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* Laboratory of Parasitology, Université Libre de Bruxelles, Belgium; and
Laboratory of Animal Physiology, Institut de Biologie et de Médecine Moléculaire, Université Libre de Bruxelles, Gosselies, Belgium
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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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The present work was undertaken to better understand the early steps of
the in vivo immune response to bacterial superantigens. These compounds
form a large group of proteins produced by several bacterial strains
(such as Staphylococcus aureus, Mycoplasma
arthritidis, and Yersinia pseudotuberculosis) that have
been recognized as pathogenic both in human and animals (reviewed in
Refs. 6, 7, 8). Purified bacterial superantigens share with
other microbial-derived compounds the ability to induce a potent
inflammatory response in vivo, causing toxic shock syndrome. Numerous
studies have shown that the immunostimulatory properties of these
compounds are related to their ability to interact simultaneously with
selected V
regions of the clonally distributed TCR and with a
conserved domain of the MHC class II (MHC II) molecule, causing the
immune stimulation of a large fraction of T cells in vivo. The presence
of additional, conserved receptors, distinct from TCRs or MHC II
molecules, able to interact with staphylococcal enterotoxin B (SEB) has
also been described (9, 10). In particular, a recent study
suggested that bacterial superantigens were able to induce an in vivo
acute inflammatory response (as judged by leukocyte accumulation at
extravascular sites) in immunodeficient mice lacking T lymphocytes or
deficient for MHC II expression (11). Thus, in contrast to
most PAMPs studied to date, it is not clear whether bacterial
superantigens directly activate cells of the innate immune system, such
as DC, in vivo. The purpose of this work was to analyze the maturation
status of splenic DC subpopulations in vivo following administration of
staphylococcal superantigens. Our observations indicate that
superantigens induce DC maturation in vivo with an unique cell surface
phenotype. Using a panel of genetically deficient mice, we demonstrate
that their ability to induce DC maturation requires T cells and MHC II
expression. The DC-activating properties of T cells were confirmed by
the analysis of in vivo responses to nonmicrobial T cell mitogens,
suggesting that polyclonal activation of T cells in vivo may lead to
activation of the innate immune response.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Six- to 8-wk-old female BALB/c mice were purchased from Harlan
Nederland (Horst, The Netherlands). Mice were maintained in
pathogen-free conditions in our animal facility. Mutant mice
(IL-4-/-, IL-12 p40-/-,
TNF-
-/-, IFN-
-/-,
CD1d-/-, CD4-/-,
CD8-/-, CD40-/-, and
RAG) were either purchased from The Jackson Laboratory (Bar Harbor, ME)
or bred in our own animal facility. Highly purified staphylococcal
enterotoxin A, SEB, and toxic shock syndrome toxin-1 (TSST-1) were
purchased from Toxin Technology (Sarasota, FL). LPS (Escherichia
coli serotype 0127:B8) was from Sigma-Aldrich (St. Louis,
MO).
In vivo treatment
Mice were injected i.v. into the lateral tail vein with the
indicated doses of LPS, SEB, TSST-1, OVA peptide, or anti-CD3
Abs solubilized in pyrogen-free NaCl (0.9%). Control animals were
injected with the same volume of diluents or with hamster control
IgG.
Purification of low-density spleen cells
Spleens were digested with collagenase (CLSIII; Worthington Biochemical, Freehold, NJ), further dissociated in calcium-free medium and separated into low- and high-density fractions on a Nycodenz gradient (Nycomed, Oslo, Norway), as previously described (12).
Cytofluorometric analysis
Cells were analyzed by flow cytometry with a FACSort cytometer
(BD Biosciences, Mountain View, CA). The cells were preincubated with
saturating doses of 2.4G2 (a rat anti-mouse FcR mAb; American Type
Culture Collection (ATCC), Manassas, VA) for 10 min before staining to
prevent Ab binding to FcR and further labeled with PE-coupled N418
(anti-CD11c) or, as indicated, biotinylated or FITC-coupled Abs
including RA3-6B2 (anti-CD45R/B220), RM4-5 (anti-CD4), 53-6.7
(anti-CD8), AF6-120.1 (anti-I-Ab), and
3.23 (anti-CD40), all from BD PharMingen (San Diego, CA). DEC-205
(anti-CD205), 14.4.4-S (anti-I-Ek,d),
16-10A1 (anti-CD80), and GL1 (anti-CD86) are available through
the ATCC and were purified and labeled in our laboratory. Cells were
gated according to size and scatter to eliminate dead cells and debris
from analysis.
