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Therapeutic Immunology Group, Sir William Dunn School of Pathology, Oxford, United Kingdom
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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E
7. They seem to represent a subset of
transcripts shared with Th2 cells, suggesting that transplantation
tolerance and normal immunoregulation may represent a unique form of
Th2-like differentiation. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The immune system is primarily orchestrated by a range of T cells with
diverse functions. CD8+ T cells are largely
involved with cytotoxicity, while CD4+ T cells
primarily direct the differentiation and effector functions of other
cells through both cell surface interactions and secreted mediators.
CD4+ T cells have been subdivided into at least
three functional Th subsets on the basis of their cytokine production.
Th1 cells are associated with cell-mediated immunity and
characteristically produce IL-2, IFN-
, and TNF-. Th2 cells
regulate humoral immunity, can moderate Th1 responses (6),
and produce cytokines IL-4, IL-5, and IL-10. Both Th1 and Th2 cells can
act as effectors for acute skin graft rejection (7). A
third, as yet poorly defined, subpopulation, variously termed Tr1
(8), Th3 (9), or regulatory T
(Treg)6
cells (10), has been shown capable of regulating T cell
proliferation in vitro and autoimmune pathology in vivo, in the latter
case seeming to require cytokines such as IL-10 and TGF- (8, 10). In addition, there is accumulating evidence of a
subpopulation of CD4+ thymocytes and spleen cells
that is thought to be required to regulate anti-self responses in
normal animals and that is able to suppress autoimmune diseases after
adoptive transfer in vivo (11) and to inhibit T cell
proliferation in vitro (12). It is not clear how, or even
if, these natural regulatory T cells are related to
Tr1/Th3/Treg cells, although they share some features
(13), including the expression of CD4, CD25
(12), CTLA4 (14), and low CD45RB
(15).
We currently have little understanding of the molecular mechanisms by
which regulatory T cells act. The suppression of T cell proliferation
that is observed with some regulatory T cell populations in vitro may
in some cases be via anti-inflammatory cytokines such as IL-10
(8) and TGF- (15), possibly acting to
down-modulate APC functions (16), while in other cases it
has been shown to be dependent on direct contact, possibly through cell
surface molecules such as CTLA4 (14) or surface TGF-
(17). We have shown that naive CD4+
T cell proliferation and IFN- production can be suppressed not only
by Tr1-like but also by Th2 and Th1 clones in vitro (13).
In the case of Th1 cells this can be completely reversed by inhibitors
of NO synthase, but these had no effect on suppression by Th2 or
Tr1-like clones (13). Therefore, there is no clear
consensus about what defines immune regulation in vitro or how this
relates to a regulatory T cell population in vivo, but we hypothesized
that there might be common patterns of gene expression that distinguish
effector T cells from regulatory populations that are independent of
their source or test system.
In this work we describe how we used serial analysis of gene expression (SAGE) (18) to search for gene transcripts that are selectively expressed in various regulatory T cell clones (hereafter collectively identified as Treg) compared with the other T cell subpopulations. We then identified, using quantitative real-time RT-PCR, whether any of the markers associated with these cultured Treg clones were also present on natural CD4+CD25+ regulatory T cells in the spleens of naive mice. We additionally showed that some, but not all, of these candidate Treg markers were indeed increased in tolerant grafts compared with rejecting grafts, and that there was generally a close correlation between genes expressed in tolerant and syngeneic grafts, suggesting a link between allogeneic and self-immune regulatory processes.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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A1(M) x RAG-1-/- and A1(M).CBA TCR-transgenic mice (7), CBA/Ca, CBK (CBA transgenic for Kb) (19), and B10.BR mice were bred and maintained in specific pathogen-free conditions at Sir William Dunn School of Pathology (Oxford, U.K.). Skin grafting was conducted as described previously (1). Tolerance was induced in CBA/Ca recipients by giving three i.p. injections of anti-CD4 and anti-CD8 nondepleting Abs (1 mg each of YTS 177.9.6 and YTS 105.18.10 per injection) over a period of 1 wk starting on the day of transplantation (1). After >100 days mice received a second B10.BR skin transplant that was harvested 7 days postgrafting. All procedures were conducted in accordance with the Home Office Animals (Scientific Procedures) Act of 1986.
