- Email: [email protected]
Flow cytometric analysis of T cell proliferation in a mixed lymphocyte reaction with dendritic cells
Journal of Immunological Methods 275 (2003) 57 – 68 www.elsevier.com/locate/jim
Flow cytometric analysis of T cell proliferation in a mixed lymphocyte reaction with dendritic cells Xuan Duc Nguyen a,*, Hermann Eichler a, Alex Dugrillon a, Christoph Piechaczek b, Michael Braun c, Harald Klu¨ter a a Institute of Transfusion Medicine and Immunology, Red Cross Blood Service of Baden-Wu¨rttemberg - Hessen, Faculty of Clinical Medicine Mannheim, University of Heidelberg, Friedrich-Ebert-Str. 107, 68167 Mannheim, Germany b Miltenyi Biotec GmbH, Bergisch Gladbach, Germany c Coulter-Immunotech Diagnostics, Krefeld, Germany
Received 31 January 2002; received in revised form 21 November 2002; accepted 9 December 2002
Abstract Background: Dendritic cells (DCs) are the most potent antigen-presenting cells. They can be generated in vitro from CD14+ cells, and also from CD34+ progenitor cells. Although T cell proliferation using [3H] thymidine incorporation assay has been used widely to check DC function, this technique only provides limited information about the T cell proliferation. Here, we describe a novel method for quantitative analysis of T cell proliferation using flow cytometry. Materials and methods: DCs were generated from CD14+ cells from six healthy blood donors. Monocytes were isolated using positive selection with magnetic cell sorting (MACS) and then cultured with IL-4, GM-CSF, IL-1h, IL-6, TNF-a and PGE2 to yield fully mature DCs. Allogeneic naive T lymphocytes with known mismatches in HLA classes I and II were cocultured with DCs. Naive T cells without DC stimulation served as negative controls. T cells were harvested on days 0, 3, 5, 7, 9, 11 and analysed by flow cytometry. CD3-ECD and CD4-fluorescein isothiocyanate (FITC) or CD8-FITC antibodies were used to distinguish T cell subsets, whereas T cell activation was measured by assessment of HLA-DR, CD45RO, CD25 and CD71 expression. For T cell quantification, fluorescent microparticles were used. Dead cells were excluded with 7-AAD. The bromdeoxyridine (BrdU)-incorporation ELISA procedure was also performed in order to compare with the T cell proliferation assay with regard to absolute cell counts and CD71 expression. Results: The initial T cell concentration on day 1 was 203.9 F 39.7 (173 – 265) CD3+/CD4+ cells/Al and 184.5 F 41.6 (148 – 260) CD3+/CD8+ cells/Al. The maximal T cell proliferation was recorded on day 7 with a five- to tenfold T cell expansion which resulted in 1994.9 F 383 (1446 – 2404) CD3+/CD4+ cells/Al and 944 F 303.7 (560 – 1483) CD3+/CD8+ cells/Al. Furthermore, activation markers of both cell lineages were upregulated and reached maxima on days 7 (CD71) and 9 (CD25, HLA-DR). T cell count/Al as well as CD71 expression both correlated significantly with BrdU incorporation. Conclusion: Flow cytometric analysis permits simple, precise and rapid quantification of T cell proliferation in a mixed lymphocyte reaction with DCs. Activation, proliferation and cell viability can be simultaneously determined. CD71 is particularly well suited as an activation marker for the
Abbreviations: PBMNC, peripheral blood mononuclear cells; DCs, dendritic cells; Th, T helper cell; CTL, cytotoxic T lymphocyte; mAb, monoclonal antibody. * Corresponding author. Tel.: +49-621-3706-8219; fax: +49-621-3706-876. E-mail address: [email protected] (X.D. Nguyen). 0022-1759/03/$ - see front matter D 2003 Elsevier Science B.V. All rights reserved. doi:10.1016/S0022-1759(03)00002-4
58
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
simultaneous measurement of T cell proliferation. Thus, specific T cell subsets involved in antigen-specific proliferation can be evaluated in detail. D 2003 Elsevier Science B.V. All rights reserved. Keywords: Dendritic cells; T cell proliferation and activation; Flow counts
1. Introduction Dendritic cells (DCs) are the most potent antigenpresenting cells and unique in their ability to stimulate naive and memory T cells (Steinman, 1991; Hart, 1997; Banchereau and Steinman, 1998). Recently, in vitro stimulation and vaccination with dendritic cells has opened up a new field of tumor immunotherapy for the treatment of different malignancies (Lotze, 1997; Schadendorf and Nestle, 2001; Nguyen et al., 2002). Various protocols have been described for in vitro generation of DCs from CD14+ or CD34+ progenitor cells (Jonuleit et al., 1997; Thurner et al., 1999; Min et al., 2000). In order to check the ability of DCs to induce proliferation of allogeneic T cells, tritiated thymidine ([3H] TdR) incorporation assays have been widely used. This method is based on the measurement of [3H] TdR incorporation into the DNA of proliferating cells in S phase (Frome et al., 1996). Alternatively, nonradioactive bromdeoxyridine (BrdU)-incorporation assays have also been established. These assays are indirect methods to establish cell proliferation by the determination of DNA synthesis in proliferating cells. Although these methods are sensitive they do not provide information on cell viability and in particular do not indicate which subsets of cells are in fact proliferating in response to the in vitro stimuli. Notably, many subpopulations of DCs with different functions have been shown to act on naive T cells to differentiate into different T cell subsets (Kalinski et al., 1999; Arprinati et al., 2000; Jonmuleit et al., 2001). Consequently, more detailed information on cell viability or responding T cell subsets are necessary in addition to proliferation. In the present study, mixed lymphocyte reactions with DCs were observed over time, and an alternative method using flow cytometry for precise quantification of proliferating T cells was established. Different activation markers were investigated in order to find out the best activation marker correlating with cell proliferation. CD71, which is the transferrin receptor of a
type II membrane glycoprotein and absent from resting blood leucocytes, has been extensively investigated as a potential activation marker. This receptor is expressed by activated cells and upregulated on all cell types entering in proliferation (Testa et al., 1993; Caruso et al., 1997; Barten et al., 2001). Responding T cells were assessed with regard to their viability and activation during allogeneic stimulation. Our aim was to obtain simultaneously information on viability, activation and proliferation of immunoresponsive T cells.
2. Materials and methods 2.1. Separation of CD14+ cells and T cells Peripheral blood mononuclear cells (PBMNC) were obtained from buffy coats and monocytapheresis products of six healthy blood donors. Monocytapheresis was performed as recently described (Nguyen et al., 2002). PBMNC from buffy coats were isolated by Ficoll density-gradient centrifugation (Lymphosep, C.C. Pro, Neustadt, Germany). Differential blood counts were measured using an automated hematology analyzer (Cell-Dyn 3200, Abbott, Wiesbaden, Germany). CD14+ cells were positively separated by high-gradient magnetic sorting using the MIDImagnetic cell sorting (MACS) and Clini-MACS techniques according to the protocol provided by the manufacturer (Miltenyi Biotec, Bergisch Gladbach, Germany). The use of Clini-MACS for CD14+ cell selection from PBMNC of monocytapheresis products was performed under conditions of Good Manufacturing Practice (GMP). Briefly, PBMNC were incubated with saturating concentrations of CD14 MicroBeads for 15 min at 6 jC and washed in PBS containing 2 mM EDTA and 0.5% human serum albumin. Labelled and positively enriched cells were eluted after removal of the columns from the magnetic device. The purity of the CD14+ cell fraction, as assessed by flow cytometry, was consistently over 90%.
