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Carlos and Marguerite Mason Transplantation Research Center and Department of Surgery, and
Emory Vaccine Center and Department of Microbiology and Immunology, Emory University School of Medicine, Atlanta, GA, 30322
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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As these strategies move closer to the clinic, one cause for concern is the effect that tolerance induction protocols might have on protective immunity to foreign pathogens. Disruption of the CD40/CD40L pathway has been shown to inhibit effective Ab responses in several settings (10, 20, 21, 22, 23). Furthermore, blockade of the CD40 and/or CD28 pathways as a method to achieve allogeneic bone marrow engraftment leads to the selective deletion of donor-reactive T cells (15, 16). Administration of reagents blocking these costimulatory interactions could therefore potentially impair both antiviral humoral and cellular responses. Alternatively, the presence of acute viral infections and their associated immune responses might interfere with the induction of allogeneic tolerance due to up-regulation of alternative costimulatory pathways, enhancement of inflammatory cytokines and stimuli, increased availability of T cell growth factors, or activation of viral Ag-specific T cells that also have specificity for donor MHC. In support of the latter possibility, several mouse pathogens stimulate allogeneic effector T cells following a primary infection (24, 25, 26).
Welsh et al. (6) recently showed that acute infection with lymphocytic choriomeningitis virus (LCMV)6 inhibits long-term skin graft survival and donor-specific hyporesponsiveness induced by administration of donor splenocytes and anti-CD40L (27). Interestingly, this effect was observed following LCMV and pichinde virus (PV) infections, but not following infection with other viruses (e.g., vaccinia virus (VV) and mouse CMV (MCMV)). Previous reports established that primary infection of C57BL/6 (B6) mice with LCMV induces a substantial number of H-2d-reactive T cells at the peak of the antiviral response (28). The ability of virus-infected mice to generate an alloresponse was subsequently mapped to the MHC locus, but independent of class II (24). Further studies have characterized T cell clones having dual specificity for H-2d alloantigens and syngeneic infected targets, as well as clones specific for alloantigen but not syngeneic infected targets (24). One possible explanation for the ability of LCMV to induce early graft rejection is the generation of activated T cells specific for both viral Ags and donor MHC. Conversely, at least two mouse pathogens that have been shown to induce allogeneic responses during primary infection, VV and MCMV, do not induce early graft rejection, suggesting that cross-reactivity may be either not required or not sufficient to prevent allograft survival. The role of TCR cross-reactivity in the virus-induced rejection of allogeneic tissue therefore remains unclear.
In this study we extend previous results to show that LCMV infection disrupts the beneficial effects of combined blockade of the CD40 and CD28 costimulatory pathways. Furthermore, we extend these findings to a mixed chimerism tolerance induction model. We have recently reported that administration of donor bone marrow, costimulation blockade, and the hematopoietic stem cell selective toxin busulfan around the time of transplant leads to indefinite skin graft survival, deletion of donor-reactive T cells, and the induction of high levels (>50%) of mixed hematopoietic chimerism (18). We report in this work that acute infection with LCMV causes prompt graft rejection, failure to generate mixed chimerism, and an inability to delete donor-reactive CD4 T cells. Rejection can be mediated by either CD4+ or CD8+ T cells, and a delay of infection following transplantation has no effect on tolerance induction.
We present a detailed quantitative assessment of allogeneic and
antiviral responses using a graft-vs-host disease (GVHD) model and MHC
tetramer analysis, as well as ELISPOT and intracellular IFN-
staining. Although LCMV infection allows for the generation of
costimulation blockade-resistant alloresponses, cross-reactivity of
LCMV-specific CD8 T cells to H-2d alloantigen is
minimal at a single cell level. Although mice receiving LCMV infection
without a skin graft do generate high numbers of alloreactive cells at
the peak of the response, their presence is largely associated with
acute infection, as they are reduced 50- to 100-fold by day 30
postinfection, while normal LCMV-specific T cells are reduced 10- to
12-fold over the same time period. To explore alternative reasons for
the virus-induced abrogation of tolerance induction, we assess
dendritic cell activation in the spleen following transplantation and
concurrent viral infection. We show that LCMV drives dendritic cell
maturation, regardless of the presence of costimulation blockade. We
propose that one possible explanation for the LCMV-induced rejection of
skin allografts is that infection leads to dendritic cell maturation
and an increased capacity for stimulating CD28/CD40-independent T cell
responses.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Adult male 6- to 8-wk old BALB/c, B6, and C3H/HeJ mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Mice were infected with 2 x 105 PFU LCMV Armstrong injected i.p. Virus stocks were grown and quantitated as previously described (29).
