International Immunology Advance Access originally published online on January 13, 2006
International Immunology 2006 18(3):415-423; doi:10.1093/intimm/dxh382
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The proliferative response of CD4 T cells to steady-state CD8+ dendritic cells is restricted by post-activation death
The Walter and Eliza Hall Institute of Medical Research, 1G Royal Parade, Parkville, Victoria 3050, Australia
Correspondence to: K. Shortman; E-mail: shortman{at}wehi.edu.au
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Abstract |
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CD8+ splenic dendritic cells (DCs) from steady-state mice are less effective than the CD8– DC subset in their capacity to stimulate CD4 T cell proliferation in culture. However, we found that the two DC subtypes were equally potent at activating CD4 T cells, based on up-regulation of CD69 and CD25 expression. Also, we found no difference in the rate of T cell death prior to entry into the first division. We then tracked carboxyfluorescein diacetate succinimidyl ester-labeled T cells and employed a quantitative model to assess in detail the CD4 T cell expansion process in response to stimulation with CD8+ or with CD8– DCs. The time required for most T cells to replicate their DNA prior to the first division was similar in both DC cultures. However, progression of the CD4 T cell population through subsequent divisions was reduced in CD8+ DCs compared with CD8– DC culture. This was associated with an increased loss of viable T cells at each division. Post-activation, division-associated T cell death is therefore a major factor in the reduced response of CD4 T cells to CD8+ DCs.
Keywords: dendritic cells, modelling, T cells, T cell proliferation
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Introduction |
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Dendritic cells (DCs) are potent antigen-presenting cells with the unique capacity to initiate primary T cell responses (1). In their activated state, DCs instruct T cells to develop into effector cells, and regulate the nature of the immune response by biasing the activated T cells toward a Th1, Th2 or regulatory T development. In steady-state, quiescent DCs also have the capacity to induce and maintain tolerance to self-components, both in the thymus and in the periphery (2–5). However, DCs consist of a series of discrete subtypes, and these make specialized contributions to these general DC functions.
Four DC subtypes can be isolated from the spleen of non-infected steady-state mice. These include plasmacytoid pre-DCs and three subsets of conventional DCs, distinguished by surface phenotype as CD8+4– (CD8+), CD8–4+ (CD4+) and CD8–4– double negative (DN); the latter two are often grouped together as CD8– DCs. These conventional DCs share many properties, including dendritic morphology and the expression of a range of molecules required for the activation of naive T cells. However, they differ in their capacity to regulate immune responses. The CD8+ DCs, when fully activated, are the most potent producers of IL-12 (6–8), and strongly bias CD4 T cells toward an inflammatory Th1 response with extensive IFN
production. In contrast, studies from this laboratory have shown that freshly isolated, quiescent CD8+ DCs have the lowest capacity to stimulate CD4 T cell proliferation and IFN production in culture, compared with either of the CD8– DC subtypes (9, 10). The differential in the response to steady-state CD8+ compared with CD8– DCs was shown to be independent of antigen and DC dose, or DC survival in culture. It appeared to depend on the initial surface interactions between the DCs and the CD4 T cells. However, the key surface molecules responsible for this difference in response have not yet been identified.
In this study, we shift focus from the differences between the quiescent DC subsets themselves to the differences in the proliferative response of the CD4 T cells once activated. Differences in T cell proliferation in response to the different DC subtypes might reflect differences in (i) the number of T cells receiving an activation signal, (ii) the time taken for division to commence, (iii) the rate of T cell division once activated or (iv) the rate of T cell death before or after activation. Some previous experiments from this laboratory, using an allogenic system, indicated there was a higher level of apoptotic CD4 T cells in cultures stimulated with CD8+ DCs compared with CD8– DCs, pointing to T cell death as one important factor in the differential proliferative response (9). Using an antigen-specific system with a uniform population of TCR-transgenic T cells and a carboxyfluorescein diacetate succinimidyl ester (CFSE)-based system of analysis of the T cell expansion process (11–13), we now demonstrate that the differences in the CD4 T cell response are only apparent after cell division commences, and that an increased rate of T cell death during each division cycle is an important factor limiting the response to CD8+ DCs.
