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The Journal of Immunology, 1998, 160: 3689-3697.
Copyright © 1998 by The American Association of Immunologists

Phorbol Esters Induce Differentiation of Human CD34+ Hemopoietic Progenitors to Dendritic Cells: Evidence for Protein Kinase C-Mediated Signaling1

Thomas A. Davis*,{dagger}, Abha A. Saini*, Patrick J. Blair*, Bruce L. Levine*, Nancy Craighead*, David M. Harlan*,{dagger}, Carl H. June*,{dagger} and Kelvin P. Lee2,*,{dagger}

* Immune Cell Biology Program, Stem Cell Biology Branch, Naval Medical Research Institute, Bethesda, MD 20889; and {dagger} Uniformed Services University of the Health Sciences, Bethesda, MD 20889


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The intracellular signals that mediate the differentiation of pluripotent hemopoietic progenitors to dendritic cells (DC) are largely undefined. We have found that the phorbol ester PMA by itself induced 47% ± 8.7% of input human CD34+ hemopoietic progenitors to differentiate into cells with morphology and surface Ag phenotype characteristic of DC by day 7 of culture. Functionally, PMA-generated DC processed and presented whole soluble Ag and also induced resting T cell proliferation and Ag-specific CTL effector function. Unlike cytokine-driven DC differentiation, PMA suppressed proliferation and induced cell death (in part via apoptosis) in cells that did not differentiate to DC. The effects of PMA were blocked by inhibitors of protein kinase C activation, suggesting a central role for this signaling molecule. PMA-mediated signaling also induced expression of the RelB transcription factor, an NF-{kappa}B family member implicated in DC differentiation. These findings suggest that phorbol esters activate protein kinase C, which then initiates the terminal component of an intracellular signaling pathway(s) involved in the DC differentiation of CD34+ hemopoietic progenitors.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Antigen uptake, processing and presentation by professional APCs are requisite steps in both the activation of CD4+ T cells and the initiation of the primary immune response. Among the professional APC, dendritic cells (DC)3 are thought to play the pivotal role in Ag presentation to, and activation of, naive T cells (1, 2). In humans, DC are thought to arise from the CD34+ hemopoietic progenitor cell population (3, 4, 5) and can be generated in vitro using combinations of cytokines, such as granulocyte-macrophage CSF (GM-CSF), TNF-{alpha}, IL-4, and stem cell growth factor (SCF) (3, 6, 7, 8, 9), or by CD40 cross-linking (10). Cytokine treatment of more differentiated CD14+ peripheral blood monocytes also gives rise to DC (11, 12, 13, 14, 15).

Despite these in vitro culture systems, the specific molecular mechanisms of lineage commitment of CD34+ progenitors to DC are not well defined. Because cytokine receptor stimulation activates complex signaling cascades that initiate multiple responses, it is difficult to separate the components specifically involved in lineage commitment from those involved in cell proliferation, for example. In addition, differentiation to multiple lineages is simultaneously induced; in the case of CD34+ cells GM-CSF plus TNF-{alpha} ± SCF not only generates DC but also macrophages and neutrophils as well (3, 6, 7, 8, 9). This lack of specificity hampers the identification of signaling pathways involved in the differentiation of a specific lineage. At the other end of the signaling pathways, studies examining nuclear events have implicated rel/NF-{kappa}B-responsive genes and the RelB transcription factor in terminal DC differentiation (16, 17, 18).

One component of the IL-4-, TNF-{alpha}-, and CD40-mediated signaling pathways is the activation of protein kinase C (PKC) (19, 20, 21, 22, 23). Although not described for DC, PKC activation induces differentiation in other hemopoietic cell lineages (24, 25, 26, 27, 28, 29, 30, 31, 32). Activated PKC can phosphorylate/activate a number of downstream signaling molecules (33, 34), including RelB and other members of the rel/NF-{kappa}B transcription factor family (35, 36, 37) and members of the ras/raf-1/mitogen-activated protein kinase cascade (34).

One strategy to study signal transduction in DC differentiation is to bypass membrane proximal events with agents that directly activate intracellular molecules suspected to be part of relevant cytokine signaling pathways. PMA is a stable analogue of 2,3-diacylglycerol that activates the classical ({alpha}, ß1, ß2, {gamma}) and new ({delta}, {epsilon}, {eta}, {theta}, µ) isoforms of PKC (33). In the present study, we show that PMA by itself induces primary human CD34+ bone marrow (BM) progenitor cells to differentiate into functional DC. The effects of PMA may be blocked by PKC inhibitors, suggesting that activation of this signaling molecule is required. PMA-mediated signaling also induced the expression of the RelB transcription factor, suggesting a pathway by which genetic events involved in DC differentiation are initiated.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Purification of CD34+ BM cells

Human vertebral body BM was procured from cadavers as part of an existing tissue procurement protocol that has been previously described (38). Briefly, lumbar vertebral BM was obtained from the bone matrix by sterile technique and placed in culture support media. Low density mononuclear cells were separated over Ficoll-Hypaque. CD34+ BM progenitor cells were purified by positive immunomagnetic selection using a biotinylated mAb specific for the CD34 Ag (K6.1) that was linked to magnetic Dynabeads (Dynal, Great Neck, NY). After three to four cycles of magnetic attraction, the beads were disassociated from the cells with an excess of biotin (Life Technologies, Grand Island, NY) and separated from the cells magnetically. Cells by this procedure were >95% CD34+ by flow cytometric analysis using noncross blocking Ab (HPCA-2 PE, Becton Dickinson, San Jose, CA).

