| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Department of Immunology, Erasmus University, Rotterdam, The Netherlands
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
) from the BB-DP rat in the MLR was
investigated. Lower numbers of splenic DC were isolated from BB-DP rats
than from control Wistar rats. In the preautoimmune phase, DC of the
BB-DP rat had a lower surface MHC class II expression (and in
preliminary data, a lower CD80 expression), ingested more bacteria, and
had a lower stimulatory potency in the syngeneic (syn)MLR as compared
with control DC. During disease development, the MHC class II
expression further decreased, and a low stimulatory activity became
evident in the allogeneic (allo)MLR. With regard to the expansion of
suppressor/regulatory T cells, a lower percentage of RT6+ T
cells but higher percentages of CD45RClow T cells were
induced by BB-DP DC in synMLR, but not in alloMLR. An increase in the
CD4/CD8 T cell ratio was observed in both the syn- and alloMLR due to a
relative weak expansion of CD8+ T cells with DC of the
BB-DP rat. Resident pM isolated from BB-DP or Wistar rats were
equally effective in suppressing the DC-driven synMLR. In conclusion,
splenic DC from the BB-DP rat have a lower accessory cell function
already at young age, before the development of disease, and expanded
different subsets of effector/suppressor T cells in vitro as compared
with those from Wistar rats. The dysfunction of DC from BB-DP rats is
likely to be caused by their relative immaturity as indicated by their
low class II and costimulatory molecule expression and relatively high
phagocytic activity. |
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The local accumulation of DC early in the development of disease suggests a pivotal role of these cells in the pathogenesis of EAD. In patients with EAD, a defective maturation and function of DC has been reported: in diabetic patients, there is a loss of MHC class II+ Langerhans cells from the epidermis at the onset of diabetes (10) and a reduction in the number of DC generated from monocytes, the precursor population in the peripheral blood (11). These monocyte-derived DC had a low stimulatory capability in autologous (auto)MLR (11), which was also described for unfractioned APC populations from the blood of diabetic patients (12, 13, 14, 15). Furthermore, monocyte-derived DC from patients with Graves’ disease and from diabetic patients showed a reduction in cellular interactions, i.e., the cells formed less homotypic clusters and clusters with T cells as compared with DC from healthy controls (11, 16).
A defective maturation and function of APC has also been
described in the NOD mouse. APC precursors in the bone marrow showed a
weakened maturational response to the growth factors IL-3 and CSF-1
(17), and splenic APC demonstrated a low stimulatory capacity in the
syngeneic (syn)MLR (18, 19). Hence, these findings are—as in the
human—indicative for a maturational defect of APC underlying the poor
stimulatory capability. Indeed, immature DC have been recognized as
relative poor T cell stimulators as compared with mature and activated
DC (20, 21). Apart from intrinsic maturational defects, a reduced
function of APC can also be caused by an increased production of
suppressive factors from other monocyte-derived cells, i.e.,
macrophages (M). M from NOD mice and BB rats are able to produce
high amounts of suppressive factors such as prostaglandins and nitric
oxide (22, 23, 24, 25, 26, 27, 28, 29).
Optimal APC function is required for appropriate stimulation of
suppressor forces (12, 18). The reduced accessory cell function in
auto/synMLR was regarded, both in the NOD mouse and in the patients, as
a sign of a faulty capability to expand suppressor/regulatory T cells.
The advantage of studying the BB rat is the presence of a well-defined
suppressor/regulatory T cell population expressing the Ag RT6. These T
cells play an important regulatory role in the pathogenesis of EAD in
the BB-DP rat (30). BB-DP rats lack RT6+ T cells and
injections of these cells into BB-DP rats prevents EAD (31).
