Efficient priming of antigen-specific cytotoxic T lymphocytes by human cord blood dendritic cells

Mariolina Salio1, Nicolas Dulphy1, Joelle Renneson1, Mark Herbert2, Andrew McMichael2, Arnaud Marchant2 and Vincenzo Cerundolo1

1 Cancer Research UK Tumour Immunology Unit and 2 Human Immunology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, OX3 9DS, Oxford, UK

Correspondence to: M. Salio; E-mail: msalio{at}molbiol.ox.ac.uk or V. Cerundolo; E-mail: vincenzo.cerundolo{at}imm.ox.ac.uk
Transmitting editor: E. Simpson


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Previous studies have suggested that defective immune responses in early life may be related to the immaturity of neonatal antigen-presenting cells. To test this hypothesis, we assessed the capacity of neonatal dendritic cells (DC) to prime and polarize in vitro human naive antigen-specific T cells. We report that mature cord blood DC efficiently prime an oligoclonal population of antigen-specific CD8 T cells, capable of cytolytic activity and IFN-{gamma} secretion. In contrast, cells primed by immature cord blood DC do not acquire cytolytic activity and secrete lower amounts of IFN-{gamma}. Upon priming by either immature or mature DC, neonatal T cells acquire markers of activation and differentiation towards effector-memory cells. Our results demonstrate that, if appropriately activated, neonatal DC can prime efficient cytotoxic T lymphocyte (CTL) responses. Furthermore, these findings have important implications for the development of vaccine strategies in early life and for the reconstitution of a functional CTL repertoire after bone marrow transplantation.

Keywords: neonatal immunity, T cell polarization, tetramer


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Neonates and young children show higher morbidity and mortality rates from infectious diseases than adults. The higher susceptibility of neonates to pathogens as well as their poor responses to vaccines has been related to the immaturity of the neonatal immune system. The mechanisms underlying this immaturity are still unclear. It has been shown that neonatal T cells weakly proliferate in response to anti-CD3 stimulation, and have Th2-biased responses, lower up-regulation of CD40 ligand (CD40L) upon activation and limited cytotoxic activity (1). In addition, studies on virus-specific cytotoxic T lymphocytes (CTL) indicate that CD8 responses are reduced in numbers in neonates as compared to adults and cells secrete less IFN-{gamma} (24). However, in the presence of strong adjuvants or co-stimulatory signals, neonatal CD4 and CD8 T cells are capable of adult-level responses (510). The reduced ability of young infants to develop Th and CTL responses could be due to the immaturity of dendritic cells (DC) that are the main antigen-presenting cells, unique in their ability to efficiently prime and polarize naive CD4 and CD8 cells (11). Indeed, it has been reported that human neonatal monocyte-derived DC have defective production of IL-12 and expression of co-stimulatory molecules, which may account for the Th2 bias and the inefficient T cell stimulatory capacity (1216). However, other investigators have differentiated functional and mature DC from human cord blood progenitors (1719), and it has been recently shown that mouse neonatal DC are capable of activating CTL responses both in vitro and in vivo (20). However, the analysis of human antigen-specific CTL priming by cord blood DC has been hampered by the low frequencies of naive T cells. To address this question we set up an in vitro priming model, which we have previously used to expand naive cells from adult blood (21,22). We report that human neonatal DC efficiently prime naive melan-A-specific CD8+ T cells, capable of cytolytic activity and IFN-{gamma} secretion. This observation has important implications for vaccination in early life and for the reconstitution of the CTL repertoire after bone marrow transplantation. In addition, our results provide a cellular mechanism to account for the recently reported in utero expansion of Trypanosma cruzi- and human cytomegalovirus (HCMV)-specific CTL responses (5,23).


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Cell lines and cultures
The medium used throughout was RPMI 1640 supplemented with 2 mM L-glutamine, 1% non-essential amino acids, 1% pyruvate, 50 µg/ml kanamycin, 5 x 10–5 M 2-mercaptoethanol (Gibco) and 10% FCS (Hyclone) or 5% pooled human serum AB+ (UK National Blood Service). JY is an HLA-A2+ lymphoblastoid cell line. Melanoma lines Na8 (HLA-A2+, melan-A) and HBL (HLA-A2+, melan-A+) were a gift of Dr Elisabetta Padovan (Kantonspital, Basel, Switzerland). Recombinant human IL-2 and IL-4 were produced in our laboratory as described (24).