CD34+ cell culture for DC

CD34+ BM cells were cultured in 96-well plates containing complete culture medium (Iscove’s modified Dulbecco’s medium (Life Technologies)) supplemented with 10% heat-inactivated FCS (HyClone, Logan, UT), 100 mM L-glutamine, and 100 U/ml penicillin/streptomycin solution (Life Technologies) for 7 days at a concentration of 5 x 104 cells/ml. Cultures were stimulated with 10 ng/ml PMA (Sigma, St. Louis, MO) in the presence and absence of GM-CSF plus IL-3, IL-6, and SCF and then incubated at 37°C in a humidified 5% CO2-in-air atmosphere for 7 days. Cytokines were used at the follow concentrations: 2 ng/ml GM-CSF, 5 ng/ml IL-3, 5 ng/ml IL-6, 120 ng/ml SCF, and 10 ng/ml TNF-{alpha} (all from R&D Systems, Minneapolis, MN). Where indicated, staurosporine (Calbiochem, San Diego, CA) was added at 0.1 ng/ml, a dose sufficient to block PMA-induced proliferation of human T cells. Mezerein was used at 100 nM, and bisindolylmaleimide I was used at 5 µM (both from Calbiochem). At the end of the culture period, both nonadherent cells and adherent cells were resuspended (3 mM EDTA), washed, and concentrated by centrifugation.

In experiments involving CD34+ viability, cells (5 x 104 in 0.5 ml of complete culture medium) were incubated for 24, 48, 72, and 96 h with PMA in the presence or absence GM-CSF plus IL-3, IL-6, and SCF and then assayed for cell viability by trypan blue dye exclusion.

Flow cytometric analysis and mAbs

Adherent and loosely adherent cells from the day 7 PMA-treated CD34+ BM cultures were harvested with 3 mM EDTA, washed twice, and resuspended in staining medium (PBS plus 5% FCS, 2% BSA, and 0.1% sodium azide). Phenotypic analysis of cells (2 x 105) was performed by flow cytometry using saturating concentrations of the following mAbs: CD1a (clone SK9), CD4 (SK3), CD13 (L138), CD14 (MOP9), CD34 (8G12), CD80 (B7-1) (all from Becton Dickinson), CD83 (HB15, a gift from Dr. T. Russel, Coulter, Miami, FL), CD86 (HF2.3D1, a gift from Dr. G. Gray, Genetics Institute, Cambridge, MA), MHC class I (H58A), and MHC class II (H42A) (both from VMRD, Pullman, WA). Appropriate conjugated, isotype-matched Abs were used as controls. To exclude subcellular particles, 10,000 cells were analyzed on a Coulter XL (Hialeah, FL) flow cytometer through a viable cell gate as determined by forward light scatter (FLS) and right-angle light scatter (RALS) parameters. The cytometer was calibrated using autocomp beads and software that were supplied by the manufacturer. The software used was Coulter XL software that was supplied and installed by the manufacturer.

T cell activation

Human peripheral blood leukocytes were obtained by leukophoresis from normal healthy adult donors. T cells were purified by negative selection as previously described (39) and resuspended in RPMI 1640 (Life Technologies) supplemented with 10% heat-inactivated FCS, 2 mM L-glutamine, 100 U/ml penicillin, 100 mg/ml streptomycin, and 20 mM HEPES. T cells were cultured at 1 x 105 cells/well with media alone (no stimulus), 3 µg/ml staphylococcal enterotoxin B (SEB) (Toxin Technology, Sarasota, FL), 5 µg/ml Con A (Calbiochem, La Jolla, CA), or 10 ng/ml PMA plus anti-CD28 (mAb 9.3, 1.0 µg/ml) in the absence (media) or presence of the indicated number of gamma-irradiated (3000 rad 137Cs) day 7 PMA-generated DC. Cultures were incubated for 3 days at 37°C in a humidified 5% CO2-in-air atmosphere. T cell proliferation was assessed after 0.5 mCi/well [3H]methyl thymidine (New England Nuclear, Boston, MA) had been added for the final 18 h of culture. Cells were harvested using a 96-well cell harvester, and [3H]methyl thymidine incorporation was measured using a Betaplate scintillation counting system (Pharmacia/LKB, Gaithersburg, MD). All determinations were performed in quadruplicate, and data are expressed as the mean cpm ± 1 SD. After 24 h, culture medium was collected and assayed for IL-2 by ELISA (R&D Systems); this data is expressed as pg of IL-2/ml per 5 x 105 T cells from the appropriate culture condition.

