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*
Wistar Institute, Philadelphia, PA 19104; and
Institute for Human Gene Therapy, University of Pennsylvania Medical School, Philadelphia, PA 19104.
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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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, or B220
expression, and no Ig rearrangements were detected. No thymocytes
positive for the T markers Thy-1 or CD2 were detected, and the thymus
was relatively hypocellular. In addition, the PU.1 deficiency resulted
in late embryonic lethality due to unknown causes. These results
strongly suggest that both lymphoid and myeloid cells require
PU.1. A second strain of PU.1-/- mice was independently generated by McKercher et al. (5). Where examined, the embryonic phenotype of the two mutations is nearly identical. If, however, McKercher et al. maintained PU.1-/- pups on antibiotics, they could survive up to 2 wk. Surviving animals were found to contain increasing numbers of thymocytes expressing CD4 and CD8 and TCR. By day 12 after birth, many PU.1-/- thymi contained thymocyte numbers within fivefold of wild type, and were relatively normal with respect to expression of CD4, CD8, and TCR. Mature phenotype T cells were also found in the spleen. Interestingly, older antibiotic-treated mice also contained small but significant numbers of B220, rag-expressing cells, possibly aberrant B progenitors, and rare immature and nonfunctional neutrophils. The basis for the differences in the two models is currently unexplained. Possibilities include strain differences and/or differences in targeting constructs. In light of these discrepancies in phenotype, we have reexamined T cell development in PU.1-/- fetal thymus.
The thymus is seeded on an ongoing basis with hemopoietic progenitors derived from fetal liver or bone marrow. These incoming progenitors do not self-renew within the thymus but rather become rapidly committed for differentiation and expansion to the T and associated (NK, thymic dendritic) cell lineages. One hypothesis of lineage commitment suggests that initial specification events induce the expression of particular transcription factors that replace or supplement transcription factors already present in the pluripotent precursors (6). The key role played by large transcription factor families in lineage-specific gene expression supports this hypothesis. Transcription factor families couple similar DNA-binding specificities with differences in transactivation ability and protein-protein interactions. Divergent patterns of expression within a family can provide the necessary regulation of a specific developmental program. A second feature of this hypothesis is the combinatorial nature of gene regulation that requires multiple transcription factors to work in concert to express a given gene. Specific protein-protein interactions within a higher order protein complex can have as much influence on gene expression as DNA-binding specificity. The ets family of transcription factors is a good example in support of this hypothesis (7). The ets family is composed of at least 17 different members with similar DNA-binding specificities due to related DNA-binding domains. The nonmammalian family members have been implicated in a variety of developmental programs including steroid hormone-induced metamorphosis in Drosophila (8, 9) and oocyte pattern formation in frogs (10). In birds and mammals, ets proteins play an important role in regulating hemopoiesis (3, 7, 11).
Studies from many groups have characterized early T cell development
and commitment with respect to a variety of cell surface markers. The
earliest, uncommitted progenitors are known to express CD44,
c-kit, and lack expression of TCR, CD4, CD8, and other
T-specific markers. Commitment of these progenitors to T, NK, and
lymphoid dendritic cell lineages has been correlated with expression of
a variety of markers. For example, Hattori et al. have determined that
expression of Fc
RII/III on E12.5-E14.5 thymocytes is correlated with
the expression of T-specific factors, including TCF-1, GATA-3, and
intracellular CD3
(12, 13). Carlyle and Zuniga-Pflucker
(14, 15) have correlated early thymic expression of the
NK1.1 marker with transition to the T/NK bipotential stage. Finally,
up-regulation of CD25 and down-regulation of CD44 are associated with
commitment to the 
ß T cell lineage to the exclusion of other cell
types, including 
T cells, NK cells, and thymic dendritic cells
(16, 17, 18). Using this battery of markers, we have
characterized the extent of differentiation and commitment among
PU.1-deficient fetal thymocytes. We find that the majority of
PU.1-deficient fetal thymocytes are blocked at an apparently
uncommitted progenitor stage. However, we also find that PU.1-deficient
thymus contains an extremely rare population of T committed and early
differentiating T progenitors, which go on to develop mature,
functionally competent thymocytes with time in fetal thymic organ
culture.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PU.1-deficient mice were of mixed 129/C57BL/6 background (backcrossed to 129 a maximum of three times) and were bred in our own facility.