Generation of Th1, Th2, Treg, and Tskin CD4+ T cell clones
Spleen cells were taken from a primed A1(M) x RAG-1-/- TCR female transgenic mouse that had been grafted three times previously with male tail skin, and 5 x 105 cells were cultured with mitomycin C-treated male CBA/Ca stimulators in 2 ml of RPMI 1640 and 10% FCS plus either 50 U/ml mouse rIL-2 (to generate the R2.2 Th1 line) or 200 U/ml mouse rIL-4 (to generate the R2.4 Th2 line) as previously described (13). Cells were cloned by limiting dilution in the presence of Ag. The Tr1D1 clone was generated from naive A1(M) x RAG-1-/- spleen cells according to the method of Groux et al. (8). Briefly, 5 x 105 cells were cultured with 5 x 106 mitomycin C-treated male CBA/Ca spleen cells in 2 ml of RPMI 1640 containing 10% FCS and 50 ng/ml IL-10 (Genzyme, Cambridge, MA) for 7 days, at which time spent medium was removed and fresh stimulator cells and medium containing IL-10 were added. After three cycles of polarization in IL-10, viable cells were harvested and cloned at limiting dilution on anti-CD3-coated plates (50 µg/ml 145.2C11) in the presence of mitomycin C-treated female CBA/Ca cells and 20 U/ml IL-2. Cells were expanded with IL-2 (20 U/ml), IL-4 (20 U/ml), and mitomycin C-treated male spleen cells every 2 wk. The clone A1MP was similarly generated from A1(M).CBA naive spleen cells by stimulation with mitomycin C-treated male CBA/Ca spleen stimulators in the presence of 10 µg/ml anti-CLTA4 mAb (clone 4F10; BD PharMingen, San Diego, CA) plus 20 U/ml each of IL-2 plus IL-4. After cloning on anti-CD3 as described above, the cells were maintained and expanded using female mitomycin C-treated spleen cells together with 100 nM DBY-Ek peptide (REEALHQFRSGRKPI) (20), IL-2 (20 U/ml), and IL-4 (20 U/ml) every 2 wk. The Tskin lines were generated by removing secondary challenge male CBA/Ca skin grafts from female A1(M).CBA mice that had previously accepted male skin for >60 days, cutting them into small pieces, digesting with trypsin at 37°C for 1 h, and removing dead cells on nylon wool. Viable lymphocytes were enriched by Histopaque-1083 centrifugation, followed by AutoMACS (Miltenyi Biotec, Auburn, CA) positive selection of CD4+ T cells according to the manufacturer’s instructions. These T cells were then maintained and expanded using female mitomycin C-treated spleen cells together with 100 nM DBY-Ek peptide, IL-2 (20 U/ml), and IL-4 (20 U/ml) every 2 wk.
MACS enrichment of T cell clones
T cell cultures were stimulated for 7 or 14 days with mitomycin-treated male spleen cells from CBK (Kb+) transgenic mice. Viable cells were collected by Histopaque-1083 centrifugation, washed, labeled with biotin-anti-mouse Kb conjugate (clone 28-8-6; BD PharMingen), washed, incubated with MACS streptavidin microbeads (Miltenyi Biotec), and run through the AutoMACS using the slow depletion program. The negative fraction was then further purified using automated positive selection with MACS anti-mouse CD4 microbeads (Miltenyi Biotec). The purity of all fractions was monitored by four-color FACS immunostaining, and the T cell fraction was in all experiments >97% CD4+Kb- cells.
MACS enrichment of spleen CD4+CD25+ cells
Normal CBA/Ca mice were first depleted of
CD8+ T cells in vivo by administration of 1 mg
each of YTS 156.7 and YTS 169.1.2 Abs (21), and after 1
day the spleen cells were harvested and erythrocytes were lysed by
isotonic shock. The spleen cell suspension was labeled with a mix of
anti-mouse Ig
(187.1) and MHC-II (M5/114) mAbs, followed by
sheep anti-rat coupled Dynabeads (Dynal Biotech, Oslo, Norway) and
magnetic depletion according to the manufacturer’s instructions. The
unbound cells were then labeled with biotin-conjugated anti-mouse
CD25 (BD PharMingen) in the presence of Fc block (BD PharMingen),
incubated with streptavidin MACS microbeads (Miltenyi Biotec), and
separated according to the manufacturer’s recommendations on an
AutoMACS using the two-column positive selection program. The
positive fraction was used as the
CD4+CD25+ fraction and was
generally >90% pure by FACS analysis. The negative fraction was
further purified by adding excess streptavidin beads, followed by
further negative selection on the AutoMACS, and finally a positive
selection of the CD4+ cells using CD4-conjugated
MACS microbeads. This fraction was usually >98%
CD4+CD25- cells. To obtain
activated CD4+CD25+ and
CD4+CD25- cells these
purified fractions were incubated overnight with 1 µg/ml plate-bound
anti-CD3 (KT3) at 37°C.
SAGE libraries
Viable cells were harvested from cultures of Th1 (clone R2.2),
Th2 (clone R2.4), Treg (clone Tr1D1), or Tskin
cells, 7 days after Ag stimulation, by density gradient centrifugation
on Histopaque-1083, and in the case of the Treg and
Tskin lines CD4+ cells were purified
by two-step MACS separation and the cell pellets were snap-frozen.