For the proliferation of autologous T cells to tetanus toxoid (TT), irradiated DC from day 7 PMA-treated CD34+ cell cultures were plated in triplicate wells of 96-well flat-bottom plates at concentrations ranging from 1.6 x 102 to 2 x 104 cells/well. Purified autologous T cells (1 x 105) from the BM donor in RPMI 1640 plus 10% heat-inactivated AB human serum (Normlcera-Plus; NABI, Miami, FL), 2 mM L-glutamine, 100 U/ml penicillin and 100 µg/ml streptomycin) were added to the DC-containing wells with 10 µg/ml preservative-free TT (Connaught Labs, Ontario, Canada). Cultures were incubated for 7 days at 37°C in a humidified 5% CO2-in-air atmosphere. T cell proliferation was assayed as described above. Data are presented as the mean cpm ± 1 SD of triplicate cultures.

Cytotoxic T cell assay

Purified allogeneic CD28+ T cells (4 x 105 cells/ml) were cultured for 7 days in six-well culture plates (5 ml/well) with irradiated allogeneic DC (0.5–2 x 105 cells/well) that were derived from day 7 PMA-treated CD34+ cell cultures. The autologous and allogeneic CD28+ T cell blasts used as target cells were cultured in RPMI 1640 supplemented with 10% heat-inactivated AB human serum, 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin and activated with immobilized anti-CD3 and anti-CD28 mAbs (conjugated to beads, 1 bead/cell). After 7 days of culture, T cell blasts (1 x 107/ml) were labeled with 200 µCi of 51Cr (Na51CrO4; New England Nuclear) for 1 h at 37°C and extensively washed before use. Effector T cells were assayed at the E:T ratios indicated in round-bottom microtiter plates (Costar, Cambridge, MA) with 1 x 104-labeled target cells/well. Spontaneous and maximal release samples were prepared by adding the target cells to wells containing RPMI 1640 alone or a final concentration of 2% Triton X-100. Plates were incubated for 4 h, and supernatants were collected with absorption cartridges (Skatron Instruments, Sterling, VA) for radioactive counting in a gamma counter (Pharmacia/LKB). The percentage of specific lysis was calculated using the following formula: ([experimental 51Cr release - spontaneous release]/[maximal release - spontaneous release]) x 100. The spontaneous release of target cells was <20% of maximal release.

Terminal deoxynucleotidyl transferase (TdT)-mediated deoxyuridine triphosphate nick end labeling (TUNEL) assay for apoptosis

DNA fragmentation in apoptotic cells was assayed as previously described (40). Briefly, 5 x 105 cells after 24, 48, 72, and 96 h of culture were fixed with 1% paraformaldehyde solution for 10 min on ice and then washed twice in PBS, permeabilized with 1 ml of 70% ethanol, and stored at -20°C for 2 h to 3 days. Following a single wash in PBS, cells were resuspended in 50 µl of a TdT reaction mixture (0.1 M cacodylic acid, 1 mM CoCl2, 1 mM DTT, and 50 µl of BSA) containing 0.5 nM biotin 16-deoxyuridine triphosphate (Boehringer Mannheim, Indianapolis, IN) and 10 U of TdT (Boehringer Mannheim) for 30 min at 37°C. After washing with PBS, 2.5 µg/ml FITC-avidin (Life Technologies, Gaithersburg, MD) was added to a staining solution (4x SCC, 0.1% Triton X-100, and 5% nonfat dry milk), and samples were incubated for 15 min at room temperature. Samples were analyzed by cytofluorometric analysis using an ELITE-ESP (Coulter) flow cytometer. Using the parameters of RALS vs log-FITC intensity, 10,000 gated cells were enumerated. Apoptotic cells were defined as FITC-positive cells with low FLS and RALS. Samples lacking TdT served as negative controls.

RT-PCR

Total mRNA from unstimulated (T = 0), cytokine-treated (GM-CSF plus IL-3, IL-6, and SCF), and cytokine plus PMA-treated CD34+ cells was isolated at the times indicated using RNAzol (Cinna Biotecs, Friendswood, TX) according to the manufacturer’s instructions. First strand cDNA was made from equal amounts of mRNA with a First Strand Synthesis Kit (Stratagene, La Jolla, CA) as per the manufacturer’s instructions. Five microliters of first-strand cDNA was used as a template in parallel PCR reactions, with one set using oligonucleotide primers specific for human RelB (sense GGGGAGAGCAGCACCGAGGCCAGCAAGACG, antisense AGCTCTGATGTGTTTGTGGATTTCTTGTCA) and the second set using primers for human ß-actin (sense TGACGGGGTCACCCACACTGTGCCCATCTA, antisense CTAGAAGCATTGCGGTGGACGATGGAGGG, Stratagene). Cycle parameters of 92°C x 1', 54°C x 1', and 72°C x 2' were used for 15, 25, and 35 cycles. Twenty microliters of each reaction was run on a 1% agarose gel, transferred to nylon, and hybridized against either 32P-labeled RelB (cDNA, a gift of Dr. U. Siebenlist, National Institute of Allergy and Infectious Diseases, National Institutes of Health) or actin probes. Hybridization was quantified by densitometry on a PhosphorImager 445 (Molecular Dynamics, Sunnyvale, CA). We found that 25 cycles were within the logarithmic amplification range for actin.

