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*
Departments of Molecular and Experimental Medicine and Immunology, Scripps Research Institute, La Jolla, CA 92037; and
The Burnham Institute, La Jolla, CA 92037
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Because DCs are unique in their exceptional ability to stimulate T
cells, particularly naive T cells, it is not unreasonable to propose
that this function might be orchestrated by a unique set of
transcription factors. A few candidate DC-critical factors have been
identified. Studies have shown that RelB, a member of the NF-
B
family, is expressed specifically in certain populations of DCs
(16, 17, 18). RelB-deficient mice have very reduced numbers of
thymic medullary and splenic DCs, as well as a lack of thymic medullary
epithelial cells (17, 19). Similarly, Ikaros transcription
factor-null and dominant-negative mutant mice display defects in both
myeloid- and lymphoid-derived DC populations (20). The
ets family transcription factor PU.1 is expressed in
multiple hematopoietic lineages, as well as in human
CD34+ progenitor cells, and can regulate the
expression of many myeloid and lymphoid genes (21, 22). Previously we have shown that mice deficient in PU.1
display specific defects in the development and maturation of multiple
hematopoietic lineages (23). In this study we demonstrate
that PU.1 is normally expressed in mouse hematopoietic
progenitor-derived CD11c+ MHC class
II+ DCs, as well as in human PBMC- and
CD14+ cell (monocyte)-derived MHC class
II+CD1a+CD14-
DCs. In contrast to cells from wild-type mice, when hematopoietic
progenitor cells from PU.1 null mice were cultured in a
variety of DC-supportive/inductive growth factors, no cells with
characteristic DC appearance or cell marker phenotype were produced.
Furthermore, although development of a thymic cortex and medulla
occurred and T cells were generated in all 8- to 12-day-old PU.1 null
mice, DEC-205+ and MIDC-8+
thymic DCs could not be detected. Thus, PU.1 is required for the
differentiation of the distinct myeloid-derived and thymic,
lymphoid-derived populations of DCs in mice.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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C57BL/6 x 129 PU.1 gene-disrupted mice were produced and identified as previously reported (23).
Culture medium and cytokines
Culture medium included RPMI 1640 (Life Technologies, Grand
Island, NY) supplemented with 10% FCS (HyClone, Logan, UT), with
5 x 10-5 M 2-ME (Sigma, St. Louis, MO) 1%
L-glutamine (Life Technologies), 1%
penicillin/streptomycin (Life Technologies), termed complete RPMI
(cRPMI), and IMDM (Life Technologies) supplemented with 10% FCS
(HyClone), with 5 x 10-5 M 2-ME (Sigma),
1% L-glutamine (Life Technologies), 1%
penicillin/streptomycin (Life Technologies), termed complete IMDM
(cIMDM). The following human and mouse cytokines were used: recombinant
mouse (rm)-GM-CSF (R&D Systems, Minneapolis, MN), rm-TNF-
(R&D
Systems), recombinant rat (rr)-stem cell factor (SCF, Amgen, Thousand
Oaks, CA), rm-IL-3 (R&D Systems), recombinant human (rh)-GM-CSF
(Amgen), rh-IL-4 (R&D Systems), rh-IL-6 (R&D Systems), rh-M-CSF (a kind
gift from Dr. David Hume, University of Brisbane, Queensland,
Australia), and rh-TNF- (R&D Systems).
Multiparameter flow cytometry analysis for cell surface Ags
Three- and four-color flow cytometric analysis was performed as described for mouse and human cells (23) using a Becton Dickinson FACScalibur and CellQuest software (Becton Dickinson, San Jose CA). All Abs used for flow cytometric analysis were directly conjugated with fluorochromes of choice. Abs used for mouse cell analyses included I-Ab, CD11b, CD11c, Gr-1, CD3, B220 (all from PharMingen, San Diego, CA), and F4/80 (Caltag, Burlingame, CA). NLDC-145 (DEC-205) and MIDC-8 were both used as culture supernatants (hybridomas were a kind gift of Dr. G. Kraal, Free University of Amsterdam, Amsterdam, The Netherlands). Anti-human Abs included CD45, HLA-DR, CD1a, CD3, CD4, CD8, CD14, CD33 (all reagents from Becton Dickinson), and CD83 (Immunotech, Coulter, Miami, FL).
