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The Edward Jenner Institute for Vaccine Research, Compton, Berkshire, United Kingdom
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Recent evidence suggests that generation of some CTLs requires CD40-CD40 ligand (CD40L)3 interactions (5, 6, 7). CD40L is a member of the TNF gene family and is expressed on mast cells, autoreactive B cells, eosinophils, and activated platelets and is up-regulated on activated CD4+ T cells. CD40, the receptor for CD40L, is expressed on a number of cell types including dendritic cells (DC), B cells, macrophages, and endothelial cells. CD40-CD40L interactions play a role in both humoral and cellular responses (for review, see Ref. 8) including isotype switching of B cells, generation of B cell memory, and activation and maturation of macrophages and DC. More recently, CD40-CD40L interactions have been shown to be critical for the generation of some helper-dependent CTLs (for review, see Ref. 9 ; Refs. 10, 11, 12, 13) though other CTLs can be induced by CD40-independent mechanisms (4, 14, 15). The CD40-dependent pathway is regulated not only by interactions of the CD8+ T cell with the Ag-specific CD4+ T helper cell via lymphokines, (14) but also by interactions of the helper cell with the Ag-bearing APC (5, 6, 7). The helper cell is involved in activating the APC, generating a "sensitized/modulated" APC able to prime naive CD8+ T cells to become CTLs. Interactions between CD40 on the APC and CD40L on the helper cell are critical for a dominant component of this process (5, 6, 7). Triggering of CD40 by agonistic anti-CD40 Ab (aCD40) or incubation with Ag-specific CD4+ T helper cells can modulate DC to drive naive CD8+ T cells, while ligation of CD40 on DC restores CTL activity in CD4+ depleted mice. In addition, mice treated with anti-CD40L Ab had impaired induction of Ag-specific CTLs (16). These data indicate that CD40-CD40L interactions play a key role in modulating APC function so that these cells can prime CD8+ T cells in vivo.
The precise mechanism by which ligation of CD40 on DC is able to
provide the necessary signal(s) to empower a DC to generate CTLs is not
fully understood. Maturation effects of CD40 triggering of DC have been
described including up-regulation of costimulatory molecules such as
CD80, CD86, and CD54 (17, 18, 19) and secretion of cytokines
such as IL-12, TNF-
, and IL-8 (17, 18, 20). However, it
is not clear whether these factors are responsible for the capability
of the DC to drive an Ag-specific CD8+ T cell response.
To investigate the mechanism of CD40-dependent generation of CTLs by DC, we have developed an in vitro system in which bone marrow-derived DC can prime CD8+ cells from naive (i.e., unprimed) C57BL mice to the known CD8+ epitope SIINFEKL from OVA. We compare the ability of two modulators, aCD40 or LPS, that are known to mature and activate DC, to generate SIINFEKL-specific CTLs. We show that a high concentration of LPS is not essential for the generation of CD40-dependent CTLs. However, we are unable to rule out the possibility that very low concentrations of LPS (<20 pg/ml) may act in synergy with CD40 ligation. We also show that the precise mechanism by which CD40-CD40L interaction provides help for the generation of CTLs is likely to be an as-yet-undefined factor.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Female 6- to10-wk-old C57BL/6 mice were purchased from Charles River Breeding Laboratories (Wilmington, MA) through the Institute for Animal Health (Compton, Berkshire, U.K.). Transgenic OT-I mice expressing a TCR specific for the peptide SIINFEKL from OVA presented in association with H-2Kb were generously donated by Dr. N. Kukutsch (Nuffield Department of Surgery, John Radcliffe Hospital, Oxford, U.K.) and Dr. M. Merkenschlager (Medical Research Council Clinical Sciences Center, Hammersmith Hospital, London, U.K.). The EL4 thymoma and the MC57G fibroblastoma cell lines (both H-2b) were maintained in RPMI supplemented with 10% FCS, 2 mM of L-glutamine, and combined antibiotics.