Western blot analysis

CD34+ cells (1 x 107) were cultured in PMA alone for 7 days. Cell lysates were then made from this culture (4 x 106 viable cells) and from 6 x 106 unstimulated (day 0) cells. Samples containing equal amounts of protein were separated by SDS-PAGE (4% stacking/7.5% resolving), electroblotted to nitrocellulose, probed with Abs against RelB (C-19, Santa Cruz Biotechnology, Santa Cruz, CA), and visualized by chemoluminescent detection (ECL, Amersham Life Sciences, Buckinghamshire, U.K.). We determined m.w. from a relative mobility log10 (m.w.) plot of standard m.w. markers run on the same gel.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Phorbol ester induces differentiation of CD34+ hemopoietic progenitors to DC

By day 3, 40 to 60% of input CD34+ progenitor cells cultured in 10 ng/ml (1.6 x 10-8 M) PMA became large, adherent single cells, whereas culturing in the inactive analogue 4{alpha}-phorbol had no effect. By day 7, 47% ± 8.7% of the input number of cells remained. These cells were motile and adherent, with most cells displaying round or stellate morphologies that projected multiple neurite and veiled processes in culture and hair-like cytoplasmic projections on cytopreparations (Fig. 1GoA). Immature or mature granulocytes/macrophages were not seen. This morphology was stable over more than 30 days of culture. The nonphorbol PKC agonist mezerein also induced DC morphology. Conversely, the addition of the PKC-specific inhibitor bisindolylmaleimide I completely blocked the effects of PMA. Morphologic changes induced by PMA were observed over a dose range of 0.1 to 100 ng/ml and were not affected by pretreatment with cytokines (GM-CSF plus IL-3, IL-6, and SCF, which induce CD34+ cell expansion with myeloid but not DC differentiation (38)), for 0, 24, or 48 h before PMA addition (data not shown).



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  		FIGURE 1. Phenotype of PMA-treated human CD34+ BM progenitor cells. A indicates the typical morphology of CD34+ cells cultured in PMA alone for 7 days (upper left panel, arrows indicate elongated neurite processes; lower left panel, cytopreparation of these cells (Wright stained, x200 magnification) or cultured in the nonphorbol PKC activator mezerein (upper right panel) or in PMA plus the PKC inhibitor bisindolylmaleimide I (lower right panel). B shows the surface Ag phenotype characterization of day 7 PMA-treated CD34+ cell cultures. Total cells in culture (nonadherent and adherent) were analyzed by flow cytometric analysis. Isotype-matched controls are indicated by a heavy solid line, while Ag-specific mAbs are depicted by a dotted line. Results are representative of three different experiments. C shows CD14 expression on CD34+ BM-derived cells cultured for 7 days in both 10 ng/ml PMA alone and GM-CSF plus IL-3, IL-6, and SCF with or without 10 ng/ml PMA. After 7 days of culture, adherent and nonadherent cells were harvested, pooled, stained, and analyzed cytofluorometrically for CD14 Ag expression. Results are representative of three different experiments.

 
Phenotypic analysis of day 7 PMA-induced cells (adherent and nonadherent) demonstrated that the entire population expressed MHC class I and II, CD13, CD83, CD80, and CD86 (Fig. 1GoB), consistent with the previously reported phenotype of DC (41, 42, 43, 44, 45, 46). The cells were negative for CD1a and the monocyte marker CD14. The addition of GM-CSF plus IL-3, IL-6, and SCF did not overcome the differentiating effects of PMA as measured by CD14 expression (Fig. 1GoC), consistent with the morphologic findings above.

PMA-generated DC from CD34+ progenitor cells are functional APCs

The functional hallmark of DC is their potent ability to activate T cells (1, 4, 5, 6, 13, 47, 48). We found that irradiated PMA-generated DC alone induced an alloproliferative response in purified resting human peripheral blood T cells (DC/T cell ratio of 1:100) (Fig. 2GoA). PMA-generated DC also stimulated T cell proliferation and IL-2 production in response to the superantigen (superAg) SEB and the mitogen Con A. Polyclonal activation by the latter two agents most likely accounts for the higher level of proliferation vs DC+T cells alone. T cell viability is demonstrated by their response to PMA plus anti-CD28 mAb alone and the lack of contaminating accessory cells as a result of the low proliferative response to Con A alone.