BB-diabetic resistant (DR) rats do not develop diabetes and thyroid
autoimmunity because of the presence of RT6+ T cells, but
depletion of these cells induces EAD (32). Thus, the BB rat is an
excellent model to study putative functional defects of DC in relation
to a defective suppressor/regulatory T cell expansion. We hypothesized
that the loss of suppressor/regulatory T cells in BB-DP rats, which is
already evident at young age, might be at least in part due to a
defective function of DC before the development of EAD. Therefore, the
function of splenic DC from BB-DP rats of different ages in allogeneic
(allo)MLR (with Lewis T cells) and synMLR (with Wistar and BB-DP T
cells) was studied. In particular, we were interested in BB-DP rats of
ages 3–7 wk, i.e., before the DC infiltration into the thyroids and
pancreas islets and before EAD development. We focussed on the in vitro
expansion of RT6+ T cells within a normal T cell population
from Wistar and Lewis rats. Because CD8+ cells and
CD45RClow cells have also been associated with regulatory T
cells in the BB rat (33, 34), CD4/CD8 ratios of the in vitro expanded T
cells were calculated and CD45RC expression was examined. DC of the
BB-DP rat proved to be less capable in generating RT6+ and
CD8+ T cells, and this could be due to an immature
phenotype. For this purpose, the uptake of dextran molecules and
bacteria was studied (20, 21), together with the expression of MHC
class I and II, costimulatory and adhesion molecules, the DC marker
OX-62, and the DC/M markers, ED1 and ED7. Finally, we studied the
ability of BB-DP M to reduce accessory cell function by adding
resident peritoneal M (pM) to synMLR driven by splenic DC of the
Wistar rat.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Male and female BB-DP rats were bred in the Experimental Animal
Center of the Erasmus University (Rotterdam, The Netherlands) and
control Wistar rats were purchased from Harlan (Zeist, The
Netherlands). All rats were kept under controlled light conditions
(12/12 h light/dark cycle) throughout this study. A standard pelleted
diet (0.35 mg iodine/kg; AM-II, Hope Farms BV, Woerden, The
Netherlands) and tap water were provided ad libitum. Between 12 and 20
wk of age, 90% of our BB-DP rats developed 
-colloid Abs detected in
serum, and 70–80% of the rats became diabetic. BB-DP rats were daily
tested for glucosuria (Gluketur test sticks; Boehringer Mannheim BV,
Almere, The Netherlands). For all experiments, age matched BB and
Wistar rats were used.
Cell preparations
Splenic DC were enriched according to the method of Knight et al. (35), with slight modifications. Briefly, spleens from BB-DP and Wistar rats were minced and digested for 1 h at 37°C in RPMI 1640 medium (Life Technologies, Breda, The Netherlands) with 25 mM glutamax-1 and 25 mM HEPES (hereafter referred to as RPMI+) containing: 125 U/ml collagenase (type III; Worthington Biochemical, Freehold, NJ) and 0.1 mg/ml DNase (Boehringer Mannheim BV). The remaining tissue was teased through a 105-µm filter, and the erythrocytes were removed by lysis. Finally, the separated cells were washed and cultured in RPMI+ supplemented with 10% inactivated FCS (FCSi), penicillin (100 U/ml; Seromed, Biochrom, Berlin, Germany), and streptomycin (0.1 mg/ml; Seromed). After an overnight culture period in culture flasks (Costar Europe, Badhoevedorp, The Netherlands; 37°C, 5% CO2 incubator), DC were isolated from the nonadherent cells by using a 14.5% (w/v) Nycodenz (Nycomed Pharma As, Oslo, Norway) density gradient (800 x g for 20 min). Low-density cells were collected from the interphase and washed. This cell fraction demonstrated a dendritic morphology, a strong MHC II expression (see Results), and a weak acid phosphatase (AP) activity in 70–95% of cells in both rat strains.
Resident pM were harvested by lavage of the peritoneal cavity of
BB-DP and Wistar rats with 10 ml ice-cold PBS, pH 7.4, containing 50
U/ml heparin. Cells were washed, and, when present, erythrocytes were
lysed. The peritoneal fluid contained, in both rat strains, 50–63%
ED2+ and AP+ cells as determined on cytospin
preparations.
