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*
Cancer Research Campaign Institute for Cancer Studies and
Medical Research Council, Centre for Immune Regulation, University of Birmingham, Birmingham, United Kingdom;
Medical Research Council Human Immunology Unit, Institute of Molecular Medicine, John Radcliffe Hospital, Oxford, United Kingdom; and
University School of Medicine, Shigenobu, Ehime, Japan.
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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,
TNF-
) responses, measured by intracytoplasmic staining after peptide
stimulation, also were detectable in CD45RO+ and
RA+ subsets as well as CD28+ and
CD28- subsets. Of other markers that were heterogeneous in
both lytic and latent epitope populations, CCR7 gave the best
discrimination of functionality; thus, CCR7+ cells
consistently failed to give an IFN- or TNF- response, whereas
many CCR7- cells were responsive. Our data are consistent
with effector functions having a broad distribution among
phenotypically distinct subsets of "effector memory" cells that
have lost the CCR7 marker. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Although the primary effectors of a CD8+ T
cell-mediated response in man are phenotypically identifiable as
activated cells expressing the CD45RO isoform and activation markers
such as CD38, CD69, and HLA-DR (3, 4, 5), the assignment of
Ag-specific memory to a phenotypically distinct subset of circulating
CD8+ T cells has proved controversial. Even
before single-cell assays allowed Ag-specific memory cells to be
individually identified in lymphocyte preparations ex vivo, it was
suggested that the Ag-experienced human CD8+ T
cell pool could be distinguished from the naive pool on the basis of
increased CD11a/CD18 (LFA1) adhesion molecule expression
(6). However, in terms of other putative markers of
differentiation such as CD45RO, the other isoform CD45RA (once thought
to be unique to naive cells), CD27, CD28, CD57, and CD62L, the
LFA1high population is quite heterogeneous
(7, 8, 9). This led to numerous suggestions that these
phenotypically distinct subsets were functionally distinct, and in
particular that an immediate effector function such as cytotoxicity was
confined to particular CD8+ subsets, especially
those defined as CD45RA+
CD27- (7, 8, 10) and/or
CD28- (11, 12, 13), whereas the
capacity for immediate cytokine (IFN-, TNF-) release was more
widespread. In this latter context, a more recent study based on the
chemokine receptor CCR7 as a discriminatory marker has
suggested that the capacity for immediate IFN-/TNF- release is
restricted to a CCR7- subset of Ag-experienced
CD8+ T cells (14). However, these
studies focused on all of the cells within a particular
CD8+ subpopulation and not on cells responsive to
a particular Ag. Accordingly, effector functions were assayed
in a non-Ag-specific manner. Cytotoxicity was assessed by
anti-CD3 mAb -stimulated (redirected) lysis of Fc receptor-positive
target cells and immediate cytokine production by PMA/ionomycin
stimulation, approaches which may well not be representative of
Ag-specific responses.
The identification of viral peptide epitopes against which the human
CD8+ T cell response to common pathogens is
directed has opened the way for new assays, based on staining with HLA
class I/peptide tetramers (15) and on peptide-induced
cytokine release (16, 17), which allow epitope-specific T
cells in memory to be identified at the single-cell level. The present
work focuses on one such pathogen, the EBV, a persistent
-herpesvirus that establishes both lytic and latent infections in
vivo and where both types of infection elicit detectable
CD8+ responses (18). Studies on
patients with primary EBV infection, manifesting as infectious
mononucleosis (IM),4 have shown
that the primary EBV-specific response accounts for much if not all of
the highly expanded pool of activated CD45RO+
CD8+ T cells seen in the blood during this
disease, with the response to lytic cycle Ags being amplified to
10-fold higher levels than that to latent Ags (3). Those
expansions, visualized by tetramer staining, also are reflected
functionally when IM effectors are tested on peptide-loaded targets in
ex vivo cytotoxicity assays (19, 20). In the subsequent
life-long virus carrier state (both in post-IM patients and in
individuals who had asymptomatic primary EBV infection), small numbers
of both lytic and latent Ag-specific cells are maintained and together
may account for 1–3% of the circulating CD8+
pool, with lytic epitope reactivities again usually in the majority
(21). Interestingly, not all of these cells appear to be
functionally equivalent because estimates of epitope-specific T cell
numbers in the blood of healthy carriers from ELISPOT assays of
peptide-induced IFN- release are generally 20–40% of the numbers
assessed by tetramer staining (21). It is not known how
differences in the capacity for IFN- production might relate to
phenotypic heterogeneity seen within EBV-specific memory populations,
nor whether any of these cells might also be capable of a second
immediate effector function, such as epitope-specific target cell
lysis. As with all persistent viruses, it might be argued that at any
one time, chronic antigenic stimulation might be driving a small number
of memory cells to an activated effector phase like that seen in the
acute primary infection and that any detectable cytotoxicity would be
restricted to cells with this phenotype.