Determination of surface Ag expression by cytofluorometry
Expression of cell surface Ags on DC and T cells was confirmed
by staining with mAbs and flow cytometric analysis using a FACSCaliber
immunocytometry system (BD Biosciences, Oxford, U.K.). DC were stained
with Abs from BD PharMingen (San Diego, CA), unless stated
otherwise, against CD11c (PE-conjugated clone HL3), and MHC I
(FITC-conjugated clone AF6-88.5), MHC II (FITC-conjugated clone 2G9),
CD80 (FITC-conjugated clone 16-10A1), CD86 (FITC-conjugated clone GL1),
CD40 (FITC-conjugated clone HM40-3), CD54 (FITC-conjugated clone 3E2)
and Ly-6G (Gr-1; FITC-conjugated clone RB6-8C5), and CD19
(FITC-conjugated clone 1D3), F4/80 (FITC-conjugated clone CI:A3-1;
Serotec, Oxford, U.K.). DC were also stained for MHC I
(H-2Kb)-SIINFEKL presentation using clone 25-D1
(21) which was a kind gift from Dr. C. Reis e Sousa,
Imperial Cancer Research Fund (London, U.K.). Purified 25-D1 was
biotinylated using a kit from Pierce according to the manufacturer’s
instructions. This Ab was detected using streptavidin-APC (BD
PharMingen) and staining was performed in the presence of rat
anti-mouse Fc
RIII/II (clone 2.4G2) to block FcR binding. T
cells were stained for expression of CD69 (FITC-conjugated clone
H1-2F3, Harlan SeraLab, Loughborough, U.K.),
CD4+ T cells (L3T4; FITC-conjugated RM4-4 or
Cy-chrome-conjugated clone H129.19), CD8a (Ly-2; PE-conjugated clone
53-6.7) and CD8b (Ly-3.2; PE-conjugated clone 53-5.8). Appropriate
isotype controls were obtained from BD PharMingen.
Generation of DC from bone marrow precursors
DC were generated from bone marrow precursors by incubation with GM-CSF (10 ng/ml; R&D Systems, Abingdon, U.K.) as previously described (22). Briefly, a single cell suspension was made from bone marrow removed from the femurs and tibias of young female mice and cultured in IMDM with glutamax supplemented with 10% FCS and 5 x 10-5 M 2-ME and GM-CSF (10 ng/ml). After 2–3 days incubation, medium plus nonadherent cells were removed and fresh medium with GM-CSF was added. At day 6, nonadherent cells obtained from these cultures were considered to be immature bone marrow-derived DC. They were removed, washed, and placed in 6-well plates for 12–72 h in the presence of fresh GM-CSF and various modulators. DC phenotype was confirmed by staining with Abs and subsequent flow cytometric analysis. DC preparations were purified by sorting CD11c+ cells (clone N418) using maxi-MACS columns (Miltenyi Biotec, Bergisch Gladbach, Germany). Typically cultures were >98% positive for CD11c.
Modulators of bone marrow-derived DC
The following modulators were added to DC cultures for 12–72 h: aCD40 (2 µg/ml; clone IC10; R&D Systems; and clone FGK45 generously provided by A. Rolink, Basel Institute for Immunology, Basel, Switzerland); LPS (800 ng/ml; Sigma-Aldrich, St. Louis, MO); OVA (4 mg/ml; grade VII, Sigma-Aldrich); and SIINFEKL (1 µM; Sigma-Genosys, Cambridge, U.K.). DC were washed twice before additional experiments were performed. Before use in functional assays, DC were irradiated (2500 rad) and washed twice.
Enrichment of CD8+ and CD4+ T cells
Single-cell suspensions were prepared from spleens of naive C57BL/6 mice. RBC were depleted by lysis in Tris-buffered 0.83% (w/v) ammonium chloride and debris was removed by centrifugation. The splenocyte suspension was initially depleted of CD19+ (clone 1D3) cells using maxi-MACS beads and columns (Miltenyi Biotec, Germany). CD4+ T cells (clone GK1.5) were sorted from the remaining CD19- cell suspension also using the Miltenyi system resulting in a cell population that was 96% pure for CD4+ T cells. CD8+ T cells were then sorted from the CD19+- and CD4+-depleted splenocytes using an anti-mouse CD8+/Ly-2 mAb (clone 53-6.7), yielding populations of 98–99% purity. Some cell preparations were passed through a second MACS column for increased purity.