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Methods |
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Animals
The normal source of DCs was 5- to 10-week-old female C57BL/6J Wehi mice bred under specific pathogen-free conditions at the Walter and Eliza Hall Institute animal facility (Melbourne, Victoria, Australia). In antigen-specific experiments, the source of CD4 T cells was 6- to 10-week-old ovalbumin (OVA)-specific, MHC class II-restricted, TCR-transgenic mice (OT-II mice) (14). In case of allogeneic responses, CD4 T cells from 6- to 10-weeks-old CBA/J mice were used.
Reagents and antibodies
CFSE was obtained from Molecular Probes (Eugene, OR, USA). Demecolcine was purchased from Sigma–Aldrich (St Louis, MO, USA). The fluorescent mAbs used for analyzing CD4 T cell activation were biotin-conjugated anti-CD69 (H1.2F3) and anti-CD25 (PC61). These antibodies were purified and labeled as published elsewhere (15). They were revealed using streptavidin–Cy5.
DC isolation
The procedure was as described in detail elsewhere (16) and involved collagenase/DNase digestion of spleen fragments, selection of light density cells and then immunomagnetic bead depletion of non-DCs. This preparation, 70–80% DCs, was then stained for CD11c, CD4 and CD8 markers and positively sorted for CD11c+ DCs of the appropriate subtype, on a MoFlo (Cytomation, Fort Collins, CO, USA) or a FacsDiVa instrument (BD Bioscience, Mountain View, CA, USA). In most experiments, the CD4 and DN DC subsets which induce similar high T cell proliferation were pooled and referred to as CD8– DCs.
Isolation of CD4 T cells
Subcutaneous and mesenteric lymph nodes from two to five mice were pooled and CD4 T cells purified by immunomagnetic bead depletion using anti-B220 (RA3-6B2), anti-GR-1 (RB6-8C5), anti-erythrocytes (TER-119), anti-Mac1 (M1/70), anti-CD25 (PC61) and anti-CD8
(53-6.7). In some experiments, the CD4 T cell preparation was washed in PBS/0.1% BSA, made to 10 x 106 cells ml–1, labeled with 10 µM CFSE for 10 min at 37°C with frequent agitation, then washed once before being used in proliferation assays.
Proliferative responses of CD4 T cells in culture
Purified CD4 T cells (50 000) were cultured together with syngeneic DCs (0–5000). Replicate cultures were in 200 µl medium in the U-bottom wells of 96-well culture plates. The culture medium was modified RPMI 1640 containing 10% FCS. When OT-II T cells were used with C57BL/6 DCs, synthetic OVA 323–339 peptide(0–100 ng ml–1, Auspep, Melbourne, Australia) was added to the culture medium. The cultures were then harvested on day 3 or according to a time course. In some experiments where stated, demecolcine at 20 ng ml–1 was added to the culture medium to block T cell division. In all cases, CD4 T cell proliferation was determined after the indicated time of culture, using either flow cytometry to count live CFSE-labeled cells which had divided (17) or by a 6-h pulse of [3H]thymidine ([3H]TdR) followed by cell harvesting and measurement of radioactivity incorporated into cellular DNA (9). For CFSE experiments, a known number of Calibrite beads (Becton Dickinson) were added to each culture prior to harvest. These beads can be distinguished from cells by flow cytometry and allow the number of cells in culture to be determined by a ratio method (12). Propidium iodide (PI) was also added to a final concentration of 10 µg ml–1 to allow live cells to be gated. For quantitative analysis, cell numbers in each division were converted to starting cell ‘cohorts’ by dividing by 2division number as described (11). Normal distributions were fitted to cohort numbers in each division, excluding division 0, using a Microsoft Excel fitting program. Mean division numbers and total area of the fitted distributions were determined and plotted to estimate the division times and rates of cell death per division.
Cultured CD4 T cell transfer in vivo
Ly5.2 OT-II CD4 T cells cultured for 3 days in the presence of CD8+ DCs (CD8+ primed OT-II) or CD8– DCs (CD8– primed OT-II) and 100 ng ml–1 OVA peptides were washed and dead cells removed by density centrifugation. Primed OT-II (2 x 106) were then injected intravenously into Ly5.1 recipient mice. Spleens from recipient animals were assessed for the presence of primed OT-II cells 19–46 h later, using Ly5.2 and TcR V2 markers and flow cytometric analysis.