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Allogeneic PBMNC from heparinized whole blood were isolated by Ficoll density-gradient centrifugation (Lymphosep, C.C. Pro). Naive T helper (Th) cells and cytotoxic T lymphocytes (CTLs) were separately selected. Highly purified (>90%) CD4+/CD45RA+ and CD8+/CD45RA+ cells were obtained by depletion of all non-T cells (B cells, monocytes, NK cells, memory T cells, cytotoxic or helper cells, dendritic cells, early erythroid cells, platelets and basophils) by the above magnetic sorting technique using monoclonal antibodies against CD11b, CD16, CD19, CD36, CD56, CD45RO and CD8 or CD4. CD3+/CD4+ and CD3+/CD8+ were defined as Th cells and CTLs, respectively. 2.2. DC generation from CD14+ cells Culture medium 1: X-VIVO 15 (Bio Whittaker, Verviers, Belgium), 1% heat-inactivated human AB serum (Sigma, Deisenhofen, Germany), 1% penicillin – streptomycin, 1000 U/ml GM-CSF (Leukomax, Sandoz, Nu¨rnberg, Germany), 1000 U/ml IL-4 (R&D System, Wiesbaden, Germany). Culture medium 2: X-VIVO 15, 1% heat-inactivated human AB serum, 1% penicillin – streptomycin, 2000 U/ml GM-CSF, 1000 U/ml IL-4. Culture medium 3: X-VIVO 15, 1% heat-inactivated human AB serum, 1% penicillin – streptomycin, 1000 U/ml GM-CSF, 1000 U/ml IL-4, 1900 IU/ml IL1h, 1000 IU/ml IL-6, 1100 IU/ml TNF-a (R & D System) and 1Ag/ml prostaglandin E2 (PGE2; Sigma). Isolated CD14+ cells were cultured for 10 days at a cell concentration of 2 106 cells/ml in standard culture flasks (NUNC, Wiesbaden, Germany) with culture medium 1 at 37 jC in a 5% CO2-containing atmosphere. On days 3 and 5, a volume of culture medium 2 equal to one-third of the original flask volume of culture medium 1 was added. For the maturation of DCs, the culture medium was freshly replaced with culture medium 3 on day 7. The cell concentration was reduced at 1 106 cells/ml. At day 10, fully matured DCs were harvested for T cell activation assays and flow cytometric analysis. 2.3. Allogeneic mixed lymphocyte reaction Allogeneic Th cells and CTLs were separately cultured with DCs. Naive Th cells/ml or CTLs/ml
59
(2.5 105) were cultured for 11 days with 2.5 104 matured DCs in X-VIVO 15 medium supplemented with 10% heat-inactivated AB serum and 1% penicillin –streptomycin in 24-well culture plate (NUNC) at 37 jC in a humidified 5% CO2-containing atmosphere. As a negative control, naive Th cells or CTLs were cultured without DCs. On days 0, 3, 5, 7, 9 and 11, (in two experiments also on day 1) cultured T cells were harvested and washed once with PBS containing 0.5% BSA and 2 mM EDTA and resuspended in 2 ml. 2.4. Flow cytometry All flow cytometry analysis was performed on an EPICS XL-MCL machine (Coulter-Immunotech, Krefeld, Germany) equipped with an argon laser tuned at 488 nm. System II Version 3.0 software was used for data acquisition and evaluation. EXPO 32 software (Coulter-Immunotech) was used for the reanalysis of data. Compensation of the four channel fluorescence was precisely adjusted using Cyto-Compk reagents and Cyto-Trolk control cells (Coulter-Immunotech). 2.5. Analysis of DCs All samples were stained using a direct method. A 100-Al sample of DCs (1 106/ml) was incubated for 15 min at room temperature with 10 Al of each monoclonal antibody (mAb) in PBS containing 0.5% BSA. The following mAbs were used: CD1a (BL6)-phycoerythrin (PE), CD83 (HB15A)-fluorescein isothiocyanate (FITC), CD45 (Immu19.2)-FITC, CD14 (RMO52)-PE, CD80 (MAB104)-FITC, CD86 (HA5.2B7)-PE, HLA-ABC (B9.12.1)-FITC, HLADR (B8.12.2)-FITC and -PE, CD40 (MAB 89)-PE, CD3 (UCHT1)-ECD, CD4 (13B8.2)-FITC, CD8 (B9.11). Appropriate isotype controls were used at the same protein concentration as the test antibody. All mAbs were purchased from Immunotech, Marseille, France. 2.6. Analysis of T cell proliferation 2.6.1. BrdU-incorporation ELISA procedure DNA synthesis in proliferating cells was determined by measuring BrdU incorporation in the commercial Cell Proliferation ELISA System (Roche Molecular Biochemicals, Mannheim, Germany). The
60
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
assay was performed according to the instructions of the manufacturer. Briefly, 180 Al/well of the cell culture was resuspended in triplicate in 96-well plates. A 20 Al/well BrdU labelling reagent (final concentration, 10 AM) was added. At 6 h, the cells were harvested by centrifugation at 300 g for 10 min and were fixed in ethanol solution, followed by incubation with peroxidase-labeled anti-BrdU for 90 min. After removal of the antibody conjugate and three washes, 100 Al/well tetramethyl-benzidine (TMB) was added and the mixtures incubated until color development (15 min). Absorbance values were measured at 450 nm using an ELISA reader (Tecan, Crailsheim, Germany). Culture medium alone was used as a control for nonspecific binding.
then ready for measurement. Fluorescent microparticles were warmed to room temperature, premixed and 100 Al of Flow-Countk Fluorespheres added to two specimens immediately before the measurement. The absolute count of target cells was calculated on the basis of the known bead concentrations using the following equation:
2.6.2. Flow cytometric analysis of T cell proliferation and activation The flow cytometer settings were established for linear amplification of light scatter and logarithmic amplification of fluorescence channels. The threshold was defined in forward scatter with FS 100 to exclude cell debris. Flow cytometric analysis using four different fluorochromes were performed. The following mAbs were used: CD45RA (ALB11)-PE, CD45RO (UCHL-1)-PE, CD25 (B1.49.9)-PE, CD71 (YDJ1.2.2)-PE, CD38 (T16)-PE, CD69 (TP1.55.3)PE, CD3 (UCHT1)-ECD, CD4 (13B8.2)-FITC, CD8 (B9.11)-FITC. All mAbs were purchased from Immunotech. CD3 /CD4 or CD3 /CD8 antibodies were used separately to distinguish Th cells or CTLs from the other cell populations. T cell activation was measured by the assessment of HLA-DR, CD45RO, CD25, CD38, CD69 and CD71 expression. Dead cells were excluded with 7-AAD. For the absolute cell count, fluorescent microparticles (Flow-Countk Fluorespheres, Coulter-Immunotech) of defined concentration were used. The assayed concentration of Flow-Countk Fluorespheres was described in each lot at delivery. Using a reverse pipetting technique, 100 Al of each sample was incubated for 15 min at room temperature with 10 Al aliquots of each mAb (e.g. CD3-ECD, CD4-FITC, CD71-PE and 7-AAD). Corresponding isotypic mAbs were used to define the cut-off which had to be less than 1%. After incubation the cells were resuspended in PBS containing 0.5% paraformaldehyde and were
The correct assayed concentration of fluorespheres was used as a calibration factor in the software program and the EPICS XL flow cytometer calculated the absolute cell count for the specimen automatically. The absolute cell counting was determined twice.
Absolute cell counts=Al ¼
total number of cells counted total number of fluorespheres counted flow count concentration
2.7. Comparison of flow cytometric analysis of T cell proliferation with BrdU-incorporation assay Additional experiments were performed to compare the flow cytometric method with the standard BrdU-incorporation ELISA procedure (n = 4). DCs and T cells (cultured DCs/Th cells or DCs/ CTLs or T cells alone) were cultured at increasing ratios of DC/T cells (1:10, 1:33, 1:100, 1:300). On day 6, T cells were harvested for the BrdU assay. On the same day and 6 h later, T cells of the same culture were harvested for flow cytometric analysis of absolute cell counting and the expression of CD71. 2.8. Statistical analysis Means, standard deviations and coefficients of variation were calculated where indicated. Statistically significant differences were assessed using one-way analysis of variance (MANOVA) or a paired t-test. The Pearson test was used for the calculation of correlations. Differences between data were considered statistically significant at p < 0.05. All tests were performed with commercially available software for personal computer (SPSS for Windows NT, SPSS software, Munich, Germany).
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
3. Results 3.1. DC generation from CD14+ cells CD14+ cells were cultured for 10 days in a specific medium and developed to fully mature DCs with a typical dendritic morphology. Characteristically, numerous cytoplasmic projections were present. The expression of CD83 was 76 F 13.9% (57 – 94). The mean yield of mature CD83+ DCs was 8 F 7% (range 1.6– 19%) of the plated cells. Flow cytometric analysis showed that these cells expressed little CD1a, but abundant quantities of CD80, CD86, CD40, HLA-DR and MHC class I. Furthermore, 17.3 F 11.1% (range 7– 29%) of the cultured cells expressed CD4, but were negative for CD3 and CD8. 3.2. Allogeneic mixed lymphocyte reaction 3.2.1. Cell morphology The DCs were tested for their ability to induce proliferation of allogeneic CD4+/CD45RA+ and CD8+/CD45RA+ cells in a mixed lymphocyte reaction. CD4+/CD45RA+ and CD8+/CD45RA+ cells were purified prior to the mixed lymphocyte reaction and cultured separately with DCs. DCs induced proliferation of responder T cells (Th cells as well as CTLs) when compared with responder cells alone. After 12 h, DCs formed clusters with T cells, whereas this phenomenon could not be observed in cultures of T cells alone. On day 3, cluster formation increased. When viewed by inverted microscopy, cocultured T cells apparently changed their morphology. They appeared to be larger and were heterogeneous. From day 5, the culture dish well was filled with proliferated cells which increased maximally on day 7. On subsequent days, clusters of dead cells were observed. 3.3. Flow cytometric analysis of T cell proliferation and activation Flow cytometry was used to determine activating and proliferating T cells in a mixed lymphocyte reaction with DCs. CD3+/CD4+ or CD3+/CD8+ cells were separately analysed. Fig. 1 shows the simultaneous flow cytometric analysis of cell viability (row a), percentage of CD3+/CD4+ cells (row b) and CD71 activation (row c) on days 0, 7 and 11.