Culture of mouse progenitor-derived DCs, macrophages, and isolation of CD11c+ cells
Neonatal and 4–8 wk of age liver and/or bone marrow cells were
obtained as described (24). For some experiments, low
density mononuclear cells were isolated by density gradient
centrifugation as previously described (24). Culture
methods used to generate myeloid DCs from the bone marrow of mice were
those established by Lutz et al. (25), with slight
modifications. Cultures were started at 20 ng/ml of rm-GM-CSF and at
day 3, 5 ml of culture medium was removed and 5 ml of fresh
rm-GM-CSF-containing medium was added (as per Lutz et al.
(25)) and at days 4 and 7–8, 5 ml of the spent medium was
removed and 5 ml of fresh rm-GM-CSF (10 ng/ml) containing medium was
added. On day 10 nonadherent cells were removed by vigorous pipetting
and resuspended with 10 ng/ml rm-GM-CSF and 1400 ng/ml rm-TNF- in
tissue culture-treated 100-mm2 dishes (Becton
Dickinson). Nonadherent cells were harvested 1 day later for
CD11c+ cell isolation. In addition to these
factors, IL-3 (100 U/ml) was added to PU.1 null cell cultures. PU.1
null hematopoietic cells have previously been shown to be dependent
upon this factor (24). Because wild-type cultured mouse
myeloid DCs express CD11c (2, 3) at high levels before and
after culture, whereas contaminating granulocytes and monocytes do not,
cultured CD11c+ cells were isolated for
determination of PU.1 protein expression by using an anti-CD11c mAb
and streptavidin magnetic microbeads in conjunction with a magnetic
isolation apparatus as specified by the manufacturer (VarioMacs,
Miltenyi Biotec, Auburn CA). Mouse macrophages were obtained by plating
30 x 106 bone marrow cells on
100-mm2 tissue culture plates (Becton Dickinson)
in 10 ml of cRPMI, incubated for 5 h at 37°C in a humidified
incubator, then nonadherent cells were removed, and fresh medium
containing 5000 U/ml of rh-M-CSF was added and nonadherent cells were
removed the next day. Cultures were fed every 3–4 days by one-half
medium changes; cells were used on day 12.
Generation of human PBMC-derived macrophages, DCs, and CD14+ cell-derived DCs
Human DCs were generated from whole PBMCs or
CD14+ cells isolated from PBMCs. All blood
donations were obtained from qualified donors under protocols approved
by The Scripps Research Institute’s Human Subject Committee.
Procedures used to generate DCs were those established by Thurner et
al. (26). The procedure was slightly modified in that in
some cases cultures were conducted to 7 or 12 days. In brief, PBMCs
were isolated by density gradient centrifugation using Ficoll-Paque
(Pharmacia, Uppsala, Sweden) from heparinized whole blood obtained from
healthy donors, washed twice in 0.9% sodium chloride solution after
separation, resuspended in cRPMI, and plated at 50 x
106 cells/ml to generate DCs from whole PBMCs as
established (26). DCs were harvested on days 8–12 after
culture in 800 U/ml of rh-GM-CSF (Amgen) and 1000 U/ml of rh-IL-4 (R&D
Systems) to promote DC differentiation from PBMCs or
CD14+ cells. To generate DCs from
CD14+ cells, these cells were first isolated from
PBMCs using anti-CD14 Ab and a VarioMACS apparatus as specified by
the manufacturer. To increase purity, positively selected
CD14+ cells were run through two
VS+ separation columns (Miltenyi Biotec). The
purity and efficiency of the CD14+ cell
separation was evaluated by staining the positively selected and
negatively depleted cell fractions with CD45, CD19, CD14, CD3, CD4,
CD8, CD83, and CD33 by four-color flow cytometry.
CD14+ purity of cells isolated after positive
selection was found to be 
98%. These cells were cultured identically
to whole PBMCs for DC differentiation.
To produce human macrophages, 50 x 106 PBMCs were plated on tissue culture plates in 10 ml of cRPMI, incubated for 5 h at 37°C in a humidified incubator, nonadherent cells were removed, fresh medium containing 5000 U/ml of rh-M-CSF was added, and nonadherent cells were removed the next day. Cultures were fed every 3–4 days by one-half medium changes, and cells were used on days 7–12.