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  		FIGURE 2. Allogeneic naive T cell proliferation induced by PMA-generated CD34+->DC. A shows the induction of proliferation and IL-2 production in resting allogeneic human peripheral blood T cells by PMA-generated DC from CD34+ BM progenitor cells. 1 x 105 T cells/well were cultured with media alone (no stimulus), SEB (3 µg/ml), Con A (5 µg/ml), or PMA (10 ng/ml) plus anti-CD28 (9.3, 1.0 µg/ml) in the absence (media) or presence of gamma-irradiated (3000 rad 137Cs) day 7 PMA-induced DC (1 x 103). Results are representative of three different experiments and are expressed as the mean cpm ± 1 SD of quadruplicate cultures. After 24 h, culture medium was collected and assayed for IL-2 by ELISA; data are expressed as pg/ml per 1 x 105 T cells indicated above the appropriate culture condition. B indicates the proliferation of allogeneic T cells in response to titrated numbers of DC. Day 7 {gamma}-irradiated DC were generated from CD34+ cells as described above and added to 1 x 105 purified allogeneic T cells at the numbers indicated. Two different T cell donors were assayed. T cell proliferation is expressed as the mean cpm ± 1 SD of quadruplicate cultures. Proliferation of 1 x 105 T cells alone and 1 x 105 DC alone are also shown.

 
The allogeneic T cell proliferative response was able to be titrated by the number of added DC (Fig. 2GoB). T cell proliferation could be induced by as few as three DC (38-fold greater than the T cells used alone as a control (5129 ± 3869 vs 136 ± 78 cpm for donor A)) with the half-maximal dose of ~400 to 800 DC. As shown above, DC or T cells alone had minimal proliferation.

Induction of allogeneic T cell proliferation by PMA-generated DC does not rule-out an effect by PMA carryover from the original CD34+ cell culture nor does it demonstrate the ability to process and present whole Ag. To assess the presence of residual phorbol ester, PMA-generated DC were cultured with autologous T cells in the absence or presence of SEB (Fig. 3GoA). Whereas robust and titratable T cell proliferation was induced by PMA-generated DC plus SEB, minimal proliferation was seen with autologous T cells plus DC alone at an eightfold higher DC/T ratio than that used for the maximal T cell plus DC and SEB response. This lack of proliferation makes it unlikely that T cell proliferation is due to phorbol ester carryover.



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  		FIGURE 3. Autologous T cell proliferation induced by PMA-generated CD34+->DC pulsed with superAg or soluble whole Ag. A indicates SEB-induced proliferation of autologous T cells in response to titrated numbers of DC. Day 7 gamma-irradiated DC were generated from CD34+ cells as described above and added to 1 x 105 purified autologous T cells at the numbers indicated in the presence of 3 µg/ml SEB. Data are presented as the mean cpm ± 1 SD of triplicate cultures. B indicates the proliferation of autologous T cells following soluble Ag processing (TT) and presentation by PMA-derived DC. Irradiated DC from day 7 PMA-treated CD34+ cell cultures were plated in triplicate wells of 96-well flat-bottom plates at concentrations ranging from 1.6 x 102 to 2 x 104 cells/well. Purified autologous T cells (1 x 105) from the BM donor were added to the DC-containing wells with 10 µg/ml preservative-free TT. T cell proliferation is presented as the mean cpm ± 1 SD of triplicate cultures.

 
SEB-mediated T cell activation is not MHC restricted and does not require Ag processing. We found that PMA-generated DC could also process and present whole soluble Ag (TT) to autologous T cells (Fig. 3GoB). The proliferative response was generated in a total population of resting peripheral blood T cells (not tetanus-reactive T cell clones) that was not preactivated with TT. As described above, PMA-generated DC did not induce the proliferation of autologous T cells in the absence of Ag.

In addition to T cell proliferation, PMA-generated DC were capable of inducing Ag-specific cytotoxic T cell effector function (Fig. 4Go). Allogeneic T cells stimulated for 7 days by PMA-derived DC were capable of lysing target cells (T cell blasts) generated from the original CD34+/DC donor but had no activity against autologous (original T cell donor) or third party targets.



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  		FIGURE 4. Cytotoxic T cell activity induced by PMA-generated CD34+->DC. Two x 105 (black symbols), 1 x 105 (gray symbols), and 5 x 104 (white symbols) irradiated PMA-induced DC were plated with 2 x 106 purified allogeneic T cells for 7 days. The cytotoxic activity of these T cells was then assayed against allogeneic target cells from the original CD34+/DC cell donor (circles), autologous target cells derived from the original T cell donor (triangles), and unrelated donor target cells (squares). The percentage of specific lysis was calculated using the following formula: ([experimental 51Cr release - spontaneous release]/[maximal release - spontaneous release]) x 100. The spontaneous release of target cells was <20% of maximal release.