61
The absolute cell count of each gate (viable cells, CD3+/CD4+, CD4+/CD71+) was calculated automatically using beads of known concentration. The same technique was used for analysis of CD3+/CD8+ cells (data not shown). T cell proliferation was determined by absolute cell counting. Fig. 2a and b shows the development of absolute cell count/Al of Th cells and CTLs over time, respectively. With DC stimulation, the maximal T cell proliferation showing typical characteristics with a high intensity of side and forward scatter was recorded on day 7, at which time a five- to tenfold T cell expansion was observed, whereas T cells alone decreased continuously. Significant differences in Th and CTL cell numbers between days 0, 3, 7 and 11 were confirmed by the MANOVA test with p < 0.0001. The average standard deviation of flow cytometric quantification analysis was 2.2% (coefficient of variation = 2.2%). Th cell activation was determined by analysis of the expression of CD25, CD71, CD45RO and HLADR. Fig. 3a and b shows the development of the activation molecules of Th cells over time. In accordance with the absolute cell numbers, the maximal expression of CD71 was recorded on day 7. In addition, the expression of CD71 began to increase as early as on day 1 before cell proliferation (7.6 F 2.4% versus 0.3 F 0.3% on day 0). This activation molecule showed a strong correlation with the absolute Th cell count. The expression of HLA-DR, CD25 also increased continuously and reached a maximum on day 9 (Fig. 3a). Cell viability decreased continuously after day 7. The expression of CD45RA (naive Th cells) and CD45RO (memory Th cells) developed inversely. During the stimulation period, Th cells lost their CD45RA expression and developed into memory Th cells with a high expression of CD45RO (Fig. 3b). No significant changes of the activation molecules of T cells without DC stimulation were observed during the culture period, but cell viability decreased earlier (data not shown). Other activation molecules, such as CD38 and CD69 (an early activation marker) were found not to be helpful and were not further assessed (n = 2). These markers increased even on T cells alone without DCs after 3 days in culture (data not shown).
62
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Fig. 1. Flow cytometric protocol for analysis of CD3+/CD4+ cell proliferation in mixed lymphocyte reaction with DCs on days 0, 7 and 11. In row (a), dead cells were excluded using 7-AAD (emission spectrum of 650 nm). The rectilinear gate was set to include viable cells. All viable cells were gated back in row (b) and analysed for CD3+/CD4+ cells. In row (c), only viable CD3+/CD4+ cells appeared for analysis of CD71 expression. In an extra diagram of the same protocol (diagram not depicted in figure), beads with a broad fluorescence emission range (510 – 700 nm) were easily gated in a forward scatter versus the fourth fluorescence channel. The absolute cell count of each gate (viable cells, CD3+/CD4+, CD4+/CD71+) was automatically calculated relative to beads of known concentration and given as cell counts/ Al. Thus, cell viability, absolute cell counting and activation were determined simultaneously. At the beginning of cell culture (day 0), almost 100% of the cells were viable. The flow cytometric analysis of the morphology showed a small population of CD3+/CD4+ cells without expression of CD71. On day 7, more than 95% of cells were still viable with a high intensity of side scatter (also with a high intensity of forward scatter, diagram not shown in figure). The population of CD3+/CD4+ cells appeared larger. In parallel, CD71 expression increased strongly. On day 11, most of CD3+/CD4+ cells were dead correlating with a low value of CD71 (analysis of CD3+/ CD8+ cells not shown as figure).
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
63
Fig. 2. The course of absolute T cell count/Al during the whole stimulation period with DCs (n = 6) shown. Allogeneic CD3+/CD4+ (Th cells) and CD3+/CD8+ cells (CTLs) were purified prior to the mixed lymphocyte reaction and cultured separately with DCs. The analysis of both cell lineage was successively performed. Th and CTL cell count (Fig. 1a and b, respectively) increased strongly and reached a maximum on day 7 ( p < 0.00001 and p < 0.001, respectively) whereas T cells alone decreased continuously. The error bars indicate mean F 2 S.D.
According to the activation of CTLs, they showed a similar development like Th cells (data not depicted as figure). The maximal expression of CD71 was recorded on day 7 with 82.4 F 11 (64.3 – 94%). A strong correlation between the expression of this activation molecule and the absolute cell count was also observed (r = 0.87, p < 0.01). The expression of
HLA-DR, CD25 increased continuously and also reached a maximum on day 9 with 14.1 F 1.8 (11.8 –16%) and 83.8 F 14.6 (58 – 93%), respectively ( p < 0.0001 for both markers, in comparison with day 0). In contrast, cell viability decreased continuously from day 7 with 68.4 F 8.1 (58.2 – 81.4%) on day 9, and only 40.8 F 15.2 (24.3 –63.3%) on day 11.
64
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Fig. 3. The course of viability and activation of CD3+/CD4+ cells during the stimulation period with DCs (n = 6) depicted. Each marker (CD25, CD71, CD45RO, CD45RA and HLA-DR) was simultaneously determined with cell viability (7-AAD) and CD3/CD4 marker. Box (a) shows an increasing expression of CD25, CD71 and HLA-DR with p < 0.0001. CD71 expression reached a maximum on day 7. In comparison to CD25 and HLA-DR expression, each of which did not reach a maximum before day 9, CD71 correlated best with the absolute cell count (r = 0.95, p < 0.001). Box (b) shows the development of cell viability and CD45RA and CD45RO expression over time. The expression of CD45RA (naive Th cells) and CD45RO (memory Th cells) developed inversely. During the stimulation period, Th cells lost CD45RA and developed into memory Th cells with a high expression of CD45RO ( p < 0.0001) which did not significantly change after day 9. From day 7, cell viability decreased constantly and more than the half of the cells were dead on day 11 (data of CD3+/CD8+ cells not shown). The error bars indicate mean F 2 S.D.
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
65
Fig. 4. Comparison of the BrdU-incorporation ELISA procedure (a) and flow cytometric analysis of absolute Th cell count (b) and CD71 expression (c). Different ratios of DC/Th cells were cultured and harvested on day 6. Similar to the development of CD3+/CD4+ cell concentrations and the expression of CD71, BrdU incorporation in proliferating cells decreased constantly with an increasing DC/Th cell ratio. Th cell count/Al as well as CD71 expression showed a significant correlation with BrdU incorporation (r = 0.89, p < 0.001; and r = 0.94, p < 0.001, respectively). The A450 value of Th cells alone was lower than 0.05 (data of CTLs not shown). The error bars indicate mean F 2 S.D.
66
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
3.4. Comparison of T cell proliferation analysis with the BrdU-incorporation ELISA procedure The BrdU-incorporation ELISA procedure was compared with the T cell proliferation assay with regard to absolute cell counts and CD71 expression. Fig. 4a shows decreasing BrdU incorporation with increasing DC/Th cell ratios. Similar trends for Th cell counts/Al and CD71 expression data were also observed (Fig. 4b and c). Th cell count/Al as well as CD71 expression both correlated significantly with BrdU incorporation (r = 0.89, p < 0.001; and r = 0.94, p < 0.001, respectively) (data of DC and CTLs measurement not shown).
4. Discussion The concept of in vitro stimulation and vaccination with DCs has led to a new field of immunotherapy for cancer (Lotze, 1997; Schadendorf and Nestle, 2001; Nguyen et al., 2002). Recent publications suggest that subsets of DCs differ in their antigen presentation characteristics (Kalinski et al., 1999; Arprinati et al., 2000; Jonmuleit et al., 2001). In order to check DC function, [3H] TdR incorporation assays have been used for the measurement of T cell proliferation in mixed lymphocyte reactions (Jonuleit et al., 1997; Thurner et al., 1999; Min et al., 2000). Using [3H] TdR or BrdU-ELISA incorporation assays, T cell proliferation is measured by incorporation of thymidine analogues into the DNA of proliferating cells. Thus, monitoring of DNA synthesis is an indirect parameter of cell proliferation. Although this technique is sensitive, it only provides information on the overall proliferative response and gives no insight into specific cell subsets involved. Furthermore, the incorporation of thymidine analogues does not always correlate with true increases in DNA content and cell proliferation (Neckers et al., 1995). It can be assumed that various DC subpopulations exist having distinct functions with respect to T cell activation or inhibition. Therefore, it is much more important to investigate the T cell response in the presence of DCs. Caruso et al. (1997) investigated the correlation between activation markers on stimulated T cells and their proliferation using flow cytometric analysis and [3H] TdR
incorporation assays. Both assays are complementary and a combination of both methods was recommended to obtain a clearer understanding of that events leading to efficient cell-mediated immune response (Caruso et al., 1997). The most accurate method to measure cell proliferation in vitro is to determine viable cells directly. In the present study, we developed a novel flow cytometric method for quantitative determination of T cell proliferation in mixed lymphocyte reactions with DCs using a combination of immunophenotyping and quantification. Our aim was to obtain rapid and precise information on cell viabilitity, activation and proliferation simultaneously in a single measurement. Furthermore, we sought to identify the activation marker correlating most closely with cell proliferation. Accordingly, the activation markers CD25, CD69, CD71 and HLA-DR (Caruso et al., 1997; Barten et al., 2001) were investigated. As T cells are nonadherent in culture, they were easily resuspended for single cell analysis by flow cytometry. The cultured DCs proved to be strong stimulators of naive T cells with high expression of costimulator molecules. As early as 12 h after initiation of the coculture, DCs and T cells formed a cluster. CD3+/CD4+ and CD3+/CD8+ cells began to increase their activation markers and, in addition, the expression of CD71 molecule increased earlier than cell proliferation. On day 7, maximal expansion of CD3+/ CD4+ and CD3+/CD8+ cells were recorded. In association with cell expansion over time, maximal expression of CD71 was observed on day 7. This activation marker showed the strongest correlation with absolute T cell counts. In fact, CD71 is upregulated on all cell types entering proliferation (Testa et al., 1993). In order to confirm this result, a BrdUincorporation ELISA procedure was performed. Using different ratios of DCs and T cells, the BrdU-incorporation ELISA procedure and flow cytometric analysis of cell proliferation and CD71 expression showed a strong correlation. Nevertheless, the [3H] TdR incorporation assay is the gold standard method with which to determine cell proliferation. The maximal expression of the activation markers CD25 and HLADR of both T cell lineages was observed on day 9. However, cell counts and cell viability decreased strongly by this time. Thus, it is apparently not feasible to investigate T cells after day 7.