T cells from BB-DP, Wistar, and Lewis rats were enriched using a nylon wool column. In short, spleens were minced and teased through a 105-µm filter, and the erythrocytes were removed by lysis. Cells were washed and loaded onto a nylon wool column (3 g; Polyscience, Eppelheim, Germany) packed into a 60 ml plastic syringe. After 1 h in 5% CO2 incubator, T cells (80–90% CD5+ cells) were harvested by collecting the effluent.
MLR
For the MLR, splenic DC from BB-DP and Wistar rats were
irradiated with 2,000 rad and added at various ratios to T cells (fixed
number of 150,000 T cells/well) in flat-bottom 96-wells plates (Nunc,
Roskilde, Denmark). Subsequently, these MLR were cultured for 3 or 5
days in RPMI 1640 containing 50 mM HEPES buffer (Life Technologies),
10% FCSi, 110 µg/ml sodium pyruvate (Merck, Munich, Germany), 0.5%
(v/v) ß-mercapto-ethanol (Merck), and antibiotics. To determine the
effect of pM on accessory cell function, splenic Wistar DC were
mixed with Wistar T cells (150,000 T cells/well) at a ratio of 1:20,
and pM of either BB-DP or Wistar rats were added at a DC:M ratio
of 1:50, 1:10, and 1:2. As a control, accessory cell function of the
pM alone were determined. Next, cells were cultured for 5 days.
In the MLR, T cell proliferation was measured via tritiated thymidine ([3H]TdR) incorporation (0.5 µCi/well during the last 16 h of total culture period). Finally, cells were harvested on filter papers, and radioactivity was counted in a liquid scintillation analyzer (LKB Betaplate, Wallac, Turun, Finland). For phenotyping (see below), T cells were collected and washed in PBS containing 0.5% BSA (Sigma, Axel, The Netherlands) and 20 mM sodium azide (Sigma).
Abs
The following mAbs were used: anti-MHC class I (1:400, MRC
OX-18; Serotec, Oxford, England), anti-MHC class II conjugated to
phycoerythrin (PE) (1:400; MRC OX-6, Serotec), anti-B7-1 (1:500,
CD80; Research Diagnostics, Flanders, NJ), anti-B7-2 (1:500,
CD86; Research Diagnostics), anti-rat DC-PE (undiluted, MRC OX-62;
Serotec) anti-LFA-1 (1:10, CD11a; Serotec), anti-ICAM-1
(1:50, CD54; Serotec), anti-very late Ag-4 (VLA-4) (1:10, CD49b;
Serotec), ED1 (1:400, monocytes, M), ED7 (1:10, monocytes, M;
both ED Abs were a kind gift from Prof. Dr. C. D. Dijkstra,
Department of Cell Biology, Free University, Amsterdam, The
Netherlands), anti-CD5 (1:100, MRC OX-19; Serotec) anti-CD4
conjugated to biotin (1:4, Domain 1; Serotec), anti-CD8-PE (1:4,
MRC OX-8; Serotec), anti-CD45RC (1:100, MRC OX-22; Serotec), anti
RT6.1-FITC (1:10, 3G2, for staining Lewis T cells), and
anti-RT6.2-Biotin (1:10, GY 1/12, for staining Wistar T cells; both
anti-RT6 Abs were a kind gift from Dr. H. Groen, Department of
Histology and Cell Biology, University of Groningen, Groningen, The
Netherlands).
Cell staining
Cells were added in round-bottom 96-wells plates (Nunc) at a
concentration of 
105 cells/well and washed twice in
PBS/0.5% BSA/20 mM sodium azide. Pelleted cells were resuspended in 20
µl solution with labeled primary Abs, incubated for 10 min., and
followed by two washing steps. Using unconjugated Abs, a second step
was incorporated with rabbit anti-mouse-FITC Abs (Dako, Glustrup,
Denmark) with 1% normal rat serum. For the visualizing of
biotin-conjugated Abs, streptavidin-tricolor (Caltac Laboratories, San
Francisco, CA) was used. For cell analysis, 10,000 events were recorded
with a FACS (FACScan, Becton Dickinson, Sunnyvale, CA). Dead cells,
recognized by their uptake of propidium iodide and their specific
forward- and side-scatter pattern, were excluded from analysis. For
determination of background staining, cells were incubated with either
labeled irrelevant Abs or with secondary Abs or with
streptavidin-tricolor alone.