Peptides and tetramers
Melan-A26–35 ELAGIGILTV is an analogue of the 26–35 epitope with an improved HLA-A2 binding affinity (25). Peptides were purchased from Genosys (Sigma) and were purified by HPLC. Melan-A/HLA-A2 tetrameric complexes were synthesized as previously described (26,27). Tetramers were tested against positive CTL clones and background levels of staining (<0.02%) were defined by staining the peripheral blood mononuclear cells (PBMC) of HLA-A2 healthy donors.

Generation and stimulation of DC
Cord blood samples were obtained from the John Radcliffe Hospital maternity unit, upon written consent and approval by the ethical committee. Samples were screened for HLA-A2 expression by FACS analysis. Monocytes were purified by positive selection using anti-CD14-conjugated magnetic microbeads (Miltenyi). The recovered cells were >95% CD14+ as determined by flow cytometry with the anti-CD14 antibody TIB228 (ATCC, Rockville, MD). DC were generated as previously described (28) by culturing monocytes in RPMI/10% FCS supplemented with 50 ng/ml granulocyte macrophage colony stimulating factor (Leucomax; Novartis Pharma) and 1000 U/ml IL-4 for 5 days; 5% human serum was used instead of FCS in some experiments, but we did not observe any qualitative or quantitative difference in the priming experiments. Cells (3 x 105/ml) were stimulated by addition of either 1 µg/ml lipopolysaccharide (LPS; from Salmonella abortus equi; Sigma), mock transfected or CD40L-transfected J558 cells [at a 1:5 ratio; provided by P. Lane, Birmingham, UK (29)]. The CD14 fraction was frozen and thawed on the day of the in vitro priming, a few hours before addition to the autologous DC.

T cell priming
DC were pulsed for 3 h with melan-A26–35 peptide in serum-free medium. Cells were thoroughly washed and incubated with the autologous CD14 fraction at a 1:5 ratio in RPMI/5% human serum. Recombinant human IL-2 was added from day 4 to 7 at 10 U/ml. Cells were then expanded with 500 U/ml IL-2 and analyzed at day 10–15.

FACS analysis
Cells were stained in PBS with phycoerythrin (PE)- or allophycocyanin (APC)-labelled melan-A tetramer at 37°C for 20 min, washed at room temperature and incubated on ice with one of the following antibodies: CD8–FITC (Dako), CD8–PerCP (Becton Dickinson), 2B4–FITC (Immunotech), CD8–APC, CD27–FITC, CD45RO–APC, CD45RA–FITC, CD62L–PE, CLA–FITC, CCR7–FITC, CD95 FITC, CD38–APC, class II–FITC and CD28–APC (all from PharMingen); TCR-BV-specific antibodies were purchased from Serotec. The samples were analysed on a FACSCalibur (Becton Dickinson) using CellQuest Software. Lymphocytes were gated according to FSC/SSC profile and dead cells were excluded by staining with propidium iodide (Sigma).

Expression of melan-A and HLA-A2 was tested using mouse mAb A103 (Novocastra) and BB7.2 (ATCC) respectively, followed by a PE-conjugated affinity purified goat anti-mouse antibody (Southern Biotechnology Associates).

For intracellular cytokine detection, 106 cells were labelled with melan-A tetramer and subsequently stimulated with 20 µM melan-A peptide in RPMI/10% FCS or left unstimulated (30). Control cells were either unstimulated or treated with 10–6 M phorbol myristate acetate (PMA; Sigma) and 0.5 µM ionomycin (Sigma). Brefeldin A (Sigma) was added at a final concentration of 5 µM during the second hour of stimulation. Cells were harvested after a total of 6 h, washed, fixed and permeabilized in FACS Permeabilizing Buffer (Becton Dickinson) according to the manufacturer’s instructions. Staining was performed with anti-CD8–PerCP (Becton Dickinson), anti-IFN-{gamma}–FITC, tumor necrosis factor (TNF)-{alpha}–FITC, IL-2–FITC, IL-4–PE, IL-10–PE and IL-13–PE (PharMingen).