Activation and proliferation assays of CD8+ and CD4+ T cells
CD4+ T cells or CD8+ T cells were purified as described above and set up in 96-well U-bottom plates (1 x 105/well). DC (1 x 104/well) treated with a combination of Ag, aCD40, and LPS were placed in triplicate into wells with or without T cells. ConA (5 µg/ml; Sigma-Aldrich) was added to some wells of T cells (i.e., without DC). Cells were double-stained for CD69 and either CD4+or CD8+ 1–18 h later. Proliferation was assayed by adding [3H]thymidine (1 µCi/well: Amersham Pharmacia Biotech, Buckinghamshire, U.K.) to individual plates at 24, 48, 72, 96, and 120 h after the DC were added to the T cells. Cells were harvested 16 h after the addition of [3H]thymidine using a TOMTEC cell harvester (Tomtec, Orange, CT) and [3H]thymidine incorporation into DNA was measured as counts per min on a Wallac 1450 MicroBeta Trilux beta counter (Gaithersburg, MD).
Generation of CTL responses
Spleen cells or purified CD8+ T cells (2 x 107) were cultured for 5–6 days in the presence of DC (1–2 x 106) in upright T25 flasks in 5 ml of IMDM supplemented with 10% FCS and 5 x 10-5 M 2-ME. Additional medium (up to 2 ml) was sometimes added on day 3–4 of the culture period. Splenocytes from OT-I mice were used to generate CTLs in a similar manner. IL-2 (2 ng/ml; R&D Systems) was added to cultures where stated.
Measurement of cytolytic activity
Cytolytic activity was measured using 51Cr sodium chromate-labeled MC57G or EL4 cells with or without the addition of the OVA-derived peptide, SIINFEKL (1 µM). Effector cells were washed twice and serial dilutions were incubated in 96-well round bottom microtiter plates with 1 x 104 target cells for 6 h at 37°C. Maximal release was induced by the addition of 2.5% Triton X-100 to wells of target cells without effectors. The percentage of specific lysis was calculated as follows: 100 x [(cpm experimental release) - (cpm spontaneous release)]/[(cpm maximal release) - (cpm spontaneous release)]. Assays were performed in IMDM with 5% FCS, 5 x 10-5 M 2-ME and 7 mM of HEPES.
RNase Protection Assay (RPA)
Total cellular RNA was extracted from DC with TRIzol Reagent
(Sigma-Aldrich) according to the manufacturer’s protocol. Cytokine
transcript levels were determined by RPA using RiboQuant kits (BD
PharMingen) with the following probe sets: mCK-1b, mCK-2b, mCK-3b,
mCK-4, and mCK-5. Briefly, total cellular RNA (2 µg) was hybridized
to a [-32P]UTP-labeled antisense
riboprobe for 16 h at 56°C. After digestion of single-stranded
RNA by RNase A and proteins by proteinase K, protected RNA fragments
were separated on a denaturing sequencing gel, followed by
phosphorimaging. As internal controls for the amount of RNA loaded onto
the gel, each riboprobe set contained antisense
[32P]-L32 and -GAPHD.
Measurement of endotoxin and IL-12p70
Endotoxin was measured by the kinetic Limulus
amebocyte lysate assay (Charles River Endosafe, Margate, Kent,
U.K.) in an ELISA plate reader (Spectra MAX 340, Molecular
Devices, Wokingham, U.K.). The detection limit of the assay was
2 pg/ml of endotoxin. The levels of endotoxin were determined for
the medium, FCS, GM-CSF, PBS, aCD40, SIINFEKL, and OVA and are recorded
as the concentration present in the medium used for maturing the DC or
generating CTL. IL-12 was measured in 0–72 h supernatants of DC
cultures (5 x 106 DC/4 ml). DC were
untreated or stimulated with aCD40 or LPS as described above. The IL-12
p70 heterodimer was detected by a two-site ELISA using a murine IL-12
p70 immunoassay kit from BioSource International (Camarillo, CA)
according to the manufacturer’s instructions. The detection limit of
the assay was 4 pg/ml IL-12 p70.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We chose to work with CD8+ T cells from the spleens of naive or unprimed C57BL/6 mice representing a repertoire of cells with physiological Ag-specific precursor frequencies and affinities. First, it was necessary to establish that these CD8+ T cells were not activated and did not respond to DC polyclonally. The purified CD8+ T cells cultured alone did not express the early activation marker CD69 after 18 h in culture (data not shown). CD69 expression increased slightly to 1.8% when the T cells were incubated with untreated DC and to 3.5–5.3% when incubated with DC pulsed with combinations of aCD40, LPS, and OVA for up to 18 h (data not shown). In contrast, 78.1% of the CD8+ T cells expressed CD69 when they were incubated with ConA. CD69 expression was time-dependent, peaking at 5–6 h after stimulation and remaining constant for up to 18 h. These results showed that the CD8+ T cells were initially quiescent and did not respond to the DC polyclonally. It remains to be determined whether the small number of cells activated by the modulated DC represent a subset of T cells preactivated in vivo.