Skin grafting
Full thickness skin grafts (
1 cm2) were
transplanted on the dorsal thorax of recipient mice and secured with a
Band-Aid (Johnson & Johnson, Arlington, TX) for 7 days. Graft
survival was then followed by daily visual inspection. Rejection was
defined as the complete loss of viable epidermal graft tissue.
Statistical analyses were performed using a Mann-Whitney
U test.
Bone marrow preparation and treatment protocols
Skin graft recipients were treated with 500 µg each of hamster anti-murine CD40L Ab (MR1) and human CTLA4-Ig (provided by D. Hollenbaugh, Bristol-Myers Squibb, Princeton, NJ) administered i.p. on the day of transplantation (day 0) and on postoperative days 2, 4, and 6. CD4- and CD8-depleted experimental groups received 100 µg of rat anti-mouse CD4 (GK1.5) or rat anti-mouse CD8 (TIB105) i.p. on days -3, -2, -1, and weekly until harvest. Mice treated with busulfan (Busulfex; Orphan Medical, Minnetonka, MN) received 600 µg on postoperative day 5. Bone marrow was flushed from tibiae, femurs, and humeri, and red blood cells were lysed using a Tris-buffered ammonium chloride solution. Cells were resuspended in saline and injected i.v. at 2 x 107 cells/dose on postoperative days 0 and 6.
CFSE labeling and adoptive transfers
Labeling of naive or immune B6 T cells and adoptive transfer into irradiated BALB/c recipients were performed as previously described (30). Harvested splenocytes were analyzed by flow cytometry.
Intracellular IFN- assay
Intracellular IFN- expression in response to restimulation
with LCMV peptides was analyzed essentially as described
(31). In the case of irradiated recipients of CFSE-labeled
cells, harvested splenocytes were incubated for 5 h with
LCMV-infected or uninfected MC57 fibroblasts in the presence of
brefeldin A (GolgiPlug; BD PharMingen, San Diego, CA). In the GVHD
assay, peptide-specific IFN- production was assessed by
restimulating with uninfected IC21 macrophage cells pulsed with the
appropriate LCMV peptide at 0.1 µg/ml. After surface staining, cells
were permeabilized and stained for IFN- expression using the
Cytofix/Cytoperm kit (BD PharMingen) according to the manufacturer’s
instructions.
IFN- ELISPOT assays
Allospecific T cell responses were measured by IFN- ELISPOT
assay. Three-fold dilutions of recipient splenocytes
(H-2k or H-2b) were
stimulated overnight with 5 x 105
irradiated donor splenocytes (H-2d) per well in
ester-cellulose-bottom plates (Millipore, Bedford, MA) that had
been previously coated with IFN- capture Ab. To measure
LCMV-specific responses, splenocytes were restimulated overnight with
infected L929 (H-2k) or MC57
(H-2b) cells. Plates were coated and developed as
previously described (30).
