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*
Surgical Research Laboratory, Department of Surgery, Harvard Medical School, Brigham and Women’s Hospital, Boston, MA 02115;
The Dumont-University of California at Los Angeles Transplant Center, University of California at Los Angeles School of Medicine, Los Angeles, CA 90095; and
Institute of Medical Immunology, Charité, Humboldt University Berlin, Germany
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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50% of recipients. Thymus was
required for generation of regulatory T cells under the cover of CD4
mAb therapy and for the ability of these cells to confer infectious
tolerance. However, thymus was not mandatory to maintain an
infectious-permissive environment in cohorts of adoptively transferred
recipients. Intragraft expression of IL-2, IL-4, and IL-10 genes was
diminished in euthymic and thymectomized tolerant hosts. However,
grafts in the latter group showed significant IFN-
gene expression,
suggesting a less efficient down-regulation of Th1-like cells in the
absence of regulatory cells. Indeed, exogenous challenge with rIL-2 or
freshly alloactivated spleen cells recreated rejection in
thymectomized, but not euthymic, hosts, suggesting that a state of
cytokine-responsive anergy contributes to the "noninfectious" form
of tolerance in thymectomized rats. The infection-tolerant state did
not result from "graft adaptation," and regulatory T cells
restricted for the original alloantigen were exposed to its continuous
stimulation. The effective memory for suppression was dependent upon
persistent donor-specific alloantigen stimulation; it disappeared
within 3 weeks after its removal. Hence, both central and peripheral
immune mechanisms, orchestrated by the tolerizing alloantigen,
contribute to the infectious tolerance pathway in CD4 mAb-treated rat
transplant recipients. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The infectious tolerance, as originally described by Dr. H. Waldmann’s group, was induced and perpetuated in adult thymectomized (ATX)3 mouse recipients of nonvascularized skin allografts treated with a mixture of nonlytic CD4 and CD8 mAbs (2, 3, 4). We have recently shown, however, that cardinal features of such a classic peripheral form of tolerance may be also applied to euthymic Ag-primed rat recipients of vascularized transplants (6). Hence, 1) treatment of sensitized rats with RIB-5/2, a CD4-nondepleting mAb, abrogated accelerated (<36 h) rejection and produced indefinite cardiac allograft survival; 2) tolerant cells in mAb-treated hosts could disable naive or even alloimmune cells, so that they failed to trigger rejection; and 3) donor-specific and organ-nonspecific tolerance could be transferred by spleen cells alone, and with no concomitant therapy, into new cohorts of cardiac or renal test allograft recipients. Furthermore, we have documented that regulatory CD4+ Th2-like IL-4-producing cells are instrumental for the maintenance of the infectious tolerance pathway (7). Given the phenomena associated with this clinically relevant rat transplant model, we have now performed studies aimed at exposing mechanisms underlying the induction and maintenance of infectious tolerance, with particular emphasis on the role of the thymus and alloantigen. We provide evidence that the induction of infectious tolerance in rat recipients is thymus dependent, consistent with the role of the thymus in generating regulatory T suppressor cells (8). Once established, these immunoregulatory T cells remain critically dependent upon the continuous presence of a tolerizing donor-specific alloantigen and may now operate in ATX test recipients. Hence, both central and peripheral immune mechanisms for induction and maintenance to CD4 mAb-induced infectious tolerance must exist.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Inbred male adult (10–12-wk-old) rats weighing 200 to 250 g were used (Harlan Sprague Dawley, Indianapolis, IN). Lewis (LEW, RT1l) served as recipients of cardiac allografts from (LEW x BN)F1 (LBNF1) hybrids. Brown Norman (BN) (RT1n) rats were used as skin donors. Wistar-Furth (WF, RT1u) rats were employed as third-party donors for specificity experiments. Full-thickness skin was sutured bilaterally to appropriate defects in the chest of prospective rat recipients. Hearts were transplanted to the abdominal great vessels by standard microvascular techniques. Their function was monitored daily by palpation, and rejection was defined as the day of cessation of heart beat.