Total RNA was isolated using thiocyanate buffer (4 M guanidinium
thiocyanate, 20 mM NaOAc, 0.1 mM DTT, and 0.5% sodium
lauroyl sarcosine), and RNA was pelleted through a 5.7-M CsCl cushion.
First-strand cDNAs were prepared from 1 µg of total RNA from each of
the cell lines using Superscript II (Life Technologies, Gaithersburg,
MD) annealed with SMARTII oligonucleotide
(5'-AAGCAGTGGTAACAACGCAGAGTACGCGGG-3') and the anchoring primer
(5'-GACTCGAGTTGACATCGAGG(T)20V-3'; Clontech
Laboratories, Palo Alto, CA). The cDNAs were preamplified with the
forward (5'-AGTGGTAACAACGCAGAGTAC-3') and reverse
(5'-GACTCGAGTTGACATCGAG-3') primers using the Advantage-GC cDNA PCR
enzymes (Clontech Laboratories) with 1 M GC-Melt, following the
manufacturer’s protocol. cDNAs were subjected to 16 cycles of
preamplification at 94°C for 30 s and at 68°C for 7 min. The
preamplification steps were monitored by RT-PCR using various
housekeeping and cytokine cDNAs as tests. SAGE was applied to these
samples. The SAGE lymph node libraries were generated from 40 µg of
total RNA, prepared the same way as described above. The
poly(A)+ fraction was purified using
oligo(dT)25 Dynabeads (Dynal Biotech).
Double-stranded cDNAs were generated using the cDNA synthesis kit from
Roche (Lewes, U.K.). SAGE was performed on all these cDNAs using
NlaIII as the anchoring enzyme, BsmF1 as the
tagging enzyme, and SphI as the cloning enzyme, as
previously described (18). DNA sequencing was performed
using the 377 ABI automated sequencer (PE Applied Biosystems, Foster
City, CA). Sequence analysis software SAGE 3.04 was provided
by K. W. Kinzler (Johns Hopkins Oncology Center, Baltimore, MD). A
conservative estimate of the differential up-regulation of each gene
within the given library compared with a pool of the other three
libraries was calculated using a Bayesian statistics model developed by
S. Altschul (22, 23) using the function
f(x) =
xc(1 -
x)c, where c = 3,
i.e., (4, 4). The differential ratio for each tag that could then be
assigned with 95% confidence was obtained by iteration.
Immunofluorescence analysis and Abs
Cells were Ag-stimulated for 14 days in the presence of the
appropriate cytokines (as described above), and viable cells were
collected from Histopaque-1083 centrifugation. The cells were then
stimulated with anti-CD3 (100 ng/ml 145.2C11 absorbed to 24-well
plates), anti-CD3 plus anti-CD28 (clone 37.51 absorbed to wells
at 1 µg/ml), or PMA (50 ng/ml) plus ionomycin (500 ng/ml) or were
left unstimulated in RPMI 1640/10% FCS at 37°C overnight. For FACS
staining, CD4-CyChrome (CD4-PerCP; BD PharMingen) and anti-E
(CD103-biotin, M290; BD PharMingen) were used to label live T cells in
PBS containing 0.1% NaN3, 1% BSA, 10 µg/ml Fc
block (BD PharMingen), and 5% heat-inactivated normal rabbit serum at
4°C. The cells were then washed, fixed in 2% paraformaldehyde,
permeabilized in PBS containing 0.5% saponin, and stained with
Alexa-488 (Molecular Probes, Eugene, OR)-conjugated rabbit
anti-Ly116 C-terminal peptide (NVPGNVYKNHPGEIV; AbCam, Cambridge,
U.K.), anti-CTLA-4-PE (4F10; BD PharMingen), and
streptavidin-allophycocyanin (BD PharMingen) in the dark at 4°C.
Four-color analysis was performed using a FACSort (BD Biosciences,
Oxford, U.K.) with dual laser (488 and 633 nm) excitation in
combination with data acquisition and cross-beam color compensation
using CellQuest 3.1 software (BD Biosciences). The analysis gate was
set on the forward and side scatters to eliminate cell debris and dead
cells.