Cytokine determination
The concentration of IL-12 p75 and p40 in DC supernatants was measured by ELISA (PharMingen). The sensitivity of the assay was 400 pg/ml.

Spectratyping and CDR3 size analysis
CD8+ T cells were positively selected with MACS beads (Miltenyi), with a purity >95% as assessed by flow cytometry. Total RNA was extracted from 3 x 106 CD8+ cells in guanidine thiocyanate buffer (RNA-Bee; Biogenesis). The cDNA was then prepared with MMLV reverse transcriptase (Invitrogen) as previously described (31).

The nomenclature of BV and the BV- or BC-specific primers used has been previously described (32,33). The 24 PCR reactions for each cDNA were performed according to Garderet et al. (32). Amplification products were labelled in an extension reaction, so-called run-off reaction, using a BC-specific Fam-fluorescent probe. The fluorescent run-off products, together with a Tamra-fluorescent DNA weight marker (Genescan 500; Applied Biosystems), were loaded on sequence gel in an automated DNA sequencer (Perkin Elmer). CDR3 sizes and fluorescent intensities were analysed with Genescan software (Perkin Elmer).

Cytolytic activity
Cytolytic activity was assessed using a Cr-release assay. JY EBV or melanoma cells were labelled with 51Cr for 90 min at 37°C and washed twice. Labelled cells were peptide pulsed for 1 h, washed and added (5000 cells/well) to graded numbers of CD8+ purified effector cells. CD8+ cells were purified from the in vitro priming with Miltenyi CD8 microbeads according to the manufacturer’s instructions. Chromium release was measured in the supernatant, which was harvested after 5 h of incubation at 37°C. Total release was determined in the presence of 5% Triton X-100. The percent specific lysis was calculated as follows: 100 x (experimental – spontaneous release)/(total – spontaneous release). Each value was calculated as the average of triplicates.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Melan-A tetramer+ CTL precursors detectable in newborn blood can be efficiently primed by autologous DC
To assess the ability of human neonatal DC to prime antigen-specific CD8+ T cells we used as a model antigen melan-A/MART1, a melanoma antigen which encodes for the HLA-A*0201-restricted immunodominant epitope 26–35 (34). We have previously reported that melan-A tetramer+ cells can be detected in the peripheral blood of most healthy blood donors (35,36). These cells have the characteristics of naive cells, both functionally and at the molecular level (37), and can be efficiently primed only by professional antigen-presenting cells, such as mature DC (21). This large pool of antigen-specific T cells is already detectable in newborns (37) and we found melan-A tetramer+ cells in all HLA-A2+ newborns tested, with frequencies ranging from 0.02 to 0.12%, while all HLA-A2 newborns had frequencies below the threshold of 0.02% (Fig. 1). These cells were CCR7+, CD27+, CD45RA+, CD45RO, consistent with their naive phenotype [data not shown and (37)].



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Fig. 1. Melan-A tetramer+ cells can be detected ex vivo in HLA-A2+ newborn peripheral blood. Fifteen cord blood samples, of which nine were HLA-A2+, were stained with melan-A tetramer–PE and CD8–FITC. The frequency of CD8+ tetramer+ cells is shown. One million events per sample were acquired in the live gate.

