrophages in vitro. Journal of Virology 70, 2650-2653.[Abstract]
Baker, J. C. (1995). The clinical manifestations of bovine viral diarrhea infection. Veterinary Clinics of North America Food Animal Practice 11, 425-445.
Bielefeldt Ohmann, H. (1988). BVD virus antigens in tissues of persistently viraemic, clinically normal cattle: implications for the pathogenesis of clinically fatal disease. Acta Veterinaria Scandinavica 29, 77-84.[Medline]
Bielefeldt Ohmann, H., Rønsholt, L. & Bloch, B. (1987). Demonstration of bovine viral diarrhoea virus in peripheral blood mononuclear cells of persistently infected, clinically normal cattle. Journal of General Virology 68, 1971-1982.[Abstract]
Brownlie, J., Clarke, M. C. & Howard, C. J. (1984). Experimental production of fatal mucosal disease in cattle. Veterinary Record 114, 535-536.[Medline]
Bruschke, C. J. M., Weerdmeester, K., Van Oirschot, J. T. & Van Rijn, P. A. (1998). Distribution of bovine virus diarrhoea virus in tissues and white blood cells of cattle during acute infection. Veterinary Microbiology 64, 23-32.[Medline]
Cook, J. K. A., Jones, B. V., Ellis, M. M., Jing, L. & Cavanagh, D. (1993). Antigenic differentiation of strains of turkey rhinotracheitis virus using monoclonal antibodies. Avian Pathology 22, 257-273.
Davis, W. C., Brown, W. C., Hamilton, M. J., Wyatt, C. R., Orden, J. A., Khalid, A. M. & Naessens, J. (1996). Analysis of monoclonal antibodies specific for the TcR. Veterinary Immunology and Immunopathology 52, 275-283.[Medline]
Ehtisham, S., Sunil-Chandra, N. P. & Nash, A. A. (1993). Pathogenesis of murine gammaherpesvirus infection in mice deficient in CD4 and CD8 T cells. Journal of Virology 67, 5247-5252.[Abstract]
Ellis, S. A., Staines, K. A., Stear, M. J., Hensen, E. J. & Morrison, W. I. (1998). DNA typing for BoLA class I using sequence-specific primers (PCR-SSP). European Journal of Immunogenetics 25, 365-370.[Medline]
Fray, M. D., Clarke, M. C., Thomas, L. H., McCauley, J. W. & Charleston, B. (1998). Prolonged nasal shedding and viraemia of cytopathogenic bovine virus diarrhoea virus in experimental late-onset mucosal disease. Veterinary Record 143, 608-611.[Medline]
Fredriksen, B., Press, C. M., Sandvik, T., Ødegaard, S. A. & Løken, T. (1999). Detection of viral antigen in placenta and fetus of cattle acutely infected with bovine viral diarrhea virus. Veterinary Pathology 36, 267-275.[Abstract]
Grommé, M., Uytdehaag, F. G. C. M., Janssen, H., Calafat, J., Van Binnendijk, R. S., Kenter, M. J. H., Tulp, A., Verwoerd, D. & Neefjes, J. (1999). Recycling MHC class I molecules and endosomal peptide loading. Proceedings of the National Academy of Sciences, USA 96, 10326-10331.
Hanby-Flarida, M. D., Okragly, A. J. & Baldwin, C. L. (1996). Autologous mixed leucocyte reaction and the polyclonal activation of bovine /
T cells. Research in Veterinary Science 61, 65-71.[Medline]
Houe, H. (1995). Epidemiology of bovine viral diarrhea virus. Veterinary Clinics of North America Food Animal Practice 11, 521-547.