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Results |
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CD8+ DCs induce restricted CD4 T cell proliferation compared with other DC subsets
To extend earlier observations where only CD8+ and DN DCs were compared (9), we first assessed the capacity of the three splenic DC subsets from C57BL/6 mice to induce allogeneic CBA CD4 T cell proliferation, using a [3H]TdR incorporation readout. As shown in Fig. 1A, the earlier observations were confirmed and the response to both the DN and CD4+ DCs was shown to be higher than the response of CD8+ DCs. The CD4+ and DN DC subtypes were then pooled and referred as CD8– DCs in subsequent experiments, for comparison with the CD8+ DCs.
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To obtain more quantitative data, we shifted to an antigen-specific response system in which OVA-specific TCR-transgenic CD4 T cells were used (referred to as OT-II cells). These CD4 T cells were depleted of CD25-expressing cells to avoid effects of activated or regulatory T cells. The OT-II cells were then cultured with CD8+ or CD8– DCs in the presence of OVA peptide in the culture medium. The proliferation readout was the count of CFSE-labeled OT-II cells which had divided. As shown in Fig. 1B, this gave a result in accordance with the previous system, the CD8+ DCs inducing a 2- to 3-fold lower CD4 T cell response than the CD8– DC subtype. The actual time that proliferation commenced varied from one experiment to another, but the reduced T cell response to CD8+ DCs was consistent. This result was unaltered when exogenous IL-2 was added to the culture medium, indicating this was not a limiting factor.
CD8+ and CD8– DCs induce a similar CD4 T cell activation
The relatively poor stimulatory capacity of CD8+ DCs compared with CD8– DCs might have been due to a reduced capacity of the CD8+ DC subtype to activate the CD4 T cells. We therefore looked at the expression of the early T cell activation marker CD69 and the IL-2R alpha chain CD25, on OT-II cells during a time course experiment (Fig. 2). OT-II cells labeled with CFSE were cultured with each DC subset at a ratio of 10 : 1 and proliferation, as well as expression of CD69 and CD25, were followed over the early stages of culture. After 2 days, OT-II cell proliferation was lower in the CD8+ DC culture (Fig. 2A), confirming previous results. However, the proportion of OT-II cells expressing CD69 (Fig. 2B) and CD25 (Fig. 2C) in the CD8+ and CD8– DC cultures was similar during the time course of the experiment. OT-II cells cultured in medium containing OVA peptide only did not proliferate, nor did they show significant expression of CD69 or CD25. Thus, despite a lower T cell proliferative response, all T cells in culture with CD8+ DCs received a proliferative stimulus sufficient to activate CD69 and CD25 expressions.
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Undivided CD4 T cells in culture with CD8+ or CD8– DCs die at the same rate
It was possible that CD8+ DCs induced a form of activation-dependent T cell death before the cell proliferation started, so lowering the number of T cells available to enter division. Accordingly, we analyzed by flow cytometry the number of viable CD4 T cells during a time course experiment. We focused our attention on the early time points of the culture, when CD4 T cells are activated but still undivided. The number of live (PI exclusion) CSFE-labeled OT-II cells cultured with OVA peptide alone, or with OVA peptide together with CD8+ or CD8– DCs, all decreased at a similar rate between 0 and 24 h of culture (Fig. 3A and its enlargement). This suggested that DC subsets did not influence the rate of death of OT-II cells before they entered in division. However, whereas number of live OT-II cultured without DCs continued to decrease after 24 h, the number of live OT-II cells in the culture with the two DC subtypes and OVA peptide began to increase at roughly the same time. By 66 h, OT-II cells in culture with CD8– DCs had proliferated more than those with CD8+ DCs. A similar rate of death of the undivided OT-II cells was observed in all cultures, in the presence and even in absence of OVA peptide (Fig. 3B). Thus, a differential in the undivided T cell death rate was not involved in the differential proliferation induced by the two DC subsets.