Uptake of FITC-dextran
The uptake of FITC-dextran (Sigma) by DC was performed according to the method by Sallusto et al. (20). In short, DC were added to round-bottom 96-wells plates (Nunc) at a concentration of 105 cells/well in RPMI+, 10% FCSi, and antibiotics. FITC-dextran was added at a final concentration of 1 mg/ml for 0, 15, 30, and 60 min. After incubation, the cells were put on ice and washed thrice. Finally, the cells were stained with OX-6-PE (Serotec) and analyzed on a FACScan.
Phagocytosis of FITC-labeled Escherichia coli
For studying the uptake of E. coli bacteria by DC, a phagotest (Becton Dickinson) was used according to the instructions of the manufacturer with some minor modification. Briefly, 20 µl precooled E. coli bacteria were added to suspension with DC (106 cells/ml in RPMI+, 10% FCSi, and antibiotics). The controls remained on ice and the phagocytosis assay was incubated for 30' and 60' at 37°C in an incubator under rotation (4 rpm). Cells were put on ice, washed, and 100 µl quenching solution was added. Cells were washed thrice and stained with OX-6-PE (Serotec) and analyzed on a FACScan.
Statistical analysis
The results are presented as means ± SEM, and statistical analysis of the data was performed with the Mann-Whitney U test.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The percentages of isolated splenic DC from BB-DP rats were significantly lower than from Wistar rats (1.0 ± 0.7% vs 1.4 ± 0.6%, p = 0.003, n = 6, age 3–20 wk). The purity of DC, relevant for performing further functional studies (see below), was measured by dendritic morphology and MHC class II expression, and did not differ for both rat strains (range 70–95%, data not shown).
In the preautoimmune phase, DC from BB-DP rats (age 3–7 wk)
demonstrated a lower surface MHC class II expression than those from
Wistar rats (mean fluorescence intensity (MFI) for Wistar and BB-DP
rats, respectively, was 1586 ± 510 and 1084 ± 386,
p = 0.02, n = 23). During development
and expression of disease (age 8–20 wk), surface MHC class II
expression further decreased on BB-DP DC (MFI for Wistar and BB-DP
rats, respectively, was 1452 ± 642 and 533 ± 124,
p = 0.006, n = 19; see also Fig. 1
). DC from both rat strains had a weak
AP activity located in a spot near the nucleus and did not differ in
the percentage of positive cells and expression of the following
surface markers: MHC class I molecules (MFI for Wistar and BB-DP rats,
respectively, was 214 ± 61 and 231 ± 57, n
= 13); adhesion molecules LFA-1 (73 ± 10 and 77 ±
13, n = 3), ICAM-1 (116 ± 19 and 114 ± 15,
n = 4), and VLA-4 (65 ± 6 and 61 ± 11,
n = 3); DC/M markers ED1 (57 ± 14 vs 55
± 17, n = 3) and ED7 (80 ± 9 vs 72 ± 9,
n = 3). DC from both rat strains were negative for CD4
and CD8. The rat DC marker OX-62 only showed a weak expression in our
hands on both Wistar DC and BB-DP DC (data not shown).
|
Uptake of dextran and E. coli bacteria
Incubation with FITC-dextran resulted in an increased uptake in time by the DC and this tended to be higher for the BB-DP rat (age 3–20 wk) after 60 min. (MFI for Wistar and BB-DP rats, respectively, was 67 ± 9 and 81 ± 14, n = 6).
During incubation with FITC-E. coli bacteria, DC showed an
increased phagocytosis and after 60 min. significantly more bacteria
were taken up by DC from the BB-DP rat (age 3–20 wk) as compared with
the Wistar rat (Table I). DC from BB-DP
rats of different ages (3–7 wk and 8–20 wk) did not differ in the
uptake of the bacteria (data not shown). In contrast with DC, >85% of
Wistar pM, which are professional phagocytes, were loaded with
FITC-E. coli bacteria after 60 min (data not shown).
|
After 3 days in syn- and alloMLR, no differences in the
proliferation of T cells were observed with DC from either BB-DP or
Wistar rats (data not shown). After 5 days, optimal proliferation was
observed, and in the synMLR with Wistar T cells a significantly lower
uptake of [3H]TdR by the T cells (low proliferation) was
found with DC from BB-DP rats compared with Wistar rats for both age
groups of 3–7 and 8–20 wk (Fig. 2).