Abs, staining, flow cytometric analysis
Thymic lobes were mechanically dissociated and stained using
standard procedures. mAbs were: anti-CD4 FITC (Caltag, South San
Francisco, CA), anti-CD8 Tricolor (Caltag), and anti-CD3 PE
(Life Technologies, Rockville, MD) or anti-CD4-FITC,
anti-CD8-FITC (Caltag), anti-CD25-PE (PharMingen, San Diego,
CA), biotinylated-CD44 (a gift of Dr. E. Pure), and Streptavidin Red
670 (Life Technologies), anti-Thy-1 FITC (Becton Dickinson),
Sca-1-PE (PharMingen), NK1.1-FITC (PharMingen), HSA-FITC (PharMingen),
anti-FcRII/III (a gift of S. Carding). Flow cytometric analysis
was performed on a FACScan and analyzed using CellQuest software
(Becton Dickinson). In some cases, flow cytometry was used to quantify
the relative cell yield from each thymic lobe. In these cases, thymic
lobes were dissociated in equal volumes, 200 µl, which provides a
cell density within the limit of accurate detection by the cytometer.
Samples were collected for a fixed time, generally 1 min, with the
sample tube-washing system disabled. Samples were gated as usual for
live and dead cells using forward and side scatter, and the number of
live events compared between different samples. The reliability of the
technique was confirmed by direct counting of trypan blue excluding
cells using a hemacytometer (
10% of the samples were directly
counted).
Thymocyte fractionation and RT-PCR
Thymi were harvested from C57BL/6 mice and single cell suspensions prepared by manual disruption. Thymocyte populations were fractionated via magnetic bead separation as per the manufacturer’s instruction (MACS, Miltenyi Biotec, Auburn, CA) as follows. Thymocytes were stained with B cell (anti-B220) and myeloid (anti-CD11b) specific Abs (PharMingen) and subjected to two rounds of negative selection. The remaining thymocytes were stained with anti-CD4 and anti-CD8 Abs followed by two rounds of positive selection to generate a mixed population of CD4+CD8+ (double-positive), CD4+8-, and CD4-8+ (single-positive) cells. The thymocytes were then stained with anti-CD25 and subjected to two rounds of positive selection to generate the CD25+, CD4-, CD8- population. Finally, the remaining cells were stained with anti-CD44 and subjected to two rounds of positive selection to generate the CD44+, CD25-, CD4-, CD8- population. CD4+ or CD8+ splenic T cells were also selected in a similar manner. All populations were confirmed >95% pure by flow cytometry (data not shown). RNA was prepared using RNAzol (Teltest, Friendswood, TX) following the manufacturers instructions. RNA was also prepared from whole spleen to serve as a positive control. Semiquantitative PU.1 RT-PCR was performed as described previously (19) using the following primers: PU.1, 5'-GAGTTTGAGAACTTCCCTGAG-3', 5'-TGGTAGGTCATCTTCTTGCGG-3'; ETS-1, 5'-GGAATTCCTGAATACACAGTATAGTGAG-3', 5'-CGGTCGACTCCTGTGTAGCCAGCCAG-3'; FLI-1, 5'-GGAATTCGGGTCAATGTGTGGAATATTGG-3', 5'-CGGTCGACCCCAGGGTTTGCTAGGCG-3'; HPRT, 5'-CACAGGACTAGAACACCTGC-3', 5'-GCTGGTGAAAAGGACCTCT-3'. PCR conditions were established for each transcript to ensure that analysis was undertaken in the linear amplification range. For the abundant HPRT message, 25 amplification cycles were used. For the transcription factor cDNAs, 28 rounds of amplification were used. RT-PCR for the myeloid-specific transcript CD11b was used to confirm the purity of the fractionated thymocyte subset RNA (data not shown). No template reactions also served as a negative control (data not shown).