Real-time quantitative RT-PCR
Total RNA from grafted tissues or from purified populations from
the T cell clone cultures was prepared using the SV Total RNA isolation
system (Promega, Madison, WI), followed by DNase I treatment. Rever
transcription was performed using the proStar kit with random hexamers
(Stratagene, Cedar Creek, TX). From a total volume of 50 µl/cDNA, 2.5
µl were used in the PCR reactions. Real-time quantification was
performed using gene-specific, fluorogenic probes and the Universal
MasterMix kit (PE Applied Biosystems) in a final volume of 25 µl. The
reaction mixture contains all primers at 300 nM and the probe at 200
nM. The enzyme was heat-activated for 10 min at 95°C. A two-step PCR
procedure of 15 s at 95°C and 60 s at 60°C was applied
for 40 cycles. PCR and TaqMan analysis were performed using the
ABI/PRISM 7700 sequence detector system (PE Applied Biosystems). The
multiplex PCR reactions were performed using VIC-labeled CD3
or hypoxanthine-guanine phosphoribosyltransferase (HPRT) probes and
FAM-labeled test probes, as shown in Table II
. Standard curves
of cDNAs from the R2.2, R2.4, or Tr1D1 clones were used to calibrate
the threshold cycle to amounts of test and normalizing cDNAs on each
96-well plate run. Normalized values for mRNA expression were
calculated as (1000 x test mean)/(normalizer mean), except where
stated otherwise. All samples were run in triplicate. Significance
between values for skin graft groups was calculated by a Mann-Whitney
U test.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We derived Th1, Th2, and Treg
CD4+ T cell clones with identical specificity for
a male (H-Y)-derived peptide (REEALHQFRSGRKPI) in association with
H2-Ek (DBY-Ek)
(20) from the A1(M) TCR-transgenic mouse on the
RAG-1-/- background as previously described
(Table I). Both the Th1 (R2.2) and Th2
(R2.4) clones were stable over many months and produced the appropriate
cytokines whether stimulated with male spleen cells, peptide-pulsed
bmDCs, CD3 cross-linking, or PMA plus ionomycin (data not shown). We
have shown previously that both Th1 and Th2 clones elicit rapid
rejection of male skin grafts after adoptive transfer to T
cell-depleted recipients (7). Tr1-like Treg
cells were derived from the spleen cells of naive A1(M) x
RAG-1-/- mice following the protocol of Groux
et al. (8). Alternatively, spleen cells from nonresponding
A1(M).CBA female mice were repeatedly stimulated by male Ag in the
presence of anti-CTLA4 mAb (13). We had previously
found that this enhanced the proliferation of Treg clones
without apparently modifying the phenotype (13). This line
was cloned on anti-CD3 (clone A1MP), then maintained on 100 nM
DBY-Ek peptide. Finally, we generated T cell
lines against DBY-Ek peptide from male skin grafts that had
been permanently accepted by A1(M).CBA females (here called
Tskin lines). All these CD4+
Treg clones and the Tskin cell lines from
tolerated skin grafts shared a similar phenotype
(CD25+ and CTLA-4++),
secreted IL-10 and variable IL-4 but no IFN- (13), and
could suppress the proliferation and IFN- production of naive or Th1
cells in vitro (13) (data not shown).
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It was important to demonstrate that cultured Treg
clones could suppress skin graft rejection in vivo, as there is still
no proven surrogate assay for regulatory T cell activity in vitro. We
adoptively transferred each of the Th1, Th2, or Treg clones
into RAG-1-/- female recipients of a male skin
graft. As expected, both the Th1 and Th2 clones caused rapid rejection
of the male, but not control female, skin grafts (Fig. 1). This result confirmed what we had
previously demonstrated in T cell-depleted mice (7) and
also work by others (24, 25), i.e., that both Th1 and Th2
responses are capable of causing acute graft rejection. More
significantly, the Treg clone (Tr1D1) failed to reject four
of five male grafts but was subsequently able to suppress the rejection
of fresh male grafts given to these same mice together with sufficient
new Th1 cells to cause rapid rejection in controls (Fig. 1).
Previously, we have also demonstrated that the A1MP clone was able to
suppress the rejection of male B10.BR grafts by adoptively transferred
CBA/Ca spleen cells (13). Therefore, we have evidence that
two independently derived CD4+ T cell clones with
similar phenotype and growth characteristics in vitro are able to act
as regulatory T cells to suppress skin graft rejection in
vivo.
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SAGE libraries were first produced from the Th1, Th2, and
Treg (Tr1D1) CD4+ T cell clones and
the Tskin line 7 days after their last stimulation. The
intention was to minimize the APC contribution and to bias the search
to stable lineage markers rather than transitory activation Ags. Our
first SAGE analysis of genes in Treg cultures
(13) had found an association with mast cells that was a
consistent finding with all the independent Treg clones and
lines described in this work, so the analysis presented here is from
new libraries derived from highly purified (>99%)
CD4+ Treg and Tskin cells.