Similar results were obtained in the synMLR with T cells from the BB-DP
rat, although the overall counts were much lower due to the reduced
proliferative capacity of these T cells (Fig. 2). Here, the number of
experiments were too low (n = 2) to perform statistical
analysis.
|
). In contrast, BB-DP DC induced significantly less proliferation of
Lewis T cells than Wistar DC in the age group of 8–20 wk (Fig. 2).
With DC isolated from lymph nodes, similar results in the alloMLR for
the different strains were observed (data not shown). Expansion of T cell subsets in DC-driven MLR
No differences in the expansion of T cell subsets were observed on
day 3 in the syn- and alloMLR (data not shown). On day 5 of these MLR,
a significant increase in the CD4/CD8 ratio with DC from the BB-DP rat
was observed (Table II). In the synMLR
with T cells from the BB-DP rat, similar observations were made: the
CD4/CD8 ratio with DC from Wistar and BB-DP rat was 0.7 ± 0.2 and
2.1 ± 0.5, respectively (n = 2). In synMLR,
significant lower percentages of CD45RC+ and
RT6+ T cells were induced by BB-DP DC as compared with
Wistar DC. This was not observed in the alloMLR using Lewis T cells
(Table II). An example of CD4, CD8, and RT6 expression on Wistar T
cells cultured with DC from Wistar and BB-DP rats is given in Fig. 3.
|
|
Different concentrations of pM were cocultured with DC and T
cells from the Wistar rat to study their effect in the synMLR. Culture
of DC together with T cells (DC:T cell ratio of 1:20) gave >40,000
counts, whereas culture with pM alone with T cells resulted in a
much lower T cell proliferation (Fig. 4).
The addition of pM in the synMLR at a DC:M ratio of 1:50 and
1:10, resulted in a similar inhibition of T cell proliferation for both
rat strains. No significant inhibitory effect on the DC-driven MLR was
found using lower numbers of pM (DC:M ratio of 1:2; Fig. 4).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
This study demonstrates that splenic DC from the BB-DP rat show signs of immaturity as indicated by the following. First, a low expression of surface MHC class II molecules and also of CD80 together with an enhanced uptake of bacteria is shown. These abnormalities occurred already in the preautoimmune period (3–7 wk of age) and, with respect to MHC class II expression, further decreased in the autoimmune period (8–20 wk of age). Although not analyzed in this study, determination of intracellular class II expression would help to substantiate immaturity by differences in the distribution of MHC class II molecules (20, 21). Second, a lower T cell stimulatory capacity in the synMLR for all periods studied, and also in the alloMLR in the autoimmune period is shown.
In addition, BB-DP splenic DC were also less in number upon isolation, which may be taken as a sign for a reduced generation/maturation of DC from their precursors. Further studies with monocyte-derived and bone marrow-derived DC and the effect of maturation stimuli like LPS are planned, as these would best answer the issues regarding defective DC maturation and function in the BB-DP rat.
The described DC dysfunction before the onset of autoimmunity, i.e., before the accumulation of DC in the thyroids and pancreas islets of the BB-DP rat, is suggestive for an innate defect, which may form the basis for the development of EAD and aggravates during aging/disease. Interestingly, similar impaired function of DC has recently been described in the preclinical phase of human diabetes (43).