Cell preparations and flow cytometry
MHC class I tetramers were prepared and refolded with
2-microglobulin and the appropriate peptide as
described previously (31). Analyses of splenocytes of
irradiated recipients of CFSE-labeled T cells were conducted using
fluorochrome-conjugated Abs (rat IgG2a PE, rat IgG2b PE, anti-CD4
PE, anti-CD8 PE; BD PharMingen) and APC-labeled tetramers. For
intracellular staining, cells were labeled with anti-CD8 PE and rat
IgG2b APC or anti-IFN- APC (BD PharMingen). Peripheral blood was
analyzed by staining with fluorochrome-conjugated Abs (rat IgG2a APC,
anti-CD4 APC, mouse IgG2a FITC,
anti-H-2Kd FITC, mouse IgG1 FITC,
anti-V5 FITC, rat IgG2b FITC, anti-V11 FITC; BD
PharMingen), followed by red blood cell lysis and washing with a
whole-blood lysis kit (R&D Systems, Minneapolis, MN). Splenic dendritic
cells were enriched on an Optiprep column (Nycomed, Oslo, Norway) as
previously described (32) and analyzed using
fluorochrome-conjugated Abs (ham IgG PE, anti-CD11c PE, ham IgM
FITC, anti-CD40 FITC, anti-CD54 FITC, rat IgG2a FITC,
anti-CD80 FITC, anti-CD86 FITC, mouse IgG2a FITC, anti
H-2Kd FITC, anti-I-Ab
FITC; BD PharMingen). Flow cytometry was performed using a FACSCaliber,
with CFSE fluorescence data being collected on the FL1 (FITC) channel.
Data were analyzed using CellQuest software (BD Biosciences,
Braintree, MA).
Cell lines
The fibrosarcoma cell line MC57 (H-2b+) and the liver-derived cell line L929 (H-2k+) were grown and passaged in RPMI 1640 supplemented with 10% FBS, antibiotics, and 2-ME.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Recent evidence has indicated that some viral infections
(e.g., LCMV and PV) inhibit the prolongation of skin allograft
survival mediated by blockade of the CD40 pathway and administration of
donor splenocytes (27). We sought to assess whether acute
LCMV infection could alter skin graft survival time when the CD28 and
CD40 T cell costimulatory pathways were blocked simultaneously. We
previously reported that C3H/HeJ mice receiving a BALB/c skin allograft
enjoy substantial prolongation of graft survival when treated with
CTLA4-Ig and anti-CD40L for a short time course at the time of
engraftment, with median survival times (MSTs) often exceeding 100 days
(9). In this experiment, C3H/HeJ mice receiving BALB/c
skin allografts and costimulation blockade survived >80 days. In
contrast, mice receiving the same procedure and treatment, along with a
concomitant infection of 2 x 105 LCMV
Armstrong on or near the day of transplant, rejected their grafts
promptly (MST = 20 days; Fig. 1
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, depletion of
CD4+ T cells did not alter the ability of C3H/HeJ
mice to reject BALB/c skin grafts following infection with LCMV
(MST = 18 days). Depletion of CD8+ T cells
resulted in a slight delay of skin graft rejection (MST = 26
days). Depletion of both subsets simultaneously resulted in long-term
allograft survival (MST > 60 days), indicating both that the
depletions were effective and that the virus was not directly harmful
to the allograft. These results parallel and extend the observations previously made, indicating that LCMV induces accelerated graft rejection in the face of combined blockade of the CD28 and CD40 T cell costimulatory pathways. This suggests that the mechanism of graft failure in the previous report is not likely due to the CD40-independent up-regulation of B7 molecules. Furthermore, they suggest that either CD4+ or CD8+ T cells are sufficient to mediate LCMV-induced skin graft rejection in this setting.
Acute LCMV infection prevents the establishment of partial hematopoietic chimerism, deletion of alloreactive T cells, and the induction of donor-specific tolerance
We next sought to determine whether LCMV infection had the same effect in a more robust tolerance induction model. Specifically, we sought to determine whether acute LCMV infection could disrupt the costimulation blockade-mediated establishment of mixed hematopoietic chimerism and donor-specific tolerance. Recent work in our lab has demonstrated that administration of donor bone marrow following treatment with the selective stem cell toxin busulfan, together with blockade of the CD40/CD28 costimulatory pathways, result in high levels of chimerism, deletion of donor-reactive T cells, and indefinite donor-specific tolerance (18).