CD40-activated DC pulsed with Ag are able to activate CD8+ T cells
We then examined the ability of DC to induce proliferative
responses of either CD4+ or
CD8+ T cells. DC were pulsed with various
combinations of aCD40, LPS, and OVA and incubated with either
CD4+ or CD8+ T cells for
2–6 days (Fig. 1
). Although all DC
stimulated some proliferation of CD8+ T cells in
the first three days of culture, only DC modulated with OVAaCD40 and
OVAaCD40LPS induced a strong response, peaking at day 5 (Fig. 1A). Proliferation of the T cells remained low for DC pulsed
with OVA and OVALPS while untreated DC and DC pulsed with aCD40 did not
sustain proliferation after day 3. DC pulsed with LPS had a similar
effect to untreated DC (data not shown). These data indicated that to
induce strong proliferative responses from CD8+ T
cells both OVA and aCD40 were required to activate DC, while high doses
of LPS did not synergize with OVA. In contrast,
CD4+ T cells proliferated in response to
stimulation by a variety of DC, although the responses were greatest
when DC were modulated with OVAaCD40 or OVAaCD40LPS (Fig. 1B). Only untreated DC did not induce a continued
proliferative response of CD4+ cells beyond day
3, whereas proliferation continued in cultures stimulated with DC
pulsed with combinations of aCD40, LPS, and OVA. DC pulsed with LPS,
aCD40, or OVA alone (data not shown) stimulated similar proliferative
responses to DC pulsed with OVALPS (Fig. 1B). DC pulsed with
OVAaCD40LPS induced the highest response, followed by DC pulsed with
OVAaCD40, OVALPS and OVA, aCD40, and LPS, in descending order.
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CD40-activated DC pulsed with Ag are able to generate CTLs
Although DC modulated with OVAaCD40 were able to induce
proliferation of CD8+ T cells, it was necessary
to show that these cells became Ag-specific CTL. Specific cytolytic
activity could only be detected when naive CD8+ T
cells were stimulated with DC modulated with OVA and aCD40 (Fig. 2, B–E). Neither untreated DC
nor DC pulsed with either aCD40 or OVA were able to generate CTLs from
unprimed CD8+ T cells. The addition of exogenous
IL-2 was not required to generate CTLs (Fig. 2, E–F). These
data showed that there was an absolute requirement for aCD40 triggering
of DC and Ag to generate DC able to prime CTLs from normal naive
polyclonal CD8+ T cells. However, modulation of
DC is not required to generate CTLs from CD8+ TCR
transgenic T cells (OT-I) specific to SIINFEKL. DC pulsed with OVA
alone were able to stimulate cells to become cytolytic (Fig. 2A).
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Although these experiments clearly show the ability of DC to
cross-prime and that CD40 triggering is involved in the generation of
CTLs, we wanted to establish the role of endotoxin. At the
concentration of OVA used in most experiments to pulse DC, the OVA
contributed 70 ng/ml endotoxin, thus indicating that DC pulsed with OVA
were in fact modulated with OVA and LPS. However, the endotoxin
concentration of SIINFEKL peptide used to pulse APC was below the limit
of detection (<2 pg/ml). DC pulsed with peptide could only generate
CTL when they were also modulated with aCD40 and not with LPS (Fig. 3). Further investigation showed that
endotoxin was not detectable in the PBS and GM-CSF used, but that
endotoxin was present in the tissue culture medium (8.7 pg/ml), aCD40
(8.1 pg/ml), and FCS (3.3 pg/ml). These results indicated that DC were
possibly matured in the presence of a low concentration of endotoxin
(<20 pg/ml) and that unprimed CD8+ T cells were
stimulated by DC in the presence of endotoxin (12 pg/ml). Therefore, we
are unable to exclude the possibility that low levels of LPS synergize
with aCD40 to modulate DC to prime CTLs.