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CD4 T cells stimulated by CD8+ or CD8– DCs plus OVA peptide enter the first division cycle with similar kinetics
The reduced T cell proliferative response to CD8+ DCs might have been the result of a delay in entry into division, although this was not apparent in Fig. 3. We therefore determined accurately the time required for the OT-II cells to enter the first division after stimulation with CD8+ or CD8– DCs plus OVA peptide. For this, we used a previously described system employing the cell cycle inhibitor demecolcine, which inhibits cells in metaphase of the cell cycle (18, 13). Thus, the cells are able to replicate their DNA but do not undergo cell division. As a consequence, DNA replication is only possible from cells entering their first division. Cultures containing demecolcine were pulsed for 1 h with [3H]TdR at varying times to detect DNA replication. As shown in Fig. 4, OT-II cells cultured in the presence of either CD8+ or CD8– DCs subset initially incorporated a similar level of [3H]TdR, suggesting that the number of cells able to initiate DNA synthesis as part of the first division cycle was similar in both cultures. Furthermore, the peak time of maximal [3H]TdR incorporation was very similar in both CD8+ and CD8– DC cultures, although it varied a little from one experiment to another. The mode of the time to first division was 55.7 ± 2.7 h in the CD8+ DC cultures, and 58.2 ± 2.7 h in the CD8– DC cultures. However, after this period, the incorporation of [3H]TdR was a little higher in the CD8– DC culture, suggesting that this subset recruited a few more of the ‘late entry’ OT-II cells.
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Quantitative effects of DC subsets on CD4 T cell proliferation kinetics
The foregoing results indicated little difference in the ability of CD8+ versus CD8– DCs to initiate activation events leading up to the entry of the CD4 T cells into their first cell division cycle. Thus, the proliferation differences we observe must be occurring later, either by affecting T cell viability or division times in subsequent divisions. To explore these possibilities, we applied quantitative methods previously described (11, 13). CFSE-labeled OT-II cells were stimulated with CD8+ or CD8– DCs in the presence of OVA peptide and harvested and analyzed by flow cytometry at various times after 2 days of culture. To assist the quantitation, a known number of beads were added to each sample before harvest (12) and PI was added to assist in monitoring the live cells. An example of two time samples is given in Fig. 5. In agreement with previous observations (19), cells divided asynchronously, so that at any time point, cells could be found spread across a range of divisions in both cultures. Despite the fact that OT-II cells in the CD8+ and CD8– DC cultures take a similar time to begin the first division cycle and a similar number initiate DNA replication, at 48 h more cells were found consistently in the undivided peak in the CD8+ DC cultures. However, a proportion of the cells in culture with CD8+ DCs had progressed through two divisions. Interestingly, although at the same time some OT-II cells in culture with CD8– DCs had progressed through same number of divisions, most cells were found in division 1 rather than in the undivided peak. Moreover, the total number of cells in divisions 1 and 2 were higher in the CD8– DC culture than in the CD8+ DC culture. After 66 h of culture, some OT-II cells in both DC cultures had progressed through five divisions, with most cells being found in divisions 3 and 4. However, the total number of cells in these divisions was reduced in the CD8+ DC culture, confirming a lower proliferation of OT-II cells. Since the time required to initiate DNA replication seemed to be similar in both DC cultures, the reduced progression through divisions observed in the CD8+ DC cultures could be the result of a longer cell cycle time, or due to a progressive loss of dividing cells or a combination of both.