As seen in Fig. 2A, 5/5 B6
mice receiving BALB/c skin and bone marrow, as well as busulfan and
costimulatory blockade treatment, had >200-day skin graft survival in
100% of mice tested. Conversely, 5/5 mice receiving the same treatment
concomitantly with an acute LCMV infection rejected their grafts
promptly (MST = 14 days). These results are representative of
three separate experiments. As in the previous model, predepletion of
CD8+ T cells demonstrated that
CD4+ T cells could mediate graft rejection,
although in a somewhat delayed fashion. Following depletion, no
CD8+ T cells could be detected in the peripheral
blood, while simultaneous depletion of both subsets during infection
resulted in indefinite graft survival, indicating that the depletions
were effective (data not shown).
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B). By day 125, >60% of peripheral blood leukocytes were
H-2Kd+ in all the mice (n = 5).
Chimerism was seen in all lineages tested, including
CD4+, CD8+,
B220+, CD11b+, and
GR-1+ cells (data not shown). Conversely, mice
receiving the same treatment along with LCMV at the time of engraftment
never developed detectable long-term chimerism. Predepletion of
CD8+ T cells (Fig. 2B) did not alter
the ability of the infection to abrogate chimerism.
Donor-specific tolerance following bone marrow engraftment and
treatment with costimulation blockade is due at least in part to
deletion of alloreactive T cells (15, 16). To determine
whether LCMV-induced skin graft rejection was associated with impaired
peripheral deletion of donor-reactive T cells, we compared the use of
V11 and V5.1/2 by CD4+ T cells from B6
recipients in the uninfected group (accepted both bone marrow and skin
grafts) and from the infected groups (rejected bone marrow and skin
grafts). BALB/c mice delete V11 and V5-bearing T cells in the
thymus due to their high affinity for endogenous retroviral
superantigens (mouse mammary tumor virus (MMTV)) presented by I-E MHC
class II molecules. B6 mice do not express I-E and thus use V11 on
5–7% of CD4+ T cells and V5.1/2 on
3–5% of CD4+ T cells. In this experiment,
uninfected mice treated with costimulation blockade, bone marrow, and
busulfan following skin engraftment showed decreased percentages of
V11+CD4+ and
V5+CD4+ T cells in the
peripheral blood by day 28 posttransplant. At day 60 posttransplant,
expression of these cell populations was nearly undetectable in the
peripheral blood, comprising similar percentages of the total
CD4+ population as those found in BALB/c mice. In
contrast, mice receiving 2 x 105 PFU LCMV
Armstrong at the time of engraftment failed to delete
V5+CD4+ and
V11+CD4+ T cells at any
time posttransplant (Fig. 2, C and D).
Failure to delete these cell populations occurred regardless
of the presence of CD8+ T cells. This
correlates with earlier observations noting an LCMV-induced
inhibition of peripheral deletion of alloreactive T cells following
disruption of the CD40/CD40L pathway (33).
These results indicate a role for LCMV in overcoming the tolerizing effects of combined costimulation blockade and bone marrow administration. We show rapid rejection of skin and hematopoietic allografts following acute infection, preventing the induction of donor-specific tolerance. We have also performed heterotopic heart allografts using the same treatment, and again LCMV inhibits the generation of donor-specific tolerance (data not shown). This effect cannot be attributed to either CD8+ or CD4+ T cells alone, as either subpopulation appears capable of inducing rapid CD40/CD28-independent graft rejection following acute infection. As predicted by graft survival, donor-reactive T cells are not deleted in infected mice, whereas uninfected mice receiving the tolerizing regimen delete donor MMTV superantigen-reactive T cell subpopulations within 60 days.
LCMV infection does not abrogate established tolerance
Based on previous studies (27) and our own
conclusions concerning the peripheral deletion of alloreactive T cells,
we considered it unlikely that a delayed infection with LCMV could
induce rejection of skin or bone marrow grafts in tolerant chimeric
mice. To test this hypothesis, mice were infected with LCMV 4–5 wk
following transplantation and tolerance induction. 5/5 mice were >20%
chimeric in the peripheral blood at the time of infection. Following
infection, we monitored skin graft survival and the development of
chimerism. As seen in Fig. 3, A and B, skin grafts on mice receiving a delayed
LCMV infection survived indefinitely, while hematopoietic chimerism
developed normally as compared with uninfected controls.