The model of infectious tolerance
LEW rats were sensitized with BN skin grafts (day -7), followed 1 wk later by transplantation of LBNF1 hearts (day 0). These cardiac allografts are rejected in an accelerated manner in <36 h (6, 7, 9). However, intermittent treatment with RIB-5/2 (5 mg/rat i.v. at days -7, -4, -1, 0, +3, +4, +7, +10, +14, and +21), a mouse antirat CD4-nondepleting mAb (10), results in permanent cardiac allograft acceptance, with features characteristic for the infectious tolerance pathway (6, 7). Skin grafts are rejected despite RIB-5/2 therapy (6). Rats maintaining their transplants for >100 days were used in the present study.
Host immune manipulations
ATX was performed following partial median sternotomy in
otherwise untreated rats at 10–12 wk of age. These were then
sensitized with skin grafts 4–8 wk later, followed 1 wk later by
transplantation of donor-strain hearts. Some animals underwent ATX at
>100 days posttransplant. To test for tolerance induction, long-term
allograft recipients were challenged with a secondary donor-specific
(LBNF1) or third-party (WF) heart transplant. In an
adoptive transfer assay, 100 x 106 erythrocyte-free
spleen cells were administered i.v. into sublethally gamma-irradiated
(450 rads) syngeneic rats (one donor spleen/two test recipients). These
were then challenged 24 h later, unless specified otherwise, with
donor-specific test cardiac allografts. In an attempt to break
tolerance, long-term engrafted hosts were injected i.v. with 1) spleen
cells (100 x 106 cells/rat once a week for 3 wk) from
sensitized rats undergoing accelerated cardiac allograft rejection, and
2) human rIL-2 (20,000 U (1.4 µg)/day for 5 days; Collaborative
Research, Bedford, MA). Finally, long-term LBNF1 cardiac
allografts "parked" in tolerant LEW recipients for >100 days were
retransplanted into normal LEW rats. The "graft-free" hosts were
then challenged with a fresh LBNF1 heart at different times
after removal of the original transplant.
RT-PCR analysis
For evaluation of intragraft cytokine gene expression,
competitive template RT-PCR was performed (11). Total RNA was prepared
from the grafts and reverse transcribed into cDNA. A cDNA equivalent of
5 ng total RNA was amplified in a 25-µl reaction volume containing
250 µM of each deoxynucleotide triphosphate, 10 µM of the primer
pair, 2.5 µl 10-fold PCR buffer, and 0.5 U Taq DNA
polymerase (Perkin-Elmer/Cetus, Emeryville, CA). As control DNA
fragment, a synthetic gene containing nonhomologous DNA with binding
sites for 5' and 3' cytokine-specific primers was constructed. The gene
was cloned into pBluescript II SK+ (Stratagene, La Jolla,
CA). The sense and antisense primer sequences derived from rat CD3,
CD25, IL-2, IFN-, IL-4, IL-10, and ß-actin cDNA have been
described (11). For quantification, cDNA samples were adjusted to equal
input concentrations based on their ß-actin cDNA content. For this
purpose, 10-fold serial dilutions of the control fragment (500–5 pg)
were mixed with identical volumes of cDNA prepared from the tissue
RNAs; these were then coamplified by using ß-actin-specific primers.
After electrophoresis of the PCR products, the cDNA samples were
adjusted to equal ß-actin cDNA content by ethidium bromide
fluorescence. The adjusted cDNA samples and specific primers were then
subjected to another coamplification with 10-fold dilutions of the
control fragment using primers specific for each cytokine. The level of
cytokine gene expression was expressed in AU (arbitrary unit)/µl
cDNA. One AU was defined as the lowest concentration of the control
fragment that yielded a detectable amplification product at the
conditions used.