 
Phorbol ester-induced CD34+->DC differentiation does not involve proliferation

In contrast to cytokine-driven systems, PMA-induced CD34+->DC differentiation did not involve proliferation (Fig. 5GoA). Moreover, PMA inhibited the proliferation induced by GM-CSF plus IL-3, IL-6, and SCF even when cells were pretreated 24 or 48 h before the addition of PMA. PMA completely suppressed the colony formation of CD34+ cells seeded in semisolid media and GM-CSF plus IL-3, IL-6, SCF, and erythropoietin, which is consistent the proliferation findings and suggests that this suppressive effect is directly mediated at the single-cell level (data not shown). In addition, 40 to 60% of CD34+ progenitors (depending upon the donor marrow) that were plated as single cells in 96-well plates developed typical DC morphology following PMA treatment (vs 0% cultured in media or GM-CSF plus IL-3, IL-6, and SCF), demonstrating again that there is a direct effect at the single-cell level and a lack of a small contaminating population of CD34- DC precursors.



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  		FIGURE 5. CD34+ cell proliferation and viability in response to PMA. A shows CD34+ cell proliferation. Purified CD34+ cells were cultured with the specified cytokines in the absence or presence of 10 ng/ml PMA. Where indicated, hours indicate the length of pretreatment with cytokines before the addition of PMA. Results are expressed as the mean cpm ± 1 SD of three independent experiments using three different donors of CD34+ cells. B shows CD34+ cell viability. CD34+ cells (5 x 104) were incubated for 24, 48, 72, and 96 h with PMA in the presence or absence GM-CSF plus IL-3, IL-6, adn SCF and then assayed for cell viability by trypan blue dye exclusion. Where indicated, hours indicate the length of pretreatment with cytokines before the addition of PMA. Results are expressed as the mean viability ± 1 SD of three independent experiments using three different donors of CD34+ cells.

 
The observation that only ~50% of the input number of cells are recovered after 7 days suggested that PMA had a negative effect on CD34+ survival. Cultures treated with PMA were only 48% viable vs 85% in media alone at 48 h (Fig. 5GoB), demonstrating that cell death was not simply due to lack of growth factors. CD34+ cells cultured in GM-CSF plus IL-3, IL-6, and SCF remained >84% viable throughout the culture period. As with proliferation, pretreatment with cytokines did not affect subsequent PMA-induced cell death. Since PMA treatment is not directly cytotoxic for B and T lymphocytes (49), the abrupt decline in cell viability compared with media alone suggested that these cells were undergoing programmed cell death. The TUNEL flow cytometric assay of programmed cell death-associated DNA fragmentation demonstrated that 14 to 17% of GM-CSF plus IL-3, IL-6, and SCF-treated cells were undergoing apoptosis throughout the entire culture period (Table IGo). In contrast, 39 to 42% of the gated viable cells were apoptotic after 48 h in cultures with PMA alone or PMA plus cytokines.


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  		Table I. Induction of programmed cell death in CD34+ BM progenitors by PMA in the absence or presence of exogenous hemopoietic cytokines1

 
Phorbol ester-induced CD34+->DC differentiation involves PKC activation

Although phorbol esters are conventionally regarded as PKC agonists, it is possible that PMA-induced DC differentiation does not involve PKC activation. Besides PKC, phorbol esters bind to two other proteins: the Caenorhabditis elegans protein unc-11 and the p21rac-guanosine triphosphatase activating protein n-chimaerin (50, 51). In addition to bisindolylmaleimide I, we examined whether the PKC inhibitor staurosporine could block the effects of PMA on CD34+ cells. One characteristic effect of PMA is the suppression of cytokine-induced CD34+ cell proliferation. As can be seen in Table IIGo, staurosporine completely inhibited this suppressive effect of PMA and restored CD34+ cell proliferation to cytokine plus staurosporine levels. In addition, staurosporine blocked PMA-induced cell morphology changes (data not shown).


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  		Table II. Effect of the PKC inhibitor staurosporine1

 
PMA-mediated signaling induces expression of the RelB transcription factor

The differentiation of CD34+ cell to DC most likely occurs through cascades of new gene expression. Previous studies suggesting that this expression is mediated in part by the RelB transcription factor acting on rel/NF-{kappa}B-responsive genes led us to examine whether PMA-mediated signaling induced RelB expression in CD34+ cells. By semiquantitative RT-PCR, relB expression is very low in unstimulated (T = 0) CD34+ cells (Fig. 6GoA, evident upon longer exposure). Interestingly, these cells also express an alternatively spliced form of relB lacking exon 5. Treatment with either cytokines alone (GM-CSF plus IL-3, IL-6, and SCF) or PMA plus cytokines initially up-regulates relB expression to equivalent levels at 24 h. However, by 48 h, relB expression declines in cultures with cytokine alone, whereas expression remained high in cultures with PMA plus cytokines. Consistent with relB gene expression is the up-regulation of RelB protein by PMA stimulation (Fig. 6GoB). There are very low levels of RelB detectable in unstimulated (day 0) CD34+ cells (seen on longer exposures). Stimulation with PMA alone substantially up-regulates the expression of both the p68 and p45 forms of RelB (36) by 7 days. In addition, there is a faint doublet of ~50 kDa that has not been previously reported.