Howard, C. J., Clarke, M. C. & Brownlie, J. (1985). An enzyme-linked immunosorbent assay (ELISA) for the detection of antibodies to bovine viral diarrhoea virus (BVDV) in cattle sera. Veterinary Microbiology 10, 359-369.[Medline]
Howard, C. J., Clarke, M. C. & Brownlie, J. (1989). Protection against respiratory infection with bovine virus diarrhoea virus by passively acquired antibody. Veterinary Microbiology 19, 195-203.[Medline]
Howard, C. J., Clarke, M. C., Sopp, P. & Brownlie, J. (1992). Immunity to bovine virus diarrhoea virus in calves: the role of different T-cell subpopulations analysed by specific depletion in vivo with monoclonal antibodies. Veterinary Immunology and Immunopathology 32, 303-314.[Medline]
Howard, C. J., Collins, R. A., Sopp, P., Brooke, G. P., Kwong, L. S., Parsons, K. R., Weynants, V., Letesson, J.-J. & Bembridge, G. P. (1999). T-cell responses and the influence of dendritic cells in cattle. Advances in Veterinary Medicine 41, 275-288.[Medline]
Jensen, J. & Schultz, R. D. (1991). Effect of infection by bovine viral diarrhea virus (BVDV) in vitro on interleukin-1 activity of bovine monocytes. Veterinary Immunology and Immunopathology 29, 251-265.[Medline]
Johnson, D. W. & Muscoplat, C. C. (1973). Immunologic abnormalities in calves with chronic bovine viral diarrhea. American Journal of Veterinary Research 34, 1139-1141.[Medline]
Ketelsen, A. T., Johnson, D. W. & Muscoplat, C. C. (1979). Depression of bovine monocyte chemotactic responses by bovine viral diarrhea virus. Infection and Immunity 25, 565-568.[Medline]
Knight, S. C. & Macatonia, S. E. (1991). Effect of HIV on antigen presentation by dendritic cells and macrophages. Research in Virology 142, 123-128.[Medline]
MacHugh, N. D., Bensaid, A., Howard, C. J., Davis, W. C. & Morrison, W. I. (1991). Analysis of the reactivity of anti-bovine CD8 monoclonal antibodies with cloned T cell lines and mouse L-cells transfected with bovine CD8. Veterinary Immunology and Immunopathology 27, 169-172.[Medline]
Mayernik, D. G., Ul-Haq, A. & Rinehart, J. J. (1983). Differentiation-associated alteration in human monocytemacrophage accessory cell function. Journal of Immunology 130, 2156-2160.
Meyers, G. & Thiel, H.-J. (1996). Molecular characterization of pestiviruses. Advances in Virus Research 47, 53-118.[Medline]
Moennig, V. & Liess, B. (1995). Pathogenesis of intrauterine infections with bovine viral diarrhea virus. Veterinary Clinics of North America Food Animal Practice 11, 477-487.
Muscoplat, C. C., Johnson, D. W. & Teuscher, E. (1973). Surface immunoglobulin of circulating lymphocytes in chronic bovine diarrhea: abnormalities in cell populations and cell function. American Journal of Veterinary Research 34, 1101-1104.[Medline]
Najar, H. M., Ruhl, S., Bru-Capdeville, A. C. & Peters, J. H. (1990). Adenosine and its derivatives control human monocyte differentiation into highly accessory cells versus macrophages. Journal of Leukocyte Biology 47, 429-439.[Abstract]
Planz, O., Ehl, S., Furrer, E., Horvath, E., Brundler, M.-A., Hengartner, H. & Zinkernagel, R. M. (1997). A critical role for neutralizing-antibody-producing B cells, CD4+ T cells, and interferons in persistent and acute infections of mice with lymophocytic choriomeningitis virus: implications for adoptive immunotherapy of virus carriers. Proceedings of the National Academy of Sciences, USA 94, 6874-6879.
Potgieter, L. N. D. (1995). Immunology of bovine viral diarrhea virus. Veterinary Clinics of North America Food Animal Practice 11, 501-520.
Rhodes, S. G., Cocksedge, J. M., Collins, R. A. & Morrison, W. I. (1999). Differential cytokine responses of CD4+ and CD8+ T cells in response to bovine viral diarrhoea virus in cattle. Journal of General Virology 80, 1673-1679.[Abstract]
Schlesier, M., Krause, S., Drager, R., Wolff-Vorbeck, G., Kreutz, M., Andreesen, R. & Peter, H.-H. (1994). Monocyte differentiation and accessory function: different effects on the proliferative responses of an autoreactive T cell clone as compared to alloreactive or antigen-specific T cell lines and primary mixed lymphocyte cultures. Immunobiology 190, 164-174.[Medline]
Sopp, P., Hooper, L. B., Clarke, M. C., Howard, C. J. & Brownlie, J. (1994). Detection of bovine viral diarrhoea virus p80 protein in subpopulations of bovine leukocytes. Journal of General Virology 75, 1189-1194.[Abstract]
Sopp, P., Kwong, L. S. & Howard, C. J. (1996). Identification of bovine CD14. Veterinary Immunology and Immunopathology 52, 323-328.[Medline]
Spriggs, M. K. (1996). One step ahead of the game: viral immunomodulatory molecules. Annual Review of Immunology 14, 101-130.[Medline]
Tortorella, D., Gewurz, B. E., Furman, M. H., Schust, D. J. & Ploegh, H. L. (2000). Viral subversion of the immune system. Annual Review of Immunology 18, 861-926.[Medline]
Welsh, M. D., Adair, B. M. & Foster, J. C. (1995). Effect of BVD virus infection on alveolar macrophage functions. Veterinary Immunology and Immunopathology 46, 195-210.[Medline]
Whittall, J. T. D. & Parkhouse, R. M. E. (1997). Changes in swine macrophage phenotype after infection with African swine fever virus: cytokine production and responsiveness to interferon- and lipopolysaccharide. Immunology 91, 444-449.[Medline]
Received 24 January 2001;
accepted 14 March 2001.