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We examined factors that may be influenced by CD40 triggering of
the DC compared with that of LPS modulation of the DC to define
mechanism(s) responsible for the ability of
aCD40-modulated DC to generate CTLs.
First, the expression of MHC I was examined (Table I; Fig. 4). DC modulated by aCD40 did not
up-regulate MHC I (H-2Kb) compared with untreated
DC or DC treated with LPS (Table I). We then investigated whether MHC
I-SIINFEKL presentation was up-regulated, using the mAb 25-D1
(21), which recognizes
SIINFEKL-H-2Kb complexes. DC loaded with SIINFEKL
showed high levels of staining with this Ab (Fig. 4). However, the
median fluorescence of DC pulsed with OVA is much lower and was not
altered when the DC were pulsed with OVA and aCD40. When expression of
MHC I-SIINFEKL on the surface of the DC was examined throughout the
time course of the coculture of the DC with CD8+
T cells, no difference was observed between DC pulsed with OVA and
aCD40 or OVA and LPS at any time point (data not shown). These data
indicate that an increase in the level of presentation of peptide does
not account for the ability of DC modulated by aCD40 to generate
CTLs.
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CD40 and LPS modulate surface expression of MHC II and costimulatory molecules
The expression of MHC II and costimulatory molecules of untreated
DC and DC pulsed with either aCD40 or LPS was examined (Table I
). CD40
triggering of DC increased MHC II, CD80, CD86, CD54, and CD40 as
reported previously (17, 18, 19), but DC modulated by LPS
showed higher levels of the same cell surface markers. It was found
that on all DC types, the expression of the costimulatory molecules
declined rapidly (within 24–48 h) during culture with the
CD8+ T cells (data not shown). Although DC pulsed
with LPS showed higher levels of expression of the costimulatory
molecules compared with DC modulated by aCD40, there was no significant
difference between the rate of decline of the expression of these
molecules on either type of DC. These results indicated that if any of
these cell surface markers played an essential role in the ability of
DC to induce CTL responses, DC modulated by LPS should be a more potent
inducer of CTLs than DC modulated by aCD40. As LPS alone was not able
to modulate DC to prime CTLs, we concluded that it was unlikely that
any of these molecules were the defining factor(s) for the
CTL-generating ability of DC modulated by aCD40. However, the increased
levels of MHC II, CD80, CD86, CD40, and CD54 may still be important for
activation of CD8+ T cells especially during the
first 24 h of culture with DC.
Cytokine profiles of DC modulated with LPS and aCD40
We finally examined the cytokine profiles of various DC by
performing RPAs. This allowed an investigation of RNA synthesized by DC
modulated by aCD40, LPS, OVA, OVAaCD40, and OVALPS for 18 h (Table II). DC pulsed with SIINFEKL or
SIINFEKLaCD40 showed similar cytokine expression patterns as
untreated DC and DC pulsed with aCD40, respectively (data not shown).
None of the DC preparations expressed RNA for IL-2, IL-3, IL-4, IL-7,
IL-9, IL-11, IL-13, G-CSF, GM-CSF, and lymphotactin (LT
) and
expressed barely detectable levels of IL-5, IL-18, IFN-, IFN-,
TGF2, TGF3, and eotaxin. Another group of cytokines was expressed
constitutively, although at different levels, regardless of the stimuli
added to the DC, including IL-15, TNF-, TNF, TGF1, and
migratory inhibitory factor. Only one protein, lymphotoxin,
which is involved in lymph node development and germinal center
formation, appeared to be down-regulated by DC pulsed with OVAaCD40.
Another group of proteins was up-regulated in the DC by the different
stimuli to varying degrees including IL-6, IL-12p35, IL-12p40, IL-1,
IL-1, IL-1R antagonist, M-CSF, LIF, stem cell factor, RANTES,
macrophage-inflammatory protein (MIP)-1, MIP-1, T cell
activation, gene 3, (IFN-inducible protein-10 and MIP-2. Some
modulators of the DC induced barely detectable levels of IL-10 and
monocyte chemotactic protein-1.