We also made SAGE libraries from the draining lymph nodes of equivalent
A1(M) x RAG-1-/- TCR-transgenic mice 7 days
after a second-set challenge with male skin, as this would act as a
control to subtract many housekeeping and non-Treg cell gene
transcripts from our T cell clone-derived libraries. We sequenced a
minimum of 10,000 tags in each of the six libraries (Th1, Th2,
Treg, CD4+ Treg, and
Tskin cell lines and lymph nodes), providing a total of
97,690 tags corresponding to 5,257 different unique gene tags detected
three times or more in the combined libraries. Our intention was to
obtain sufficient tags to identify a reasonable number of candidate
genes with a statistically significant differential expression between
libraries. The differential analyses are depicted graphically in Fig. 2 as pairwise scatter plots.
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The comparison of the Th1 and Th2 libraries (Fig. 2
a)
highlighted eight tags overexpressed in Th1 cells and 14 in Th2 cells.
In the case of the Th1 clone these tags mapped to transcripts known to
be associated with Th1 cells, including RANTES, Ly116
(26), the transcription factor lymphoid enhancer binding
factor (LEF)-1, and IL-2R. The preferential Th1 expression of
the first three was confirmed by TaqMan quantitative real-time RT-PCR
(data not shown). Additionally, we identified transcripts previously
associated with NK cells: Ly6-C.2 and gp49A. The Th2 clone also
generated appropriate up-regulated tags, including IL-10, GATA-3, and
IL-1RII, that were also confirmed by TaqMan RT-PCR (data not shown).
Transcripts for both IgM H and Ig L chain constant regions
(presumably germline in the RAG-1-/- cells and
unlikely to be from contaminating B cells, as these could not be
detected by immunofluorescence and no MHC-II-associated tags were
present) may be in response to cytokines such as IL-4 and IL-5 that we
know are made by the Th2 cells (data not shown). Other Th2 transcripts
identified by SAGE included glutaredoxin and Ly6E.
Differentially expressed gene transcripts in Th2 and Treg cells
To identify Treg-specific gene transcripts we compared
the highly purified CD4+ Treg clone Tr1D1 with
both Th1 (Fig. 2b) and Th2 (Fig. 2c) SAGE
libraries. The first point to note is that while Treg cells
express a number of different genes compared with Th1 cells, for
example, granzyme A, prepro-enkephalin (ppENK), OX40, Unigene cluster
mm46382, and integrin 7, these were shared,
but generally at a lower frequency, with the Th2 clone, suggesting that
despite their different abilities to promote or suppress graft
rejection, the Th2 and Treg cells are closely related.
However, a considerable number of Th2-expressed genes were
down-regulated on the Treg cells, including the Ig germline
transcripts, 2-microglobulin (but not MHC
class I), Egr-1, and GATA-3, with a further down-regulation of IL-2R
from Th1 to Th2 to Treg. It may be that this specific gene
down-regulation, particularly of the two transcription factors GATA-3
and Egr-1, is associated with the loss of graft rejection ability, as
they may be required to initiate appropriate patterns of effector gene
expression in the Th2-committed lineage (27). The only two
tags unique to this CD4-purified Treg library were a tag
(CATGCGCCGCGGCT) that we could not assign to any known transcript and a
histone-associated gene (H2A.1; Fig. 2c).
The comparison shown in Fig. 2d of the unmanipulated
Treg culture and that from highly purified
CD4+ Treg cells demonstrate that even
as few as 5% mast cells can dominate the mRNA pool, as shown by high
frequencies of mast cell genes (mast cell protease 5, carboxypeptidase
A3, tryptophan hydroxylase, and CD63). Interestingly, the tags for
GATA-3, ST2L, and Egr-1 were found within the whole Tr1D1 cultures but
were clearly associated only with the mast cell-containing SAGE
library. We have discussed this association between Treg and
mast cells in more detail previously (13). Similarly, it
can be seen that the mix of cell types from draining lymph nodes of
grafted A1(M)xRAG-1-/- mice, despite being
numerically dominated by CD4+ T cells, is
strongly biased to tags from macrophages (and possibly germline
B-lineage cells), with a high expression of MHC-II -, -, and
invariant chains. The fact that these tags were not observed in the T
cell clone libraries makes it unlikely that the present analysis is
contaminated with any residual APCs.
Therefore, the strongest candidates for known genes that may be
positively associated with the Tr1D1 clone were ppENK, granzyme A, GM2
ganglioside activator protein (GM2a), cystatin F, integrin
7, OX40, the glucocorticoid-induced TNFR
superfamily member 18 (GITR; also known as TNFRsf18) (28),
and the cytochrome P450 enzyme Cyp11a, which is the rate-limiting step
in glucocorticoid synthesis (29). Significantly, all these
candidates were similarly expressed when the SAGE libraries of the
Tr1D1 clone and the Tskin line from the tolerated grafts
were compared (Fig. 2f). Indeed, there was an overall very
high correlation between the SAGE data from these two independent
sources of mRNA, confirming the repeatability of the SAGE methodology
and also suggesting the two cell types are functionally related.