Western blotting
Cell lysates were prepared and SDS-PAGE was performed as described (27). Polyclonal anti-PU.1 and anti-RelB Abs were obtained from Santa Cruz Biotechnology (Santa Cruz, CA), and anti-actin Ab from Sigma was used at 1:500, 1:500, and 1:200 dilution, respectively. Secondary anti-rabbit Ab was obtained from Santa Cruz Biotechnology. CruzMarker protein standards (Santa Cruz Biotechnology) were included on gels to determine the size of immunoreactive proteins.
TUNEL assay and immunohistology of thymus
Fresh thymi of neonatal and older normal and PU.1 null mice were immersed in OCT compound (Miles, Elkhart, IN), snap-frozen in liquid nitrogen, and stored at -70°C. OCT-preserved tissue was cut on a cyrostat into 5–6 µm sections and processed for both TUNEL analysis and immunohistology as described (28).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Because PU.1 has been shown to play a critical role in regulating
both the development of multiple lineages and the function of mature
hematopoietic cells (23, 27), we examined normal mouse
bone marrow-derived and human PBMC-derived DCs for PU.1 expression. Low
density cells were isolated from wild-type mouse bone marrow and
cultured in GM-CSF plus TNF-, or M-CSF, cytokines that promote
either DC or macrophage generation, respectively (see Materials
and Methods). To ensure a relatively pure population of mouse DCs
for PU.1 analysis, CD11c+ cells were isolated
from DC cultures after 11–12 days using immunomagnetic beads. This
strategy was utilized because CD11c is present on essentially all mouse
myeloid DC populations (3, 4). These isolated cells were
recultured for 24 h under DC-promoting conditions to allow
recycling of Ab-bound CD11c. As can be seen from flow cytometry
analysis, cells obtained from cultures promoting macrophage
differentiation were class IIdull
(I-Ab), and CD11c-, a cell
surface phenotype characteristic of macrophages, whereas cells obtained
from cultures promoting DC differentiation were
CD11c+, and class IIhigh, a
cell surface phenotype characteristic of DCs (Fig. 1
A). Cells from both
macrophage and DC cultures were F4/80+, and
Gr-1-, CD3-, and
B220- (data not shown). Both populations were
also examined for the presence of RelB protein (Fig. 1C), a
transcription factor known to be expressed in DCs but not macrophages
(16, 17). Expression of RelB was consistent with the flow
cytometry analysis demonstrating that DCs were generated under our
culture conditions.
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B shows an
example of flow cytometry analysis of cells from macrophage and DC
cultures. Cells cultured under conditions that promote macrophage
differentiation were HLA-DR+ and
CD14+ (Fig. 1B),
CD1adull, and CD86- (data
not shown), consistent with a macrophage phenotype, whereas cells from
DC cultures were HLA-DRbright,
CD14- (Fig. 1B),
CD1abright, and CD86+ (data
not shown), consistent with a DC phenotype. Both human PBMC- and
CD14-derived DCs demonstrated similar cell surface phenotypes; however,
only analysis of CD14-derived DCs is shown (Fig. 1B). Cells
from both macrophage and DC cultures were negative for the T cell
marker CD3 and the B cell marker CD19 (data not shown).
The human and mouse myeloid DCs that were described above were examined
for PU.1 expression. As shown in Fig. 1D, Western blot
analysis clearly demonstrated the presence of PU.1 protein in human and
mouse DCs. Low-level B cell contamination, a possible contributor of
PU.1 protein, was ruled out by RT-PCR analysis for CD19 (data not
shown). Thus, we have demonstrated that myeloid DCs generated from
either mouse bone marrow progenitors or human peripheral blood cells
express the transcription factor PU.1.
PU.1-deficient mouse hematopoietic progenitor cells fail to generate DCs in culture
Hematopoietic progenitor cells derived from normal mice have been
shown to be capable of differentiating into mature, functional DCs
under appropriate culture conditions (Refs. 25, 30, 31 , and above). To determine whether PU.1-deficient
hematopoietic progenitors were able to generate such cells in vitro, we
isolated and pooled low density mononuclear cells from the liver and
bone marrow of PU.1 null neonates. This step was required to obtain
sufficient hematopoietic progenitors because the bone marrow of PU.1
null mice is vastly reduced in all hematopoietic cells, due at least in
part to osteopetrosis (24, 32). These progenitors were
cultured using the method of Lutz et al. (25) with the
addition of IL-3 to the culture medium in some cases, because in its
absence all PU.1-deficient cells died (Table I and Ref. 24). Our previous
work has shown that PU.1 null cells require IL-3 for survival and
growth, and fail to express detectable cell surface receptors for and
proliferate to GM-CSF (24). Following the culture period,
the cells were analyzed by flow cytometry for their expression of DC
and myeloid markers, the results of which are summarized in Table I.
Only PU.1 null cells maintained in IL-3 and GM-CSF survived, and cells
compatible with a DC phenotype were not detectable in any cultures
established from PU.1 null mice. Lastly, PU.1 null cells cultured under
these conditions expressed Gr-1 (Table I) and morphologically were
similar to immature neutrophils, as reported previously (24, 27).
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Our in vitro studies demonstrated the inability of PU.1 null
hematopoietic progenitors to produce myeloid DCs under cytokine
conditions appropriate for wild-type cells. Recently, it has been shown
that populations of lymphoid lineage-derived DCs, including those
present in the thymus, can develop in vivo in GM-CSF-cytokine and
-receptor null mice (33, 34). Given the absence of GM-CSF
receptor expression in PU.1 null mice, we next addressed whether the
loss of PU.1 expression affected the development of thymic DCs.
Rudimentary thymus from fetal (35) and neonatal
(23) PU.1 null mice is devoid of T cells. However, if mice
are kept alive using intensive antibiotic therapy, thymic development
is observed to begin between 5 and 8 days after birth
(23). CD4 and CD8 single- and double-positive cells are
found in approximately normal proportions, although their number is 5-
to 10-fold reduced compared with normal littermates (23).
Histologic examination of the thymus of a 10-day-old PU.1 null mouse
clearly demonstrated cortical and medullary areas (Fig. 2, A, D,
F, and H; PU.1 null) and the presence of double
positive and single positive CD4 and CD8 cells by flow cytometry (data
not shown). Previously, we demonstrated that PU.1 was required for
development of monocytes/macrophages in the blood, spleen, and liver
(23). Consistent with these results,
F4/80+ cells were not detected in the thymus of a
10-day-old PU.1 null mouse (Fig. 2, compare A and
B). Immature thymocytes have a rapid turnover, due to the
majority of thymocytes not undergoing positive selection and
subsequently undergoing apoptosis (28). In the cortex of a
normal thymus these apoptotic cells are rapidly engulfed by
F4/80+ macrophages (28). The
presence of a few apoptotic cells, identified by the TUNEL method,
could be seen in the thymic cortex of a 10-day-old wild-type mouse
(Fig. 2C). In contrast, the thymic cortex and medulla of a
PU.1 null littermate contained tremendously elevated numbers of
apoptotic (dark red-purple staining) cells (Fig. 2D),
presumably the result of the absence of clearance by macrophages. Also,
visible in the cortical and medullary regions was intensely blue-purple
counterstained (hematoxylin) cellular debris. Therefore, it appeared
that phagocytic cells normally responsible for clearance of dead and
dying cells in the thymus were highly reduced, absent, and/or not
functional in PU.1 null mice.
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E),
we were unable to detect medullary staining for DEC-205 in thymus
sections from a PU.1 null mouse (Fig. 2F). As reported
(38, 39, 40), DEC-205 was expressed on thymic cortical
epithelial cells in both wild-type and PU.1 null mice (Fig. 2, E and F). MIDC-8, which recognizes intracellular
granules in thymic medullary DCs (41), was also used for
immunohistology. As was seen for DEC-205, MIDC-8+
cells were readily apparent in normal thymus but not the PU.1 null
thymus (Fig. 2, G and H). Thus, our combined data
from PU.1 null mice demonstrate ongoing T cell differentiation and the
presence of a thymic cortex and medulla. However, PU.1 null thymi lack
F4/80+ macrophages, phagocytic clearance of dead
and dying cells, and detectable DEC-205+,
MIDC-8+ DCs. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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DCs have been shown to derive from a bone marrow progenitor by both allogeneic bone marrow transplantation studies and in vitro culture techniques. These types of experiments have identified a MHC class II-negative mouse progenitor shared by DCs, neutrophils, and monocytes/macrophages (8), although more recently a distinct DC-CFU cell has been demonstrated in human bone marrow (44). Despite some lack of certainty in their lineal relationship, it is clear that DCs share many similarities with myeloid cells, although key differences also exist. The demonstration that peripheral blood monocytes and postmitotic neutrophil precursors can both be induced to differentiate to a DC phenotype is further evidence for this close relationship (12, 13, 14). Because PU.1 is expressed in bone marrow-derived myeloid lineages, it was not unexpected to find PU.1 expression in DCs derived from mouse bone marrow progenitor cells as well as in human peripheral blood cell/monocyte-derived DCs. These DC populations are considered to be of myeloid origin, unlike the thymic DCs and some splenic DC populations, which are believed to originate from a lymphoid-committed precursor cell (10, 11, 34, 45). To our knowledge, this study represents the first direct demonstration of expression of PU.1 in human and mouse myeloid DCs.
Populations of DCs are heterogeneous in their phenotype, but are broadly grouped into those derived from a myeloid (bone-marrow) progenitor and those derived from a lymphoid progenitor. The latter have been identified in the mouse thymus by repopulation studies using purified thymic progenitor cells, and indicate the existence of a precursor cell capable of developing into T cells or DCs (10, 11). Although we have not demonstrated directly that PU.1 is expressed in thymic DCs, the results of our studies would indicate that it plays a vital role in the development of this population. We found no DEC-205+, MIDC-8+ cells in the thymus of 10- to 12-day-old PU.1 null mice, although CD4+, CD8+ single- and double-positive T cells were present. Similarly, T cells but not DEC-205+ cells were found when spleens from 12-day-old PU.1 null mice were analyzed by flow cytometry and immunohistology (B. E. Torbett and C. D. Surh, unpublished results). Although PU.1 expression is high in B cells, it has not been found at any stage of definitive T cell development yet examined. If PU.1 is expressed in thymic DCs, perhaps its down-regulation represents a critical decision point in the determination of thymic T cell vs thymic DC differentiation. This question must remain open at present. Because the markers we examined represent markers for mature DCs, we cannot exclude the possibility that committed DC precursors have been generated in PU.1 null mice but are unable to fully differentiate in the absence of PU.1 due to other PU.1-related defects.
We observed failure of both lymphoid-derived and myeloid-derived DC
development in the absence of PU.1. This suggests either that PU.1 may
be required for specification of the DC developmental program in a very
early shared progenitor (such as the human CD34+
cell), or that PU.1 is required for expression of multiple
phenotype-defining genes that are similarly expressed in both the
lymphoid and myeloid-derived DCs. One possibility is that the absence
of the PU.1 regulated GM-CSF receptor expression (24, 46)
in PU.1 null myeloid cells contributes to the loss of myeloid DC
development in vitro (34). However, both myeloid and
lymphoid DCs were found in GM-CSF and GM-CSF receptor null mice
(34). A similar loss of both myeloid and lymphoid DCs has
been observed in mice that express a nonfunctional or a
dominant-negative form of the Ikaros zinc-finger transcription factor
(20). Another transcription factor, the NF-B family
member RelB, was found to be expressed in interdigitating DCs
(16) in which it is expressed at high levels
(18). RelB gene-disrupted mice failed to develop myeloid
DCs, but also had very low numbers of thymic, lymphoid DCs as a result
of abnormal thymic architecture (17, 18). Although we have
directly demonstrated PU.1 expression in myeloid DCs, its expression in
lymphoid DCs is unresolved and we can only speculate as to whether the
absence of PU.1 has a direct or an indirect effect on thymic DC
development. Unlike RelB-deficient mice (17), however, the
thymic architecture of PU.1 null mice appears intact and medullary
epithelial cells were present based on UEA-1 immunostaining (C. D.
Surh, unpublished results).
In summary, we have shown that PU.1 is expressed in and is crucial for development of myeloid DCs. Lymphoid DC development in the mouse thymus is also disrupted in the absence of PU.1, although the extent and nature of the disruption will require further investigation. Nevertheless, it is clear that PU.1 is required for normal development of both major mouse DC populations. Our data from the PU.1 null mouse model suggest that the formation of a thymic cortex and medulla, and the homing and expansion of T cell progenitors, can occur in the absence of accessory cells such as macrophages and DCs.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Bruce E. Torbett, Departments of Molecular and Experimental Medicine and Immunology, MEM55, Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037. E-mail address:
3 Abbreviations used in this paper: DC, dendritic cell; rm, recombinant mouse; rh, recombinant human; cRPMI, complete RPMI.
Received for publication August 29, 1999. Accepted for publication November 29, 1999.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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B/Rel family. Cell 80:331.[Medline]
. J. Exp. Med. 182:1111. regulate expression of the granulocyte-macrophage colony stimulating factor receptor gene. Mol. Cell. Biol. 15:5830.[Abstract]
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J. Back, A. Dierich, C. Bronn, P. Kastner, and S. Chan PU.1 determines the self-renewal capacity of erythroid progenitor cells Blood, May 15, 2004; 103(10): 3615 - 3623. [Abstract] [Full Text] [PDF]
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E. Ichikawa, S. Hida, Y. Omatsu, S. Shimoyama, K. Takahara, S. Miyagawa, K. Inaba, and S. Taki Defective development of splenic and epidermal CD4+ dendritic cells in mice deficient for IFN regulatory factor-2 PNAS, March 16, 2004; 101(11): 3909 - 3914. [Abstract] [Full Text] [PDF]
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Y.-H. Chen, M. D. Layne, S. W. Chung, K. Ejima, R. M. Baron, S.-F. Yet, and M. A. Perrella Elk-3 Is a Transcriptional Repressor of Nitric-oxide Synthase 2 J. Biol. Chem., October 10, 2003; 278(41): 39572 - 39577. [Abstract] [Full Text] [PDF]
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S. L. Throm and M. J. Klemsz PU.1 regulates glutathione peroxidase expression in neutrophils J. Leukoc. Biol., July 1, 2003; 74(1): 111 - 117. [Abstract] [Full Text] [PDF]
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H. Tsujimura, T. Tamura, C. Gongora, J. Aliberti, C. Reis e Sousa, A. Sher, and K. Ozato ICSBP/IRF-8 retrovirus transduction rescues dendritic cell development in vitro Blood, February 1, 2003; 101(3): 961 - 969. [Abstract] [Full Text] [PDF]
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J. Aliberti, O. Schulz, D. J. Pennington, H. Tsujimura, C. R. e Sousa, K. Ozato, and A. Sher Essential role for ICSBP in the in vivo development of murine CD8alpha + dendritic cells Blood, January 1, 2003; 101(1): 305 - 310. [Abstract] [Full Text] [PDF]
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M. G. Manz, D. Traver, T. Miyamoto, I. L. Weissman, and K. Akashi Dendritic cell potentials of early lymphoid and myeloid progenitors Blood, June 1, 2001; 97(11): 3333 - 3341. [Abstract] [Full Text] [PDF]
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F. Colucci, S. I. Samson, R. P. DeKoter, O. Lantz, H. Singh, and J. P. Di Santo Differential requirement for the transcription factor PU.1 in the generation of natural killer cells versus B and T cells Blood, May 1, 2001; 97(9): 2625 - 2632. [Abstract] [Full Text] [PDF]
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 D. G. Gilliland The Diverse Role of the ETS Family of Transcription Factors in Cancer: Commentary re: B. Davidson, Ets-1 Messenger RNA Expression Is a Novel Marker of Poor Survival in Ovarian Carcinoma. Clin. Cancer Res., 7: 551-557, 2001. Clin. Cancer Res., March 1, 2001; 7(3): 451 - 453. [Full Text]
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 D. Traver, K. Akashi, M. Manz, M. Merad, T. Miyamoto, E. G. Engleman, and I. L. Weissman Development of CD8{alpha}-Positive Dendritic Cells from a Common Myeloid Progenitor Science, December 15, 2000; 290(5499): 2152 - 2154. [Abstract] [Full Text]
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