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B6 mice received BALB/c skin grafts and bone marrow, along with
costimulatory blockade and busulfan treatment. Control mice received
the same treatment regimen following receipt of syngeneic bone marrow
and skin grafts. On day 28 posttransplant, mice were infected with
LCMV. Eight days later splenocytes were harvested, restimulated for
5 h with LCMV peptides in the presence of brefeldin A, and stained
for intracellular IFN- expression. The peptides tested were
nucleoprotein (NP)396–404, gp33–41, gp276–286, NP205–214,
and the class II-restricted peptide gp61–80. All epitopes tested
generated large numbers of Ag-specific T cells in the spleen by day 8
postinfection in animals receiving syngeneic skin and bone marrow
grafts. In mice receiving allogeneic grafts, the number of antiviral T
cells in the spleen 8 days postinfection was moderately lower for each
epitope tested (1.5- to 2-fold), possibly due to the influx of APCs not
expressing H-2b. However, no substantial deletion
of any particular epitope could be detected, nor was there any apparent
change in epitope hierarchy between the mice receiving syngeneic grafts
and the mice receiving allogeneic grafts (see Fig. 4). These results suggest that there is a
moderate reduction of antiviral responses following the induction of
mixed chimerism but fail to implicate TCR cross-reactivity of any
specific epitope to alloantigen.
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To directly address the question of whether LCMV-specific CD8 T cells were also alloreactive, we used a previously described GVHD model (34). T cells from LCMV-immune B6 mice (>30 days postinfection) were labeled with the fluorescent dye CFSE (Molecular Probes, Eugene, OR) and injected i.v. into irradiated (1800 rad) allogeneic BALB/c hosts. In this model, allogeneic T cells responding to Ag lose fluorescence with each successive division, allowing for quantitation and analysis of highly divided alloreactive cells by flow cytometry. By using LCMV-immune mice as donors, we could assess whether LCMV-reactive T cells also divided in response to alloantigen by direct staining with the Db/NP396–404, Db/gp33–41, and Kb/gp34–43 class I MHC tetramers. Splenocytes were harvested 72 h posttransfer, stained with anti-CD8 Abs and tetramers, and analyzed by flow cytometry.
CD8+ T cells from both naive and immune mice
divided significantly in response to alloantigen, with large numbers of
cells from both groups reaching at least eight divisions. In contrast,
CD8+ T cells from either group injected into
irradiated syngeneic recipients did not divide more than three times
(data not shown). Therefore, we gated undivided and maximally divided
(four to eight divisions) CD8+ T cells and
analyzed tetramer binding in each population (Fig. 5). LCMV-specific CD8 T cells were
readily detectable within the undivided population in the recipients of
LCMV-immune T cells for each tetramer tested. However, we failed to
detect discernible staining above background for any of the tetramers
in the maximally divided population (Fig. 5). The results are
summarized in Table I. As a control to
verify that our failure to detect tetramer binding was not simply a
result of TCR down-modulation in highly divided cells, we stained for
expression of TCR. We did not observe decreased expression of the
TCR in the maximally divided cells by this assay (data not shown).
Furthermore, a previous study has established that proliferating
LCMV-specific CD8 T cells in lymphopenic hosts do not show decreased
binding to MHC tetramers (35).
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expression, and analyzed by flow cytometry. As
seen in Fig. 6, no IFN- expression was
observed above background except in the undivided CD8 T cells
stimulated with infected MC57 cells. We further analyzed responses to
four LCMV epitopes in the same manner. In these experiments, rather
than stimulating with infected cells, harvested splenocytes were
restimulated with MC57 cells pulsed with LCMV peptides (NP396–404,
gp33–41, gp276–286, NP205–214). None of these peptides induced
IFN- production above background in the divided, alloreactive CD8 T
cells. In contrast, LCMV-specific CD8 T cells could be readily detected
in the undivided population following restimulation with these peptides
(data not shown). One caveat to these experiments is the high
background IFN- production in the highly divided cells, presumably
due to the continued cycling and low-level stimulation of these cells
during brefeldin A incubation.