 
Since recent studies indicate that neonatal DC are less efficient than their adult counterpart in stimulating allogeneic T cells (1214,16) we investigated whether melan-A tetramer+ cells could be primed by autologous cord blood DC as observed in adult donors (21). The phenotype of monocyte-derived cord blood DC used for these experiments, before and after maturation with LPS and CD40L, is shown in Table 1. As shown in Fig. 2, at high peptide dose, melan-A tetramer+ cells were expanded by both immature and mature cord blood DC, the most potent stimulus being CD40L-matured DC, followed by LPS-matured cells. In contrast, when pulsed with lower concentrations of melan-A26–35 peptide (10 ng/ml), mature DC were 20-fold more efficient than immature DC in expanding antigen-specific CTL (data not shown). Melan-A-primed CTL acquired markers of activation and a phenotype similar to effector cells at an intermediate stage of differentiation (38) (Fig. 3A): they became CD45RO+, CD95+, 2B4 + and mostly HLA class II+, while they remained CD38+ and CD27+. CCR7 and CD28 expression was lost on most of the cells. There was no expression of the inhibitory receptors DX9, DX31, CD94/NKG2A, ILT2 and CD158 (data not shown). When primed by CD40L-matured cord blood DC, 22% of the cells expressed CLA and 39% CD62L (Fig. 3B), homing receptors for inflamed skin, whose expression is known to be IL-12 dependent (39,40). Although we have observed similar expansions of melan-A specific CTL upon priming by mature adult DCs, the latter induced higher percentages of CD62L and CLA expression (21). As previously reported (12) this difference may be accounted for by the 4–6 times lower secretion of IL-12 p75 by CD40L-matured cord blood DC, as compared to adult DC, in spite of similar amounts of IL-12 p40 (data not shown).


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Table 1. Phenotype of cord blood DC used for in vitro priming
 


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Fig. 2. Efficient expansion of melan-A tetramer+ cells from newborn PBMC by autologous DC. Immature (top panels), LPS-matured (middle panels) or CD40L-matured cord blood DC (bottom panels) were pulsed with 10 µg/ml melan-A26–35 peptide and incubated with autologous peripheral blood lymphocytes at a 1:5 ratio. After 10 days, cultures were stained with melan-A tetramer–APC and CD8–PerCP and the percentages of CD8+ tetramer+ cells are shown. One representative experiment of four is shown.

 


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Fig. 3. Phenotype of melan-A-specific CTL primed by cord blood DC. (A) LPS-matured cord blood DC were pulsed with 10 µg/ml melan-A26–35 peptide and incubated with autologous peripheral blood lymphocytes at a 1:5 ratio. After 10 days, cultures were stained with melan-A tetramer–PE, CD8–PerCP and the following antibodies: CD28–FITC, CD27–APC (a), CCR7–FITC, CD27–APC (b), CD27–APC, 2B4–FITC (c), CD45RA–FITC, CD45RO–APC (d), CD95–FITC (e) and HLA-DR–FITC, CD38–APC (f). The profiles refer to tetramer+ cells only, but the quadrants were set on the entire population. (B) CLA and CD62L expression was analysed on melan-A CTL primed by immature (top panels), LPS-matured (middle panels) or CD40L-matured cord blood DC (bottom panels).

 
All together, our results demonstrate that when appropriately activated cord blood DC can be powerful antigen-presenting cells.

Functional analysis of melan-A tetramer+ CTL primed by cord blood DC
To assess the antigen responsiveness of the tetramer+ cells, cord blood DC-primed cultures were stimulated with melan-A26–35 peptide in a 6-h assay. As shown in Fig. 4(A), melan-A tetramer+ CTL mainly released IFN-{gamma} and low levels of TNF-{alpha}, showing a type I polarization pattern. The percentages of IFN-{gamma}-secreting cells and the level of the cytokine increased after priming by CD40-matured cord blood DC, as observed with adult DC. Tetramer+ CTL did not secrete IL-4, IL-10, IL-13 or IL-2 (Fig. 4A and data not shown) in any of the experiments, not even when primed by immature cord blood DC. To address the type of polarization induced in the entire population, the cytokine profile was analysed in response to PMA and ionomycin. Upon priming by immature and LPS-matured cord blood DC, CD8+ cells mainly released IFN-{gamma} and low levels of TNF-{alpha} (Fig. 4B). CD8 cells secreted IL-2, IL-4, IL-13 and minimal amounts of IL-10, in addition to IFN-{gamma} and TNF-{alpha} (Fig. 4B and data not shown). CD40L-matured cord blood DC induced a strong type I polarization, both of the CD8 and CD8+ cells.