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
The determination of CD38 and CD69 expression as measures of proliferation and activation have been recommended by other groups (Funaro et al., 1990; Maino et al., 1995; Cesano et al., 1998), but these markers did not prove to be useful in our study. We found that even naive T cells without DC stimulation strongly expressed CD38 and CD69 during the culture period. Interestingly, we observed that the viability of T cells without DC stimulation decreased more rapidly. DCs as stimulators protected T cells from early cell death. In fact, costimulation has a protective effect on T cell survival, probably due to the upregulation of anti-apoptotic mechanisms (Boise et al., 1995). Fluorescent microparticles were used to determine cell concentrations simultaneously without additional cell loss or other cell preparation steps. These microparticles were easily distinguishable from other events of interest. We have previously demonstrated that absolute cell counts determined using fluorescent microparticles were stable during the processing time (Schlenke et al., 1998). A coefficient of variation of 2.2% in the present study indicates the accuracy of the measurements. Many of the cultured DCs were CD4 positive but CD3 negative. Caux et al. (1992, 1995) showed that DCs cultured from CD34+ cells strongly expressed CD4. Against this background, the use of CD3-ECD for the analysis was indispensable in order to distinguish CD3+/CD4+ cells from DCs. This point has to be considered since there are many DC subpopulations differing in their expression of surface molecules, whereas no cell surface marker is unique to DCs (Kalinski et al., 1999; Arprinati et al., 2000; Jonmuleit et al., 2001). Priming of naive T cells by DC stimulation of T cells showed a high level of proliferation. Thus, the use of 7-AAD was important to exclude dead cells. In conclusion, this novel flow cytometric T cell assay offers a simple, rapid and precise quantification of T cell proliferation in mixed lymphocyte reactions with DCs. A simultaneous determination of absolute cell counts, expression of surface activation molecules and cell viability can be performed without any additional cell preparation. CD71 is a suitable activation marker which can be simultaneously determined. Further characterizations of additional surface markers involved in proliferation and activation processes are
67
feasible. This method will open broad field of applications as it is easy to perform and accurate.
Acknowledgements The authors are indebted to Daniella Griffiths for expert editorial assistance in the preparation of this manuscript.
References Arprinati, M., Green, C.L., Heimfeld, S., Heuser, J.E., Anasetti, C., 2000. Granulocyte-colony stimulating factor mobilizes T helper 2-inducing dendritic cells. Blood 95, 2484. Banchereau, J., Steinman, R.M., 1998. Dendritic cells and the control of immunity. Nature 392, 245. Barten, M.J., Gummert, J.F., van Gelder, T., Shorthouse, R., Morris, R.E., 2001. Flow cytometric quantification of calcium-dependent and -independent mitogen-stimulation of T cell functions in whole blood: inhibition by immunosuppressive drugs in vitro. J. Immunol. Methods 253, 95. Boise, L.H., Minn, A.J., Noel, P.J., June, C.H., Accavitti, M.A., Lindsten, T., Thompson, C.B., 1995. CD28 costimulation can promote T cell survival by enhancing the expression of Bcl-XL. Immunity 3, 87. Caruso, A., Licenzi, S., Corulli, M., Canaris, A.D., De Francesco, M.A., Fiorentini, S., Peroni, L., Fallacara, F., Dima, F., Balsari, A., Turano, A., 1997. Flow cytometric analysis of activation markers on stimulated T cells and their correlation with cell proliferation. Cytometry 17, 71. Caux, C., Dezutter-Dambuyant, C., Schmitt, D., Banchereau, J., 1992. GM-CSF and TNF-a cooperate in the generation of dendritic Langerhans cells. Nature 360, 258. Caux, C., Massacrier, C., Dezutter-Damuyant, C., Vanbervliet, B., Jacquet, C., Schmitt, D., Banchereau, J., 1995. Human dendritic langerhans cells generated in vitro from CD34+ progenitors can prime naive CD4+ T cells and process soluble antigen. J. Immunol. 155, 5427. Cesano, A., Visonneau, S., Deaglio, S., Malavasi, F., Santoli, D., 1998. Role of CD38 and its ligand in the regulation of MHCnonrestricted cytotoxic T cells. J. Immunol. 160, 1106. Frome, E.L., Smith, M., Littlefield, G., Neubert, R., Colyer, S., 1996. Statistical methods for the blood beryllium lymphocyte proliferation test. Environ. Health Perspect. 105, 957. Funaro, A., Spagnoli, G.C., Ausielo, C.M, Alessio, M., Roggero, S., Delia, D., Zaccolo, M., Malavasi, F., 1990. Involvement of the multilineage CD38 molecule in a unique pathway of cell activation and proliferation. J. Immunol. 145, 2390. Hart, D.N.J., 1997. Dendritic cells: unique leucocyte populations, which control the primary immune response. Blood 90, 3245. Jonmuleit, H., Schmitt, E., Steinbrink, K., Enk, A.H., 2001. Dendritic cells as a tool to induce anergic and regulatory T cells. Trends Immunol. 22, 394.
68
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Jonuleit, H., Kuhn, U., Mu¨ller, G., Steinbrink, K., Paragnik, L., Schmitt, E., Knop, J., Enk, A.H., 1997. Pro-inflammatory cytokines and prostaglandins induce maturation of potent immunostimulary dendritic cells under fetal calf serum-free conditions. Eur. J. Immunol. 27, 3135. Kalinski, P., Hilkens, C.M.U., Wierenga, E.A., Kapsenberg, M., 1999. T cell priming by type-I and type-2 polarized dendritic cells: the concept of a third signal. Immunol. Today 20, 561. Lotze, M.T., 1997. Getting to the source: dendritic cells as therapeutic reagents for the treatment of patients with cancer. Ann. Surg. 226, 1. Maino, V.V., Suni, M.A., Ruitenberg, J.J., 1995. Rapid flow cytometric method for measuring lymphocyte subset activation. Cytometry 20, 127. Min, Y.H., Lee, S.T., Choi, K.M., Hahn, J.S., Ko, Y.W., 2000. Surface expression of HLA-DM on dendritic cells derived from CD34-positive bone marrow haematopoietic stem cells. Br. J. Haematol. 110, 385. Neckers, L.M., Funkhouser, W.K., Treel, J.B., Cossman, J., Gratzner, H.G., 1995. Significant non-s-phase DNA synthesis visualized by flow cytometry in activated and in malignant human lymphoid cells. Exp. Cell Res. 156, 429.
Nguyen, X.D., Eichler, H., Sucker, A., Hofmann, U., Schadendorf, D., Klu¨ter, H., 2002. Collection of autologous monocytes for dendritic cell vaccination therapy in metastatic melanoma patients. Transfusion 42, 428. Schadendorf, D., Nestle, F.O., 2001. Autologous dendritic cells for treatment of advanced cancer—an update. Recent Results Cancer Res. 158, 236. Schlenke, P., Frohn, C., Klu¨ter, H., Saballus, M., Hammers, H.J., Zajac, S.R., Kirchner, H., 1998. Evaluation of a flow cytometric method for simultaneous leucocyte phenotyping and quantification by fluorescent microspheres. Cytometry 33, 310. Steinman, R.M., 1991. The dendritic cell system and its role in immunogenicity. Annu. Rev. Immunol. 9, 271. Testa, U., Pelosi, E., Peschle, C., 1993. The transferrin receptor. Crit. Rev. Oncog. 4, 241. Thurner, B., Ro¨der, C., Dieckmann, D., Heuer, M., Kruse, M., Glaser, A., Keikavoussi, P., Ka¨mpgen, E., Bender, A., Schuler, G., 1999. Generation of large numbers of fully mature and stable dendritic cells from leukapheresis products for clinical application. J. Immunol. Methods 223, 1.