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LCMV facilitates the CD28/CD40-independent generation of
alloreactive IFN--producing cells
To better characterize the generation of allogeneic and antiviral
T cell responses following LCMV infection, splenocytes were monitored
for their ability to produce IFN- after restimulation in vitro by an
ELISPOT assay. In this experiment, C3H/HeJ mice receiving BALB/c skin
grafts generated 3–4 x 105 allospecific
T cells in the spleen by day 8 posttransplant, and these cell numbers
dropped slightly at day 15. Treatment with costimulation blockade
completely abolished the allogeneic response at both time points. Mice
receiving skin grafts and costimulation blockade concurrent with an
acute LCMV infection generated small numbers (9 x
104) of allospecific cells in the spleen by day
8. By day 15, these mice had overcome the immunosuppressive effects of
costimulation blockade and had generated an alloresponse comparable to
untreated controls (2.5 x 105). As
reported previously, acute LCMV infection in the absence of a skin
graft resulted in the generation of some allospecific IFN--producing
cells by day 8 (3 x 105). By day 15,
this effect had diminished markedly to 4 x
104 IFN-+ cells per
spleen (Fig. 7).
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1.2 x 107 IFN--producing T
cells in the spleen, whereas mice that received concurrent combination
blockade and a BALB/c skin graft, while still generating a large
response, had an 3-fold drop in the number of LCMV-specific cells in
the spleen (4 x 106). By day 15, spleens
from LCMV-infected mice showed a 3- to 4-fold decrease in the number of
LCMV-specific cells. In mice receiving combination blockade, the drop
was somewhat greater (Fig. 7).
To assess whether LCMV-infected mice generated memory to alloantigen,
we infected B6 mice and quantitated the number of allospecific cells in
the spleen at the peak of the infection (day 8) and following the
development of immune memory (>30 days postinfection) by IFN-
ELISPOT. LCMV-infected mice generated allospecific T cells (7.29
x 105 ± 1.78 x 105,
n = 3) at the peak of the infection, but by day 30
postinfection, the number of these cells in the spleen dropped 50- to
100-fold (1 x 104 ± 9.9 x
102, n = 3). In contrast, the
number of T cells specific for several known immunodominant and
subdominant LCMV epitopes (NP396–404, gp33–41, gp276–286,
NP205–214) dropped 10- to 12-fold in the spleen over the same period
(1.52 x 107 ± 1.13 x
106 to 1.40 x 106 ±
1.38 x 105, n = 3 for both
groups). This level of LCMV memory is similar to previous reports
(31).
We conclude that LCMV infection stimulates the activation of at least a subset of allogeneic T cells by CD40/CD28-independent mechanisms, thereby overcoming the immunosuppressive effects of costimulation blockade and leading to early graft rejection. Based on our CFSE and ELISPOT results, we propose that the frequency of virus-specific T cells also bearing TCR specificity to alloantigen is low.
LCMV infection induces the CD28/CD40-independent maturation of splenic dendritic cells
We next sought to explore other potential mechanisms whereby LCMV
infection could abrogate transplant tolerance and stimulate the
activation of alloreactive T cells. Our previous experiments studying
deletion of V subsets made it clear that in the presence of LCMV
infection, CTLA4-Ig and anti-CD40L are unable to initiate the
deletion of alloreactive T cells. A possible explanation was that LCMV
infection was able to influence the induction and/or up-regulation of T
cell costimulatory pathways by APCs. Furthermore, LCMV might induce the
expression of molecules or survival factors that prevented deletion of
alloreactive T cells. To test the merit of this hypothesis, we analyzed
the effects of LCMV infection on costimulatory molecule and MHC
expression by CD11c+ dendritic cells in the
spleen.