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Fig. 4. Cord blood DC prime antigen-specific CTL secreting type I cytokines. Melan-A tetramer+ cells primed by autologous cord blood DC were re-stimulated with 20 µg/ml melan-A26–35 peptide (A) or PMA and ionomycin (B) as described in Methods. Intracellular cytokine staining is shown for CD8+ tetramer+ cells (A) or for the whole population (B). One representative experiment of four is shown.

 
The cytolytic activity of melan-A tetramer+ CTL was assessed against an HLA-A2+ EBV-B cell line and melanoma lines expressing or not the melan-A antigen. As shown in Fig. 5, CTL primed by immature cord blood DC did not acquire cytolytic activity against peptide-pulsed EBV-B cells, despite high numbers of antigen-specific cells (Fig. 5A). In contrast, cells primed by mature cord blood DC were highly cytolytic and recognized both peptide-pulsed EBV-B cells and melan-A+ melanoma cells (Fig. 5B and D).



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Fig. 5. Mature cord blood DC prime antigen-specific CTL endowed with cytolytic activity. Melan-A CTL were expanded by autologous immature (A), LPS-matured (B) or CD40L-matured cord blood DC (C and D) pulsed with 10 µg/ml melan-A26–35 peptide. At day 14, CD8+ cells were purified and tested in a 51Cr-release assay against JY, an HLA-A2+ EBV line, pulsed with 10 µg/ml melan-A26–35 peptide (solid squares) or unpulsed (open squares). The frequencies of melan-A tetramer+ cells after CD8+ enrichment were 11 (A), 16 (B), 4 (C) and 24% (D), and these numbers were used to define the E:T ratio for each panel. In (D), two melanoma lines were also tested: Na8 (open circles, HLA-A2+, melan-A) and HBL (solid circles, HLA-A2+, melan-A+). Cells used in (C) and (D) derived from two different donors. One representative experiment of four is shown.

 
Clonal composition of melan-A tetramer+ CTL primed by cord blood DC
We determined the usage of TCR BV chains of melan-A tetramer+ CTL by spectratype analysis of newborn CD8+ T cells unprimed, primed by unpulsed or pulsed CD40L-matured cord blood DC (Fig. 6A). As previously described (32), unprimed CD8+ T cells showed a Gaussian distribution of TCR CDR3 length within all BV families (Fig. 6A and data not shown). The profile of CD8+ T cells primed by unpulsed CD40L DC showed some expansions in certain BV families, compatible with the strong stimulatory capacity of these cells. In contrast, important oligoclonal expansions of CD8+ T cells expressing BV 3, 7 and 16 were observed upon priming by pulsed CD40L DC (Fig. 6A, cf. left, middle and right panels). The magnitude of these expansions and the size of the CDR3 regions are different from those observed upon priming by unpulsed DC, clearly showing a specific T cell response to the melan-A peptide. The specificity of the oligoclonal expansion was confirmed by FACS analysis of the same cultures, using BV-specific antibodies combined with melan-A tetramer staining (Fig. 6B).




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Fig. 6. Mature cord blood DC expand an oligoclonal melan-A population. (A) Spectratype analysis performed on MACS-purified CD8+ cells unprimed (left panels), primed by unpulsed (middle panels) or pulsed (right panels) CD40L cord blood DC. The frequency of melan-A tetramer+ cells in the pulsed sample after CD8+ enrichment was 24%. Only the BV families in which an expansion was detected are shown. (B) The same cultures expanded by CD40L cord blood DC either unpulsed (left) or pulsed with melan-A peptide (right) were stained with melan-A tetramer–PE and FITC–BV-specific antibodies to confirm the specificity of the expansion.