Evaluation of host alloantibody responses
Donor-specific IgM and IgG alloantibody responses were
determined in recipient serum, as described (6). Briefly, cervical BN
lymph node target cells (1.5 x 105) were
incubated with serially diluted (1:4 
1:64) heat-inactivated serum
samples for 30 min at 4°C. To stain for IgM and IgG, the washed cells
were then reacted with a mixture of FITC-conjugated goat Ab specific
for the Fc portion of rat IgG and phycoerythrin (PE)-conjugated goat Ab
specific for the µ chain of rat IgM (Jackson ImmunoResearch, West
Grove, PA). After staining, the cells were washed, fixed in 1% neutral
buffered formalin, and analyzed on an EPICS C cell sorter (Coulter,
Hialeah, FL). The use of paired secondary Abs allowed simultaneous
assessment of IgG (FITC channel) and IgM (PE channel). Their levels
were expressed as mode channel fluorescence.
Statistical analysis
The graft survival curves were analyzed by the Kaplan-Meier
test, and statistical significance in graft survival between
experimental groups was determined by log rank 
2
test. Values were expressed as the mean ± SEM, with differences
considered statistically significant if p <
0.05.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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As infectious tolerance was originally induced and perpetuated in
thymectomized mice (2, 3, 4, 5), we first asked whether ATX may affect the
infectious tolerance pathway in our rat transplantation model. First,
we confirmed our earlier findings (12) by demonstrating that thymic
extirpation at 4–8 wk before the skin allografting did not influence
host sensitization, as cardiac allografts were rejected uniformly in
<36 h, similarly to euthymic sensitized rat recipients (Fig. 1
A). However, as shown
in Figure 1A, ATX in naive animals 4–8 wk before skin
graft-induced sensitization profoundly influenced the tolerogenic
effects of CD4-targeted therapy (mean survival time (MST) = 53.2 days,
p < 0.0001, as compared with euthymic controls).
Moreover, unlike in euthymic rats, only about 50% of ATX and RIB-5/2
mAb-treated animals maintained well functioning cardiac allografts for
>100 days; the remainder rejected their transplants within 17 days
(Fig. 1A). The permanent graft acceptance rates were
similar in groups of rat recipients, which received heart transplant at
5 wk (60%), 7 wk (50%), or 9 wk (50%) following thymectomy.
Long-term ATX and CD4 mAb-treated hosts developed donor-specific
tolerance, which could be manifested by acute rejection (7–8 days) of
third-party (WF) and acceptance (MST = >70 days) of donor-matched
(LBNF1) secondary heart grafts (n =
2 rats/group). Thus, although pretransplant thymectomy diminished the
overall efficacy of CD4-targeted therapy, donor-specific tolerance may
still have been induced in about 50% of CD4 mAb-conditioned ATX
hosts.
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D). These new euthymic, lightly gamma-irradiated but
otherwise untreated test animals rejected their transplants within 7
days (MST = 6.3 days), similarly to control rats (Fig. 1B), which were either infused with the same number
of nontolerant recipient-type spleen cells (MST = 6.8 days) or did
not receive any cells at all (MST = 10.2 days). Hence,
pretransplant thymectomy prevented the development of the infectious
tolerance pathway in engrafted rat recipients. ATX prevents induction but not maintenance of infectious tolerance-mediating regulatory T cells
Because the regulatory T cells develop gradually in CD4
mAb-treated hosts and become fully operational at >100 days
posttransplant, we hypothesized that thymic extirpation at such a late
phase should not affect the allograft outcome. Indeed, cardiac
allografts continued to function for >50 days in a group of RIB-5/2
mAb-treated recipients, despite ATX performed at 100–120
days posttransplant (n = 4; data not shown).