How do our findings relate to other studies on DC and APC function in
the BB-DP rat? In a similar study, but using another subline of BB-DP
rats, Tafuri et al. (44) reported the same level of MHC class II
expression and accessory cell activity of splenic DC from BB-DP and
Wistar rats. However, these authors only studied young rats (age 3–7
wk) and tested their DC in alloMLR but not in synMLR. Also, we could
not demonstrate DC defects in alloMLR at such young age. In other
studies, using nonpurified APC populations (45, 46), the data are
sometimes confusing, and comparison with our data is difficult. In
contrast with our findings, Bellgrau et al. (45) reported, using
unfractioned lymph node cells, a normal synMLR, although these APC
induced a lower response in the alloMLR. It is not clear from their
report whether experiments were performed with either young or old
rats. Also Elder et al. (46) described that alloMLR driven by
unseparated BB-DP spleen cells was lower, as compared with those driven
by Wistar rat APC. But, higher T cell stimulations with BB-DP DC could
be obtained depending on the conditions of the experiment and the
concentration of the stimulator cells (46). In addition, it must be
noted that the unfractioned APC populations used in the above-cited
studies may contain M, which are likely to affect the proliferation
of the responder T cell population (see present data and Refs. 22, 23,
and 28).
Both in diabetic patients and in NOD mice, the failure of DC to
sufficiently stimulate T cells in auto/synMLR has been considered as
reflecting a decreased capability of these cells to expand
regulatory/suppressor T cells (12, 18). Indeed, in the present study,
the reduced accessory cell function of the immature splenic DC from the
BB-DP rat resulted in the following conditions. First, a low
generation of RT6+ T cells in synMLR is found. The
RT6+ population has been reported to contain cells that
protect BB rats against the development of EAD (31, 47). RT6 is a
glycosyl-phosphatidylinositol-linked surface molecule present on
70% of mature peripheral T cells (48, 49). The exact mechanism of
protection is not clear but the RT6 molecule generates an activation
signal, which may influence T cell function (50). Second, an increased
CD4/CD8 ratio, due to a primary expansion of CD4+ T cells,
not only in synMLR (when BB-DP T cells or Wistar T cells were used as
responder cells), but also in allogeneic responses (Lewis T cells), is
found.
Taken together our data showing a low in vitro expansion of RT6+ T cells and of CD8+ T cells (in relation to CD4+ T cells) by immature BB-DP splenic DC supports our hypothesis that DC with a defective T cell stimulatory function, which can also be found in human type 1 diabetes, have an inability to generate sufficient suppressor forces (see Introduction). However, some words of caution are necessary.
First, in our experiments with BB-DP DC, we have not made a comparison with splenic DC and/or T cells of BB-DR rats to link the maturation defect of DC to EAD development and to avoid minor histocompatability differences in synMLR. A DR variant of our BB-DP Rotterdam strain does not exist, but in a few preliminary experiments we compared accessory cell function of splenic DC of BB-DP and BB-DR rats of the Worcester strain. Interestingly, DC of both Worcester strains did not show the defects as observed with DC of the Rotterdam strain. In this respect, it is of note that our BB-DP strain has a relatively high prevalence of thyroiditis (51, 52). We did find other signs of immaturity of DC from the Worcester sublines: a lower homotypic cluster capability as compared with the Wistar rat, and the BB-DP rat was more affected than the BB-DR rat. At present, these data are subject for further study.
Second, suppressor activity has not only been recognized in the RT6+ and CD8+ T cell population, but also in a population of CD45RClow T cells (33, 47), and, within the CD4+ T cells, these cells have been recognized as Th2-like cells, producing IL-4 (33). In our study, DC from the BB-DP rat induced not less, but more of these "suppressor" CD45RClow T cells in vitro as compared with the Wistar rat. Furthermore, the CD45RChigh/CD45RClow ratios of both CD4+ and CD8+ T cell subsets did not differ in the MLR for both rat strains using either Wistar or Lewis T cells (data not shown). On the other hand, not only CD45RClow T cells may act as suppressor/regulatory cells; also CD4+ CD45RChigh Th1-like cells can act as regulator cells suppressing graft rejection in the Lewis rat (53). Hence, protective cell populations other than CD8+, RT6+, or CD45RClow T cells are likely present in the BB-DP rat. In addition, transplantation of thymic tissue from DR rats into BB-DP rats prevented the development of diabetes without restoration of the RT6+ T cell population (54). Nagata et al. (55) reported the prevention of diabetes in BB-DP rats with a CD4+CD8- and RT6- T cell line, but it is not clear whether this cell line may obtain the RT6 phenotype after transfer. The present studied showed that DC from the BB-DP rat expanded different subsets of effector and suppressor/regulatory T cell populations in vitro as compared with Wistar control rats. The question remains whether BB-DP DC are indeed less capable of expanding T cells that suppress EAD in vivo. This question may be solved by studying the effect of in vitro expanded T cells (in synMLR) on the development of EAD after transfer in BB-DP rats.