Fetal thymic organ cultures (FTOCs)4
FTOCs were performed according to procedures previously described (20). Embryos were generated by heterozygous crosses, timed from the first day of plug observation (day 0.5), and embryos dissected and placed in ice-cold RPMI plus 10% FCS. A small piece of liver was dissected from each embryo, dissociated, and stained using anti-CD11b Ab and flow cytometry to identify mutants. The reliability of the CD11b staining to detect mutants was confirmed by Southern analysis of embryo DNA (data not shown). During the staining and analysis (<2 h), all control and mutant littermate embryos were held on ice. Dissected thymic lobes were either pooled and dissociated for staining or cultured in RPMI (Life Technologies) plus 10% FCS (Cansera, Ontario, Canada), glutamine (Life Technologies), gentamicin, and suspended on nuclepore filter rafts (Costar, Cambridge, MA) supported by gelfoam (Upjohn, Kalamazoo, MI). Culture medium was partially replaced every 3 days.
Proliferation assays
The number of cells plated was normalized such that each well
received 2 x 104 TCRhigh thymocytes. A
total of 2 x 105 irradiated spleen cells were added
to provide costimulation, and cells were exposed to plate-bound
anti-CD3 Ab for 48 h, then pulsed for 12 h with
[3H]thymidine and harvested and counted. Since
PU.1-deficient thymuses contained fewer TCRhigh thymocytes,
many more cells were added. To control for this, background levels of
3H incorporation were determined for equivalently high
numbers of wild-type thymocytes, and found to be indistinguishable from
controls.
Statistics
Differences between two means were evaluated using Student’s t test (Microsoft Excel, Redmond, WA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The ets family members, PU.1, Ets-1, Ets-2, Spi-B, Fli-1, and
Elf-1, are all expressed at some point during thymocyte development
(7). Ets-2 is ubiquitously expressed, but transgenic
overexpression leads to abnormal thymic histology, among other defects
(21, 22, 23, 24). Fli-1 is expressed at high levels in the spleen,
thymus, and lung with lower levels of expression in skeletal muscle
(25). Spi-B is expressed in fetal thymus and spleen with
low level expression found in some T cell lines (26).
Targeted disruption of PU.1 leads to defects in early stages
of lymphoid development as discussed (4, 27), while
disruption of Ets-1 leads to reduced mature thymocyte and
peripheral T cell numbers, increases in T cell apoptosis, and decreased
T cell activation (28, 29). Targeted disruption of
Fli-1 results in thymic hypocellularity (30).
Therefore, we examined the expression of PU.1,
Ets-1, and Fli-1 at different stages of thymocyte
development. To produce subsets at different developmental stages,
thymocytes were fractionated based on expression of CD4, CD8, CD25, and
CD44, and RNA was prepared. Expression levels of the ets family
transcription factors PU.1, Ets-1, and
Fli-1 were determined by semiquantitative RT-PCR (Fig. 1
). PU.1 was found to be expressed in the
most immature
CD4-8-25-CD44+
thymocytes, at lower levels on
CD4-8-CD25+ thymocytes, and was
undetectable in more mature mixed populations of
CD4+8+, CD4-8+, or
CD4+8- thymocytes or splenic T cells. Thus
PU.1 expression is inversely correlated with T cell
developmental progression. Expression of Ets-1, however, had
a nearly opposite pattern of expression with no detectable expression
in the most primitive CD4-8-25-
CD44+ thymocytes, followed by up-regulated expression as
thymocytes become committed to the T cell lineage. Fli-1 was
found to be expressed early, and increased as T cells matured. Both
Ets-1 and Fli-1 expression peaked in the mixed
CD4+8+, CD4+8-,
CD4-8+ thymocyte populations and remained high
in circulating splenic T cells.