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Modelling CD4 T cell proliferation
The modelling software only enabled us to perform accurate analysis once T cells had been through four divisions. Thus, early time points (48 h) were not used in this analysis. From CFSE data similar to those in Fig. 5, but from different experiments with more late time points, we were able to determine the number of viable (PI excluding) cells in each division at each time point of the experiment, by reference to the added beads. These numbers were converted to cell cohorts by dividing by 2division number to reveal how many cells of the original population were now found in each division (11). In Fig. 6A, the number of cohorts are plotted as a function of the division number. Inspection of the time series shows that the number of divisions reached by cells stimulated with CD8– DCs was consistently greater than for the CD8+-stimulated group when harvested at the same time. It is also clear that the total number of cohorts (the sum of cells from all divisions) was diminishing more rapidly from the CD8+ DC-stimulated group, suggesting greater levels of cell death per division. These conclusions are confirmed by plotting the mean division number for each cohort against the time of harvest (Fig. 6B) as previously described (11). A linear regression analysis was performed as described previously (13). The slope of the resulting lines gives an indication of the average subsequent division time. As shown in Fig. 6B, the slope of the line obtained in the CD8+ DC culture was lower than the one in the CD8– DC culture, suggesting that, as a population, OT-II cells in culture with CD8+ DCs were slower to progress through divisions. The calculated apparent division times for the two cultures were 9 h in the CD8– DC group and 12 h in the CD8+ DC group. However, this value depends on both the actual cell cycle time of the individual cells and the frequency with which cells are lost by death. To illustrate the rate of loss of T cells as they progress through each division, we fitted normal distributions to the cohort plots shown in Fig. 6A [excluding division 0 as described in (11)] and ascertained the area under each for all harvest times. These areas gave the total viable cell numbers. A plot of the viable cell cohort number against mean division number at the time of harvest is shown in Fig. 6C. This plot clearly shows that the loss of cells associated with each division was much greater in the CD8+ DC culture than with the CD8– DC culture. This suggested that the proportion of OT-II cells that die while traversing each consecutive cell division was greater when the cells were in culture with CD8+ DCs.
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To support this finding, we analyzed the proportion of dead (PI positive) OT-II cells in the CD8+ and CD8– DC cultures after 3 days of culture. We found that 61 ± 9% of OT-II cells in culture with CD8+ DCs were dead cells whereas only 37 ± 5% of the cells were dead in the CD8– DC cultures. There was also substantial cell debris in the cultures, presumably from dead cells, but this was gated out during the analysis and could not be quantified. Co-staining using CFSE and PI or annexin-V was performed in an attempt to measure the number of dead or apoptotic cells associated with each division, but this was unsuccessful. We assumed that cell death occurred rapidly and dying cells quickly lost their CFSE label.
Do the recovered expanded CD4 T cells differ in survival in vivo?
It was possible that the surviving CD4 T cells recovered from the culture with CD8+ DCs would be fated to die after further expansion, and therefore would not survive in vivo. To test this, OT-II cells were harvested from day 3 of CD8+ or CD8– DC cultures, dead cells removed and the cells transferred intravenously into Ly 5 disparate recipients. Spleens were assessed 19–46 h later. As shown in Fig. 7, there were no significant differences in the number of OT-II cells recovered when equal numbers of viable OT-II cells primed by CD8+ or CD8– DCs were transferred into recipient animals.
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The recovered viable T cells were also tested for CD5 expression, a marker of a novel form of unresponsive T cells (20), as well as for CD25 expression. All cells expressed CD25, as expected from their activated state (Fig. 2C). However, only marginal CD5 staining was observed with no consistent difference between OT-II cells stimulated by the CD8+ or CD8– DCs (data not shown). Overall, we concluded that there was no obvious difference in surface phenotype between in those CD4 T cells which survived after stimulation in culture with the two DC subtypes.
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Discussion |
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The results presented here confirm our previous conclusions that, despite many similarities in surface levels of MHC class II and co-stimulatory molecules, the CD8+ DC subset from steady-state mice induces a limited proliferative response in CD4 T cells, compared with both the CD8– DC subtypes (9, 10). By focussing on the fate of the responding T cells in an antigen-specific TCR-transgenic T cell system, we have obtained some insight into the basis of this difference in response. The use of CFSE-labeled T cells provided us with detailed data on CD4 T cell division and proliferation. To analyze the data, we used a stochastic mathematical model, previously only applied to CD4 T cells stimulated in culture with anti-CD3 (13, 19).
The first question we asked was whether there was any difference between the CD8+ and CD8– DC subsets in providing an activation signal to the CD4 T cells in the cultures. Based on the expression of the activation markers CD69 and CD25, both DC subsets were equally potent and most T cells in both cultures were activated early in culture.
The second question was whether CD8+ DCs caused direct CD4 T cell killing or enhanced CD4 T cell death prior to division. Detailed viable cell counting over the first 48 h showed no sign of this, CD4 T cells in both DC cultures showing the same exponential death rate as CD4 T cells cultured alone, with or without the antigen.