Moreover, such a late thymectomy did not affect the ability of spleen
cells to tolerize new sets of test recipients. We next asked whether
these regulatory T cells, which require thymus for their induction, may
then function in a thymus-free environment of test recipients. As shown
in Figure 1C, transfer of spleen cells from RIB-5/2
mAb-treated tolerant euthymic hosts prompted acceptance of cardiac
allografts in new cohorts of ATX "test-tube" rats. Moreover, we
found that spleen cells from secondary ATX long-term hosts retained
their suppressive effects upon sequential transfer to
tertiary ATX test animals (MST = >50 days; n = 5;
data not shown). These results imply that the thymus is critical for
generation of regulatory T suppressor cells under cover of
peritransplant CD4 mAb therapy and for the ability of cells to confer
tolerance in an infectious manner. However, the presence of the thymus
is not mandatory to maintain an infectious-permissive environment in
new generations of adoptively transferred test recipients.
ATX allows the development of clonal anergy in CD4 mAb-treated rats
Our recent data demonstrating the persistence of donor reactive T
cells in long-term tolerant hosts rule out clonal deletion as a
possible mechanism of tolerance maintenance in ATX hosts (13).
Moreover, as thymectomy prevented the development of regulatory T
cells, we then asked whether clonal anergy may have contributed to the
tolerance maintenance in ATX recipients. To address this issue, we have
analyzed intragraft Th1/Th2-type cytokine gene programs in transplant
recipients by employing competitive template RT-PCR. As shown in Figure 2, unlike in untreated hosts undergoing
accelerated graft rejection, intragraft IL-2, IL-4, and IL-10 gene
expression was diminished in both euthymic and ATX + CD4
mAb-treated long-term (>100 days) recipients; grafts from ATX tolerant
rats showed significant IFN- gene expression. In an attempt to break
tolerance in our model, long-term allograft recipients were then
challenged with exogenous rIL-2 or with freshly alloactivated spleen
cells. Indeed, as shown in Table I, this
treatment recreated graft rejection in ATX but not in euthymic hosts.
The challenge with rIL-2 was more effective than that utilizing
alloimmune cells (MST = 7.0 days and 24.7 days, respectively;
p < 0.0001). These results imply that unlike in
euthymic hosts, a state of cytokine-responsive anergy contributes to
the maintenance of cardiac allografts in ATX + CD4
mAb-treated rats.
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We have previously shown that accelerated rejection in
sensitized rats is accompanied by a strong systemic anti-donor IgM
response, which peaks around the time of allograft loss and then
switches to IgG alloantibody response at 7–10 days (6). The
tolerogenic RIB-5/2 mAb regimen depresses the level and duration of IgM
and prevents the switch from IgM to IgG (6, 7). We have now asked how
pretransplant ATX, which diminishes the overall efficacy of
CD4-targeted therapy and prevents the development of regulatory T
cells, will influence host circulating alloantibody responses. As shown
in Figure 3, both IgM and IgG
alloantibody levels were virtually abolished throughout in ATX
recipients regardless of the functional status of the allograft itself.
Thus, although depression of humoral alloreactivity is important for
the acquisition of infectious tolerance pathway in euthymic rats,
diminished systemic IgM/IgG levels do not correlate with the allograft
outcome in ATX hosts.