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To determine precisely the stage at which T cell development is arrested in PU.1-/- animals, embryos were harvested at E14.5, and E16.5 and thymic cellularity and phenotype assessed. When thymic lobes from E14.5 embryos were pooled and counted, mutant thymus contained on average 0.5 x 105 cells per lobe, compared with 1.65 x 105 cells per lobe in wild-type littermate controls. Cell numbers were observed to expand in vivo in wild-type embryos, so that by E16.5 wild-type lobes contained 6.7 ± 1.9 x 105 cells per lobe, while mutants still contained just 0.2 ± 0.1 x 105 cells per lobe. Therefore, at E16.5, wild-type and mutant cell numbers significantly differed (p = 0.001) by over 30-fold.
The expression of the markers Thy-1, Sca 1, NK1.1, HSA, CD44, CD25,
FCRII/III, CD4, and CD8 were determined using flow cytometry.
Neither wild-type nor PU.1-/- thymocytes expressed CD4 or
CD8 at E14.5, as expected (data not shown). The majority of
PU.1-/- thymocytes were smaller on average than wild-type
thymocytes as determined by forward scatter (Fig. 2A). These smaller
PU.1-/- thymocytes were Thy-1 and CD25 negative, and
expressed intermediate levels of CD44 (second column, Fig. 2B). A similar population of Thy-1-
CD25- CD44+ thymocytes was found in wild-type
thymus at E14.5, and these were enriched in the smaller cell gate
(third column, Fig. 2B). The majority of wild-type
thymocytes at E13.5 also showed a similar Thy-1-
CD25- and CD44+ phenotype (fourth column, Fig. 2B). Besides this major population, 2% of
PU.1-/- thymocytes were larger by forward scatter. Among
these larger cells, the majority expressed Thy-1 and about half of
these expressed CD25.
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C). Again, a similar population was observed in wild-type
thymus, which was especially enriched if thymocytes were gated for the
smaller size (Fig. 2C). HSAbright
c-kitint cells were also enriched in wild-type
thymus at E13.5 (Fig. 2C). Again, PU.1-/-
thymi contained a low but reproducible population of HSAlow
c-kitbright cells consistent with further
developmental maturation.
About half of wild-type thymocytes at E14.5 express FcRII/III, which
has been shown to correlate with the beginning of T differentiation
(12, 13, 31) (Fig. 2D). In
PU.1-/- thymus, about 3% of thymocytes were found to
express FcRII/III. In wild-type thymus, the smaller cell gate was
depleted of FcRII/III-expressing cells (Fig. 2D), as were
E13.5 wild-type embryos. Sca-I expression was observed on a subset of
E14.5 wild-type thymus, but was not seen in PU.1-deficient thymus and
was dramatically depleted on small wild-type cells.
NK1.1 is expressed on committed T/NK progenitors in fetal thymus, and
also on mature NK T cells, which have been shown to arise early in
ontogeny (16). We found that PU.1-/- thymi
contained a small fraction of NK1.1+ thymocytes, similar to
that observed in E14.5 wild-type controls gated for smaller size (Fig. 2E).
When individual E14.5 thymic lobes were examined, Thy-1+CD25+ cells were present in just one out of three lobes. This suggests that the small percentage of Thy-1+CD25+ cells in pooled samples was due to increased development in one or a few lobes, rather than a low percentage of development present in all lobes.
Individual lobes were also examined in the case of E16.5 thymi. The
majority of thymocytes expressed the Thy-1-
CD25- FcRII/III- Sca I-
NK1.1- CD4- CD8-
CD44+ HSAbright c-kitint
phenotype described above for PU.1-/- thymocytes at
E14.5. In addition, in two cases a small percentage (8–11%) of
HSAlowCD8+ cells was observed (one shown in
Fig. 3). Although by E16.5 many of the
wild-type cells were clearly expressing CD4 and CD8, we were still able
to distinguish a small population of HSAbright,
CD4-8- wild-type cells phenotypically similar
to the predominant PU.1-/- phenotype (boxed population
shown in Fig. 3).