In the present study, we have developed the work looking for relationships between EBV-specific memory CD8+ phenotype and effector capacity in three ways: 1) by comparing memory populations derived from the highly amplified primary response to EBV lytic cycle Ags with those derived from the smaller response to latent cycle Ags; 2) by using intracellular staining as a more rapid assay of the epitope-induced cytokine response (22), where the phenotype of responsive cells can be simultaneously identified; and 3) by asking whether any of these phenotypically distinct memory populations might mediate detectable epitope-specific cytolysis in ex vivo cytotoxicity assays.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The following EBV-epitope peptides were used in this study: the A2-restricted epitopes GLCTLVAML derived from the lytic cycle protein BMLF1 (23), YVLDHLIVV derived from the lytic cycle protein BMRF1 (24), and CLGGLLTMV derived from the latent cycle protein LMP2 (25). The B8-restricted epitope peptides used in this study were RAKFKQLL derived from the lytic cycle protein BZLF1 (26), FLRGRAYGL derived from the latent cycle protein EBNA3A (27), and QAKWRLQTL derived from the latent cycle protein EBNA3A (28). All epitope peptides are abbreviated to the first three amino acids throughout this text. The peptides were purchased from Alta Biosciences (University of Birmingham, Birmingham, U.K.) and diluted in DMSO (Fisher Chemicals, Loughborough, U.K.) to a concentration of 5 mg/ml and diluted to appropriate concentrations in RPMI 1640 immediately before use.
HLA class I/peptide tetramers were synthesized as described previously (15). Briefly, HLA-A2 recombinant heavy chains containing the Bir A biotinylation motif were refolded with either of the peptides GLC, YVL, or CLG, and HLA-B8 recombinant heavy chains were refolded in the presence of either FLR, QAK, or RAK peptide. After biotinylation with Bir A (Avidity, Denver, CO) the refolded heavy chains were purified on a Superdex G75 HiLoad 26/60 prep grade FPLC column (Amersham Pharmacia Biotech, Uppsala, Sweden) followed by further purification on a Mono Q anion exchange FPLC column (Amersham Pharmacia Biotech). The purified monomers were tetramerized by addition of streptavidin R-phycoerythrin (Molecular Probes, Eugene, OR) at a molar ratio of 4:1, respectively.
Donors
Healthy laboratory donors that were HLA-A2- and/or HLA B8-positive as judged by sequence-specific primer PCR were recruited for this study. All donors were identified as either EBV-seropositive (i.e., virus carriers) or EBV-seronegative (i.e., noninfected) by standard serum Ab assays for virus capsid Ags.
Cell preparation
Donors were bled 120 ml into heparin, and their PBMCs were
isolated by centrifugation on a lymphoprep gradient (Nycomed Pharma,
Oslo, Norway) as per the manufacturer’s instructions. PBMCs were
enriched for CD8+ cells by incubation of PBMCs
with mAbs specific for CD4 (clone RFT4), CD19 (clone RFB19), 
TCR, CD14, and CD16 (Immunotech, Marseilles, France), as well as Ab
specific for glycophorin (BD PharMingen, San Diego, CA) to remove
contaminating erythrocytes. The Ab-coated cells were depleted by
multiple rounds of incubation with sheep anti-mouse Ab-coated
Dynabeads M-450 (Dynal, Oslo, Norway) as per the manufacturer’s
instructions.
Subsets of CD8 cells were additionally enriched by depletion of cells with specific markers. CD8+ CD45RA+ cells were selected by depletion of CD45RO+ cells through incubation of the enriched CD8+ population with an anti-CD45RO mAb (clone UCHL1), whereas CD8+ CD45RO+ cells were enriched by incubation of the CD8+ population with an anti-CD45RA mAb (clone SN130), followed by magnetic bead depletion.
CD62L-negative cells were enriched by depletion of CD62L-expressing cells from the CD8+ population. CD8+ cells were incubated with an anti-CD62L mAb (BD Biosciences, San Jose, CA) followed by incubation with goat anti-mouse Dynabeads M450 and magnetic separation.
CD28-negative cells were enriched for by FACS. CD8+-enriched cells were stained with an anti-CD28 mAb (BD PharMingen) followed by incubation with goat anti-mouse heavy-light chain-specific FITC-labeled Abs (Southern Biotechnology Associates, Birmingham, AL). Nonstained cells were separated from CD28-expressing cells by using a FACSVantage (BD Biosciences).
Tetramer staining assays
Staining of lymphocytes was undertaken by incubating the cells
with a pretitrated concentration (
0.5 µg/ml) of tetramer at 37°C
for 15 min. The cells then were stained for surface markers by
incubation on ice with saturating amounts of anti-human CD8
conjugated to Tricolor (Caltag Laboratories, Burlingame, CA) and
CD45RA-FITC (Beckman Coulter, Fullerton, CA), CD45RO-FITC (Dako, Ely,
U.K.), CD28-FITC (BD PharMingen), CD27-FITC (BD PharMingen), CD56-FITC
(Immuno-Kontact, Abingdon, U.K.), CD69-FITC (BD PharMingen), or
CD62L-FITC (Caltag Laboratories). To detect CCR7 expression,
tetramer-stained cells were incubated with an anti-human CCR7 mAb
(a kind gift from Millennium Pharmaceuticals, Cambridge, MA). Binding
of the CCR7-specific mAb was detected by incubation with goat
anti-mouse heavy-light chain-specific FITC-labeled Ab (Southern
Biotechnology Associates). Free anti-mouse binding sites were
subsequently blocked by incubation with normal mouse serum
and the cells then stained with anti-human CD8 Tricolor labeled Abs
(Caltag Laboratories). These stained cells then were analyzed on an
Epics flow cytometer (Beckman Coulter). Note that color compensation
between the different fluorochromes was set with single
fluorochrome-stained cells from the same test population as described
previously (29).