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  		FIGURE 6. Induction of RelB expression by PMA-mediated PKC activation. A shows the RT-PCR analysis of relB gene expression in CD34+ cells. Total mRNA was extracted from unstimulated (T = 0), cytokine-stimulated (GM-CSF plus IL-3, IL-6, and SCF), or cytokine plus PMA-stimulated CD34+ cells at the times indicated. Equal amounts of mRNA were used to generate first strand cDNA, and these were used as templates for duplicate PCR reactions with relB or ß-actin (as an internal control) primers. As shown here, 25 cycles was found to be in the logarithmic amplification range. Twenty µl of each reaction were separated on 1% agarose gels, transferred to nylon, hybridized with 32P-labeled relB or ß-actin probes, and visualized on a PhosphorImager 445. B indicates Western blot analysis of RelB protein expression in PMA-stimulated CD34+ cells. Cell lysates were made from unstimulated (T = 0) and day 7-PMA-treated CD34+ cells. Samples containing equal amounts of protein were separated by SDS-PAGE (4% stacking/7.5% resolving), electroblotted to nitrocellulose, and probed with Abs against RelB (C-19, Santa Cruz Biotechnology). Molecular weights were determined from a relative mobility log10 (m.w.) plot of standard m.w. markers run on the same gel.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We have found that phorbol ester-mediated signaling in CD34+ BM hemopoietic progenitors induces DC differentiation in ~50% of the population and cell death in the other 50%. PMA-induced cells have the surface Ag phenotype characteristic of DC, including high expression of MHC class I and II, CD13, the costimulatory ligands CD80 (B7-1) and CD86 (B7-2), and the DC lineage-specific marker CD83. These cells were negative for CD1a and the monocyte marker CD14, a phenotype also reported for the most differentiated subsets of dermal DC (52), cytokine-generated CD14+ monocytes->DC (13), and peripheral blood DC (53). The ability to differentiate into DC in response to phorbol ester was limited to CD34+ cells (freshly isolated and expanded ex vivo), as PMA caused macrophage differentiation in CD34-CD15+ cells (from ex vivo cultures) and cell death in freshly isolated CD14+ monocytes (data not shown). The fact that >99% of the input CD34+ cells were CD38+ allows us to conclude, given the absence of cell proliferation, that this population is inducible to DC differentiation by phorbol ester. CD34+CD38- cells reisolated following ex vivo expansion (38) appear equally responsive (our unpublished observations).

In contrast to cytokines or CD40 ligation, PMA induces DC differentiation only, without causing cell proliferation and the generation of cellular intermediates. In this regard, PMA-induced CD34+->DC differentiation more closely resembles cytokine-driven monocyte->DC differentiation, in which there is also no proliferation, and 20 to 90% of input cells are lost during culture (11, 13, 14).

Functionally, PMA-generated DC may activate resting T cells by presenting alloantigen and superAg, activate autologous T cells by processing and presenting whole soluble Ag, and generate cytotoxic T cell effector function. T cell proliferation could be induced in an allogeneic MLR down to a DC/T cell ratio of 1:33,333, with a half maximal DC/T ratio of 1:125 to 250, comparable with that reported for cytokine-generated DC (11, 54). The absence of autologous T cell proliferation when cultured with DC alone (no SEB or TT) demonstrates the lack of PMA carryover from the original CD34+ cultures.

The inability of cytokine pretreatment to overcome the differentiative, antiproliferative, and apoptotic effects of PMA indicates that in CD34+ cells, phorbol esters act downstream of the cell survival and proliferative components of the cytokine pathways. The ability of the PKC inhibitors bisindolylmaleimide I and staurosporine to block the effects of PMA suggests that phorbol esters are acting at the level of PKC. Although staurosporine can also block protein tyrosine kinase activity (at higher doses than those used here; 0.2 nM vs an IC50 of 6 nM for v-src (55)), the inability of staurosporine to significantly suppress CD34+ cell proliferation in response to GM-CSF plus IL-3, IL-6, and SCF (which also involves tyrosine kinase activation) suggests a minor role for protein tyrosine kinase. We have also found that the nonphorbol ester PKC agonist mezerein induces, while PKC inhibitors (staurosporine, bisindolylmaleimide I, calphostin C, and ET-18-OCH3) block, DC differentiation in a human CD34+ cell line model (manuscript in preparation). Although not described for primary CD34+ cells, PKC activation in other hemopoietic cells induces both terminal differentiation and apoptosis. PMA-induced PKC activation drives primary myeloblasts to differentiate into macrophages in the absence of DNA synthesis (24), and PKC activation by macrophage-CSF causes macrophage differentiation of granulocyte macrophage CFU (32). In leukemic cell lines, phorbol esters induce terminal differentiation of multipotential progenitors (myb-ets-transformed progenitors to eosinophils or myeloblasts, depending upon the level of PKC activity (31, 56)) and more committed progenitors (KG1, HL-60, ML-30, and HEL to macrophages and megakaryocytes (25, 26, 27, 28, 29, 30)).