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RII/III- Sca I- NK1.1-
CD4- CD8- (hereafter abbreviated
T-Lin-) CD44+ HSAbright
c-kitint. Based on the total cell counts and
the percentage of cells in this subset, the absolute number of cells
with this phenotype was calculated to be 0.3–0.4 x
105 per thymic lobe for both PU.1-/- and
wild-type embryos at E14.5. Therefore, PU.1-/- mice do
not have reduced numbers of this progenitor subset. At E16.5, the
absolute number of these progenitors has dropped to between 0.1 to
0.2 x 105 in both mutant and wild-type embryos. T cell development in PU.1-deficient FTOCs
The existence of a low percentage of Thy 1+,
CD25+, and FcRII/III+, and
NK1.1+ cells in PU.1-/- fetal thymi suggested
that some of the earliest steps of T cell development were possible in
the absence of PU.1. Since the PU.1 defect is embryonic lethal, we
tested this possibility further by incubating E14.5 FTOC for 12–16
days. The thymic lobes were harvested and analyzed by flow cytometry
for expression of CD4, CD8, and TCR. We found that 4 out of 20
(20%) mutant lobes developed >1000 thymocytes. Furthermore, in 9 out
of 20 lobes (45%), greater than 10% of thymocytes developed to the
CD4+8+ stage (Fig. 4). In addition, a few lobes developed
mature CD4+TCR+ thymocytes (see below).
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). High progressors had similar patterns of
expression of CD4 and CD8 compared with wild-type littermate controls,
but often showed higher percentages of CD4-8-
and CD25+ thymocytes. Intermediate and low progressors were
progressively more enriched for CD4-8- and
CD25+ progenitors. In addition, even high progressors
maintained a higher level of overall HSA and Thy-1 expression
consistent with relative immaturity (Fig. 5). Among those PU.1-/-
lobes that developed fewer CD4 or CD8 expressing cells, (intermediate
and low progressors) the high percentages of CD25 indicates that
development had proceeded beyond the T cell commitment stage of
development. Nonprogressors failed to develop at all and few live
thymocytes were recovered (data not shown). Of the 20 lobes shown in
Fig. 4, 3 were classified as high progressors (CD4, CD8, TCR expression
comparable with wild-type), 7 were intermediate progressors (greater
than 5% CD4+8+ thymocytes), 5 were low
progressors (<5% CD4+8+, but CD25 expression
detected) and 5 did not develop (fewer than 100 cells recovered).
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In some experiments, the appearance of TCR+ T cells
was assessed. All three mutant lobes examined showing some T commitment
also contained up to 11% CD4-8-
TCR-positive, CD4- thymocytes (data not shown).
To determine whether PU.1-deficient thymocytes were functionally
competent, cultures of E14.5 thymic lobes were harvested after 14 days
and tested for expression of CD4, CD8, TCR, and, subsequently, for
proliferative responses to anti-CD3. By normalizing the number of
wild-type and PU.1-/- thymocytes plated based on
expression of TCR, we found that PU.1-/- thymocytes
proliferated as well as wild-type thymocytes in response to
anti-CD3 cross-linking (Fig. 6).
In addition, 48 h after stimulation, culture supernatants were
harvested and assessed for IL-2 production in bioassays. IL-2-dependent
proliferation of CTLL-2 cells (>10 times over background) was observed
in response to both mutant and wild-type culture supernatants from
stimulated, but not unstimulated cells. The proliferation was inhibited
by neutralizing Abs against IL-2 (data not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The phenotype of the major PU.1-/- population,
Thy-1- CD25- FcRII/III- Sca
I- NK1.1- CD4- CD8-
(T-Lin-) CD44+ HSAbright
c-kitlow, is consistent with previous
identifications of the most immature thymic progenitors before
commitment to the T lineage (12, 13, 31, 33, 34, 35, 36).