For four-color flow cytometric analysis, cells were stained with tetramer and anti-CCR7 Abs as described above. These cells then were stained with anti-human CD8-PerCP (Becton Dickinson) in combination with allophycocyanin (APC)-labeled Abs specific for either CD45RA, CD45RO, CD62L, or CD28 (BD PharMingen). These cells then were analyzed with a FACSCalibur flow cytometer (BD Biosciences).
Cytokine secretion assays
IFN- production by cells was assayed either by ELISPOT
analysis or by intracellular cytoplasmic staining, and TNF-
production was assayed by intracellular cytoplasmic staining.
For ELISPOT analysis, serial dilutions of cells were plated on
MultiScreen Immobilon-P filtration plates (Millipore, Bedford, MA) that
had been precoated with IFN--specific mAbs from an IFN- ELISPOT
kit used as per the manufacturer’s instructions (Mabtech, Nacka,
Sweden). The cells were incubated overnight with peptide at a
concentration of 20 µg/ml or an equivalent dilution of DMSO. IFN-
production by the cells was shown by using a human IFN- ELISPOT kit
(Mabtech) and regions of secreted IFN- revealed by using an alkaline
phosphatase chromogenic substrate kit (Bio-Rad, Hercules, CA). These
regions then were counted with a dissecting microscope.
Intracellular cytokines produced by CD8+
lymphocytes were detected by initially incubating PBMCs at room
temperature for 30 min with a phycoerythrin-labeled HLA-A2 GLC or an
HLA-B8 RAK tetramer in RPMI 1640 supplemented with 10% FCS. The cells
were washed and incubated in RPMI 1640 10% FCS in the presence or
absence of GLC or RAK peptide, respectively, at a final concentration
of 10 µM, in V-bottom tubes, at 37°C in the presence of 5%
CO2 for 6 h. Brefeldin A (Sigma-Aldrich,
Poole, U.K.) was added to the cultures at a final concentration of 5
µg/ml after the first hour of incubation. The cells then were stained
with saturating amounts of anti-CD8-PerCP (BD Biosciences), washed
in PBS, and fixed in 4% formaldehyde. The cells then were washed twice
in a PBS buffer containing 0.1% saponin and 1% FCS (permeabilization
buffer), and stained with either 0.5 µg of anti-IFN--APC Ab
(BD Biosciences) or 0.5 µg of anti-TNF--APC Ab (BD
Biosciences) or 0.5 µg of anti-IgG-APC-negative control Ab,
diluted in 100 µl of permeabilization buffer for 60 min on ice. The
cells then were washed, resuspended in PBS, and analyzed on a
FACSCalibur flow cytometer (BD Biosciences). In experiments to analyze
the phenotype of the T cells that expressed cytokines, an additional
staining step was performed after the 6-h incubation step whereby the
cells were stained with one of the following Abs: anti-CD45RA-FITC
(BD Biosciences), anti-CD45RO-FITC (BD Biosciences),
anti-CD28-FITC (BD Biosciences), anti-CD62L-FITC (BD
Biosciences), or anti-CCR7-FITC (made in the laboratory of H.
Hasegawa; Ref. 30). Control experiments were performed to
show that the stimulation protocol did not alter expression of CD45RA,
CD45RO, CD28, or CCR7 on the CD8+ T cells.
Ex vivo CTL assays
CD8+ PBMC and other purified CD8+ subsets were assayed for Ag-specific cytolytic activity in standard 5-h chromium release assays. Target cells were either autologous or HLA-matched EBV-transformed B lymphoblastoid cell lines that were sensitized with an appropriate synthetic peptide at a final concentration of 5 µg/ml, or an equivalent dilution of DMSO, during labeling with 100 µCi of Na251CrO4 (Amersham Pharmacia Biotech).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We initially screened eight healthy EBV-seropositive donors with
the HLA-A2.1 and/or HLA-B8 allele by staining fresh PBMC preparations
with HLA class I/peptide tetramers refolded with one of three
A2.1-restricted epitopes (the lytic epitopes GLC and YVL and the latent
epitope CLG) or one of three B8-restricted epitopes (the lytic epitope
RAK and the latent epitopes FLR and QAK). To explore whether these
freshly isolated cells might display immediate cytotoxic function, we
enriched CD8+ T cells from PBMCs by negative
selection for CD19+, CD14+,
CD16+, CD4+, and
+ populations and then assayed these
CD8-enriched effectors on autologous target cells loaded with the
relevant epitope peptide. Fig. 1
A shows representative
results from such assays involving three different donors. Clear
evidence of epitope-specific lysis was detectable, whether using
CD8+ effectors containing 1–3% cells staining
with the A2/GLC or B8/RAK tetramer or using CD8+
effectors where the frequency of B8/FLR-staining cells was only 0.2%.
Similar results were obtained in assays involving all eight donors
tested on the six different epitopes. In many cases, significant lysis
could be detected at E:T ratios as low as 0.2 tetramer-positive cells
per 1 target cell in the assay. The specificity of these results was
confirmed in parallel studies with EBV-seronegative donors carrying the
A2.1 or B8 allele. Staining assays on PBMCs from seronegative donors
never identified significant numbers of CD8+ T
cells binding the relevant tetramers, and enriched
CD8+ preparations from these donors never gave
lysis above background when assayed on epitope-labeled targets.