The mechanisms of the defective maturation of splenic DC from the BB-DP
rat could hypothetically reside on the level of the precursor in the
bone marrow, as observed for NOD APC (17), or on other levels, such as
a lack of maturation signals. The latter is illustrated by the
transition from blood monocyte to full mature DC induced by
lymphocyte-derived cytokines and lymphocytic contacts (CD40 ligation).
Already at young age, the BB-DP rat becomes severely lymphopenic (46, 56, 57), resulting in a strong reduction in the number of especially
CD8+, CD45RC+, and RT6+ T cells
(58, 59, 60). This reduction is at least partly due to genetically encoded
defects in intrathymic T cell maturation (60, 61, 62). Hence, the T
lymphopenia of the BB-DP rat could be primary and induce a faulty
cross-talk between lymphocytes and DC leading to a poor development and
maturation of both T cells and DC. The observation that DC from BB-DP
rats (age 3–7 wk) stimulated the alloMLR equivalently to Wistar DC
(see Fig. 2), suggests that the maturation and function of BB-DP DC can
be restored with "normal" T cells. Indeed, preliminary data showed
a restoration of homotypic cluster formation of BB-DP DC after
incubation with factors produced by Con A-stimulated T cells. Whether
MHC class II and CD80 expression is also restored is now under
investigation. We favor the idea that the defective cross-talk between
DC and T cells of the BB-DP rat resides on both the intra- and
extrathymic level. Extrathymic defects in T cell maturation in the
BB-DP rat have been suggested before. With respect to the defective
intrathymic T cell maturation in the BB-DP rat, a role for thymic APC
has also been recognized: introduction of a bone marrow-derived thymic
APC fraction from DR donors into the thymus of the BB-DP rat restored
the T cell proliferative function and prevented disease. However, no
restoration of the T lymphopenia and RT6 expression was achieved (54).
M-derived factors may also play a role in the defective DC
maturation in BB-DP rats. One of the major maturation-inducing
cytokines for DC, TNF- (21, 63, 64), is produced in much lower
quantities by pM of BB-DP rats than by BB-DR rats (65). Treatment
with or induction of TNF- with CFA decreased the incidence of
diabetes in the BB-DP rat (66, 67). One could speculate whether such
treatment would induce a full maturation of BB-DP DC and subsequently
up-regulate their accessory cell function in MLR. However, preliminary
data showed that culture of splenic BB-DP DC in the presence of a
single dose of rat or human TNF- and/or combined with another
maturation-inducing cytokine human granulocyte-M CSF, had no effect
on their accessory cell function in the MLR (data not shown).
Defective function of BB-DP splenic DC could also be due to an
excessive production of suppressive factors by BB-DP M, but pM of
BB-DP and Wistar rats had similar inhibitory activities on the synMLR
driven by splenic DC of Wistar rats. These data are in contrast to some
previous reports demonstrating a stronger suppression of APC by M of
the BB-DP rat and an enhanced production of suppressive factors (22, 23, 27, 28). However, other data show the opposite: the addition of
BB-DP M enhanced the stimulatory activity of DC (44). These
conflicting data may be due to the usage of different populations of
M (either isolated from the spleen or the peritoneum) and
differences in the activation state of these M (e.g., exudate vs
resident pM, the latter being strongly affected by the isolation
procedure). In our study, cells were harvested from the peritoneum
without further purification steps, like adherence or gradient
centrifugation (22, 23, 27, 28), ensuring a minimal degree of
activation. If indeed BB-DP M have a stronger suppressive effect
(22, 23, 27, 28), this would further limit the already depressed BB-DP
DC activity.