Mice received BALB/c skin grafts and bone marrow, costimulatory
blockade therapy, and busulfan. One group was infected with LCMV
Armstrong on day 0, while the other remained uninfected. Splenocytes
were harvested on day 6 and separated based on cell density using an
Optiprep column (Nycomed) as previously described (32).
The low-density fraction, which is enriched for dendritic cells, was
harvested and stained for CD11c expression, along with MHC class I and
II, ICAM-1, CD40, CD80, and CD86. Following analysis by flow cytometry,
expression of these molecules among CD11c+ cells
was analyzed. As seen in Fig. 8, LCMV
infection resulted in the increased expression of all of these
molecules, regardless of the presence of costimulatory blockade. We
conclude that LCMV infection induces a higher activation state among
dendritic cells. We suggest that one explanation for the deleterious
effects of LCMV infection on tolerance induction could be the increased
ability of APCs to stimulate and activate alloreactive T cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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T cell
subsets. We show that infection must occur at or around the time of
transplant, as a delay of 3–4 wk in the onset of infection has no
effect on graft survival or the induction of mixed chimerism. These
studies confirm prior reports of the LCMV-mediated abrogation of skin
graft survival following administration of donor splenocytes and
anti-CD40L (27), and extend upon them by showing that
LCMV-induced graft rejection is not mediated by CD40-independent
up-regulation of B7.1 or B7.2. One concern with the use of tolerance
induction strategies is the potential to induce tolerance to concurrent
viral infections. It is of considerable interest that the immune
responses to LCMV are not rendered tolerant following the use of
costimulation blockade-based tolerance induction regimens, a finding
consistent with previous observations that LCMV T cell responses are
largely independent of CD28 and CD40 (23, 36, 37). It has been proposed that one possible explanation for the deleterious effects of LCMV infection on graft survival could be the presence of cross-reactivity to alloantigen at the level of TCR/MHC recognition during an antiviral response (27). In this scenario, antiviral responses would include some cells also bearing specificity for alloantigen. In support of this hypothesis, it has been shown that LCMV induces H-2d-specific CD8 T cells at the peak of the T cell response (28). Although we make similar observations, we find little evidence for substantial cross-reactivity of LCMV-specific CD8 T cells generated in vivo to H-2d alloantigen. A delayed primary infection (4 wk posttransplant) elicits an antiviral response with unchanged epitope hierarchy, although the numbers of activated CD8 T cells are somewhat globally diminished. Furthermore, using a sensitive single cell assay using intracellular cytokine staining and MHC tetramers, we are unable to detect the division of LCMV-immune CD8 T cells in response to alloantigen. Nevertheless, as has been previously reported, LCMV primary infection does generate alloreactive cells. These cells drop greatly in number by day 15 postinfection and are barely detectable in LCMV-immune mice (>30 days postinfection). These experiments suggest that the frequency of LCMV-specific CD8 T cells that are cross-reactive to alloantigen is low. We cannot rule out the possibility of high levels of cross-reactive CD4 T cells using this assay.
The primary mechanism by which alloreactive T cells are activated during LCMV infection remains uncertain. Studies in recent years have shown that the great majority of activated CD8 T cells generated during an antiviral response are Ag-specific (31, 38, 39). Given the high frequency of alloreactive CD8 T cells in naive mice, substantial cross-reactivity at the level of TCR/MHC interaction would not be surprising. However, we are unable to detect significant levels of allospecific activation of CFSE-labeled LCMV-specific CD8 T cells following injection into irradiated BALB/c donors. Furthermore, LCMV-induced alloreactive cells do not behave as other virus-specific populations, as they have an exaggerated death phase following the peak of the response. Both CD4 and CD8 T cell subsets in isolation are capable of preventing tolerance induction and mixed chimerism. Interestingly, disruption of costimulatory pathways during LCMV infection has little effect on CD8+ T cell responses but almost entirely prevents the generation of CD4+ antiviral T cells (40). Nevertheless, CD4+ T cells are sufficient to mediate the LCMV-induced prevention of tolerance induction to alloantigen. Although cross-reactivity to alloantigens likely exists at some level, our data suggest that this is a relatively infrequent event during LCMV infection. It is of interest to note that LCMV responses are diminished in mice receiving the tolerance induction regimen. This observation could be due to nonspecific immunosuppressive effects of allogeneic bone marrow and costimulation blockade treatment. Alternatively, the influx of H-2d+ donor APCs in the immune compartments could dilute the available Ag for stimulating an H-2b-restricted response. Further studies are warranted to assess the long-term effects of tolerance induction on immune responses to other pathogens.