Representative results from control donors, again looking for evidence
of A2.1/GLC, B8/RAK or B8/FLR reactivities by tetramer staining and by
ex vivo cytotoxicity assays, are shown in Fig. 1B.
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release; and 2) intracellular staining for IFN-
and of TNF- production within 6 h of peptide stimulation. With
the same set of EBV-epitopes, both assays clearly showed specific
responses that were restricted to EBV-seropositive donors with the
appropriate A2.1 or B8 allele (Ref. 21 and data not
shown). Phenotype of EBV epitope-specific memory populations
In the next series of experiments, we analyzed the phenotype of
epitope-specific memory cells by staining PBMCs from the above
EBV-seropositive donors with a PE-labeled tetramer, a tricolor-labeled
anti-CD8 mAb, and a FITC-labeled mAb against one of a series of
potentially relevant cell surface differentiation markers. Certain
markers were either uniformly absent or uniformly present on the
tetramer-positive population irrespective of the epitope being
analyzed. Fig. 2 presents relevant
cytometric staining profiles (gated on the CD8+ T
cells) for the same PBMC populations from EBV-seropositive donors JL,
DW, and YS used as the source of effectors for the ex vivo cytotoxicity
assays in Fig. 1
A. The tetramer-positive cells within these
populations were uniformly negative for the T cell activation marker
CD69 (Fig. 2, top) and for a second activation marker
CD38 (data not shown). The cells also were negative for CD56 (Fig. 2, middle), a marker that is usually associated with NK cells
but that also has been proposed as identifying a
CD8+ T cell subset with direct cytotoxic function
in assays of anti-CD3-redirected cytotoxicity (31).
The same tetramer-positive cells were almost entirely positive for
CD27, a marker the loss of which has been proposed to mark the
acquisition of effector function in redirected killing assays (7, 8, 10).
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shows representative
results from one of the eight EBV-seropositive donors analyzed, DW.
This donor is HLA-A2.1, B8-positive and so was screened by using two
A2.1-restricted lytic cycle epitopes, GLC and YVL, the B8-restricted
lytic cycle epitope RAK, and two B8-restricted latent cycle epitopes,
FLR and QAK. The lytic epitope-specific populations, which accounted
for 1% (GLC) and 3% (RAK) of circulating CD8+ T
cells, were clearly heterogeneous in terms of CD45 isoforms. In both
cases, some 30–35% of the tetramer-staining cells were
CD45RAhigh, and some 65–70% were
CD45RO+. In earlier work, double staining for
CD45RA and RO has clearly shown an inverse relationship between cell
surface expression levels for the two markers both within the
CD8+ population as a whole (32) and
within tetramer-positive populations (9, 32). Therefore,
we interpret the illustrated data for GLC- and RAK-specific memory in
donor DW as denoting the existence of distinct
CD45RAhighRO- and
CD45RA-ROhigh populations,
with a CD45RAlowROlow
intermediate population. The YVL lytic epitope-specific population
followed the same distribution pattern as these two epitope
specificities (data not shown). Interestingly, the smaller numbers of
CD8+ T cells in donor DW staining with the FLR-
and QAK-latent epitope-tetramers did not show this heterogeneity. These
memory populations were almost uniformly
CD45ROhigh, with little or no cells of a
CD45RAhighRO- phenotype
detected.
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Table I presents the overall data from
the eight EBV-seropositive donors analyzed. The results, presented so
as to distinguish between lytic and latent epitope-specific
populations, make clear that the trends illustrated in Fig. 3 are
consistent across donors. Thus, the latent epitope-specific response
was strongly polarized toward a CD45RA-,
CD45RO+, CD28+ phenotype.
In contrast CD8+ memory to lytic epitopes showed
a significant degree of CD45RO loss and acquisition of a
CD45RAhigh phenotype and a significant percentage
of cells that had lost CD28. Interestingly, the extent of CD45RA
positivity and CD28 negativity among lytic epitope-specific T cells
differed significantly between individual donors but was similar when
responses to different lytic epitopes were compared within an
individual. In particular, we noted that where detectable in memory,
the YVL response phenotype resembled that of the GLC response despite
differences in the size of the two populations. We also noted that
latent epitope-specific populations tended to show higher proportions
of CCR7+ and of CD62L+
cells than did lytic epitope populations.
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We then sought to analyze the relationship between memory T cell
phenotype and the epitope-specific cytotoxic activity seen earlier in
CD8-enriched effector populations. For this purpose, PBMCs were first
CD8-enriched as before with CD19, CD14, CD16, CD4, and counter
selection, and then aliquots further depleted either of
CD45RO+ cells or of CD45RA+
cells.