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Finally, we wished to determine whether persistence of donor
alloantigen is required for regulatory T cells to maintain infectious
tolerance pathway. First, we evaluated the immunogenicity of long-term
cardiac allografts in RIB-5/2 mAb-treated hosts, because the
replacement of donor APCs by recipient APCs over time may have resulted
in the loss of functional allostimulatory capacity in vivo. However,
normal LEW secondary recipients rejected acutely retransplanted
LBNF1 hearts, despite such hearts having been
"parked" in LEW-tolerant hosts for >100 days (MST ± SD
= 8.5 ± 0.5 days; n = 5; data not shown). We then
performed a series of experiments in which primary heart grafts were
removed at >100 days from CD4 mAb-treated recipients, and secondary
hearts were placed at 1, 7, 14–17, and 30–90 days thereafter (Fig. 4A). By that time,
there are no detectable RIB-5/2 mAb levels in the circulation (6). As
shown in Figure 4A, such graft-free recipients accepted
secondary donor-type transplants after a delay of up to 17 days, after
which the operational tolerance decayed with time. Indeed, only two of
four grafts survived >100 days in 30-day graft-free hosts
(p < 0.05), compared with a 90-day graft-free
environment, in which all hearts were rejected promptly within 2 wk
(p < 0.01). Moreover, spleen cells from
<17-day, but not from 90-day, graft-free hosts conferred tolerance
into new cohorts of test recipients (MST = >50 days;
n = 3 rats/group; data not shown). To confirm that the
maintenance of infectious tolerance is Ag dependent, we used another
experimental system in which spleen cells from tolerant rats were
adoptively transferred into syngeneic secondary hosts, which were then
challenged with donor-specific test cardiac allografts. As shown in
Figure 4B, tolerant cells rapidly lost their suppressive
capacity in the absence of donor alloantigen, and by 3 days they were
unable to prevent rejection of test cardiac allografts. This contrasted
with a stable tolerance achieved when tolerant cells were infused on
the day of or 1 day before transplantation (100% and >80% graft
acceptance, respectively). Hence, the effective memory for suppression
in the infectious tolerance pathway depends upon the continuous
donor-specific alloantigen stimulation.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Thymus is required for the acquisition of central tolerance to self-Ags, in which autoreactive T cells are deleted or anergized by exposure to the self-Ags presented by either bone marrow-derived or thymic stromal cells (14). Similar intrathymic mechanisms are important for inducing central tolerance to alloantigens, as evidenced by permanent allograft acceptance or tolerance following intrathymic injection of donor cells or soluble Ag (15, 16). In our own studies, administration of donor-type Ag into the recipient thymus abrogated accelerated rejection and significantly prolonged cardiac allograft survival in sensitized rats (17, 18). However, the role of the thymus in the acquisition of peripheral tolerance remains unclear. It has been shown that pretransplant ATX neither abolished donor-specific blood transfusion-mediated tolerogenic effects in fully MHC-incompatible rat renal allograft recipients (19) nor affected the infectious tolerance pathway to minor histocompatibility-nonvascularized skin grafts in mice conditioned with CD4 + CD8 mAbs (2, 3, 4). In contrast, thymus was essential for rapid and stable tolerance to MHC class I-mismatched renal allografts in miniature swine (20) and for the maintenance of tolerance to skin grafts in mixed allogeneic bone marrow mouse chimeras (21). Similarly, pretransplant ATX abrogated CTLA4Ig-induced tolerance after the blockade of CD28-B7 T cell costimulation in rat renal allograft recipients (22).
Our present data document the requirement for the thymus in the induction of infectious tolerance pathway in the accelerated rejection model. Hence, unlike in euthymic rats, only about 50% of ATX animals maintained long term functioning cardiac allografts after RIB-5/2 mAb treatment. Interestingly, however, the nature of donor-specific tolerance in both recipient groups was strikingly different. First, pretransplant ATX prevented the development of operational infectious tolerance, as evidenced by the inability of spleen cells from the original ATX + CD4 mAb-treated hosts to confer the tolerant state to new generations of test recipients. The immune mechanisms involved in thymus-dependent induction of infectious tolerance and the generation of regulatory T cells remain unclear. Perhaps, by migrating from cardiac allografts to the thymus, dendritic cells may become responsible for a central component in tolerance induction. Alternatively, thymic migrants may be instrumental for inducing peripheral tolerance to alloreactive cells in the graft. Indeed, several lines of evidence have emerged that refute the dogma of one-way trafficking through the thymus. In fact, our own studies have documented the recirculation of immunoresponsive thymocytes from the thymus to cardiac grafts and then back to the thymus in this accelerated rejection model (12). Moreover, passive transfer of thymocytes from rats rendered tolerant to vascular organ transplants into secondary naive hosts can lead to prolongation of test graft survival (23, 24). The thymus has also been identified as a source of natural suppressor cells in mice (25).