Validation of SAGE data by quantitative RT-PCR and immunofluorescence
The SAGE data from T cells was obtained 7 days after Ag stimulation to deliberately try and bias the search away from transient activation-related transcripts and toward potential stable differentiation markers, so we wished to test whether the above candidate genes would remain preferentially associated with Treg cells under both resting and recently activated conditions. We first examined the expression of OX40 by immunofluorescence but found it to be expressed on all T cell populations after activation (data not shown), behaving similarly to CD25 and CTLA4 as being common activation markers that are constitutively expressed by Treg cells.
We next looked at the integrin 7, which can be
associated with either 4 or
E chains, with the latter (also known as
CD103) generally being thought to be a marker for intraepithelial T
cells (30). We did not find recognized SAGE tags for
either of these integrin -chains within our SAGE libraries, but this
could be due to expression below the level of detection with the number
of tags sequenced or to the fact that the 3'-untranslated regions of
these genes have not been fully characterized, so the correct SAGE tag
may not be known. However, immunofluorescence demonstrated that the
Treg and Tskin cells stained
highly for 7 (data not shown) and, more
interestingly, a variable proportion of the cells were also positive
for E/CD103 (Fig. 3). We then activated the clones with
anti-CD3, anti-CD3 plus anti-CD28, or PMA plus ionomycin
(Fig. 3) and found that although there was some weak expression in the
activated Th2 clone, the Treg and Tskin cells
maintained, if not increased, their CD103 expression, while Th1 cells
remained negative under all conditions. Also of considerable
interest was the observation that the majority (61%) of
E+CD4+
cells from normal CBA spleens were contained within and made up 
20%
of the spleen CD4+CD25+
population (Fig. 3). These
E+CD4+CD25+
cells were also found to be negative when permeabilized and stained
with an Ab to the C-terminal peptide of Ly116 (Chandra) that is
considered a Th1 marker (26), which is expressed on a
different subset of 30% of
E-CD4+CD25+
cells (data not shown). This combination of E
and Ly116 staining further demonstrates the heterogeneity of the
CD4+CD25+ subset that is
thought to contain the predominant natural Treg population,
but that may also contain other memory subpopulations.
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We then proceeded to compare the expression of the remaining gene
candidates, by TaqMan quantitative RT-PCR, on resting and activated T
cell clones and fractionated subpopulations of normal spleen
CD4+ T cells (Fig. 4). Of particular interest was the
finding that ppENK was not expressed in resting Th1 or
CD4+CD25- cells, and even
after activation it remained low or undetectable, respectively, while
it was highly expressed and further up-regulated after activation in
Treg, Tskin, and normal spleen
CD4+CD25+ T cells. GM2a was
not expressed on Th1 or
CD4+CD25- cells, but it
was detected at high levels in the resting Th2, Treg, and
Tskin lines. However, these GM2a transcripts were lost on
all populations after CD3 stimulation. The GITR transcript was detected
on all resting T cell clones and CD4+
subpopulations, although it was lost after activation of the Th1 clone,
but increased after CD3 stimulation of the
CD4+CD25+ cells.
Immunofluorescence staining with a polyclonal Ab to mouse GITR (BAF
524; R&D Systems, Minneapolis, MN) was positive on Th2 and Tr1D1 clones
regardless of activation, but negative on Th1 (data not shown),
confirming the SAGE analysis. The steroid synthetic enzyme Cyp11a was
not detected on Th1 cells or normal spleen CD4+
subpopulations, but the SAGE data were confirmed by a strong signal on
resting Th2 cells and weaker signals from the Treg and
Tskin lines. While granzyme B was up-regulated after
stimulation of all T cell lines and clones, granzyme A remained
undetectable in Th1 cells. The two granzyme transcripts also differed
in their expression on normal spleen CD4+
subpopulations, with granzyme B remaining undetected, while granzyme A
was highest in the stimulated
CD4+CD25+ cells. TGF-1
mRNA, which was not identified as differential from the SAGE analysis
but has previously been associated with Treg cells
(15), was detected in all T cell populations, while
TGF-2 was at the lower limit of detection in Tr1D1 and A1MP cells.
Finally, CD3, which was used as an additional control, was similarly
present in all samples, as expected.