Fig. 4 shows representative flow
cytometric profiles of the three effector populations produced in one
of several experiments of this kind. Here, the CD8-enriched preparation
from the B8-positive donor CW contained a population of B8/RAK
tetramer-staining cells that was 25% CD45RA+ and
82% CD45RO+. Depletion of
CD45RO+ cells gave a preparation in which the
tetramer-positive cells were 100% CD45RA+, of
which the great majority were CD45RAhigh and
CD45RO- but a small minority (<10%) were
CD45ROlow. Likewise, depletion of
CD45RA+ cells gave a preparation in which the
tetramer-positive cells were 100% CD45RO+ of
which the great majority were CD45ROhigh and
CD45RA-, but with <10% showing low but
detectable CD45RA expression. Epitope-specific lysis was observed with
the CD8-enriched, CD45RA-enriched, and CD45RO-enriched effector cell
preparations. Similar results were observed in additional experiments
of this kind screening for GLC-specific lysis by total CD8,
CD45RO-depleted, and CD45RA-depleted effector preparations from the
A2.1-positive donor DCC (data not shown). Throughout these experiments,
we again usually detected significant epitope-specific lysis (>15%
51Cr release above low background levels) at
tetramer-positive E:T ratios down to 0.25:1. These studies clearly show
that when CD8+ effector function is measured
against the cognate target in ex vivo assays, cytolysis is not
restricted to the
CD45RAhighRO- fraction but
is present also in the CD45RO+ population. This
could be confirmed in several assays on latent epitope-loaded target
cells with CD8-enriched effector preparations where relevant
tetramer-staining population was naturally polarized to be 100%
CD45RO+RA-; significant
levels of killing were reproducibly observed (e.g., Fig. 1A,
B8/FLR-specific killing by effectors from B8-positive donor
YS).
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A shows the results of one
such experiment with effector populations from donor DW. Here, B8/RAK
tetramer-staining cells in the CD8+ pool were
35% CD28+ and 65% CD28-.
Depletion of a CD8-enriched preparation for CD28+
cells produced an effector population in which >93% tetramer-positive
cells were CD28-; these effectors clearly
mediated significant levels of RAK-specific lysis. The converse
experiment with CD28-enriched preparations was not possible because 1)
there is no counter marker allowing CD28+ cells
to be enriched by negative selection; and 2) in preliminary
experiments, we noticed that treatment with anti-CD28 increased
levels of killing by PBMC effectors, strongly implying that CD28
ligation could amplify effector function. Therefore, we again took
advantage of the fact that many latent epitope-specific memory
populations are naturally polarized toward CD28 positivity. Fig. 5B shows the results of an ex vivo assay with CD8-enriched
effectors from donor CW, in which 93% of the B8/FLR tetramer-staining
cells are CD28+. Strong lysis of FLR-loaded
target cells was observed. These findings strongly suggest that both
CD28+ and CD28- components
of EBV-specific CD8 memory have immediate epitope-specific cytolytic
capacity.
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Memory CD8+ T cell phenotype and cytokine release
In parallel experiments, we studied the relationship between
memory cell phenotype and the ability to secrete cytokines in response
to epitope-peptide stimulation. Initial studies with the ELISPOT assay
as a measure of overnight IFN- release gave a pattern of results
that exactly mirrored the cytotoxicity assays, positive responses being
seen in CD45RO-enriched, CD45RA-enriched, CD28+,
CD28-, CD62L- and
CCR7- populations (data not shown). We
subsequently used immunofluorescence assays in which PBMCs were first
stained with the tetramer, then exposed to the epitope peptide and
cultured for 6 h (in the presence of brefeldin A to prevent
cytokine release), stained for CD8 and the additional surface marker of
choice, and then permeabilized and stained for intracytoplasmic IFN-
or TNF-. This allowed the cytokine response to epitope peptide
stimulation to be visualized within tetramer-positive cells of defined
phenotype.
In the donors studied we found that between 30 and 90% of the
GLC-specific T cells expressed IFN- after stimulation in vitro. Fig. 6 displays the results obtained from a
representative HLA-A2-positive donor whose GLC epitope-specific memory
population was heterogeneous for CD45RO, CD45RA, and CD28 expression.
Freshly isolated PBMCs from this donor were analyzed for IFN-
production after 6 h either in culture medium as a control or in
presence of the GLC epitope peptide. The flow cytometric profiles are
obtained by gating on the tetramer-positive, CD8+
cells in the PBMC population and plotting IFN- staining
(x-axis) against expression of the phenotypic marker of
choice (y-axis). As can be seen from the three profiles
shown for unstimulated cells, there were very few tetramer-positive
cells spontaneously producing the cytokine. Fig. 6
(top) shows the response in relation to CD45RO
status. Both the CD45RO+ fraction and the
CD45RO- fraction contained responsive cells
after peptide stimulation, such that slightly below half of the cells
in each fraction became positive for cytoplasmic IFN-. Likewise,
responses were observed in CD45RA+ and CD45RA-
fractions (Fig. 6, middle) and in CD28+ and
CD28- fractions (Fig. 6, bottom) in parallel
assays. A similar pattern of results was obtained in assays on
B8-positive donors with the B8-restricted RAK lytic cycle epitope and
also in assays that used TNF- instead of IFN- as the index
cytokine (data not shown). Note that in such assays the overall profile
of CD45RO, CD45RA, and CD28 staining among tetramer-positive cells was
not altered as a result of the peptide stimulation.
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shows
representative data from an experiment that again focused on the
GLC-specific memory population of an A2-positive donor. When analyzed
with respect to CD62L (Fig. 7, top), the IFN- response
was almost entirely restricted to those tetramer-positive cells that
were CD62L-. However, in this type of
experiment, exposure to the epitope peptide reproducibly induced a
shift in the distribution of tetramer-positive cells so that
poststimulation more cells scored as CD62L-.