While the thymus was important for the induction of infectious tolerance in CD4 mAb-treated hosts, its presence in the maintenance phase was not mandatory. Indeed, ATX performed at >100 days posttransplant neither influenced otherwise permanent allograft survival nor affected the ability of adoptively transferred spleen cells to tolerize new sets of test rat recipients. This suggests that recent thymic migrants were sufficient to ensure the development of infectious tolerance in this model. Moreover, unlike in the original CD4 mAb-treated rats, thymus was not required to maintain the infectious tolerance pathway in new cohorts of test animals. Perhaps, once generated in CD4 mAb-treated rats, the regulatory T cells do not require the thymus to express their "infectiousness" and to overcome allorecognition properties of naive cells, which otherwise are fully capable of triggering rejection in test allograft recipients. Indeed, this was the case in Waldmann’s infectious tolerance model, in which naive cells were transferred into allograft-containing tolerant transgene-marked recipient mice (2). Moreover, others have shown that even in the absence of the thymus, T cells capable of transferring donor-specific tolerance may emerge in the periphery (26). We favor the notion that the infectious phenomenon represents the natural host immune mechanism, independent of the initial immunomodulation, and our ongoing experiments suggest that the ability of regulatory T cells to maintain the infectious tolerance pathway may be strictly cell dose dependent (K. Onodera et al., manuscript in preparation).
The infectious-permissive environment may be maintained by a
specific cytokine milieu, resulting from the peripheral distribution of
helper cell types. Our RT-PCR analysis has revealed that intragraft
IL-2, IL-4, and IL-10 gene expression was diminished in both euthymic
and ATX tolerant hosts. However, grafts from ATX animals showed
significant IFN- gene expression, suggesting a less efficient
down-regulation of Th1-like cells in the absence of regulatory cells.
Because down-regulation of Th1- and Th2-type cytokines represents one
of the cardinal features of clonal anergy in vivo (1), we then asked
whether addition of exogenous IL-2 may break the tolerant state in our
model. Indeed, a course of rIL-2 or infusion of freshly alloactivated
spleen cells recreated cardiac allograft rejection in ATX but not in
euthymic hosts. Thus, unlike in euthymic rats, a state of
cytokine-responsive immune anergy represents the prime mechanism
responsible for the "noninfectious" form of tolerance in ATX hosts.
In the remainder of ATX animals that rejected their transplants in an
acute rather than an accelerated manner, the microenvironmental factors
associated with nonspecific immune activation and cytokine release
disturbing a delicate immune balance may be involved.
Alloantibody responses and isotype switching involve complex interactions between T cells and B cells, which may be regulated by cytokine networks (27). This study extends our previous findings (6, 18, 28, 29, 30, 31) by demonstrating that ATX did not influence CD4 mAb-mediated effects upon host alloantibody networks in this model. As prevention of the switch from IgM to IgG alloantibody response may be critical for long-term allograft acceptance or tolerance (6, 28, 29, 30, 31), no correlation between serum IgM/IgG alloantibody levels and allograft survival in ATX recipients could be found. In fact, both graft rejection and tolerance were accompanied by equally diminished host alloantibody responses, suggesting that ATX + CD4-targeted therapy erased preexisting sensitization, which otherwise triggers a mixed cellular and humoral immune response, culminating in an accelerated allograft loss in sensitized rats. This study confirms our previous findings (6) by demonstrating that systemic IgM and IgG alloantibody responses are virtually abolished in euthymic + CD4 mAb-treated tolerant animals. It remains to be determined, however, to what extent the depression of humoral alloreactivity contributes to the absence of histopathologic abnormalities pathognomonic of posttransplant arteriosclerosis and chronic rejection in the infectious tolerance pathway.