In conclusion, the present study gives evidence that splenic DC from the BB-DP rat have an immature phenotype and an impaired function before the development of endocrine autoimmunity. Furthermore, these defective DC induce a different expansion of T cell subsets in vitro and the question remains whether these APC defects also lead to a faulty in vivo induction of tolerance/suppression. The putative mechanisms underlying this altered maturation (defective bone marrow factors, T cell defects) remain to be elucidated.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
2 Address correspondence and reprint requests to Dr. Frans G. A. Delemarre, Department of Immunology, Laboratory Ee 838, Faculty of Medicine, Erasmus University Rotterdam, P.O. Box 1738, 3000 DR Rotterdam, The Netherlands. E-mail:
3 Abbreviations used in this paper: EAD, endocrine autoimmune disease; BB-DP, biobreeding-diabetic prone; NOD, nonobese diabetic; DC, dendritic cell(s); auto, autologous; syn, syngeneic; M, macrophage; DR, diabetic resistant; allo, allogeneic; pM, peritoneal M; RPMI+, RPMI 1640 with glutamax-1 and HEPES; FCSi, inactivated FCS; AP, acid phosphatase; PE, phycoerythrin; MFI, mean fluorescence intensity.
Received for publication June 16, 1998. Accepted for publication October 16, 1998.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and IL-6. J. Endocrinol. 140:503.. J. Exp. Med. 179:109.
. J. Immunol. 145:1395.[Abstract]
This article has been cited by other articles:
|
|
 |
|
  V. Sommandas, E. A. Rutledge, B. Van Yserloo, J. Fuller, A. Lernmark, and H. A. Drexhage Low-density cells isolated from the rat thymus resemble branched cortical macrophages and have a reduced capability of rescuing double-positive thymocytes from apoptosis in the BB-DP rat J. Leukoc. Biol., October 1, 2007; 82(4): 869 - 876. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Nikolic, M. Bunk, H. A. Drexhage, and P. J. M. Leenen Bone Marrow Precursors of Nonobese Diabetic Mice Develop into Defective Macrophage-Like Dendritic Cells In Vitro J. Immunol., October 1, 2004; 173(7): 4342 - 4351. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. Bouma, W. K. Lam-Tse, A. F. Wierenga-Wolf, H. A. Drexhage, and M. A. Versnel Increased Serum Levels of MRP-8/14 in Type 1 Diabetes Induce an Increased Expression of CD11b and an Enhanced Adhesion of Circulating Monocytes to Fibronectin Diabetes, August 1, 2004; 53(8): 1979 - 1986. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Poudrier, X. Weng, D. G. Kay, Z. Hanna, and P. Jolicoeur The AIDS-Like Disease of CD4C/Human Immunodeficiency Virus Transgenic Mice Is Associated with Accumulation of Immature CD11bHi Dendritic Cells J. Virol., November 1, 2003; 77(21): 11733 - 11744. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Yan, T. Devos, L. Yu, G. Xia, O. Rutgeerts, J. Goebels, C. Segers, Y. Lin, M. Vandeputte, and M. Waer Pathogenesis of Autoimmunity After Xenogeneic Thymus Transplantation J. Immunol., June 15, 2003; 170(12): 5936 - 5946. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
F. G. A. Delemarre, P. G. Hoogeveen, M. de Haan-Meulman, P. J. Simons, and H. A. Drexhage Homotypic cluster formation of dendritic cells, a close correlate of their state of maturation. Defects in the biobreeding diabetes-prone rat J. Leukoc. Biol., March 1, 2001; 69(3): 373 - 380. [Abstract] [Full Text]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
) and Wistar rats (
) (MLR with Wistar T
cells: n = 6–7; *, p < 0.03
compared with Wistar for all DC: T cell ratios used and for all age
groups; MLR with BB T cells: n = 2; MLR with Lewis
T cells: n = 9–10; *, p <
0.03 compared with Wistar for all DC: T cell ratios in age group 8–20
wk).