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Transcripts associated with Treg cells are selectively overexpressed in tolerated grafts
We then turned to a well-established in vivo model of
transplantation to test whether there was any evidence for our
candidate Treg genes being associated with donor-specific
tolerance. CBA/Ca mice can be made tolerant of a B10.BR multiple minor
mismatched skin graft by using a short course of nondepleting CD4 and
CD8 T cell Abs. We have previously shown that in such mice tolerance is
dependent on and mediated by CD4+ T cells
(3). We performed a quantitative TaqMan RT-PCR analysis of
the mRNA expression in the transplanted tissue of genes representative
of the three T cell subsets. First we assessed the degree of T cell
infiltration by measuring CD3 in a multiplex reaction standardized
to HPRT. Very little CD3 was detected in normal skin, but by
comparison it was readily detectable (but variable and low compared
with HPRT) in all skin grafts regardless of whether they were
allogeneic or syngeneic (Fig. 5). This
allowed us to normalize all subsequent mRNA measurements in syngeneic,
tolerant, or primed skin grafts to the level of the CD3 T cell
message (Fig. 5), although we found comparable results when normalizing
to other nondifferential transcripts of similar abundance to those
tested (e.g., RANTES; data not shown).
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).
TGF-2, ppENK, GM2a, GITR, and IL-1R2 showed clear up-regulation in
tolerated compared with rejecting challenge grafts of 14.8-, 624-,
32.6-, 2.8-, and 3.4-fold, respectively, while Cyp11a was
down-regulated by 7-fold. Of particular interest was that most genes
that were up-regulated in tolerated allogeneic skin grafts were
expressed similarly in syngeneic grafted skin. Indeed, there was a high
correlation between the genes expressed in tolerant and syngeneic skin
(r2 = 0.9935; Fig. 5b), while
there was much less concordance between tolerant and rejecting grafts
(r2 = 0.1414; Fig. 5c). This
suggests that the mechanisms of transplantation tolerance induced by
CD4 and CD8 mAb treatment are similar to those involved in the
acceptance of tolerated self-tissues. This may not be surprising with
the accumulating evidence for the role of CD4+
regulatory T cells in the prevention of autoimmunity and inflammatory
bowel disease but may also reflect a role for cells of this type in
so-called protective autoimmunity (31). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We have identified a number of genes that are expressed by Th2 cells
that are further up-regulated in Treg clones and T cells
from tolerated skin grafts and that are also expressed in normal
splenic subpopulations of
CD4+CD25+ cells. The first
of these, the E/7
integrin, has generally been associated with effector cell activity of
CD8+ intraepithelial cells in the skin and gut
(30). However, it is interesting to note that
E-deficient mice develop autoimmune-like skin
lesions when crossed to susceptible backgrounds
(33), suggesting a role in immune regulation within these
tissues. The finding that
E/7 (CD103) is
expressed on 20% of the
CD4+CD25+ population, which
is thought to contain the natural regulators of autoreactivity, is
intriguing in light of studies that suggested the first wave of thymic
emigrants in young mice have a different capacity to migrate through
tissues such as the skin compared with adult naive T cells
(34), where they may become tissue-specific regulatory T
cells.
Also unexpected was the high level expression of transcripts for ppENK in Treg cells and activated CD4+CD25+ populations. Prepro-ENK belongs to the opioid family and can be cleaved into different active enkephalin peptides by proteases. Remarkably, although brain and adrenal gland are the classical sites of expression of this family of molecules, ppENK cDNA was originally cloned from stimulated T cells (35). Although Abs are available to the two main products of ppENK (Met and Leu enkephalins), and these have been reported to stain Th2 cells by immunohistology (36), we have been unable to demonstrate convincing staining of any T cells or grafted tissues with such Abs. This may be because proENK can be secreted intact and processed by a number of different proteases from other cells (e.g., mast cell carboxypeptidase) to generate a range of peptides in addition to the Met and Leu enkephalins, including the non-opioid synenkephalin moiety (37). Mechanistically, there is one report showing that Met-enkephalin peptides can inhibit induced chemotaxis of Th1 cells through receptor desensitization (38), and this may be a means for regulatory cells to act locally by inhibiting a further influx of inflammatory cells. This might relate to the observation that tolerated skin grafts or regrafts, while often having similar levels of T cell infiltration, do not have the massive influx of polymorphonuclear cells associated with rejecting skin grafts (our unpublished observation).