Flow cytometric analysis of T cell proliferation in a mixed lymphocyte reaction with dendritic cells Xuan Duc Nguyen a,*, Hermann Eichler a, Alex Dugrillon a, Christoph Piechaczek b, Michael Braun c, Harald Klu¨ter a a Institute of Transfusion Medicine and Immunology, Red Cross Blood Service of Baden-Wu¨rttemberg - Hessen, Faculty of Clinical Medicine Mannheim, University of Heidelberg, Friedrich-Ebert-Str. 107, 68167 Mannheim, Germany b Miltenyi Biotec GmbH, Bergisch Gladbach, Germany c Coulter-Immunotech Diagnostics, Krefeld, Germany
Received 31 January 2002; received in revised form 21 November 2002; accepted 9 December 2002
Abstract Background: Dendritic cells (DCs) are the most potent antigen-presenting cells. They can be generated in vitro from CD14+ cells, and also from CD34+ progenitor cells. Although T cell proliferation using [3H] thymidine incorporation assay has been used widely to check DC function, this technique only provides limited information about the T cell proliferation. Here, we describe a novel method for quantitative analysis of T cell proliferation using flow cytometry. Materials and methods: DCs were generated from CD14+ cells from six healthy blood donors. Monocytes were isolated using positive selection with magnetic cell sorting (MACS) and then cultured with IL-4, GM-CSF, IL-1h, IL-6, TNF-a and PGE2 to yield fully mature DCs. Allogeneic naive T lymphocytes with known mismatches in HLA classes I and II were cocultured with DCs. Naive T cells without DC stimulation served as negative controls. T cells were harvested on days 0, 3, 5, 7, 9, 11 and analysed by flow cytometry. CD3-ECD and CD4-fluorescein isothiocyanate (FITC) or CD8-FITC antibodies were used to distinguish T cell subsets, whereas T cell activation was measured by assessment of HLA-DR, CD45RO, CD25 and CD71 expression. For T cell quantification, fluorescent microparticles were used. Dead cells were excluded with 7-AAD. The bromdeoxyridine (BrdU)-incorporation ELISA procedure was also performed in order to compare with the T cell proliferation assay with regard to absolute cell counts and CD71 expression. Results: The initial T cell concentration on day 1 was 203.9 F 39.7 (173 – 265) CD3+/CD4+ cells/Al and 184.5 F 41.6 (148 – 260) CD3+/CD8+ cells/Al. The maximal T cell proliferation was recorded on day 7 with a five- to tenfold T cell expansion which resulted in 1994.9 F 383 (1446 – 2404) CD3+/CD4+ cells/Al and 944 F 303.7 (560 – 1483) CD3+/CD8+ cells/Al. Furthermore, activation markers of both cell lineages were upregulated and reached maxima on days 7 (CD71) and 9 (CD25, HLA-DR). T cell count/Al as well as CD71 expression both correlated significantly with BrdU incorporation. Conclusion: Flow cytometric analysis permits simple, precise and rapid quantification of T cell proliferation in a mixed lymphocyte reaction with DCs. Activation, proliferation and cell viability can be simultaneously determined. CD71 is particularly well suited as an activation marker for the
Abbreviations: PBMNC, peripheral blood mononuclear cells; DCs, dendritic cells; Th, T helper cell; CTL, cytotoxic T lymphocyte; mAb, monoclonal antibody. * Corresponding author. Tel.: +49-621-3706-8219; fax: +49-621-3706-876. E-mail address: [email protected] (X.D. Nguyen). 0022-1759/03/$ - see front matter D 2003 Elsevier Science B.V. All rights reserved. doi:10.1016/S0022-1759(03)00002-4
58
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
simultaneous measurement of T cell proliferation. Thus, specific T cell subsets involved in antigen-specific proliferation can be evaluated in detail. D 2003 Elsevier Science B.V. All rights reserved. Keywords: Dendritic cells; T cell proliferation and activation; Flow counts
1. Introduction Dendritic cells (DCs) are the most potent antigenpresenting cells and unique in their ability to stimulate naive and memory T cells (Steinman, 1991; Hart, 1997; Banchereau and Steinman, 1998). Recently, in vitro stimulation and vaccination with dendritic cells has opened up a new field of tumor immunotherapy for the treatment of different malignancies (Lotze, 1997; Schadendorf and Nestle, 2001; Nguyen et al., 2002). Various protocols have been described for in vitro generation of DCs from CD14+ or CD34+ progenitor cells (Jonuleit et al., 1997; Thurner et al., 1999; Min et al., 2000). In order to check the ability of DCs to induce proliferation of allogeneic T cells, tritiated thymidine ([3H] TdR) incorporation assays have been widely used. This method is based on the measurement of [3H] TdR incorporation into the DNA of proliferating cells in S phase (Frome et al., 1996). Alternatively, nonradioactive bromdeoxyridine (BrdU)-incorporation assays have also been established. These assays are indirect methods to establish cell proliferation by the determination of DNA synthesis in proliferating cells. Although these methods are sensitive they do not provide information on cell viability and in particular do not indicate which subsets of cells are in fact proliferating in response to the in vitro stimuli. Notably, many subpopulations of DCs with different functions have been shown to act on naive T cells to differentiate into different T cell subsets (Kalinski et al., 1999; Arprinati et al., 2000; Jonmuleit et al., 2001). Consequently, more detailed information on cell viability or responding T cell subsets are necessary in addition to proliferation. In the present study, mixed lymphocyte reactions with DCs were observed over time, and an alternative method using flow cytometry for precise quantification of proliferating T cells was established. Different activation markers were investigated in order to find out the best activation marker correlating with cell proliferation. CD71, which is the transferrin receptor of a
type II membrane glycoprotein and absent from resting blood leucocytes, has been extensively investigated as a potential activation marker. This receptor is expressed by activated cells and upregulated on all cell types entering in proliferation (Testa et al., 1993; Caruso et al., 1997; Barten et al., 2001). Responding T cells were assessed with regard to their viability and activation during allogeneic stimulation. Our aim was to obtain simultaneously information on viability, activation and proliferation of immunoresponsive T cells.
2. Materials and methods 2.1. Separation of CD14+ cells and T cells Peripheral blood mononuclear cells (PBMNC) were obtained from buffy coats and monocytapheresis products of six healthy blood donors. Monocytapheresis was performed as recently described (Nguyen et al., 2002). PBMNC from buffy coats were isolated by Ficoll density-gradient centrifugation (Lymphosep, C.C. Pro, Neustadt, Germany). Differential blood counts were measured using an automated hematology analyzer (Cell-Dyn 3200, Abbott, Wiesbaden, Germany). CD14+ cells were positively separated by high-gradient magnetic sorting using the MIDImagnetic cell sorting (MACS) and Clini-MACS techniques according to the protocol provided by the manufacturer (Miltenyi Biotec, Bergisch Gladbach, Germany). The use of Clini-MACS for CD14+ cell selection from PBMNC of monocytapheresis products was performed under conditions of Good Manufacturing Practice (GMP). Briefly, PBMNC were incubated with saturating concentrations of CD14 MicroBeads for 15 min at 6 jC and washed in PBS containing 2 mM EDTA and 0.5% human serum albumin. Labelled and positively enriched cells were eluted after removal of the columns from the magnetic device. The purity of the CD14+ cell fraction, as assessed by flow cytometry, was consistently over 90%.