The third question was whether stimulation with the different DCs caused any difference in the time taken by CD4 T cells to enter the first division cycle. Entry into the division cycle is a stochastic event, but interestingly, the time that most T cells took to begin DNA synthesis and so enter the first division was not significantly different following stimulation by either DC subset.
The fourth question was whether the number of CD4 T cells capable of proliferating in response to each DC subtype differed. From the demecolcine experiment, it seemed the time at which cells initiate DNA replication and the number of cells able to do so was similar for CD4 T cells stimulated with CD8+ DCs or with CD8– DCs. This was in line with the absence of any difference in the proportion of T cells activated and the absence of any difference in the initial pre-division death rate. However, from the CFSE profiles, more cells stayed undivided in the CD8+ DC culture. Thus, we have to consider the time to begin replicating DNA and the time to divide into two daughter cells as separate issues. It appears that the CD4 T cells replicate DNA at the same time in both DC cultures, but many CD4 T cells in the CD8+ DC cultures are either delayed in completing division or are held up and destined to die.
The difference in the response of CD4 T cells to CD8+ versus CD8– DCs became apparent and could be more accurately analyzed after T cell proliferation was initiated. Over the same time period, cultures stimulated with CD8+ DCs showed a reduced progression through successive cell divisions and fewer cells were produced. This could have two causes: a longer cell cycle time or a continuous loss of cells by death in each division. We have not determined the exact relative contribution of these two parameters. However, we have demonstrated that when CD8+ DCs are the stimulating cells, a larger fraction of the product of each T cell division was lost, presumably by cell death, before they reached the next division. An increase in cell death associated with cell division was therefore a major factor limiting the response of CD4 T cells to CD8+ DCs.
An earlier study from this laboratory using an allogenic system reported an increase in apoptotic T cell blasts when CD4 T cells were stimulated with CD8+ DCs (9). We have also detected an increase in the proportion of post-division dead cells using an allogenic system. In the present study using antigen-specific OT-II CD4 T cells, there was clearly a higher proportion of dead cells in the cultures with CD8+ DCs. However, it was difficult to detect and quantitate the differences in the levels of dead cells associated with successive divisions in our OT-II system because of the large number of pre-division dead cells in all cultures and because of the loss of the CFSE label and the disintegration of dead cells. However, our calculations based on counts of the live cell component indicated an increased loss compatible with cell death, in agreement with the earlier studies.
The increased rate of dividing CD4 T cell death when stimulated by CD8+ DCs in culture could either be due to positive ‘death signals’ or to inadequate ‘life-sustaining signals’. Several possible death signals from CD8+ DCs have been postulated in the past, including ‘veto’ signals mediated by CD8 itself (21) or Fas engagement due to Fas-ligand on the CD8+ DCs (9). However, both of these have been discounted (22, 10). A survey of the expression of the B7 family of co-stimulatory molecules on CD8+ versus CD8– DCs provided no evidence of differences in potential positive or negative signals to CD4 T cells from these molecules (10). One possible reason for a deficit in positive signals in cultures with CD8+ DCs could have been the faster death of this DC subset in culture. However, direct tests of this possibility, by adding granulocyte macrophage colony-stimulating factor to sustain DC viability, or adding in more freshly isolated DCs after 1 or 2 days of culture, all showed that DC death was not a limiting factor and that the relevant signals determining CD4 T cell fate were given early, prior to the first division (10). Although at present we do not understand the differences between CD8+ DCs and CD8– DCs which cause the difference in CD4 T cell proliferation, we now know the stage of the T cell response where the difference has its impact.
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Acknowledgements |
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The authors would like to thank Viki Milovac, Jenny Garbe, Catherine Tarlinton and Carley Young for excellent technical assistance with cell sorting and Mirja Hommel for helpful comments on the manuscript.
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Abbreviations |
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| CFSE | carboxyfluorescein diacetate succinimidyl ester |
| DC | dendritic cell |
| DN | double negative |
| OVA | ovalbumin |
| PI | propidium iodide |
| [3H]TdR | [3H]thymidine |
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Notes |
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Transmitting editor: A. Kelso
Received 23 September 2005, accepted 9 December 2005.
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