Therefore, although much of the cytokine response to peptide
stimulation appears to be mediated by the original
CD62L- fraction, it is possible that some
CD62L+ cells have also responded but have
simultaneously down-regulated the CD62L marker. By contrast, peptide
stimulation does not alter CCR7 distribution within the
tetramer-positive population and, when the same responder population as
above was analyzed for CCR7 and intracytoplasmic IFN- (Fig. 7, bottom), the cytokine response was clearly restricted to
CCR7- cells only. We observed this result in
several assays of this kind, although it should be noted that the
response never involved the entire CCR7-
population; there was always a small minority of
CCR7- cells that failed to respond. Similar
results are responsiveness in relation to CD62L and CCR7 were obtained
measuring the IFN- response to a latent cycle epitope FLR (data not
shown).
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(left) shows the results of four-color staining of
PBMCs from an A2+, B8+
donor with the PE-labeled B8/RAK tetramer, PerCP-labeled anti-CD8,
FITC-labeled anti-CCR7, and an APC-labeled mAb against the fourth
marker of choice. The first profile, gated only on
CD8+ cells, shows tetramer staining in relation
to CCR7 status. Note that (as was also apparent in Fig. 3) those
tetramer-positive cells that are CCR7+ show
levels of CCR7 staining significantly lower than shown by the
CD8+ CCR7+ population as a
whole (see Discussion). The other profiles in Fig. 8
(left), gated on tetramer-positive
CD8+ cells, show that most if not all of the
CCR7+ fraction are
CD45ROhigh, CD28+ and
CD62L+. By contrast, the
CCR7- fraction of B8/RAK tetramer-positive cells
are heterogeneous for all three of the above markers and contained
significant numbers of cells that are CD45RO-,
CD28- and CD62L-. Fig. 8
(right) shows the corresponding data for CD8 memory to a
latent cycle epitope B8/FLR, in the same donor. Again, the
tetramer-positive population could be divided into low
CCR7+ and CCR7- fractions.
In this case, where epitope-specific memory is polarized toward a
CD45ROhigh CD28+ phenotype,
then not surprisingly, both CCR7+ and
CCR7- fractions are uniformly positive for both
the CD45RO and CD28 markers. However, latent epitope-specific memory is
known to be heterogeneous in terms of CD62L status. Here, just as seen
earlier for the lytic epitope response, the CCR7+
fraction of FLR-specific memory is uniformly
CD62L+ whereas the CCR7-
fraction is composed of CD62L+ and
CD62L- cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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release (21). Our aim
was to readdress the much discussed relationship between
CD8+ T cell phenotype, type I cytokine
production, and cytotoxic capacity (7, 8, 10, 11, 12, 31) with
assays specific for the cognate epitope rather than the nonspecific
assays (PMA/ionomycin stimulation and anti-CD3 redirected
cytotoxicity respectively) that had been used to date. As others have
reported (22, 33), we found that staining for the
tetramer, CD8, and a third surface marker of choice could be combined
with intracellular staining for epitope-induced IFN- or TNF-
release, allowing the phenotype of cytokine-producing cells to be
analyzed (Figs. 6 and 7). We also found that EBV-specific
CD8+ T cells in the blood of healthy virus
carriers were capable of lysing epitope-loaded target cells at
significant levels in standard 5-h cytotoxicity assays, providing
another indicator of specific effector function (Fig. 1). Although EBV
epitope-specific lysis has been detected in ex vivo assays using
primary effector populations from IM patients (19, 20), we
had not expected the much smaller numbers of epitope-specific
CD8+ T cells in healthy donors to mediate
detectable levels of lysis. However, the result was observed on
numerous occasions at tetramer-positive target cell ratios as low as
0.2:1. Direct comparisons of the efficiency of EBV-specific lysis by
primary vs memory CD8+ cells at equivalent E:T
ratios remain to be conducted. It may be that the levels of killing
seen to date in ex vivo assays with T cells from IM patients are
underestimates of the true lytic potential because the assays have been
conducted only with cryopreserved effectors and the activated T cells
present in IM are highly susceptible to apoptosis during such
manipulations (34).
The present results make clear that these two epitope-specific rapid
effector functions, IFN-/TNF- synthesis and target cell lysis,
are present within CD8+ populations in which the
tetramer-positive cells are uniformly negative for conventional markers
of activation such as CD69 or CD38 (Fig. 2 and data not shown); they
are also negative for the cell cycle marker Ki67 (9). This
contrasts with the situation in IM where the primary EBV-specific
CD8+ T cells are actively cycling and express
activation markers (3, 4, 5). Therefore, it seems unlikely
that the functions being detected in the CD8+
pool of virus carriers reflect the reactivation of a small fraction of
the circulating EBV-specific memory pool to a highly activated
lymphoblastoid state akin to that shown by primary
CD8+ effectors in IM blood.
Our results also show that epitope-specific cytolytic function is not
restricted to the particular CD8+ subpopulations
that have been proposed as "effectors" on the basis of their
capacity for anti-CD3 redirected killing of Fc receptor-positive
targets. Such "effectors" have been variously identified with the
CD45RAhigh CD11ahigh
(8), CD45RA+
CD27- (7, 10),
CD28- (11, 12), and
CD56+ HLA-DR- (13, 31) subpopulations of circulating CD8+ T
cells. However, we detected epitope-specific lysis within both the
CD45RA+ and CD45RO+
subpopulations, and within the CD28+ and
CD28- subpopulations; furthermore, lysis was
regularly observed despite the fact that the tetramer-positive
component of the CD8+ T cell pool was uniformly
CD27+ and CD56- (Figs. 1, 2, 4, and 5).