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). The levels of both RNA transcripts
did not increase further when aCD40 or OVA were added to the DC
modulated by LPS. However, in the presence of OVA, aCD40-modulated DC
expressed higher levels of both IL-12p35 and IL-12p40, although this
may be due to the presence of Ag as OVA alone also increased levels of
IL-12p35 and IL-12p40 compared with untreated DC. It is uncertain which
component(s) of OVA influenced IL-12 RNA synthesis, although it may be
due to the LPS contaminant.
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). Throughout the culture period between
20–72 h, functional IL-12p70 was detected in the supernatants from DC
modulated by aCD40. In contrast, IL-12p70 was detected in the
supernatants of DC modulated by LPS at 4 h of culture. The level
of IL-12p70 peaked at 4 h of culture, decreased at 8 h, then
gradually increased with time up to 72 h of culture. Only a
minimal level of IL-12p70 was detected in medium containing untreated
DC at 30–48 h of culture. These results clearly show that both LPS-
and aCD40-modulated DC secreted functional IL-12 but with different
kinetics.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our data show that DC are essential for the priming of CD8+ T cells and that the DC must be matured by modulatory factors to acquire the capacity to generate CTLs. The minimum requirement for maturation of DC to CD8+ T cells to become effective killer cells is CD40 triggering. When CTLs are generated by DC modulated with aCD40 and OVA, LPS may play a role as a high concentration of endotoxin (70 ng/ml) is present in the Ag. However, LPS is insufficient to generate CTLs as they were not generated by DC modulated with OVA and a high dose of LPS. It is also clear from the experiments performed with peptide that CD40 triggering of the DC can still drive a CTL response in the presence of a very low concentration of LPS (<20 pg/ml), whereas DC pulsed with peptide and a high concentration of LPS are unable to effectively generate CD8+ cytotoxic T cells. These data suggest that CD40 triggering is the key signal for maturation of DC to generate CTLs but do not eliminate the possibility that a very low concentration of LPS may act synergistically with aCD40 to modulate DC function.
We are as yet unable to explain the mechanism by which DC modulated with aCD40 acquire the capacity to prime CTLs at the level of Ag processing and expression of costimulatory molecules. First, there was no increase in the level of MHC I-peptide presented by DC as a result of CD40 triggering of DC pulsed with OVA. In contrast, presentation of peptide by MHC I was up-regulated when DC were pulsed with high concentrations of peptide (1 µM) and aCD40 but to the same extent as DC that were pulsed with peptide and LPS (data not shown). Nevertheless, only DC pulsed with SIINFEKLaCD40 induced CTL responses, not DC pulsed with SIINFEKLLPS, indicating that an increase in MHC I-peptide presentation is not responsible for the generation of CTLs. However, we do not know whether DC pulsed with Ag and aCD40 are able to process Ag at a faster rate than DC pulsed with Ag alone or Ag and LPS. In contrast, DC modulated by aCD40 do not maintain peptide presentation for longer periods compared with DC pulsed with Ag alone or Ag and LPS as MHC I-SIINFEKL presentation by the various DC was similar throughout the culture period (data not shown). Secondly, although an increase in the expression of MHC II and the costimulatory molecules CD80, CD86, CD54, and CD40 was observed on the surface of modulated DC, these factors were not unique to DC modulated by aCD40. In addition, the expression all costimulatory molecules on the DC rapidly declined during the culture period with CD8+ T cells, regardless of whether the DC were untreated or modulated with either aCD40 or LPS.
It has been previously reported that CD40 ligation of DC prolongs survival of this cell type (17, 25). We examined the viability of the various types of DC by staining with CD11c and the early apoptotic marker annexin V (data not shown). Our results show that aCD40-treated DC have enhanced survival (50–80%) when compared with untreated DC (20–50%) over the first 48 h of coculture with CD8+ T cells. Modulation of the DC by LPS decreases survival of the DC (10% or less) within 24 h of coculture with CD8+ T cells. DC pulsed with OVA, OVAaCD40, or OVALPS also showed decreased survival (10–30%) compared with untreated DC. These data show that the DC capable of stimulating CTLs, DCOVAaCD40, do not have a prolonged life when compared with DC that are unable to generate CTLs, DCOVALPS. After 96 h in culture, few viable DC could be recovered from cultures stimulated with any of the DC. Taken together, the poor survival of OVA-pulsed DC and the down-regulation of costimulatory molecules suggest that the outcome of CTL generation is determined by events that occur early in the coculture of the DC and CD8+ T cells.