The adrenal glands are not only a source of enkephalins but also have a major influence on the immune system through the production of glucocorticoids. This is most clearly demonstrated in acutely induced autoimmune models such as experimental allergic encephalomyelitis, where the steroids produced by the adrenal glands are required for the resolution of the disease and subsequent T cell-mediated resistance to further induction (39). Both the natural steroid-dependent remission and the use of artificial steroids in vivo and in vitro seem to be associated with promoting a form of immune deviation from Th1 to Th2 (40). Therefore, it is surprising to find that Th2 and Treg cells have mRNA for the critical rate-limiting enzyme, Cyp11a, for the synthesis of glucocorticoids from cholesterol. It has recently been reported that thymocytes may also express this gene (40) and that this may play a role in thymocyte selection, although the area remains controversial. Regulatory T cells have also been suggested to modulate dendritic cell maturation and Ag presentation in favor of tolerance, and one of the most potent ways to achieve this experimentally is using the artificial steroid dexamethasone (29, 40). In addition to the potential to produce glucocorticoids, the Treg cells express the mRNA for TNFRsf18, also known as the GITR protein (41), suggesting that there may be an autocrine loop of steroid production and responsiveness. It is interesting to speculate that this might provide a mechanism for maintaining regulatory T cell activity after an initial external pulse of glucocorticoids from the adrenal glands. However, unlike most of the other Th2- and Treg-associated markers that were associated with tolerance in vivo, RT-PCR of the Cyp11a transcripts at the single time point of 7 days after regrafting suggested that syngenic and rejecting grafts had higher levels of expression than the tolerated skin. While this seems counterintuitive, it may be that the timing of Cyp11a expression is an important factor, and this will need to be further investigated.
Two of the genes, ppENK and GM2 activator, while expressed on Treg cells in vitro and in vivo, were seen to respond differently to CD3-mediated T cell activation. While ppENK was markedly increased in all Th2 and Treg populations and was found to be positive on T cell lines and fresh T cells extracted from tolerated skin grafts (data not shown), GM2 activator was lost by activation. Although it was apparently up-regulated in whole tolerated skin compared with rejecting grafts, it was not detected on freshly isolated graft infiltrating cells (data not shown). GM2 activator was also detected in normal, unstimulated CD4+CD25+ and unfractionated spleen cells but was again lost on stimulation. This suggests that ppENK may be a good marker to detect Th2 and Treg activity within local sites of tolerance, while GM2 activator may be a better marker for the Treg resting state, e.g., in spleen, which is a good source of Treg cells that suppress rejection after adoptive transfer. However, both ppENK and GM2a were increased in the tolerated skin grafts 7 days after regrafting, so there may be further complexity in the timing of regulatory gene activity in vivo. It is not clear whether GM2 activator could have a functional role in immunoregulation, but there is one report that it is able to act as an inhibitor of platelet-activating factor, which is a potent proinflammatory mediator (41).
Although the gene transcripts we used to test for by RT-PCR of whole
skin grafts were selected on the basis of a restriction to Th2 and
Treg cells, we could not be sure that there were not other
additional, non-T cell sources of these transcripts as a result of
nonspecific inflammation within the grafted tissues. Indeed, although
we only found evidence for TGF-1 being highly expressed by T cells,
we found that TGF-2 (as well as TGF-1, -3, and -4; data not
shown) was up-regulated in tolerant and syngeneic grafts. This raises
the possibility that there may be considerable interplay between
regulatory T cell-mediated tolerance and tissue protection and repair
mechanisms. Indeed, there may be a precedent for this where
encephalitogenic, autoimmune T cells have been implicated in the
processes of nerve repair (31).
In summary, we have identified gene transcripts that may be indicative of regulatory T cell populations in vitro and in vivo, and these suggest that there are multiple molecular mechanisms involved in the maintenance of tolerance in vivo.
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Footnotes |
|---|
2 D.Z. and E.A. contributed equally to this work.
3 Current address: Laboratoire Hématopoïèse et Cellules Souches, Institut Gustave Roussy, Institut National de la Santé et de la Recherche Médicale Unité 362, Villejuif, France.
4 S.P.C. and H.W. are joint senior authors.
5 Address correspondence and reprint requests to Prof. Herman Waldmann, Therapeutic Immunology Group, Sir William Dunn School of Pathology, South Parks Road, Oxford OX1 3RE, U.K. E-mail address: herman.waldmann{at}pathology.ox.ac.uk
6 Abbreviations used in this paper: Treg, regulatory T; GITR, glucocorticoid-induced TNFR superfamily member 18; GM2a, GM2 ganglioside activator protein; HPRT, hypoxanthine-guanine phosphoribosyl transferase; ppENK, prepro-enkephalin; MHC-II, MHC class II; SAGE, serial analysis of gene expression; LEF, lymphoid enhancer binding factor.
Received for publication October 24, 2001. Accepted for publication November 15, 2001.
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; n = 5) or 5 x
106 Th2 (R2.4; 
: n = 5) cells both
caused rapid rejection of the male, but not the female (
), skin
grafts. Four of five mice given 1 x 107
Treg (Tr1D1; •) cells, and those given only 0.5
x 106 Th1 (R2.2; 
) did not reject their grafts up to
day 95. These latter two groups were then regrafted with male and
female skin and given sufficient Th1 (5 x 106 R2.2)
cells to normally cause rapid rejection. The group that had initially
received insufficient Th1 cells to elicit rejection lost both their
first and second grafts rapidly (
; compared with 
mean) were removed from all the plots (