Regardless of the extent to which alloreactive cells are generated during primary LCMV infection through TCR cross-reactivity, other mechanisms clearly play an indispensable role in the LCMV-mediated circumvention of the CD28/CD40 pathways. For example, MCMV and VV both generate allogeneic responses during primary infection (26), yet infection with these viruses has been shown not to impair graft survival (27). The primary CD8+ anti-LCMV response itself has been shown to be largely independent of the CD28 and CD40 pathways (23, 36, 37). Interestingly, a recent study demonstrates that LCMV-specific responses, but not those directed toward VV, can be driven by parenchymal cells (41). This suggests that LCMV, but not VV, can lower the threshold required for full activation of effector cells. One possibility is that LCMV triggers specific innate immune mechanisms that allow for the circumvention of these pathways in generating T cell responses. Also, anti-LCMV responses may provide cytokines and growth factors that aid the generation of CD28/CD40 independent alloresponses. Another possibility is that LCMV infection induces the expression of CD40/CD28-independent costimulatory pathways. In support of this latter possibility, we show in this study that LCMV infection mediates the CD28/CD40-independent up-regulation of MHC and costimulatory molecules on dendritic cells. We hypothesize that infection with LCMV facilitates the activation of alloreactive cells in the face of costimulatory blockade through the up-regulation of alternative costimulatory molecules on the surface of APCs. In this model, the need for costimulation and activation of dendritic cells by the CD28 or CD40 pathways would be abrogated by infection with LCMV. Further studies are required to elucidate the precise mechanisms by which some viral infections (e.g., LCMV, PV) but not others (e.g., MCMV, VV) mediate early graft rejection in the face of costimulation blockade.
Infections have long been a problem during attempts to immunosuppress transplant recipients. It is likely that any type of immunosuppression used in the future will carry the risk of predisposing the recipient to opportunistic pathogens and viral infections. As protocols involving the use of costimulatory molecule-blocking reagents move closer to clinical application, it will be of vital importance that we understand the mechanisms whereby viral infections can interfere with attempts to prevent graft rejection. Of particular interest is the ability of LCMV infection, but not all viral infections, to overcome attempts at tolerance induction through the use of costimulatory blockade. Our data suggest that the prevention of tolerance induction following LCMV infection is not solely due to cross-reactivity to alloantigen. Other mechanisms likely play a role, as evidenced by our observation that LCMV infection elevates the activation and maturation level of host dendritic cells. Future studies should elucidate the precise pathways by which viral infections mediate the costimulation blockade-resistant activation of alloreactive T cells.
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Acknowledgments |
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Footnotes |
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2 M.A.W. and J.T.T. contributed equally to this work.
3 Current address: Department of Immunology, The Scripps Research Institute, La Jolla, CA 92037.
4 Current address: Department of Molecular Immunology, La Jolla Institute for Allergy and Immunology, San Diego, CA 92121.
5 Address correspondence and reprint requests to Dr. Christian P. Larsen or Dr. Thomas C. Pearson, Emory University Transplantation Immunology Laboratory, Suite 5105, WMB, 1639 Pierce Drive, Atlanta, GA 30322. E-mail addresses: clarsen@emory.org or tpearson{at}emory.org
6 Abbreviations used in this paper: LCMV, lymphocytic choriomeningitis virus; PV, pichinde virus; VV, vaccinia virus; MCMV, mouse CMV; GVHD, graft-vs-host disease; MMTV, mouse mammary tumor virus; MST, median survival time; NP, nucleoprotein.
Received for publication June 18, 2001. Accepted for publication August 31, 2001.
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