We have not identified a unique cytokine or chemokine associated with
the DC able to generate CTLs. DC synthesized cytokine transcripts that
may be involved in activation of T cells such as IL-6 and IL-1,
although levels were equal to or greater in DC modulated with LPS when
compared with DC pulsed with aCD40. DC modulated by aCD40 did not
express higher levels of the inflammatory cytokine TNF- transcript
than untreated DC or DC pulsed with LPS, although up-regulation of
TNF- in CD40-activated dendritic Langerhans cells has previously
been shown (17). DC also synthesized transcripts for a
number of chemokines including MIP-1 and MIP-1, which are known
to induce preferentially the migration of CD8+ T
cells and CD4+ T cells, respectively
(26), but levels of the transcripts were somewhat higher
in DC modulated by LPS than DC modulated with aCD40. Together, these
results show that DC modulated with LPS or aCD40 share the capacity to
trigger DC maturation, as demonstrated by the up-regulation of MHC
molecules, costimulatory molecules, chemokines, and cytokines. However,
DC modulated by LPS are less effective at CTL generation.
IL-12 production by APC may be important for the development of Th1
responses and CTL generation as it can induce IFN- synthesis which
activates CTLs (for review, see Ref. 27 ; Refs.
28, 29, 30, 31). Both human and murine DC have been shown to
produce IL-12 in response to either microbial stimuli
(32, 33, 34) or T cell-dependent interactions (35, 18, 20) in which binding of CD40 by CD40L is critical (18, 20, 36). Although bone marrow-derived DC have been shown to secrete
IL-12p70 in response to microbial stimuli within 10 min to 24 h of
stimulation (33, 37), production of IL-12p70 by DC treated
with aCD40 has only been reported when the DC have been stimulated for
>36 h (20). Our study confirms that a principal
difference between DC modulated by aCD40 compared with LPS is the
timing of release of IL-12p70. DC modulated with LPS release IL-12p70
by 4 h in culture, 16 h earlier than DC modulated by aCD40.
This is also reflected in the analysis of RNA transcript expression. At
18 h after modulation of the DC, untreated DC and DC modulated by
aCD40 expressed low levels of IL-12p40 while IL-12p35 was not detected,
suggesting that these DC could not produce functional IL-12. However,
LPS modulation of the DC increased levels of both IL-12p35 and IL-12p40
indicating that these DC could potentially produce functional
IL-12.
Although it has been shown that IL-12 production may enhance CTL generation or increase perforin production (28, 29, 30, 31), it is likely that IL-12 does not play an essential role in the generation of cytotoxic T cells because IL-12 deficient mice can generate normal allogeneic CTL (38) and CD40-dependent mucosal CTL (38, 24). Our data support this view. Although both DC treated with LPS or aCD40 produce IL-12, only the latter can generate CTL in the absence of CD4+ T cells and neutralizing anti-IL-12 Ab is unable to prevent CTL induction in our system (data not shown). Nevertheless, appropriate levels and timing of IL-12 secretion, dependent on the stimulus used to mature the DC, may play a role together with other signals in CTL generation.
Our studies also demonstrate that the modulation required to allow the
DC to acquire the capacity to drive CTL responses is dependent on the
biological status of the CD8+ T cells. To
generate SIINFEKL-specific CTLs from unprimed normal
CD8+ T cells, it was necessary to modulate the DC
with aCD40. This was not the case when using CD8+
TCR transgenic T cells (OT-I), specific for SIINFEKL. Strong CTL
responses were obtained when DC were pulsed with either peptide or OVA
alone (Fig. 2A), indicating that the DC did not require
modulation by aCD40 or a high dose of LPS. In this experiment, the
frequency of precursor T cells and their affinity for MHC I-SIINFEKL
complex is much higher than in a normal repertoire of
CD8+ T cells. We did not investigate whether the
OT-I cells were naive or contained some cells that had been activated
in vivo and were in fact memory cells, able to respond faster to Ag or
requiring less DC-derived costimuli. In the MLC, it has been shown that
modulation of DC by either LPS (30 µg/ml) or aCD40 (10 µg/ml)
enhances the ability of DC to drive CD8+ T cells
(15). However, a high frequency of T cells from both the
naive and memory pools with high and low affinities for a number of
alloantigens respond in MLCs (15, 39). We have obtained
similar results in our in vitro system (data not shown). Thus, DC (from
BALB/c mice) modulated by either LPS or aCD40 are able to enhance the
generation of allospecific CTLs from unprimed
CD8+ T cells (from C57BL/6). In both the
transgenic and MLC systems, we have not determined whether the
CD8+ T cells themselves are in fact activating
the DC. The ability of CD8+ T cells to induce
activation of DC, even in the absence of direct T cell-DC contact has
been demonstrated (4). Together, the data indicate that
CD8+ T cell activation may be influenced by the
frequency of the precursor CD8+ T cell, the
affinity of the TCR for its Ag, and whether the responding
CD8+ T cell is a naive or memory cell. These
different T cells may require different maturation states of DC to
become activated and/or cytotoxic.