What physiologic signal is being mimicked by PMA? If PMA is activating PKC, it is well established that PKC is also activated by a number of receptor-mediated signaling pathways. Of particular relevance to CD34+->DC differentiation is the fact that IL-4, TNF-{alpha}, and CD40 all induce PKC activation as part of their signal transduction pathway (19, 20, 21, 22, 23). Consistent with this observation, we have found that staurosporine preferentially inhibits DC differentiation of CD34+ progenitors in response to GM-CSF plus TNF-{alpha} (manuscript in preparation). In addition to a direct involvement in the signaling pathway, PKC may activate downstream components also used by non-PKC signaling pathways, such as the ras/raf-1/mitogen-activated protein kinase pathway activated by GM-CSF and IL-3 (reviewed in 34 or NF-{kappa}B activation by macrophage CSF and GM-CSF (reviewed in 37 . It is likely that CD34+->DC differentiation involves specific PKC isoforms that mediate specific biologic events, as has been shown for PKC-ß in HL-60 differentiation (57) and PKC-{theta} in T cell/APC interaction (58).

Ultimately, for differentiation to occur, signal transduction must cross into the nucleus and initiate specific, new gene transcription. Because PMA appears to act on a differentiation pathway downstream of cell proliferation and "closer" to the nucleus, we hypothesized that PMA induced/activated a DC-"specific" transcription factor that subsequently initiated the new gene expression involved in differentiation. For several reasons, the most likely candidate is RelB, a member of the rel/NF-{kappa}B family of transcription factors (36). Using conditional v-rel-transformed chicken BM cell lines, Boehmelt et al. have shown that rel/NF-{kappa}B-responsive gene activation plays a critical role in DC differentiation (16). RelB is expressed at high levels by murine DC (59) and by DC derived in vitro from human monocytes (but not by monocytes/macrophages themselves) (12), and RelB knockout mice have significant reductions in mature DC number and APC function (17, 18). PKC activation induces relB gene expression in T cell lines (60) and the ability to bind to DNA (NF-{kappa}B sites) in fibroblasts (35). Consistent with these reports, we found that unstimulated CD34+ BM cells have a low level of RelB genes and protein expression that is substantially up-regulated by PMA. PMA differs from the cytokine combination GM-CSF plus IL-3, IL-6, and SCF, which does not generate DC, in its ability to induce sustained relB gene expression. Studies of in vitro generation of DC from monocytes suggests that such a sustained expression is a hallmark of DC differentiation (12). More detailed kinetic studies in a cell line model of DC differentiation that we have developed reveals very rapid up-regulation of RelB protein expression (within 30 min of PMA stimulation) with sustained gene and protein expression out to at least 7 days (manuscript in preparation). RelB binding to DNA/NF-{kappa}B sites as measured by electrophoretic mobility shift assays display similar kinetics. Studies are underway to determine whether RelB is immediately downstream of PKC (i.e., directly activated by PKC phosphorylation of complexed I-{kappa}B, freeing RelB to translocate from the cytoplasm into the nucleus), or whether PKC activates other signaling pathways which then induce RelB expression. Identification of RelB-responsive genes that may be involved in DC differentiation is also underway. Although our findings suggest an important role for PKC and RelB induction in DC differentiation, we feel certain that other signaling pathways and transcription factors are involved. We have found that TNF-{alpha} synergizes with PMA, suggesting that non-PKC-mediated signaling pathways (possibly through sphingomyelin/ceramide second messengers) are involved. Certainly, the requirement for multiple cytokines to generate DC in vitro suggests that a number of signals are required.

In addition to delineating signal transduction components involved in DC differentiation, the methodology described here may represent a simplified procedure for generating pure populations of DC from freshly isolated or ex vivo-expanded CD34+ progenitor cells. This may facilitate the therapeutic use of primed or genetically modified DC in vaccination protocols against infectious and tumor-associated Ags.


   Acknowledgments
 
We thank LIFENET (Virginia Beach, VA) for the procurement of human vertebral BM samples.


   Footnotes
 
1 This work was supported by the Naval Medical Research and Development Command, Research Task No. 63706.M0095.003.1458. The views presented in this paper are those of the authors; no endorsement by the Department of Navy has been given or should be inferred. Back

2 Address correspondence and reprint requests to Dr. Kelvin P. Lee, Immune Cell Biology Program, Bldg. 17, Room 214, Naval Medical Research Institute, 8901 Wisconsin Avenue Bethesda, MD 20889-5067. E-mail address: Back

3 Abbreviations used in this paper: DC, dendritic cells; GM-CSF, granulocyte macrophage CSF; SCF, stem cell growth factor; PKC, protein kinase C; superAg, superantigen; BM, bone marrow; TT, tetanus toxoid; TdT, terminal deoxynucleotidyl transferase; RALS, right-angle light scatter; TUNEL, terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick end labeling; FLS, forward light scatter; SEB, staphylococcal enterotoxin B. Back

Received for publication August 29, 1997. Accepted for publication December 11, 1997.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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