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Allogeneic PBMNC from heparinized whole blood were isolated by Ficoll density-gradient centrifugation (Lymphosep, C.C. Pro). Naive T helper (Th) cells and cytotoxic T lymphocytes (CTLs) were separately selected. Highly purified (>90%) CD4+/CD45RA+ and CD8+/CD45RA+ cells were obtained by depletion of all non-T cells (B cells, monocytes, NK cells, memory T cells, cytotoxic or helper cells, dendritic cells, early erythroid cells, platelets and basophils) by the above magnetic sorting technique using monoclonal antibodies against CD11b, CD16, CD19, CD36, CD56, CD45RO and CD8 or CD4. CD3+/CD4+ and CD3+/CD8+ were defined as Th cells and CTLs, respectively. 2.2. DC generation from CD14+ cells Culture medium 1: X-VIVO 15 (Bio Whittaker, Verviers, Belgium), 1% heat-inactivated human AB serum (Sigma, Deisenhofen, Germany), 1% penicillin – streptomycin, 1000 U/ml GM-CSF (Leukomax, Sandoz, Nu¨rnberg, Germany), 1000 U/ml IL-4 (R&D System, Wiesbaden, Germany). Culture medium 2: X-VIVO 15, 1% heat-inactivated human AB serum, 1% penicillin – streptomycin, 2000 U/ml GM-CSF, 1000 U/ml IL-4. Culture medium 3: X-VIVO 15, 1% heat-inactivated human AB serum, 1% penicillin – streptomycin, 1000 U/ml GM-CSF, 1000 U/ml IL-4, 1900 IU/ml IL1h, 1000 IU/ml IL-6, 1100 IU/ml TNF-a (R & D System) and 1Ag/ml prostaglandin E2 (PGE2; Sigma). Isolated CD14+ cells were cultured for 10 days at a cell concentration of 2 106 cells/ml in standard culture flasks (NUNC, Wiesbaden, Germany) with culture medium 1 at 37 jC in a 5% CO2-containing atmosphere. On days 3 and 5, a volume of culture medium 2 equal to one-third of the original flask volume of culture medium 1 was added. For the maturation of DCs, the culture medium was freshly replaced with culture medium 3 on day 7. The cell concentration was reduced at 1 106 cells/ml. At day 10, fully matured DCs were harvested for T cell activation assays and flow cytometric analysis. 2.3. Allogeneic mixed lymphocyte reaction Allogeneic Th cells and CTLs were separately cultured with DCs. Naive Th cells/ml or CTLs/ml
59
(2.5 105) were cultured for 11 days with 2.5 104 matured DCs in X-VIVO 15 medium supplemented with 10% heat-inactivated AB serum and 1% penicillin –streptomycin in 24-well culture plate (NUNC) at 37 jC in a humidified 5% CO2-containing atmosphere. As a negative control, naive Th cells or CTLs were cultured without DCs. On days 0, 3, 5, 7, 9 and 11, (in two experiments also on day 1) cultured T cells were harvested and washed once with PBS containing 0.5% BSA and 2 mM EDTA and resuspended in 2 ml. 2.4. Flow cytometry All flow cytometry analysis was performed on an EPICS XL-MCL machine (Coulter-Immunotech, Krefeld, Germany) equipped with an argon laser tuned at 488 nm. System II Version 3.0 software was used for data acquisition and evaluation. EXPO 32 software (Coulter-Immunotech) was used for the reanalysis of data. Compensation of the four channel fluorescence was precisely adjusted using Cyto-Compk reagents and Cyto-Trolk control cells (Coulter-Immunotech). 2.5. Analysis of DCs All samples were stained using a direct method. A 100-Al sample of DCs (1 106/ml) was incubated for 15 min at room temperature with 10 Al of each monoclonal antibody (mAb) in PBS containing 0.5% BSA. The following mAbs were used: CD1a (BL6)-phycoerythrin (PE), CD83 (HB15A)-fluorescein isothiocyanate (FITC), CD45 (Immu19.2)-FITC, CD14 (RMO52)-PE, CD80 (MAB104)-FITC, CD86 (HA5.2B7)-PE, HLA-ABC (B9.12.1)-FITC, HLADR (B8.12.2)-FITC and -PE, CD40 (MAB 89)-PE, CD3 (UCHT1)-ECD, CD4 (13B8.2)-FITC, CD8 (B9.11). Appropriate isotype controls were used at the same protein concentration as the test antibody. All mAbs were purchased from Immunotech, Marseille, France. 2.6. Analysis of T cell proliferation 2.6.1. BrdU-incorporation ELISA procedure DNA synthesis in proliferating cells was determined by measuring BrdU incorporation in the commercial Cell Proliferation ELISA System (Roche Molecular Biochemicals, Mannheim, Germany). The
60
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
assay was performed according to the instructions of the manufacturer. Briefly, 180 Al/well of the cell culture was resuspended in triplicate in 96-well plates. A 20 Al/well BrdU labelling reagent (final concentration, 10 AM) was added. At 6 h, the cells were harvested by centrifugation at 300 g for 10 min and were fixed in ethanol solution, followed by incubation with peroxidase-labeled anti-BrdU for 90 min. After removal of the antibody conjugate and three washes, 100 Al/well tetramethyl-benzidine (TMB) was added and the mixtures incubated until color development (15 min). Absorbance values were measured at 450 nm using an ELISA reader (Tecan, Crailsheim, Germany). Culture medium alone was used as a control for nonspecific binding.
then ready for measurement. Fluorescent microparticles were warmed to room temperature, premixed and 100 Al of Flow-Countk Fluorespheres added to two specimens immediately before the measurement. The absolute count of target cells was calculated on the basis of the known bead concentrations using the following equation:
2.6.2. Flow cytometric analysis of T cell proliferation and activation The flow cytometer settings were established for linear amplification of light scatter and logarithmic amplification of fluorescence channels. The threshold was defined in forward scatter with FS 100 to exclude cell debris. Flow cytometric analysis using four different fluorochromes were performed. The following mAbs were used: CD45RA (ALB11)-PE, CD45RO (UCHL-1)-PE, CD25 (B1.49.9)-PE, CD71 (YDJ1.2.2)-PE, CD38 (T16)-PE, CD69 (TP1.55.3)PE, CD3 (UCHT1)-ECD, CD4 (13B8.2)-FITC, CD8 (B9.11)-FITC. All mAbs were purchased from Immunotech. CD3 /CD4 or CD3 /CD8 antibodies were used separately to distinguish Th cells or CTLs from the other cell populations. T cell activation was measured by the assessment of HLA-DR, CD45RO, CD25, CD38, CD69 and CD71 expression. Dead cells were excluded with 7-AAD. For the absolute cell count, fluorescent microparticles (Flow-Countk Fluorespheres, Coulter-Immunotech) of defined concentration were used. The assayed concentration of Flow-Countk Fluorespheres was described in each lot at delivery. Using a reverse pipetting technique, 100 Al of each sample was incubated for 15 min at room temperature with 10 Al aliquots of each mAb (e.g. CD3-ECD, CD4-FITC, CD71-PE and 7-AAD). Corresponding isotypic mAbs were used to define the cut-off which had to be less than 1%. After incubation the cells were resuspended in PBS containing 0.5% paraformaldehyde and were
The correct assayed concentration of fluorespheres was used as a calibration factor in the software program and the EPICS XL flow cytometer calculated the absolute cell count for the specimen automatically. The absolute cell counting was determined twice.
Absolute cell counts=Al ¼
total number of cells counted total number of fluorespheres counted flow count concentration
2.7. Comparison of flow cytometric analysis of T cell proliferation with BrdU-incorporation assay Additional experiments were performed to compare the flow cytometric method with the standard BrdU-incorporation ELISA procedure (n = 4). DCs and T cells (cultured DCs/Th cells or DCs/ CTLs or T cells alone) were cultured at increasing ratios of DC/T cells (1:10, 1:33, 1:100, 1:300). On day 6, T cells were harvested for the BrdU assay. On the same day and 6 h later, T cells of the same culture were harvested for flow cytometric analysis of absolute cell counting and the expression of CD71. 2.8. Statistical analysis Means, standard deviations and coefficients of variation were calculated where indicated. Statistically significant differences were assessed using one-way analysis of variance (MANOVA) or a paired t-test. The Pearson test was used for the calculation of correlations. Differences between data were considered statistically significant at p < 0.05. All tests were performed with commercially available software for personal computer (SPSS for Windows NT, SPSS software, Munich, Germany).
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
3. Results 3.1. DC generation from CD14+ cells CD14+ cells were cultured for 10 days in a specific medium and developed to fully mature DCs with a typical dendritic morphology. Characteristically, numerous cytoplasmic projections were present. The expression of CD83 was 76 F 13.9% (57 – 94). The mean yield of mature CD83+ DCs was 8 F 7% (range 1.6– 19%) of the plated cells. Flow cytometric analysis showed that these cells expressed little CD1a, but abundant quantities of CD80, CD86, CD40, HLA-DR and MHC class I. Furthermore, 17.3 F 11.1% (range 7– 29%) of the cultured cells expressed CD4, but were negative for CD3 and CD8. 3.2. Allogeneic mixed lymphocyte reaction 3.2.1. Cell morphology The DCs were tested for their ability to induce proliferation of allogeneic CD4+/CD45RA+ and CD8+/CD45RA+ cells in a mixed lymphocyte reaction. CD4+/CD45RA+ and CD8+/CD45RA+ cells were purified prior to the mixed lymphocyte reaction and cultured separately with DCs. DCs induced proliferation of responder T cells (Th cells as well as CTLs) when compared with responder cells alone. After 12 h, DCs formed clusters with T cells, whereas this phenomenon could not be observed in cultures of T cells alone. On day 3, cluster formation increased. When viewed by inverted microscopy, cocultured T cells apparently changed their morphology. They appeared to be larger and were heterogeneous. From day 5, the culture dish well was filled with proliferated cells which increased maximally on day 7. On subsequent days, clusters of dead cells were observed. 3.3. Flow cytometric analysis of T cell proliferation and activation Flow cytometry was used to determine activating and proliferating T cells in a mixed lymphocyte reaction with DCs. CD3+/CD4+ or CD3+/CD8+ cells were separately analysed. Fig. 1 shows the simultaneous flow cytometric analysis of cell viability (row a), percentage of CD3+/CD4+ cells (row b) and CD71 activation (row c) on days 0, 7 and 11.