Not only does this work show that specific modulation of DC is necessary to stimulate unprimed CD8+ T cells, it also shows that different modulatory factors are required by DC to activate CD4+ T cells. For both CD8+ and CD4+ T cells, proliferation was greatest when DC were modulated with OVAaCD40 and OVAaCD40LPS. Although CD8+ T cells did not respond significantly to DC modulated by high concentrations of LPS, aCD40, or Ag alone, CD4+ T cells did. The addition of high doses of LPS to DC pulsed with OVAaCD40 increased proliferation of CD4+ T cells compared with DC modulated with OVAaCD40, while it had no further effect on the proliferation of CD8+ T cells stimulated by DC treated with OVAaCD40. Another difference between the two cell types is the longevity of the proliferative responses. CD8+ T cells ceased responding to stimulation at day 6 when CD4+ T cells were still proliferating. Similar data were obtained in earlier studies of MLC responses (40, 41). It was reasoned that as CTLs were generated, killing of DC by the freshly generated CTLs hindered continued proliferation of CD8+ T cells (40). In conclusion, less specific modulation of DC appeared to be required to activate CD4+ T cells compared with CD8+ T cells and the kinetics of activation of the two cell types are different. The effects of LPS modulation of DC on CD8+ T cell activation, if any, remain to be clarified (42).
In summary, while CD40 ligation of DC is essential to mature bone
marrow-derived DC to drive OVA-specific CTL responses, we are unable to
rule out a role for very low levels of microbial stimuli.
The factors that empower a CD40-modulated DC to drive cytotoxic
responses remain elusive. We were unable to account for these critical
factors by examining Ag processing, known costimulatory molecules, and
cytokine and chemokine secretion by these DC. It is possible that a
known molecule that we have not yet examined may be involved, or that
an effective and long-lasting CTL response may be due to the production
of a soluble factor by the activated T cell rather than the DC.
Recently, other receptors and their ligands have been identified
on T cells and DC such as TNF-related activation-induced
cytokine/receptor activator of NF-
(TRANCE/RANK)
(43, 44, 45), OX-40/OX-40 ligand (46, 47),
and 4-1BB/4-1BB ligand (Refs. 48, 49, 50, 51 ; for review,
see Ref. 52). It will be interesting to investigate
whether such molecules are involved in providing cooperative signals
for DC-T cell interactions to augment CD40-dependent generation of
CTLs.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Michelle Kelleher, The Edward Jenner Institute for Vaccine Research, Compton, Berkshire RG20 7NN, U.K. E-mail address: michelle.kelleher{at}jenner.ac.uk
3 Abbreviations used in this paper: CD40L, CD40 ligand; DC, dendritic cell; aCD40, anti-CD40 Ab; RPA, RNase protection assay; MIP, macrophage-inflammatory protein.
Received for publication May 23, 2001. Accepted for publication September 28, 2001.
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References |
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) or pulsed with aCD40 (
), OVA (x), OVAaCD40 (
),
OVALPS (
), OVAaCD40LPS (
). Data for DC pulsed with LPS is not
shown but is similar to that of DC pulsed with aCD40 for
CD4+ T cells and similar to that of untreated DC for
CD8+ T cells. Data for DC pulsed with aCD40 or OVA is not
shown for CD4+ T cells but is similar to that shown for DC
pulsed with OVALPS. Results shown are the mean thymidine
incorporation ± SD, in triplicate cultures, and are
representative of two experiments.