Immunohistochemistry
Immunohistochemical techniques used in this study were described in detail by Pajak et al. (13). Spleens and lymph nodes were fixed for 3 days in Immunohistofix (Aphase, Gosselies, Belgium) followed by dehydration in a graded series of ethanol doses (30, 50, 70, 90, and 100%) for 30 min each at room temperature. Tissues were embedded in Immunohistowax (Aphase), sectioned at 3 to 6 µm, de-embedded by washing in acetone for 10 min, and transferred to PBS. The tissues sections were treated for 30 min with 1% blocking reagent (Boehringer Mannheim, Mannheim, Germany) to saturate nonspecific binding sites. The endogenous peroxidase activity was neutralized by 3% H2O2 in PBS for 60 min, and the slides were stained with the following biotinylated Abs (10 µg/ml in PBS 0.5% blocking reagent): MOMA-1 (anti-metallophilic macrophages; Serotec, Raleigh, NC), GL1 (anti-CD86; ATCC), and N418 (anti-CD11c; ATCC). Slides were then incubated as indicated with 1) avidin-biotin-peroxidase complex (Vectastain ABC kit; Vector Laboratories, Burlingame, CA) and stained with a solution of 3-amino-9-ethylcarbazole tablets (Sigma-Aldrich) or 2) avidin-biotin-alkaline phosphatase complex (Vectastain ABC kit, AK-5000; Vector Laboratories) and stained with alkaline phosphatase substrates (SK-5300, Vector Laboratories). The Vector blocking kit (Vector Laboratories) was used to block excess of biotin from the first Ab in double-stained slides. Digitized images were captured using a Ikegami CCD color camera (Ikegami Tsushinki, Tokyo, Japan) and analyzed using the CorelDraw 7 software (Corel, Ottawa, Ontario, Canada).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The effect of SEB administration on DC populations from spleen and
lymph nodes was visualized by immunohistochemistry. Most DC, identified
as CD11c-expressing cells, were found in the splenic marginal zone and
around the central arteriole of control, untreated animals (Fig. 1
a). Injection of SEB led to
the rapid (6 h post-treatment) redistribution of most
CD11c+ cells to the white pulp, in T cell-rich
areas surrounding the central arteriole (Fig. 1b). Most DC
remained localized in the white pulp 12 h post-treatment (Fig. 1c), while a gradual loss of CD11c+
cells was observed at 36 and 48 h following SEB treatment, as
determined by immunohistochemistry (note the weak CD11c-specific
labeling in Fig. 1, e and f) and flow cytometry
(Ref. 14 and data not shown). DC represent a
phenotypically heterogeneous cell population, characterized by the
differential expression of CD4 and CD8 markers (three subsets,
CD4+CD8-,
CD4-CD8+, and
CD4-CD8-, have been
described (15)). Using flow cytometry analysis, we have
observed that both CD4+ and
CD8+ DC subsets were transiently lost with
similar kinetics in response to E. coli LPS or SEB, while
the number of
CD4-CD8-CD11c+
cells was only marginally affected. Reconstitution of splenic DC
numbers was observed 2–4 days after treatment, in agreement with the
rapid turnover of these cells (data not shown).
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, g and j).
Double staining experiments identified these cells as marginal zone
macrophages expressing the MOMA-1 marker (Fig. 1, h and
k). Note that CD11c- and CD86-specific staining rarely
overlap in naive animals (Fig. 1j), while most MOMA-1
expressing macrophages were also stained by the CD86-specific Ab (see
Fig. 1k as an example). As expected, the MOMA-1 and CD11c
markers were expressed by distinct cell subpopulations (Fig. 1, i and l). In marked contrast, most
CD11c+ cells located in the white pulp in
response to SEB or LPS (known to induce DC maturation in vivo)
expressed the CD86 molecule (note the purple staining of double-labeled
CD11c- and CD86-positive cells in Fig. 1, m and
n), suggesting that these cells had undergone maturation in
vivo. The distribution of MOMA-1-positive cells was not affected by SEB
or LPS administration (data not shown). Finally, while CD11c and CD86
markers were expressed by distinct cell subpopulations in lymph node
sections from naive animals (Fig. 1o), injection of SEB led
to the maturation of DC cells of the paracortical and medullary zones
of mesenteric lymph nodes, as demonstrated by the rapid (6 h
post-treatment) induction of the CD86 markers on CD11c-positive cells
(note the purple staining in Fig. 1p). The capacity of SEB
to induce DC maturation in vivo was further confirmed by flow
cytometry. DC-enriched, low-density spleen cells were purified from
SEB- or LPS-treated mice and analyzed at different time points for the
expression of markers known to be up-regulated during DC maturation. As
previously demonstrated (16), LPS injection led to the
increased expression of the MHC II, CD80, and CD86 markers. Similarly,
SEB administration led to an increased expression of the MHC II and
CD86 markers, while it only marginally affected the CD80 marker (Fig. 2). Although up-regulation of MHC II
determinants was detectable 2 h post-treatment, all markers were
optimally expressed between 6 and 12 h after in vivo
administration of both microbial products (see Fig. 2). Reduced
expression of MHC II markers observed 2–4 days post-treatment (see
Fig. 2) is probably related to reconstitution of splenic DC populations
by immature DC or DC precursors (see Ref. 15) expressing
lower levels of MHC II molecules. Note that all CD11c-expressing DC
subsets analyzed
(CD4+CD8-,
CD4-CD8+, and
CD4-CD8-) were found to
mature in response to SEB or LPS (data not shown).
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To further analyze the DC-maturing properties of bacterial
superantigens, CD11c+ low-density spleen
cells from LPS- and SEB-treated animals were analyzed for the
expression of MHC II, CD40, CD80, CD86, and CD205 molecules by flow
cytometry. As shown in Fig. 3a, SEB injection led to the
up-regulation of most maturation markers, albeit with distinctive
efficacy when compared with LPS. Indeed, while DC that matured in
response to SEB expressed optimal levels of MHC II, CD40, and CD205
molecules, they reproducibly expressed reduced and marginal levels of
the CD86 and CD80 markers, respectively. This distinctive phenotype was
not related to an altered kinetic of DC maturation in response to SEB
(see Fig. 2 and data not shown) and was preserved in response to graded
doses of SEB. Fig. 3b shows the results of a typical
experiment in which mice were injected with graded doses of LPS or SEB
and analyzed by flow cytometry as previously described. For the sake of
simplicity, the results are expressed as the fraction of maximal
receptor expression induced by a saturating dose of LPS (100 µg).
This experiment revealed that markers often used to characterize the
mature status of DC were optimally induced at different LPS doses. In
particular, low doses of LPS were sufficient to induce MHC II, CD40,
and CD205 up-regulation, while doses in excess of 10- to 100-fold were
required to induce CD86 or CD80 expression, respectively. Injection of
SEB led to an increased expression of MHC II, CD40, and CD205
molecules, and to a moderate but significant up-regulation of the CD86
marker. Note that doses of SEB (10–100 µg) known to induce an
inflammatory response in vivo mimicking the naturally occurring
response to toxin-producing strains of S. aureus in humans
(6) were found effective in inducing DC maturation. This
experiment confirmed that DC isolated from SEB-injected mice display a
distinctive phenotype, characterized by increased expression of CD40,
CD205, and MHC II molecules but only a moderate up-regulation of the
costimulatory markers CD80 and CD86, known to interact with the
CD28/CTLA4 counterreceptors. It is noteworthy that the relative
expression of the CD205 and CD80 markers differed markedly between DC
that matured in response to LPS or SEB, suggesting that DC may respond
differently to these microbial products. Similar results were obtained
using both staphylococcal enterotoxin A and TSST-1 staphylococcal
superantigen (data not shown).
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The role of MHC II and T cell subsets in mediating the DC-maturing
properties of bacterial superantigens was studied using a panel of
genetically manipulated mice. Mutant mice were treated with the
indicated superantigen (mice in the C57BL/6 background were injected
with TSST-1, a staphylococcal superantigen known to optimally stimulate
I-Ab-expressing mouse strains, while BALB/c mice
expressing the I-Ed molecule were injected with
SEB) and LPS and analyzed as previously described for the expression of
several DC-specific maturation markers. For the sake of simplicity,
only expression of CD86 is reported in Fig. 4 (similar results were obtained for
CD40, CD80, and MHC II expression; data not shown). LPS induced DC
maturation in all mouse strains studied (Fig. 4). In marked contrast,
in vivo activation of DC by TSST-1 required both T cells and the
expression of MHC II molecules. In particular, MHC
II+/+RAG-/- (which lack
both T and B cells) mice failed to up-regulate CD86 in response to
TSST-1, suggesting that binding of bacterial exotoxins to MHC II is not
sufficient to induce DC maturation. MHC II-deficient mice also lack
CD4+ cells and were unresponsive to
TSST-1-induced DC maturation. TSST-1 was able to induce partial DC
maturation in both CD4-/- and
CD8-/- mice, indicating that both CD4- and
CD8-expressing T cell subsets were able to mediate DC maturation. Note,
however, that CD4+ T cells appeared to be more
efficient than CD8+ T cells in mediating
TSST-1-induced DC maturation, as shown by the enhanced CD86 expression
of DC from CD8-/- mice in response to bacterial
superantigen injection. NKT cells represent a subset of lymphocytes
that express a CD1d-restricted TCR repertoire, with the great majority
expressing an invariant TCR (V14-J281) paired with V8, -7,
or -2. Numerous observations suggest that these cells rapidly respond
in vivo to mitogenic or specific stimulation by producing high levels
of cytokines, including IL-4 and IFN-; therefore, these cells have
been implicated as potential effector cells in innate-like
immune responses to virus or tumors (17). These data
prompted us to test the potential role of NKT cells in inducing DC
maturation in response to SEB. As shown in Fig. 4, injection of SEB in
CD1d-/- mice led to DC maturation, suggesting
that NKT cells do not play a critical role in DC activation by
bacterial superantigens. CD40 ligation by Abs or CD40
ligand-transfected cells is sufficient to induce DC maturation in vitro
(18). However, the CD40 molecule was found dispensable for
DC maturation in vivo in response to TSST-1 (Fig. 4). Finally, absence
of IL-4, IFN-, TNF-, or IL-12 p40 did not affect the ability of
superantigens to induce DC maturation (Fig. 4). Collectively, these
data indicate that DC maturation in response to bacterial superantigen
is a consequence of in vivo T cell stimulation induced by the
simultaneous binding of the exotoxin to selected V regions of the
clonally distributed TCR and to a conserved domain of the class II
molecules.
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The previous experiments strongly suggested that polyclonal
activation of naive T cells in vivo may lead to DC maturation. To
formally demonstrate that T cell responses can induce DC maturation in
the absence of microbial products, naive mice were injected with
mitogenic anti-CD3 Abs. In keeping with our previous
observations, activation of naive T cells in vivo led to DC maturation,
characterized by DC migration (Ref. 19 and data not shown)
and up-regulation of MHC II and CD86 molecules (Fig. 5a). It is noteworthy that a
selective deficiency in CD80 up-regulation in response to a wide range
of Ab doses was observed. Use of RAG-deficient mice confirmed the role
of T cells in mediating DC maturation in response to anti-CD3
mAbs (Fig. 5b). These observations were extended to two
additional experimental models in which a sufficient number of T cells
were stimulated in vivo by an adequate Ag/MHC combination. Control mice
and mice transgenic for an OVA-specific TCR (BALB/C DO.11.10 strain)
were injected i.v. with the nominal antigenic peptide from OVA.
Transgenic, but not control, mice responded to peptide injection by
increased expression of MHC II and CD86 on CD11c-expressing cells (Fig. 6a). Similar observations were
performed in the C57BL/6 OTII OVA-specific TCR-transgenic mouse strain
(data not shown).
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b). In agreement with the previous observations, T
cell-induced DC maturation in vivo led to a distinctive DC phenotype,
characterized by a weak up-regulation of the CD80 marker (Fig. 6).
The specific role of T cell responses in the regulation of DC
maturation was ascertained by studying the effect of polyclonal B cell
responses in vivo on DC phenotype. Naive mice were injected with LPS,
anti-CD3, or anti-IgD mAbs, and the phenotype of B cells
(identified by expression of the CD45RB marker) and DC
(CD11c+ cells) was determined by multiparameter
flow cytometry (Fig. 7). All mitogenic
compounds were able to activate B cells in vivo, in agreement with
previous observations (20, 21, 22). Note that the ability of
anti-CD3 mAbs to up-regulate MHC II molecules on B cells is
related to the production of IL-4 by anti-CD3-activated T cells
(21). Of relevance to this study, anti-IgD mAbs led to
B cell activation in vivo but failed to induce DC maturation, as judged
by their inability to affect MHC II (data not shown) and CD86
expression on CD11c+ cells
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The major conclusion from the present study is that polyclonal T cell
responses in vivo, such as those induced by staphylococcal
superantigens, induce DC maturation in vivo, characterized by a
distinct phenotypic profile. Injection of staphylococcal superantigens
induced the rapid migration of splenic DC to the white pulp and the
up-regulation of MHC II, CD40, CD80, CD86, and CD205 on all DC
subpopulations tested. Despite wide variation in their protein
structure, staphylococcal superantigens share the ability to
simultaneously interact with MHC II molecules and the variable region
of the TCR V chain. Although additional receptors for these
bacterial toxins have been reported (9, 10, 11), it is clear
from this study that interaction of staphylococcal superantigens with
both MHC II and TCR V molecules is required for optimal DC
maturation in vivo (Fig. 4). Therefore, it is tempting to speculate
that polymorphic receptors (MHC II and TCR V molecules) primarily
dedicated to the recognition of Ags in the context of the adaptive
immune response act as pattern recognition receptors when confronted
with bacterial superantigens.
The mechanism by which superantigens cause DC maturation is presently
unclear. Experiments in which T cells were activated in vivo by ligands
of nonmicrobial origin support the notion that DC mature as a
consequence of an excessive in vivo T cell response. Indeed,
stimulation of a sufficient number of T but not B lymphocytes in vivo
by nominal Ag/MHC complexes (OVA peptide or allo-MHC) or
anti-CD3 mAbs led to DC maturation. Of note, injection of
bacterial superantigens (23), anti-CD3 mAbs
(24), or nominal Ag in a TCR-transgenic model
(25) cause the rapid (1–3 h post treatment, depending on
the experimental settings) activation of T cells to cytokine secretion.
In keeping with these observations, in vivo signaling studies performed
in an OVA-transgenic model recently demonstrated that most Ag-specific
T cells were activated in vivo a few minutes after Ag administration
(26). Thus, in contrast to predictions mostly based on in
vitro studies, activation of T cells in these experimental models
precedes DC maturation, and in particular up-regulation of
costimulatory molecules. This conclusion is strengthened by the
aforementioned study (26), in which activation of naive,
TCR-transgenic T cells in vivo by Ag was found to be largely CD28
independent. Collectively these observations support the notion that
naive T cells can be rapidly stimulated in vivo in the absence of
pathogen-derived danger signals and eventually initiate a series of
events leading to DC maturation. This conclusion is also supported by a
study performed with human cells which demonstrated that Ag-stimulated
T cells can positively regulate the expression of costimulatory
molecules on DC (27).
Soluble signals (inflammatory cytokines) and cell-cell interactions
(such as CD40/CD40 ligand signaling) are known to initiate DC
maturation in vitro. However, despite numerous experiments using
genetically manipulated mice, we could not single out a soluble
mediator responsible for T cell-dependent DC maturation. Indeed, SEB
induced an optimal DC maturation in mice defective for TNF-, IL-4,
IL-12 p40, and IFN-. Similarly, TSST-1 induced DC maturation in
CD40-deficient mice (Fig. 4). Therefore, it is not clear whether
bacterial superantigen-induced maturation requires DC/T cell
contacts or whether it is mediated by a functionally redundant set of
soluble mediators. In any event, the available evidence indicates that
activation of a sufficient number of naive T cells in vivo represents a
novel danger signal, functionally linked to DC maturation. In keeping
with this conclusion, it has been recently demonstrated that adaptive
stimuli (such as provided by a high frequency of Ag-reactive T cells)
can increase the number of Ag/MHC complexes displayed by DC in vivo
(28).
Although both innate and T cell-mediated danger responses activate DC, maturation of this cell population does not represent a homogeneous stereotyped biological response, as is often inferred from in vitro studies. Indeed, the phenotype of DC varied according to the origin and dose of microbial products. In particular, low doses of LPS (in the 10-ng range) and SEB (in the 1-µg range) were sufficient to elicit up-regulation of CD40, CD205, and MHC II molecules, while near-lethal doses of SEB (100 µg) were unable to effectively induce CD80 up-regulation, suggesting that expression of the CD80 costimulatory molecules may be under the strict control of the innate immune response. These observations provide evidence for a graded response of DC to maturation stimuli, suggesting that DC may display distinct phenotypes reflecting both the origin and the intensity of a danger signal (see Ref. 29). The functional consequence of these distinct DC phenotypic profiles on the in vivo adaptive immune response is difficult to establish. One of the best-characterized consequences of bacterial superantigen administration is the induction of anergy and/or apoptosis in a fraction of responding T cells (6, 7, 8); therefore, it is tempting to correlate this inadequate T cell response with the unusual phenotype of SEB-activated DC. Interaction of T cells with DC able to simultaneously provide a strong signal 1 (due to the high expression of MHC II molecules) and a low signal 2 (due to reduced expression of CD28 ligands, known to promote T cell survival (30)) may explain the altered response of these cells and their propensity to die by apoptosis. Accordingly, it has been shown that Ag presentation to T cells in vivo by immature DC induced incomplete and nonsustained T cell activation (31).
The observations reported in this study do not contradict the well-accepted notion that responses to Ags for which a low frequency of responding cells is available in the periphery is strictly dependent on costimulation delivered by DC that have matured under the influence of innate immunity-derived signals. In particular, autoreactive T cells that escaped negative selection in the thymus are present at low frequencies in the periphery, and their activation probably requires adequate costimulation only provided by mature DC. As previously discussed, the response to bacterial superantigens represents a clear exception to this rule. A second exception that bears significant clinical relevance is the response to allo-MHC determinants. The induction of a vigorous allograft rejection in the absence of inflammation has recently been reported (32). These authors elegantly demonstrated that well-healed skin and vascularized cardiac allografts were efficiently rejected by T cells developing from a transplanted marrow graft. These studies are in agreement with the observations reported herein, which suggest that a high frequency of responding T cells is sufficient to initiate a series of events (including DC migration and maturation), culminating in the induction of an adequate adaptive immune response. The observations reported herein provide a sound basis for understanding why alloreactive T cells that are continuously produced in the bone marrow of transplanted patients represent a continuous threat for a well-functioning allografts, even in the absence of additional danger signals.
In conclusion, these studies are in line with the concept that DC represent an important biological integrator of danger signals. The stimuli that activate DC maturation and thereby initiate adaptive immune responses are 1) exogenous microbial products interacting with cells of the innate immune system; 2) endogenous signals released by cells under stress, damage, or necrotic death; and, finally, 3) usual stimulation of a large fraction of naive T cells, which can only occur in response to exogenous signals such as superantigens or allo-MHC determinants. A better understanding of the signals involved in DC maturation induced by polyclonally activated T cells in vivo may ultimately lead to improved treatments for septic shock caused by bacterial superantigens and for inducing and maintaining peripheral tolerance to allografts.
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Acknowledgments |
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Footnotes |
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2 E.M. and C.D.T. should be considered equally as first authors.
3 Address correspondence and reprint requests to Dr. Oberdan Leo, Laboratoire de Physiologie Animale, Université Libre de Bruxelles, 12 rue des Professeurs Jeener et Brachet, 6041 Gosselies, Belgium. E-mail address: oleo{at}ulb.ac.be
4 Abbreviations used in this paper: DC, dendritic cell; MHC II, MHC class II; TSST-1, toxic shock syndrome toxin-1; SEB, staphylococcal enterotoxin B; TLR, Toll-like receptor; PAMP, pathogen-associated molecular pattern.
Received for publication January 7, 2002. Accepted for publication March 4, 2002.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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+ and CD8- dendritic cells to prime Th1/Th2 cells in vivo. J. Immunol. 167:4345.-unrestricted T cell unresponsiveness and defective APC functions. J. Immunol. 158:2638.[Abstract]
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 E. Andreakos, C. Smith, C. Monaco, F. M. Brennan, B. M. Foxwell, and M. Feldmann Ikappa B kinase 2 but not NF-kappa B-inducing kinase is essential for effective DC antigen presentation in the allogeneic mixed lymphocyte reaction Blood, February 1, 2003; 101(3): 983 - 991. [Abstract] [Full Text] [PDF]
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B. Adkins, Y. Bu, and P. Guevara Murine Neonatal CD4+ Lymph Node Cells Are Highly Deficient in the Development of Antigen-Specific Th1 Function in Adoptive Adult Hosts J. Immunol., November 1, 2002; 169(9): 4998 - 5004. [Abstract] [Full Text] [PDF]
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