61
The absolute cell count of each gate (viable cells, CD3+/CD4+, CD4+/CD71+) was calculated automatically using beads of known concentration. The same technique was used for analysis of CD3+/CD8+ cells (data not shown). T cell proliferation was determined by absolute cell counting. Fig. 2a and b shows the development of absolute cell count/Al of Th cells and CTLs over time, respectively. With DC stimulation, the maximal T cell proliferation showing typical characteristics with a high intensity of side and forward scatter was recorded on day 7, at which time a five- to tenfold T cell expansion was observed, whereas T cells alone decreased continuously. Significant differences in Th and CTL cell numbers between days 0, 3, 7 and 11 were confirmed by the MANOVA test with p < 0.0001. The average standard deviation of flow cytometric quantification analysis was 2.2% (coefficient of variation = 2.2%). Th cell activation was determined by analysis of the expression of CD25, CD71, CD45RO and HLADR. Fig. 3a and b shows the development of the activation molecules of Th cells over time. In accordance with the absolute cell numbers, the maximal expression of CD71 was recorded on day 7. In addition, the expression of CD71 began to increase as early as on day 1 before cell proliferation (7.6 F 2.4% versus 0.3 F 0.3% on day 0). This activation molecule showed a strong correlation with the absolute Th cell count. The expression of HLA-DR, CD25 also increased continuously and reached a maximum on day 9 (Fig. 3a). Cell viability decreased continuously after day 7. The expression of CD45RA (naive Th cells) and CD45RO (memory Th cells) developed inversely. During the stimulation period, Th cells lost their CD45RA expression and developed into memory Th cells with a high expression of CD45RO (Fig. 3b). No significant changes of the activation molecules of T cells without DC stimulation were observed during the culture period, but cell viability decreased earlier (data not shown). Other activation molecules, such as CD38 and CD69 (an early activation marker) were found not to be helpful and were not further assessed (n = 2). These markers increased even on T cells alone without DCs after 3 days in culture (data not shown).
62
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Fig. 1. Flow cytometric protocol for analysis of CD3+/CD4+ cell proliferation in mixed lymphocyte reaction with DCs on days 0, 7 and 11. In row (a), dead cells were excluded using 7-AAD (emission spectrum of 650 nm). The rectilinear gate was set to include viable cells. All viable cells were gated back in row (b) and analysed for CD3+/CD4+ cells. In row (c), only viable CD3+/CD4+ cells appeared for analysis of CD71 expression. In an extra diagram of the same protocol (diagram not depicted in figure), beads with a broad fluorescence emission range (510 – 700 nm) were easily gated in a forward scatter versus the fourth fluorescence channel. The absolute cell count of each gate (viable cells, CD3+/CD4+, CD4+/CD71+) was automatically calculated relative to beads of known concentration and given as cell counts/ Al. Thus, cell viability, absolute cell counting and activation were determined simultaneously. At the beginning of cell culture (day 0), almost 100% of the cells were viable. The flow cytometric analysis of the morphology showed a small population of CD3+/CD4+ cells without expression of CD71. On day 7, more than 95% of cells were still viable with a high intensity of side scatter (also with a high intensity of forward scatter, diagram not shown in figure). The population of CD3+/CD4+ cells appeared larger. In parallel, CD71 expression increased strongly. On day 11, most of CD3+/CD4+ cells were dead correlating with a low value of CD71 (analysis of CD3+/ CD8+ cells not shown as figure).
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
63
Fig. 2. The course of absolute T cell count/Al during the whole stimulation period with DCs (n = 6) shown. Allogeneic CD3+/CD4+ (Th cells) and CD3+/CD8+ cells (CTLs) were purified prior to the mixed lymphocyte reaction and cultured separately with DCs. The analysis of both cell lineage was successively performed. Th and CTL cell count (Fig. 1a and b, respectively) increased strongly and reached a maximum on day 7 ( p < 0.00001 and p < 0.001, respectively) whereas T cells alone decreased continuously. The error bars indicate mean F 2 S.D.
According to the activation of CTLs, they showed a similar development like Th cells (data not depicted as figure). The maximal expression of CD71 was recorded on day 7 with 82.4 F 11 (64.3 – 94%). A strong correlation between the expression of this activation molecule and the absolute cell count was also observed (r = 0.87, p < 0.01). The expression of
HLA-DR, CD25 increased continuously and also reached a maximum on day 9 with 14.1 F 1.8 (11.8 –16%) and 83.8 F 14.6 (58 – 93%), respectively ( p < 0.0001 for both markers, in comparison with day 0). In contrast, cell viability decreased continuously from day 7 with 68.4 F 8.1 (58.2 – 81.4%) on day 9, and only 40.8 F 15.2 (24.3 –63.3%) on day 11.
64
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Fig. 3. The course of viability and activation of CD3+/CD4+ cells during the stimulation period with DCs (n = 6) depicted. Each marker (CD25, CD71, CD45RO, CD45RA and HLA-DR) was simultaneously determined with cell viability (7-AAD) and CD3/CD4 marker. Box (a) shows an increasing expression of CD25, CD71 and HLA-DR with p < 0.0001. CD71 expression reached a maximum on day 7. In comparison to CD25 and HLA-DR expression, each of which did not reach a maximum before day 9, CD71 correlated best with the absolute cell count (r = 0.95, p < 0.001). Box (b) shows the development of cell viability and CD45RA and CD45RO expression over time. The expression of CD45RA (naive Th cells) and CD45RO (memory Th cells) developed inversely. During the stimulation period, Th cells lost CD45RA and developed into memory Th cells with a high expression of CD45RO ( p < 0.0001) which did not significantly change after day 9. From day 7, cell viability decreased constantly and more than the half of the cells were dead on day 11 (data of CD3+/CD8+ cells not shown). The error bars indicate mean F 2 S.D.
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
65
Fig. 4. Comparison of the BrdU-incorporation ELISA procedure (a) and flow cytometric analysis of absolute Th cell count (b) and CD71 expression (c). Different ratios of DC/Th cells were cultured and harvested on day 6. Similar to the development of CD3+/CD4+ cell concentrations and the expression of CD71, BrdU incorporation in proliferating cells decreased constantly with an increasing DC/Th cell ratio. Th cell count/Al as well as CD71 expression showed a significant correlation with BrdU incorporation (r = 0.89, p < 0.001; and r = 0.94, p < 0.001, respectively). The A450 value of Th cells alone was lower than 0.05 (data of CTLs not shown). The error bars indicate mean F 2 S.D.
66
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
3.4. Comparison of T cell proliferation analysis with the BrdU-incorporation ELISA procedure The BrdU-incorporation ELISA procedure was compared with the T cell proliferation assay with regard to absolute cell counts and CD71 expression. Fig. 4a shows decreasing BrdU incorporation with increasing DC/Th cell ratios. Similar trends for Th cell counts/Al and CD71 expression data were also observed (Fig. 4b and c). Th cell count/Al as well as CD71 expression both correlated significantly with BrdU incorporation (r = 0.89, p < 0.001; and r = 0.94, p < 0.001, respectively) (data of DC and CTLs measurement not shown).
4. Discussion The concept of in vitro stimulation and vaccination with DCs has led to a new field of immunotherapy for cancer (Lotze, 1997; Schadendorf and Nestle, 2001; Nguyen et al., 2002). Recent publications suggest that subsets of DCs differ in their antigen presentation characteristics (Kalinski et al., 1999; Arprinati et al., 2000; Jonmuleit et al., 2001). In order to check DC function, [3H] TdR incorporation assays have been used for the measurement of T cell proliferation in mixed lymphocyte reactions (Jonuleit et al., 1997; Thurner et al., 1999; Min et al., 2000). Using [3H] TdR or BrdU-ELISA incorporation assays, T cell proliferation is measured by incorporation of thymidine analogues into the DNA of proliferating cells. Thus, monitoring of DNA synthesis is an indirect parameter of cell proliferation. Although this technique is sensitive, it only provides information on the overall proliferative response and gives no insight into specific cell subsets involved. Furthermore, the incorporation of thymidine analogues does not always correlate with true increases in DNA content and cell proliferation (Neckers et al., 1995). It can be assumed that various DC subpopulations exist having distinct functions with respect to T cell activation or inhibition. Therefore, it is much more important to investigate the T cell response in the presence of DCs. Caruso et al. (1997) investigated the correlation between activation markers on stimulated T cells and their proliferation using flow cytometric analysis and [3H] TdR
incorporation assays. Both assays are complementary and a combination of both methods was recommended to obtain a clearer understanding of that events leading to efficient cell-mediated immune response (Caruso et al., 1997). The most accurate method to measure cell proliferation in vitro is to determine viable cells directly. In the present study, we developed a novel flow cytometric method for quantitative determination of T cell proliferation in mixed lymphocyte reactions with DCs using a combination of immunophenotyping and quantification. Our aim was to obtain rapid and precise information on cell viabilitity, activation and proliferation simultaneously in a single measurement. Furthermore, we sought to identify the activation marker correlating most closely with cell proliferation. Accordingly, the activation markers CD25, CD69, CD71 and HLA-DR (Caruso et al., 1997; Barten et al., 2001) were investigated. As T cells are nonadherent in culture, they were easily resuspended for single cell analysis by flow cytometry. The cultured DCs proved to be strong stimulators of naive T cells with high expression of costimulator molecules. As early as 12 h after initiation of the coculture, DCs and T cells formed a cluster. CD3+/CD4+ and CD3+/CD8+ cells began to increase their activation markers and, in addition, the expression of CD71 molecule increased earlier than cell proliferation. On day 7, maximal expansion of CD3+/ CD4+ and CD3+/CD8+ cells were recorded. In association with cell expansion over time, maximal expression of CD71 was observed on day 7. This activation marker showed the strongest correlation with absolute T cell counts. In fact, CD71 is upregulated on all cell types entering proliferation (Testa et al., 1993). In order to confirm this result, a BrdUincorporation ELISA procedure was performed. Using different ratios of DCs and T cells, the BrdU-incorporation ELISA procedure and flow cytometric analysis of cell proliferation and CD71 expression showed a strong correlation. Nevertheless, the [3H] TdR incorporation assay is the gold standard method with which to determine cell proliferation. The maximal expression of the activation markers CD25 and HLADR of both T cell lineages was observed on day 9. However, cell counts and cell viability decreased strongly by this time. Thus, it is apparently not feasible to investigate T cells after day 7.