Wherever the cytokine synthesis and cytotoxicity assays could be
applied to the same subpopulation of CD8+ T
cells, concordant results were observed. Hence, in the experiment
described in Fig. 6 we found that a significant proportion (30–50%)
of tetramer-positive cells responded to epitope stimulation by IFN-
synthesis whether these cells lay within the
CD45RA+, CD45RO+,
CD28+, or CD28-
subpopulation (Fig. 6). These findings are in line with those of an
earlier report focussing entirely the CD8+ memory
response to the A2-restricted lytic cycle epitope GLC
(33). That report also analyzed the IFN- response in
relation to CD62L status and found that all positive responses appeared
to map within the CD62L- fraction. Essentially
similar results were observed in the present work; however, they need
to be interpreted with caution because, as we show, some responses may
be mediated by cells that originally lie within the
CD62L+ fraction but that down-regulate the marker
during the assay period. More importantly, we showed that CCR7 status
was a better discriminator of competence in the cytokine assay. Thus,
there were essentially no IFN--responsive cells within the
CCR7+ fraction of the tetramer-staining
population, whereas the majority of (but not all)
CCR7- cells were responsive (Fig. 7).
These findings strengthen the view that during
CD8+ T cell differentiation, loss of CCR7
(subsequently followed by loss of CD62L) denotes the transition from a
"central memory" compartment of cells that naturally home to lymph
nodes and lack immediate effector function to an "effector memory"
compartment of cells that can migrate to tissues and do have immediate
effector function (14). We also noted that CCR7 levels in
tetramer-positive CCR7+ cells were always lower
than on the CD8+ CCR7+
population as a whole. Because the latter population is predominantly
made up of naive CD8+ T cells, recognized by their
CD45RAhigh LFA1low phenotype
(9, 14), we postulate that the transition from naive to
"central memory" compartments is associated with a reduction in
CCR7 surface levels. All EBV epitope-specific memory cells identifiable
by tetramer staining contained both CCR7+ and
CCR7- fractions; we found that the
CCR7+ fraction of this memory population is
uniformly CD45RO+, CD28+,
and CD62L+, whereas the
CCR7- fraction can be heterogeneous for all
three markers (Fig. 8). This is consistent with a view of
CD8+ T cell differentiation where, after loss of
CCR7, Ag-experienced cells can lose CD62L and/or CD28, and
(irrespective of CD62L or CD28 status) may revert to become
CD45RA+ (7, 9, 12). However, based
on the assays used in the present work, there is as yet no clear
functional difference between these phenotypically distinct subsets of
the effector memory (CCR7-) pool.
Finally, the present study highlights a hitherto unnoticed feature of
EBV-specific CD8+ memory that may help to explain
the circumstances under which heterogeneity is generated in "effector
memory" populations. There is a clear relationship between the
phenotype of CD8 memory and the identity of the EBV epitope. Thus,
latent epitope-specific T cells in the blood of healthy virus carriers
are strongly polarized toward a CD45ROhigh
CD45RA-, CD28+ phenotype.
Such findings are consistent both with early work showing that
EBV-specific memory T cells responsive to LCL stimulation in vitro
(i.e., latent Ag-specific reactivities) were concentrated within the
CD45ROhigh subpopulation (35), and
also with more recent studies that used TCR rearrangement to locate
FLR-specific memory clonotypes in the CD45RO+ but
not in the CD45RA- fraction of circulating
CD8+ T cells (36). Others also have
used EBV-latent epitope peptides as inducers of IFN- release and
have found responses only within the CD28+
CD8+ subset (37). By contrast, a
significant proportion (2–66%, mean 25%) of lytic epitope-specific T
cells in the donors studied here had a CD45RA+
phenotype, and many had also lost the CD28 marker (Fig. 3 and
Table I).
This difference is interesting in view of what is known about the
magnitude of the relevant responses. Although lytic epitope-specific
reactivities in CD8+ memory usually outnumber
those against latent epitopes, the contrast is much more dramatic
during primary infection. There the expansion of lytic epitope
responses is at least 10-fold greater than that of latent responses
(3), and this is even true for the primary response to a
lytic cycle epitope such as YVL, of which the subsequent representation
in memory can be quite low (N. Annels and A. Hislop, manuscript in
preparation). Indeed, some clonotypes within the highly amplified
primary response to the lytic cycle epitope GLC can apparently be
driven to the point of clonal exhaustion (29), whereas
immunodominant clonotypes within the less highly amplified response to
the latent epitope FLR are not (38, 39). These differences
in the degree to which responses are amplified may reflect the higher
levels of Ag load produced by lytic EBV replication in vivo compared
with that present in latently infected cells. We suggest that this
amplification of the lytic Ag-induced response over time drives some
cells within the "effector memory" to differentiate further to a
CD45RAhigh and/or CD28-
phenotype. In this context, several studies of infection with human
CMV, a 
-herpesvirus that appears to elicit even higher levels of CD8
immunity than does EBV, report that CMV-specific memory populations are
predominantly CD45RA+,
CD28-, CD27- (32, 40, 41, 42). This appears to reflect an even greater level of
virus-driven CD8+ T cell differentiation in vivo
to a point where the CD27 marker also is lost. This might also explain
the finding that in HIV-positive patients, where EBV loads can become
very high, EBV-specific CD8 memory cells may themselves acquire a
CD27- phenotype (D. van Baarle, personal
communication). In the present work, it was interesting to note that
although there were differences between individuals in the
degree to which lytic epitope-specific effector memory populations
became CD45RAhigh and/or
CD28-, within any one individual all lytic
epitope-specific populations analyzed show similar degrees of
phenotypic progression. Further studies will be needed to determine
whether the prevailing level of chronic EBV replication occurring in
virus carriers influences the memory phenotype or whether these
differences between individuals are established early on in the
immediate aftermath of the primary infection.