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The immaturity of the neonatal immune system has been considered the major obstacle to successful vaccinations in early life and may account for the higher susceptibility of newborns to infectious diseases (41). It is still unclear whether the cause of such immaturity depends on intrinsic T cell defects or is mainly due to the nature and quality of antigen presentation (1,42). As DC are the most potent antigen-presenting cells we investigated whether human cord blood DC could prime functional antigen-specific CTL. Experiments were carried out to assess the expansion and activation of melan-A-specific T cells in an in vitro priming model that we have previously used for adult DC (21). Melan-A is a melanoma antigen which encodes for the HLA-A*0201-restricted immunodominant epitope 26–35 (34). In contrast to the low frequency of most antigen-specific T cells in the pre-immune repertoire, up to 1:1000 naive CD8 T cells from A2 individuals specifically bind melan-A tetramers (35). Because of the unusual high precursor frequency, up to date this represents the only naive antigen-specific T cell repertoire accessible to direct analysis in humans. It has been unequivocally demonstrated by cellular and molecular methods that melan-A precursors are truly naive cells (3537). This large pool of T cells specific for a self-antigen is generated by thymic output of a high number of precursors (37) and this has been explained by the existence of largely cross-reactive subsets of naive CD8 T cells displaying multiple specificities (43).

We showed that mature cord blood DC efficiently primed melan-A specific T cells, which acquired markers of activated cells and were capable of cytolytic activity towards peptide-pulsed targets and tumour cells. In contrast, cells primed by immature cord blood DC did not acquire cytolytic activity, despite expression of activation markers. We confirmed the findings that cord blood DC secrete less bioactive IL-12, as compared to adult DC (data not shown), and this may explain the lower CLA and CD62L expression observed in cells primed by cord blood DC. Indeed, the expression of these homing receptors is known to be IL-12 dependent (21,39,40).

It is generally believed that neonates are more prone to Th2 immune responses, although CTL and Th1 responses can be elicited using strong known type I adjuvants (8,4446). We have observed a strong type I polarization of CD8 cells primed by CD40L-matured cord blood DC, and lower levels of IFN-{gamma} and TNF-{alpha} when cells were primed by immature cord blood DC. In contrast, CD8 cells secreted both Th1 and Th2 cytokines when primed by immature or LPS cord blood DC, but a strong Th1 polarization was induced by CD40L cord blood DC. These results suggest that although cord blood DC secrete lower levels of bioactive IL-12, this may be sufficient to elicit type I immune responses (47) or, alternatively, other cytokines may be responsible for T cell polarization in our in vitro system.

By spectratype analysis we observed an oligoclonal expansion of melan-A tetramer+ CTL primed by cord blood DC, hence confirming and extending previous results obtained in melanoma patients (48,49).

The demonstration that mature cord blood DC can efficiently expand functional CTL in vitro provides a powerful strategy to support immune reconstitution of individual antigen specificities (i.e. HCMV CTL) after cord blood transplantation. Moreover, understanding the dynamics of DC–T cell interactions is of fundamental importance in improving the success rates of cord blood transplantation and further reducing the risk of graft versus host disease, e.g. by promoting the expansion of T regulatory cells (50).

Altogether, our results suggest that the weak CTL responses observed in young infants may not be due to an intrinsic defect in the antigen-presenting cells nor in the T cells. Neonatal T cells may have more stringent requirements for co-stimulatory signals than adult T cells (1); however, when appropriately activated, neonatal DC can prime efficient CTL and Th1 responses (20,44). With regard to this, and in agreement with the study by Hermann on T. cruzi-infected newborns (5), we have recently observed that fetuses infected in utero with HCMV develop a mature and functional CD8 T cell response (23). It has been demonstrated that CD4 help is required for the generation of long-lived CTL responses (51). Since we have observed that HCMV-specific CTL present in newborn after in utero infection persist throughout the first year of life (A. Marchant, in preparation), it is likely that cord blood CD4+ T cells are capable of providing T cell help for the generation of CD8+ memory responses. The challenge for the future relies in identifying adjuvants and delivery systems that allow efficient neonatal DC activation and migration to achieve optimal DC–T cell interactions in vivo.


    Acknowledgements
 
This work was funded by grants from Cancer Research UK (CRUK Project Grant C399/A2291) and the Cancer Research Institute.


    Abbreviations
 
APC—allophycocyanin

CTL—cytotoxic T lymphocyte

DC—dendritic cell

HCMV—human cytomegalovirus

LPS—lipopolysaccharide

PBMC—peripheral blood mononuclear cell

PE—phycoerythrin

PMA—phorbol myristate acetate

TNF—tumor necrosis factor


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
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