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
The determination of CD38 and CD69 expression as measures of proliferation and activation have been recommended by other groups (Funaro et al., 1990; Maino et al., 1995; Cesano et al., 1998), but these markers did not prove to be useful in our study. We found that even naive T cells without DC stimulation strongly expressed CD38 and CD69 during the culture period. Interestingly, we observed that the viability of T cells without DC stimulation decreased more rapidly. DCs as stimulators protected T cells from early cell death. In fact, costimulation has a protective effect on T cell survival, probably due to the upregulation of anti-apoptotic mechanisms (Boise et al., 1995). Fluorescent microparticles were used to determine cell concentrations simultaneously without additional cell loss or other cell preparation steps. These microparticles were easily distinguishable from other events of interest. We have previously demonstrated that absolute cell counts determined using fluorescent microparticles were stable during the processing time (Schlenke et al., 1998). A coefficient of variation of 2.2% in the present study indicates the accuracy of the measurements. Many of the cultured DCs were CD4 positive but CD3 negative. Caux et al. (1992, 1995) showed that DCs cultured from CD34+ cells strongly expressed CD4. Against this background, the use of CD3-ECD for the analysis was indispensable in order to distinguish CD3+/CD4+ cells from DCs. This point has to be considered since there are many DC subpopulations differing in their expression of surface molecules, whereas no cell surface marker is unique to DCs (Kalinski et al., 1999; Arprinati et al., 2000; Jonmuleit et al., 2001). Priming of naive T cells by DC stimulation of T cells showed a high level of proliferation. Thus, the use of 7-AAD was important to exclude dead cells. In conclusion, this novel flow cytometric T cell assay offers a simple, rapid and precise quantification of T cell proliferation in mixed lymphocyte reactions with DCs. A simultaneous determination of absolute cell counts, expression of surface activation molecules and cell viability can be performed without any additional cell preparation. CD71 is a suitable activation marker which can be simultaneously determined. Further characterizations of additional surface markers involved in proliferation and activation processes are
67
feasible. This method will open broad field of applications as it is easy to perform and accurate.
Acknowledgements The authors are indebted to Daniella Griffiths for expert editorial assistance in the preparation of this manuscript.
References Arprinati, M., Green, C.L., Heimfeld, S., Heuser, J.E., Anasetti, C., 2000. Granulocyte-colony stimulating factor mobilizes T helper 2-inducing dendritic cells. Blood 95, 2484. Banchereau, J., Steinman, R.M., 1998. Dendritic cells and the control of immunity. Nature 392, 245. Barten, M.J., Gummert, J.F., van Gelder, T., Shorthouse, R., Morris, R.E., 2001. Flow cytometric quantification of calcium-dependent and -independent mitogen-stimulation of T cell functions in whole blood: inhibition by immunosuppressive drugs in vitro. J. Immunol. Methods 253, 95. Boise, L.H., Minn, A.J., Noel, P.J., June, C.H., Accavitti, M.A., Lindsten, T., Thompson, C.B., 1995. CD28 costimulation can promote T cell survival by enhancing the expression of Bcl-XL. Immunity 3, 87. Caruso, A., Licenzi, S., Corulli, M., Canaris, A.D., De Francesco, M.A., Fiorentini, S., Peroni, L., Fallacara, F., Dima, F., Balsari, A., Turano, A., 1997. Flow cytometric analysis of activation markers on stimulated T cells and their correlation with cell proliferation. Cytometry 17, 71. Caux, C., Dezutter-Dambuyant, C., Schmitt, D., Banchereau, J., 1992. GM-CSF and TNF-a cooperate in the generation of dendritic Langerhans cells. Nature 360, 258. Caux, C., Massacrier, C., Dezutter-Damuyant, C., Vanbervliet, B., Jacquet, C., Schmitt, D., Banchereau, J., 1995. Human dendritic langerhans cells generated in vitro from CD34+ progenitors can prime naive CD4+ T cells and process soluble antigen. J. Immunol. 155, 5427. Cesano, A., Visonneau, S., Deaglio, S., Malavasi, F., Santoli, D., 1998. Role of CD38 and its ligand in the regulation of MHCnonrestricted cytotoxic T cells. J. Immunol. 160, 1106. Frome, E.L., Smith, M., Littlefield, G., Neubert, R., Colyer, S., 1996. Statistical methods for the blood beryllium lymphocyte proliferation test. Environ. Health Perspect. 105, 957. Funaro, A., Spagnoli, G.C., Ausielo, C.M, Alessio, M., Roggero, S., Delia, D., Zaccolo, M., Malavasi, F., 1990. Involvement of the multilineage CD38 molecule in a unique pathway of cell activation and proliferation. J. Immunol. 145, 2390. Hart, D.N.J., 1997. Dendritic cells: unique leucocyte populations, which control the primary immune response. Blood 90, 3245. Jonmuleit, H., Schmitt, E., Steinbrink, K., Enk, A.H., 2001. Dendritic cells as a tool to induce anergic and regulatory T cells. Trends Immunol. 22, 394.
68
X.D. Nguyen et al. / Journal of Immunological Methods 275 (2003) 57–68
Jonuleit, H., Kuhn, U., Mu¨ller, G., Steinbrink, K., Paragnik, L., Schmitt, E., Knop, J., Enk, A.H., 1997. Pro-inflammatory cytokines and prostaglandins induce maturation of potent immunostimulary dendritic cells under fetal calf serum-free conditions. Eur. J. Immunol. 27, 3135. Kalinski, P., Hilkens, C.M.U., Wierenga, E.A., Kapsenberg, M., 1999. T cell priming by type-I and type-2 polarized dendritic cells: the concept of a third signal. Immunol. Today 20, 561. Lotze, M.T., 1997. Getting to the source: dendritic cells as therapeutic reagents for the treatment of patients with cancer. Ann. Surg. 226, 1. Maino, V.V., Suni, M.A., Ruitenberg, J.J., 1995. Rapid flow cytometric method for measuring lymphocyte subset activation. Cytometry 20, 127. Min, Y.H., Lee, S.T., Choi, K.M., Hahn, J.S., Ko, Y.W., 2000. Surface expression of HLA-DM on dendritic cells derived from CD34-positive bone marrow haematopoietic stem cells. Br. J. Haematol. 110, 385. Neckers, L.M., Funkhouser, W.K., Treel, J.B., Cossman, J., Gratzner, H.G., 1995. Significant non-s-phase DNA synthesis visualized by flow cytometry in activated and in malignant human lymphoid cells. Exp. Cell Res. 156, 429.
Nguyen, X.D., Eichler, H., Sucker, A., Hofmann, U., Schadendorf, D., Klu¨ter, H., 2002. Collection of autologous monocytes for dendritic cell vaccination therapy in metastatic melanoma patients. Transfusion 42, 428. Schadendorf, D., Nestle, F.O., 2001. Autologous dendritic cells for treatment of advanced cancer—an update. Recent Results Cancer Res. 158, 236. Schlenke, P., Frohn, C., Klu¨ter, H., Saballus, M., Hammers, H.J., Zajac, S.R., Kirchner, H., 1998. Evaluation of a flow cytometric method for simultaneous leucocyte phenotyping and quantification by fluorescent microspheres. Cytometry 33, 310. Steinman, R.M., 1991. The dendritic cell system and its role in immunogenicity. Annu. Rev. Immunol. 9, 271. Testa, U., Pelosi, E., Peschle, C., 1993. The transferrin receptor. Crit. Rev. Oncog. 4, 241. Thurner, B., Ro¨der, C., Dieckmann, D., Heuer, M., Kruse, M., Glaser, A., Keikavoussi, P., Ka¨mpgen, E., Bender, A., Schuler, G., 1999. Generation of large numbers of fully mature and stable dendritic cells from leukapheresis products for clinical application. J. Immunol. Methods 223, 1.