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Acknowledgments |
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Footnotes |
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2 A.D.H., N.H.G., and M.F.C.C. contributed equally to this work.
3 Address correspondence and reprint requests to Dr. Alan B. Rickinson, Cancer Research Campaign Institute for Cancer Studies, University of Birmingham, Edgbaston, Birmingham, B15 2TT, U.K. E-mail address: williamsd{at}cancer.bham.ac.uk
4 Abbreviations used in this paper: IM, infectious mononucleosis; APC, allophycocyanin.
Received for publication April 12, 2001. Accepted for publication June 5, 2001.
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E. V. Ravkov, C. M. Myrick, and J. D. Altman Immediate Early Effector Functions of Virus-Specific CD8+CCR7+ Memory Cells in Humans Defined by HLA and CC Chemokine Ligand 19 Tetramers J. Immunol., March 1, 2003; 170(5): 2461 - 2468. [Abstract] [Full Text] [PDF]
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D. Zhang, P. Shankar, Z. Xu, B. Harnisch, G. Chen, C. Lange, S. J. Lee, H. Valdez, M. M. Lederman, and J. Lieberman Most antiviral CD8 T cells during chronic viral infection do not express high levels of perforin and are not directly cytotoxic Blood, January 1, 2003; 101(1): 226 - 235. [Abstract] [Full Text] [PDF]
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C.-L. Lin, W.-F. Lo, T.-H. Lee, Y. Ren, S.-L. Hwang, Y.-F. Cheng, C.-L. Chen, Y.-S. Chang, S. P. Lee, A. B. Rickinson, et al. Immunization with Epstein-Barr Virus (EBV) Peptide-pulsed Dendritic Cells Induces Functional CD8+ T-Cell Immunity and May Lead to Tumor Regression in Patients with EBV-positive Nasopharyngeal Carcinoma Cancer Res., December 1, 2002; 62(23): 6952 - 6958. [Abstract] [Full Text] [PDF]
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E. M. van Leeuwen, L. E. Gamadia, P. A. Baars, E. B. Remmerswaal, I. J. ten Berge, and R. A. van Lier Proliferation Requirements of Cytomegalovirus-Specific, Effector-Type Human CD8+ T Cells J. Immunol., November 15, 2002; 169(10): 5838 - 5843. [Abstract] [Full Text] [PDF]
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P. J. Dunne, J. M. Faint, N. H. Gudgeon, J. M. Fletcher, F. J. Plunkett, M. V. D. Soares, A. D. Hislop, N. E. Annels, A. B. Rickinson, M. Salmon, et al. Epstein-Barr virus-specific CD8+ T cells that re-express CD45RA are apoptosis-resistant memory cells that retain replicative potential Blood, July 18, 2002; 100(3): 933 - 940. [Abstract] [Full Text] [PDF]
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 P C L Beverley Immunology of vaccination Br. Med. Bull., July 1, 2002; 62(1): 15 - 28. [Abstract] [Full Text] [PDF]
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H. Tomiyama, T. Matsuda, and M. Takiguchi Differentiation of Human CD8+ T Cells from a Memory to Memory/Effector Phenotype J. Immunol., June 1, 2002; 168(11): 5538 - 5550. [Abstract] [Full Text] [PDF]
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M. D. Catalina, J. L. Sullivan, R. M. Brody, and K. Luzuriaga Phenotypic and Functional Heterogeneity of EBV Epitope-Specific CD8+ T Cells J. Immunol., April 15, 2002; 168(8): 4184 - 4191. [Abstract] [Full Text] [PDF]
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V. Appay, L. Papagno, C. A. Spina, P. Hansasuta, A. King, L. Jones, G. S. Ogg, S. Little, A. J. McMichael, D. D. Richman, et al. Dynamics of T Cell Responses in HIV Infection J. Immunol., April 1, 2002; 168(7): 3660 - 3666. [Abstract] [Full Text] [PDF]
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M. Suresh, J. K. Whitmire, L. E. Harrington, C. P. Larsen, T. C. Pearson, J. D. Altman, and R. Ahmed Role of CD28-B7 Interactions in Generation and Maintenance of CD8 T Cell Memory J. Immunol., November 15, 2001; 167(10): 5565 - 5573. [Abstract] [Full Text] [PDF]
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A. D. Hislop, N. E. Annels, N. H. Gudgeon, A. M. Leese, and A. B. Rickinson Epitope-specific Evolution of Human CD8+ T Cell Responses from Primary to Persistent Phases of Epstein-Barr Virus Infection J. Exp. Med., April 1, 2002; 195(7): 893 - 905. [Abstract] [Full Text] [PDF]
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