A continuous presence of alloantigen has been identified as a critical factor in the development and maintenance of unresponsiveness both to MHC (3, 32) and non-MHC (33, 34) Ags. While the graft function is maintained, donor alloantigens are shed into the periphery, where they may be able to inactivate newly emerging T cells. The results of our retransplantation studies corroborate other published reports (35) by demonstrating that normal LEW rats rejected LBNF1 hearts despite such hearts having been parked for >100 days in CD4 mAb-conditioned LEW hosts. Hence, this tolerant state does not result from "graft adaptation," and regulatory T cells restricted for the original alloantigen are exposed to its continuous stimulation. In addition, our results from both graft-free and adoptive transfer studies document that effective memory for suppression in the infectious tolerance pathway depends upon persistent donor-specific alloantigen stimulation; it wanes at about 3 wk after its removal. Similarly, regulatory T cells lost their tolerizing potential in murine recipients, if hearts were engrafted 14 days after adoptive transfer (5). Moreover, because a functional graft was required to maintain operational tolerance in our studies, microchimerism via passenger donor-type cells or donor-derived peptides may not be sufficient for tolerance maintenance. Putative mechanisms leading to the loss of tolerance in an alloantigen-free environment may include inactivation of regulatory T cells or a failure of infectious tolerance to divert naive cells toward tolerance.
In summary, the induction of infectious transplantation tolerance in original hosts under the cover of CD4-targeted therapy is thymus dependent, consistent with the role of thymus in generating tolerance-mediating regulatory T suppressor cells. These regulatory T cells remain critically alloantigen dependent and may then "spread" tolerance in an infectious manner into a thymus-free environment of new cohorts of test recipients. A rather unstable level of clonal anergy may replace regulatory T cells in ATX long-term hosts. Hence, both central and peripheral immune mechanisms, orchestrated by the tolerizing donor-type alloantigen, contribute to the development of infectious tolerance in CD4 mAb-treated engrafted rats. This study should add further substance to the current discussion of why all clinical attempts to induce transplantation tolerance in adult patients have been so far unsuccessful.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. J. W. Kupiec-Weglinski, The Dumont-UCLA Transplant Center, 77–120 CHS, 10833 Le Conte Ave, Los Angeles, CA 90095-7054. E-mail address:
3 Abbreviations used in this paper: ATX, adult thymectomy/thymectomized; BN, Brown Norway; LBNF1, (LEW x BN)F1; WF, Wistar-Furth; MST, mean survival time; PE, phycoerythrin; AU, arbitrary unit.
Received for publication December 1, 1997. Accepted for publication February 11, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and B cell stimulatory factor-1 reciprocally regulate Ig isotype production. Science 236:944. expression correlates with increased IgG1 alloantibody response following intrathymic immunomodulation of sensitized rat recipients. Transplantation 60:1516.[Medline]
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, RIB-5/2 mAb-treated euthymic tolerant (>100 days)
hosts: 115.8 ± 13.9 days (n = 7)(*); 
,
untreated naive rats: 6.8 ± 0.3 days (n =
4); and 
, no cell transfer: 10.3 ± 2.1 days
(n = 12) (*statistical comparisons: 
, RIB-5/2 mAb-treated euthymic tolerant hosts
>100 days (n = 4). D, Cardiac
allograft survival in test euthymic rats after adoptive transfer of
spleen cells (100 x 106) from: 
, RIB-5/2
mAb-treated ATX tolerant (>100 days) hosts: 6.3 ± 0.3 days;
n = 4(*) (*statistical comparison: C vs
D, p < 0.0001).
, 30-day graft-free recipients,
76.0 ± 30.5 (n = 4). •, 90-day
graft-free recipients, 11.5 ± 0.3 (n = 4)
(*statistical comparisons: