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Mucosal Immunology Laboratory, Massachusetts General Hospital, and Harvard Medical School, Charlestown, MA 02129
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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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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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BALB/cByJ mice (8–10 wk of age) were purchased from The
Jackson Laboratory (Bar Harbor, ME) and were maintained in a specific
viral pathogen-free facility at Massachussetts General Hospital. To
examine the influence of parasitic infection on the expression of the
costimulatory molecules B7.1 and B7.2, groups of mice were infected
with 200 third-stage larvae as previously described (8)
and sacrificed at 8 days postinfection (p.i.). To follow the response
of transgenic, OVA-specific T cells in vivo, BALB/c mice were
adoptively transferred with cells from age- and sex-matched homozygous
DO.11.10 mice on a BALB/c background as described below. The adoptive
transfer recipients were either not infected or were orally inoculated
with 200 third-stage larvae 2 days after transfer. Groups of mice were
then fed and/or immunized with OVA using four different experimental
protocols as indicated in Fig. 2
. Where indicated, 25 mg of OVA
(fraction VII; Sigma, St. Louis, MO) or PBS was administered
intragastrically using a ball-tipped feeding needle. In some
experiments, mice were immunized at day 8 p.i. by the injection of
100 µg of OVA in CFA or IFA in the hind footpads.
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The expression of the costimulatory molecules B7.1 and
B7.2 was analyzed on APC populations isolated from both noninfected
mice and from mice at 8 days p.i. Spleen, mesenteric lymph nodes (MLN),
and Peyer’s patches (PP) were pooled from 5 to 15 mice/group and
pressed through nylon mesh to prepare single-cell suspensions. To
enrich for macrophages, the cell suspensions were incubated in complete
DMEM (Life Technologies, Grand Island, NY) containing 10% FCS (HyClone
Laboratories, Logan UT), 10 mM HEPES, 2 mM L-glutamine, 100
U penicillin/ml, 100 µg streptomycin/ml, 50 µM 2-ME, 0.1 mM
nonessential amino acids, and 1 mM sodium pyruvate for 1 h at
37°C, and the plastic adherent cells were removed for analysis. To
enrich for dendritic cells, each of the tissues was digested with
collagenase (200 U/ml; Worthington Biochemical, Lakewood, NJ). The
low-density cell population was obtained by centrifugation in an
OptiPrep gradient (Life Technologies). In each of the tissue
preparations, the various APC populations were identified with
FITC-labeled Abs (purchased from PharMingen, San Diego, CA) to B cells
(CD45R/B220, clone RA3-6B2), macrophages (CD11b, Mac-1
-chain,
M1/70), and dendritic cells (CD11c, HL3) or with a mixture of
FITC-labeled rat and hamster Ig isotype controls. B7.1 and B7.2 were
identified with PE-labeled anti-CD80 (16-10A1) or CD86 (GL1),
respectively. For Ab staining, the cells were incubated in PBS
containing 10 µg/ml anti-CD16/CD32 (Fc
III/IIR; PharMingen)
plus 5% normal mouse serum to block Fc receptor-mediated binding. Each
of the cell suspensions was then washed, and 1 x
106 cells were incubated with Ab in flow
cytometry buffer (PBS containing 0.05% BSA/0.01 M
NaN3) for 15 min on ice. The stained samples were
analyzed on a FACScan (Becton Dickinson, Mountain View, CA) with
CellQuest (Becton Dickinson) software. Dead cells
and debris were excluded from analysis by gates set on forward and side
angle light scatter. A total of 1000–5000 events were analyzed in the
gated B220, Mac-1, or CD11c+ populations.
Adoptive transfer protocol
DO.11.10 mice on an inbred BALB/c background
(9) were obtained from Dr. Kenneth Murphy (Washington
University, St. Louis, MO) and were maintained as a breeding colony in
a specific pathogen-free facility at Massachussetts General Hospital.
Peripheral lymph nodes (LN) (axillary, inguinal, and popliteal) and
spleen were harvested from donor mice that were age and sex matched to
the adoptive transfer recipients. Single-cell suspensions were prepared
by pressing the tissues through nylon mesh. CD8+
T cells were depleted by two rounds of treatment with an
anti-CD8 Ab culture supernatant (536.7) and rabbit complement
(Low Tox M; Life Technologies). The treated cell suspensions were
washed, and aliquots were removed for flow cytometric analysis of the
percentage of cells expressing CD4 (with CD4-CyChrome; PharMingen) and
the clonotypic transgenic TCR (identified with PE-labeled KJ1-26
(Caltag, San Francisco, CA). The cell counts were then adjusted so that
each transfer recipient received the same number of
CD4+ KJ1-26+ cells (which
varied from 2 to 5 x 106/mouse, depending
on the experiment). In some experiments the transgenic donor cells were
labeled with the fluorescent dye 5,6-carboxyfluorescein diacetate
succinimidyl ester (CFSE; Molecular Probes, Eugene, OR) as previously
described (10, 11, 12, 13). Briefly, pooled lymphocytes from
DO.11.10 transgenic mice were resuspended at 107
cells/ml in PBS containing 0.1% BSA with 10 µM CFSE for 10 min at
37°C. The labeled cells were washed twice in PBS/BSA before transfer
(via tail vein injection) to BALB/c recipients. The adoptive transfer
recipients were sacrificed 0–7 days after Ag feeding (see experimental
protocols outlined in Fig. 2
). Suspensions of cells were prepared from
the MLN and either pooled peripheral LN or the draining popliteal LN
(PLN) of individual mice and stained with CyChrome-labeled
anti-CD4, PE-KJ1-26, or a PE-mouse IgG2a isotype control. Flow
cytometric analysis was performed using a FACScan (Becton Dickinson)
with CellQuest (Becton Dickinson) software. Dead cells and debris were
excluded from analysis by gates set on forward and side angle light
scatter. Between 200,000 and 500,000 events were collected for each
sample. At least two independent experiments were performed for each of
the experimental protocols outlined in Fig. 2.
The histograms obtained for the CFSE fluorescence intensity of the
gated CD4+ KJ1-26+ T cells
were used to calculate responder frequency (R) and
proliferative capacity (Cp) as described by
Turka and colleagues (12, 13) using the formulas:
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In vitro restimulation and ELISA for cytokine production
PLN cells (2 x 106/ml) from
each mouse were cultured in triplicate in flat-bottom microtiter plates
(Costar, Cambridge, MA) in 200 µl of complete DMEM with or without
OVA (10 µg/ml). The cultures were pulsed at 72 h with 1
µCi/well of [3H]TdR (DuPont NEN, Boston, MA)
and harvested 16 h later. [3H]Thymidine
incorporation was determined by liquid scintillation counting (LS 1801;
Beckman Coulter, Fullerton, CA). Additional PLN cells from each mouse
were cultured in 24-well plates (4 x 106
cells/ml) in complete DMEM, with or without 100 µg/ml of OVA.
Cytokine secretion into the supernatants 72 h after the initiation
of the culture was determined by ELISA, as previously described
(8, 14). Immulon II plates (Dynatech Laboratories,
Chantilly, VA) were coated with capture Abs (BVD4-1D11 for IL-4 and
R4-6A2 for IFN-) overnight at 4°C, followed by blocking in PBS
with 3% FCS at 37°C for 1 h. After washing with PBS with 0.05%
Tween, the culture supernatants, or recombinant murine IL-4 or IFN-
(Genzyme, Cambridge, MA), were incubated, in triplicate, overnight at
4°C. The plates were then washed and incubated with biotinylated
secondary Abs (BVD6-24G2 for IL-4 and XMG1.2 for IFN-), followed by
peroxidase-conjugated strepavidin (Zymed, San Francisco, CA) and
developed with O-phenylenediamine (Zymed). The reaction was
stopped with 2 N H2SO4, and
the plates were read at 490 nm using an Emax Microplate reader
(Molecular Devices, Sunnyvale, CA). The concentrations of cytokine in
each sample were calculated from standard curves using SOFTmax PRO
software (Molecular Devices).
Statistical analyses were performed using a two-tailed Student’s t test. A value of p < 0.05 was considered significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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At least two signals are required to induce T cell
activation and proliferation: 1) recognition of peptide/MHC complexes
by the Ag-specific TCR, and 2) the signal delivered by costimulatory
molecules on APCs. Among the most potent costimulatory molecules are
two members of the B7 family, B7.1 and B7.2. Gause and colleagues have
shown that the Th2 response to H. polygyrus infection, like
that to other inflammatory stimuli, is B7 dependent and can be blocked
by treating mice with Abs to B7.1 and B7.2 (15). To
examine the levels of costimulatory molecules present on potential APC
populations at day 8 p.i., the point at which we administer oral
Ag, we looked at the expression of B7.1 and B7.2 on B cells (Fig. 1A), dendritic cells (Fig. 1B), and macrophages (Fig. 1C) in the
PP, MLN, and spleen of both infected and noninfected mice. LPS blasts
served as a positive control for B7 staining. Fig. 1A shows
that B7.2 is markedly up-regulated on B cells, particularly in the MLN,
at day 8 p.i., with little up-regulation in the PP and little or
none in the spleen. B7.1 expression is not greatly up-regulated at any
site. As we have previously noted (8), the proportion of B
cells in the MLN of helminth-infected mice is also dramatically
altered. By day 8 p.i., the enlarged MLN has shifted from an organ
containing 70% T cells and 30% B cells (in noninfected mice) to one
that contains 70% B cells and 30% T cells and is the primary site for
the induction of the polyclonal parasite-induced IgG1 and IgE response.
Fig. 1B shows that both B7.1 and B7.2 expression is
up-regulated on dendritic cells in the MLN and PP of infected mice, but
not in the spleen. The proportion of macrophages in the PP is
increased, but B7 expression is not altered (Fig. 1C). By
contrast, B7.2 (but not B7.1) expression is up-regulated on macrophages
in the MLN of infected mice, and both B7.1 and B7.2 are down-regulated
in the spleen. Together these results suggest that H.
polygyrus infection dramatically up-regulates costimulatory
molecule expression for each of the professional APC populations
examined. This effect is most prominent in the MLN, detectable in the
PP, and absent in peripheral lymphoid tissues such as the spleen (Fig. 1) and peripheral LN (data not shown). Therefore, H.
polygyrus infection fulfills one of the criteria expected of an
adjuvant by inducing the expression of costimulatory molecules on
APCs.
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It is difficult, in general, to follow Ag-specific
responses induced in vivo without in vitro restimulation, in part
because the numbers of cells stimulated in response to a nominal Ag
like OVA are too small to follow in vivo in normal mice. To examine the
consequences of mucosal infection for the T cell response to orally
administered Ag, we tracked the response of OVA-specific lymphocytes
directly isolated ex vivo, using the adoptive transfer system
originally described by Kearney et al. (16). In this
model, transgenic T cells are transferred into normal BALB/c mice in
numbers large enough to track in vivo with an anti-TCR-specific Ab,
but small enough to simulate the normal physiological response to Ag
(17, 18). The transgenic TCR expressed on 70–80% of the
CD4+ T cells in these mice is specific for the
chicken OVA peptide 323–339 in the context of
I-Ad (9) and is detectable with the
clonotypic anti-TCR Ab KJ1-26 (19). This model has
been used extensively to track the response to tolerogenic and
immunogenic forms of OVA in vivo (2, 7, 10, 16, 17, 18, 20, 21, 22, 23, 24, 25). Four different types of adoptive transfer experiments
were performed, as indicated in the experimental protocols outlined in
Fig. 2.
In the first set of experiments, BALB/c mice were adoptively
transferred with transgenic, OVA-specific T cells (Fig. 2, protocol A). Two days after transfer, one group of mice was
infected with H. polygyrus. At 8 days p.i., groups of both
infected and noninfected mice were immunized in the footpads with OVA
in CFA. We used two-color flow cytometric analysis to look at the
percentage of clonotype-positive cells in the PLN (Fig. 3). Very few, if any, clonotype
(KJ1-26)-positive cells are detected in the PLN of normal BALB/c mice.
After transfer, but without Ag, we can reproducibly detect a small
(0.4%) population of KJ1-26+ cells. If we
immunize with OVA/CFA in the footpad and take the draining PLN 3 days
later we see a 10-fold expansion in the OVA-specific
KJ1-26+ T cells. This clonal expansion is further
enhanced (about 2-fold) in the presence of H. polygyrus
infection. Therefore, enteric helminth infection potentiates the
response to footpad immunization with OVA in CFA.
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A) and
PLN (Fig. 4C) at 0, 2, 3, and 7 days after OVA feeding
according to the protocol in Fig. 2B. Fig. 4A shows that intragastric
administration of OVA to adoptively transferred BALB/c mice induces the
Ag-specific expansion of transgenic, OVA-specific T cells in the MLN
with a peak at day 3, followed by a decline. However, in infected mice,
the absolute number of CD4+
KJ1-26+ cells increases in the MLN of both OVA-
and PBS-fed mice. The proportion of CD4+
KJ1-26+ cells in the infected MLN is actually
lower than that in noninfected mice (Fig. 4B), but the
dramatic parasite-induced enlargement of the MLN results in greater
absolute numbers of clonotype-positive cells in this tissue. By day 7,
the numbers of CD4+ KJ1-26+
cells have declined in the MLN of normal mice but remain elevated in
infected mice. There is a statistically significant difference in the
numbers of transgenic T cells that have survived in the MLN of infected
and noninfected mice 7 days after OVA feeding
(p = 0.03). No expansion of transgenic T cells
is observed in the PLN after oral administration of Ag, and both the
numbers (Fig. 4C) and percentage (Fig. 4D) of
CD4+ KJ1-26+ cells in the
PLN decrease slightly over 7 days.
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To examine whether mucosal or peripheral
CD4+ KJ1-26+ cells
proliferate in response to OVA feeding, transgenic donor cells were
labeled with the vital dye CFSE before transfer according to the
protocol in Fig. 2C. CFSE segregates equally into daughter
cells upon cell division, allowing proliferation to be measured as a
reduction in CFSE fluorescence intensity using flow cytometric
analysis. This powerful technique provides a proliferative history of
subpopulations of cells in response to in vivo Ag challenge, without
manipulation in vitro. In the representative histograms in Figs. 5 and 6
proliferating cells are indicated by the peaks of reduced CFSE
fluorescence intensity to the left of the dotted lines. Fig. 5 shows
the proliferative response of gated CD4+
KJ1-26+ cells in the MLN and peripheral LN of
infected and noninfected mice 3 days after intragastric administration
of OVA or PBS (i.e., at the peak of clonal expansion; see Fig. 4). OVA
feeding induces the proliferation of CD4+
KJ1-26+ cells in the MLN (Fig. 5A) but
not peripheral LN (Fig. 5C). In Fig. 5B, the data
obtained from the CFSE fluorescence histograms of
CD4+ KJ1-26+ cells from the
MLN of individual mice in each of the four groups is averaged and
plotted as the mean percentage of undivided cells and the mean
percentage of cells with greater than four divisions (±SEM). Fig. 5B shows that, in the absence of infection, most (82 ±
1.4%) of the CD4+ KJ1-26+
cells in PBS-fed mice do not divide, while only 25 ± 3.5% of the
transgenic T cells are undivided in OVA-fed mice
(p = 0.0001). A total of 41 ± 1.4% of
the cells in the MLN of OVA-fed mice have undergone greater than four
divisions, compared with only 7 ± 0.9% in-PBS-fed mice
(p < 0.0001). A total of 62 ± 8% of the
CD4+ KJ1-26+ cells from the
MLN of helminth-infected, PBS-fed mice did not divide, whereas only
32 ± 1.1% had not divided after OVA feeding
(p = 0.021). The mean percentage of
CD4+ KJ1-26+ cells in the
MLN of OVA-fed, infected mice that have undergone greater than four
divisions (33 ± 0.6) resembles that seen in OVA-fed, noninfected
mice but is not significantly different from that seen in PBS-fed,
infected mice (22 ± 9). The polyclonal immune activation induced
by helminth infection is apparently inducing some nonspecific
activation and proliferation in the CD4+
KJ1-26+ cells in the PBS-fed mice, which varied
from mouse to mouse (note the larger error bars for PBS-fed, infected
mice in Fig. 5B). This Ag-nonspecific proliferation of the
transgenic T cells is consistent with the expansion of
CD4+ KJ1-26+ cells in the
MLN of both PBS- and OVA-fed mice noted in Fig. 4. By contrast, no
proliferative response to OVA feeding was observed in the peripheral LN
of either infected or noninfected mice (Fig. 5C).
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Our earlier work had shown that in vitro restimulation of
cells from the draining PLN of helminth-infected mice immunized in the
footpads with OVA in CFA resulted in a Th2-biased proliferative and
cytokine response to OVA instead of the systemic nonresponsiveness
normally induced by orally administered Ags (Ref. 8). Yet
our experiments to this point indicate that the initial peripheral and
mucosal response to orally administered OVA is the same in both
infected and noninfected mice. Therefore, we next examined the response
of helminth-infected mice given oral Ag to immunization in the footpad
with OVA plus IFA. We chose s.c. immunization with Ag in IFA because
previous reports had shown that this protocol elicits a T cell response
in the draining LN while minimizing the inflammation and Th1 bias that
would be induced by Ag in CFA (16, 23). BALB/c mice were
adoptively transferred with CFSE-labeled transgenic donor cells as in
Fig. 5. However, in this set of experiments, 5 days after oral
administration of OVA or PBS the mice were immunized in the footpads
with OVA in IFA and sacrificed 3 or 10 days later (according to the
protocol outlined in Fig. 2D). As in the experiments shown
in Fig. 5, the orally administered soluble OVA induced the
proliferation of CD4+
KJ1-26+ cells in the MLN of both noninfected and
infected mice. Footpad immunization drains predominantly to the PLN and
does not induce a response in the MLN (data not shown). A much more
pronounced proliferative response is induced in the PLN by immunization
with OVA in adjuvant (Fig. 6). By 3 days after immunization, most of
the CD4+ KJ1-26+ cells have
undergone some division. However, the histogram overlays shown in Fig. 6 demonstrate that CD4+
KJ1-26+ cells from the PLN of OVA-fed noninfected
mice (shaded) proliferate less well than those from OVA-fed infected
mice (heavy lines, no shading) or PBS-fed mice with or without
infection. About 60% of the CD4+
KJ1-26+ cells from OVA-fed noninfected mice
underwent greater than four cell divisions, compared with between 76
and 80% of the CD4+
KJ1-26+ cells for each of the other three
groups.
To exclude the possibility that helminth infection is simply altering
the kinetics of the response to OVA in IFA, we examined the
proliferation of CFSE-labeled CD4+
KJ-126+ T cells at both early (3 days) and late
(10 days) time points after footpad immunization. This data is shown in
Table I, where the events under each peak
in the histograms of CFSE fluorescence have been used to calculate the
responder frequency (the proportion of cells that participate in clonal
expansion) and the proliferative capacity (the number of daughter cells
generated by each precursor T cell) of the CD4+
KJ1-26+ T cells (Refs. 13 and
12 ; see Materials and Methods). The mean
frequency of CD4+ KJ1-26+
cells that responded by dividing in the PLN of PBS-fed, noninfected
mice was 64% (at day 3) and 59% (day 10), similar to the responder
frequency of 65% reported by Turka and colleagues (12, 13). The responder frequency was somewhat reduced in OVA-fed
mice (between 45 and 57%) but did not significantly differ from
PBS-fed controls in either infected or noninfected mice. However, the
mean proliferative capacity of CD4+
KJ1-26+ cells in the PLN of OVA-fed, noninfected
mice was significantly reduced at both 3 and 10 days postimmunization
when compared with PBS-fed controls. This reduction in proliferative
capacity was abrogated in helminth-infected mice. To directly link the
reduction in proliferative capacity measured directly ex vivo in CFSE-
labeled cells to our previous observations on the influence of helminth
infection on the response to oral Ag, we also restimulated cells from
the draining PLN, at both time points postimmunization, with OVA in
vitro. Both proliferative responsiveness (as assessed by incorporation
of [3H]thymidine) and cytokine secretion into
the culture supernatants (as measured by ELISA) were examined. At 3
days postimmunization, cells from the draining PLN proliferated poorly
when restimulated with OVA in vitro (Fig. 7E) and secreted low levels of
IFN- (Fig. 7A). Interestingly, however, although
the IFN- response was undetectable, IL-4 was detectable only in the
supernatant of PLN cells from helminth-infected, OVA-fed mice (Fig.
7C). By day 10 postimmunization PLN cells from PBS-fed
mice proliferated and secreted IFN- in response to restimulation
with OVA in vitro. In both infected and noninfected mice OVA feeding
virtually abrogates the ability of these cells to secrete IFN- in
response to OVA restimulation in vitro (Fig. 7B), in
agreement with our previous report (8). T cells from
OVA-fed, noninfected mice proliferate weakly in response to OVA in
vitro, whereas the proliferative responsiveness of cells from the PLN
of OVA-fed, helminth-infected mice was only partially reduced (Fig.
7F).
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). As can be seen in Figs. 5 and 6, as well as in previous reports (12, 13), division
of clonotype-positive cells in this model is asynchronous; some cells
have divided as many as eight times while others divide only two or
three times, or not at all. The death and elimination of cells that
have undergone multiple rounds of division, combined with the reduction
in proliferative capacity induced by the tolerizing stimulus, may
result in a cumulative loss of functional responsiveness that is
compounded by each subsequent challenge (see Discussion). By
contrast, an immunizing stimulus primes for the clonal expansion of
Ag-specific cells that are further expanded with each challenge such
that the loss of cells to clonal deletion is compensated for by cell
division and the generation of more daughter cells. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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) and
up-regulating costimulatory signals (Fig. 1), which restore the
capacity of KJ1-26+ T cells to proliferate in
response to Ag challenge to that seen in PBS-fed mice (Fig. 6). In the
presence of the parasite-induced, Th2-biased cytokine response, these
adjuvant effects result in a Th2-biased immune response to OVA upon
restimulation in vitro (Ref. 8 and Fig. 7).
Our results also have implications for the role of the gut-associated
lymphoid tissue in the induction of oral tolerance, which remains
controversial. Recent reports demonstrating peripheral T cell
activation in response to oral Ag have reinforced the view that
nonresponsiveness is due primarily to Ag uptake into the bloodstream
and the direct induction of anergy in peripheral sites
(27). However, subsequent work has demonstrated that this
peripheral up-regulation of phenotypic markers of activation (e.g.,
CD69) is not accompanied by proliferation or clonal expansion, which
occurs primarily in the MLN (10, 26). These observations
have been confirmed, and extended, in this report. Moreover, because we
have no indication of a proliferative response of cells in the
peripheral LN to orally administered OVA (Fig. 5C), we
suggest that at least some of the reduction in the proliferative
capacity of KJ1-26+ cells in the peripheral LN of
noninfected, OVA-fed mice represents the migration of cells from the
MLN into the peripheral LN in response to footpad immunization with OVA
in IFA. Other work has shown that T cell anergy is likely to be the
consequence of the preferential binding of low levels of constitutively
expressed B7 to its higher-affinity, inhibitory T cell ligand CTLA-4
(23). A recent report demonstrating that engagement of
CTLA-4 induces TGF-
secretion by CD4+ T cells
provides a link between the induction of functional
nonresponsiveness at low levels of costimulation and the generation of
regulatory cells secreting TGF-, which have been implicated in oral
tolerance (28). TGF--secreting cells have been thought
to require the unique cytokine microenvironment of the gut-associated
lymphoid tissue for their growth and differentiation (29).
It is tempting to speculate that the reduced proliferative capacity of
KJ1-26+ cells in the peripheral LN of OVA-fed
mice reflects the migration of cells from the MLN, which have already
transiently divided, but have received an inhibitory signal via CTLA-4.
These functionally nonresponsive, gut-derived cells may further dampen
the responsiveness of cells in the peripheral LN to
subsequent challenge with Ag via their secretion of
regulatory cytokines like TGF-, resulting ultimately in
systemic nonresponsiveness. This hypothesis would also be consistent
with a recent report indicating that, in the periphery, CD4 T cell
tolerance induction in vivo is not due simply to an insufficient
proliferative response to initial TCR engagement (30).
Subcutaneous administration of LPS with soluble OVA, using the same
adoptive transfer model, also elicits the clonal expansion of
Ag-specific T cells (2). LPS stimulates macrophages to
secrete the proinflammatory cytokines TNF-, IL-1, and IL-6, and
coadministration of soluble Ag with TNF- and IL-1 mimics the T cell
clonal expansion and follicular migration seen with LPS itself
(2, 32). Cholera toxin (CT) is among the most potent of
mucosal adjuvants (4, 5). Recent work has shown that
immunity to orally administered soluble Ag plus CT can be enhanced by
treatment with Flt3L, a dendritic cell growth factor (31).
The potentiation of CT’s adjuvanticity resulted from the
combined effects of DC expansion and CT’s ability to
induce the up-regulation of B7 expression (and DC maturation) through
the induction of proinflammatory cytokines, particularly IL-1 and IL-6
(31). Indeed, administration of Ag (OVA) plus IL-1 alone
(without CT) was sufficient to induce a productive immune response to a
normally tolerogenic form of Ag (31). The ability of IL-1
to act as an adjuvant to abrogate both peripheral and mucosal tolerance
to soluble Ag has been documented in other reports as well
(33). Helminth infection also induces the secretion of
proinflammatory cytokines, at least one of which, TNF-, has recently
been shown to be essential in regulating the Th2 response involved in
host protection against helminth infection (34). A recent
report has also linked the polyclonal activation induced by helminth
infection to the up-regulation of accessory cell production of IL-6.
Helminth infection was shown to enhance the survival of activated T
cells by increasing T cell proliferation and reducing
activation-induced cell death (35).
It is increasingly clear that systemic and mucosal adjuvants are microbial products like CT (31, 33), bacterial CpG DNA (6), and LPS (2, 32), which elicit the secretion of proinflammatory cytokines by cells of the innate immune system. The resultant up-regulation of costimulatory molecule expression induces a response to a normally tolerogenic form of Ag (reviewed in Ref. 36). Therefore, the innate immune system can control the generation of the adaptive immune response through the up-regulation of B7 costimulatory molecule expression on APC. We report here the novel observation that an ongoing enteric infection appears to use a similar mechanism to act as an adjuvant for the response to an orally administered soluble Ag. Because parasitic infection is endemic in developing countries, our results also have important clinical implications for strategies for oral vaccination and the development of allergic responses to food Ags.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Cathryn Nagler-Anderson, Mucosal Immunology Laboratory, Massachussetts General Hospital East, Room 3308, Building 149, 13th Street, Charlestown, MA 02129.
3 Abbreviations used in this paper: CT, cholera toxin; MLN, mesenteric lymph node(s); PLN, popliteal lymph node(s); LN, lymph node(s); PP, Peyer’s patch; CFSE, 5,6 carboxyfluorescein diacetate succinimidyl ester; p.i., postinfection.
Received for publication February 28, 2000. Accepted for publication August 31, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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(TGF-) production by murine CD4+ T cells. J. Exp. Med. 188:1849. is a critical component of interleukin 13-mediated protective T helper cell type 2 responses during helminth infection. J. Exp. Med. 190:953.This article has been cited by other articles:
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  C.-C. Chen, S. Louie, B. A. McCormick, W. A. Walker, and H. N. Shi Helminth-Primed Dendritic Cells Alter the Host Response to Enteric Bacterial Infection J. Immunol., January 1, 2006; 176(1): 472 - 483. [Abstract] [Full Text] [PDF]
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 C.-C. Chen, S. Louie, B. McCormick, W. A. Walker, and H. N. Shi Concurrent Infection with an Intestinal Helminth Parasite Impairs Host Resistance to Enteric Citrobacter rodentium and Enhances Citrobacter-Induced Colitis in Mice Infect. Immun., September 1, 2005; 73(9): 5468 - 5481. [Abstract] [Full Text] [PDF]
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D. W. Smith and C. Nagler-Anderson Preventing Intolerance: The Induction of Nonresponsiveness to Dietary and Microbial Antigens in the Intestinal Mucosa J. Immunol., April 1, 2005; 174(7): 3851 - 3857. [Abstract] [Full Text] [PDF]
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A. Boitelle, H. E. Scales, C. Di Lorenzo, E. Devaney, M. W. Kennedy, P. Garside, and C. E. Lawrence Investigating the Impact of Helminth Products on Immune Responsiveness Using a TCR Transgenic Adoptive Transfer System J. Immunol., July 1, 2003; 171(1): 447 - 454. [Abstract] [Full Text] [PDF]
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E. Williamson, J. M. Bilsborough, and J. L. Viney Regulation of Mucosal Dendritic Cell Function by Receptor Activator of NF-{kappa}B (RANK)/RANK Ligand Interactions: Impact on Tolerance Induction J. Immunol., October 1, 2002; 169(7): 3606 - 3612. [Abstract] [Full Text] [PDF]
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M. E. H. Bashir, P. Andersen, I. J. Fuss, H. N. Shi, and C. Nagler-Anderson An Enteric Helminth Infection Protects Against an Allergic Response to Dietary Antigen J. Immunol., September 15, 2002; 169(6): 3284 - 3292. [Abstract] [Full Text] [PDF]
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and 
,
OVA-fed; 
and •, PBS-fed.