Therefore, the major population of thymic progenitors present in
PU.1-/- thymus is likely blocked at an early stage of
development.
Our data further demonstrate that PU.1 is not absolutely required for
fetal T cell development, since even as early as E14.5, rare
Thy-1+, CD25+ and at E16.5, CD8+
thymocytes are detected. We found that committed T cell progenitors in
PU.1-deficient embryos occurred as a low percentage of total
thymocytes, and stochastically in rare thymic lobes, features which
likely contributed to their lack of detection in earlier experiments.
We further found that, when placed in FTOC, a significant fraction of
PU.1-/- thymi developed beyond the point of T cell
commitment and in some cases developed to maturity. These mature T
cells were shown to proliferate and produce IL-2 in response to
stimulation through the TCR. We have also detected TCR
expressing CD4- thymocytes in PU.1-/- FTOCs,
demonstrating that this lineage also does not absolutely require PU.1
activity. NK1.1 expression was observed in PU.1-/- thymi,
suggesting that thymic NK cell development may also occur. However,
further experiments are necessary to determine whether mature,
functional NK T cells are present in the PU.1 mutant, since the NK1.1
marker is present on a subset of immature progenitors. Furthermore, we
also detect significant numbers of lymphoid dendritic cells in both
fresh E16.5 thymus and after FTOC (see footnote 3). This demonstrates
that PU.1 activity is not absolutely essential for T cell development
and function.
PU.1-/- T cell development differed from wild-type in
several major aspects. First, developmental progression, and stage of
progression was variable in PU.1-/- thymuses. Two thymic
lobes isolated from the same individual differed in their extent of
development in FTOC, suggesting that differences in genetic background
do not contribute to developmental variability. Interestingly, T cell
development in neonatal PU.1 mutants (5) also showed
considerable variability. This suggests that PU.1 may contribute to
whatever process ensures consistent developmental timing of T cell
development. Second, T cell development in PU.1-deficient thymus was
delayed. Third, even when cultured PU.1-/- thymocytes
progressed comparably to wild type, recovered cell numbers were still
substantially lower. These results are most consistent with a model in
which PU.1 function is most essential in an early progenitor, before
the ß T, T, and thymic dendritic cell lineages split. For
example, if PU.1 deficiency reduced the efficiency of T cell
commitment, fewer progenitors would be able to differentiate, reducing
cellular recovery. In addition, a lowered efficiency of T cell
commitment could also explain the variability and delayed developmental
kinetics seen in PU.1-/- thymus cultured in vitro.
T cell development may also provide direct evidence to support the overlapping functional nature of transcription factor families during development. We examined the expression of selected ets family transcription factors during thymocyte development. PU.1, Ets-1, and Fli-1 were chosen because gene targeting has demonstrated that all three play a role in the T lineage (7). Fli-1 was expressed at all stages of thymocyte development examined, both pre- and post-commitment. Its early expression pattern is consistent with its probable function in an early thymic progenitor, contributing to the reductions in thymic cellularity observed in Fli-1 gene disruption experiments (30). Since Fli-1 is also expressed later, it may also play a role in other aspects of T cell function. PU.1 was expressed in noncommitted thymocytes, but was down-regulated at the point when T cells become committed, consistent with its role in early progenitors prior to T cell commitment. In contrast, Ets-1 has a later expression pattern consistent with its importance for the survival and/or maturation of single-positive thymocytes and peripheral T cells, and in T cell activation, as shown by gene targeting (28, 29). The promoters for both PU.1 and Ets-1 have functionally important ets family-binding motifs that are thought to play a role in auto-regulation (7, 37, 38, 39). Perhaps low level gene expression of Ets-1 is initiated by PU.1, which then leads to a true auto-regulatory loop for high level expression. Thus, overlapping expression of transcription factor families may explain the "chicken or egg" paradox of initiating gene expression. Since T cell development can occur in the absence of PU.1, Ets-1, or Fli-1, these transcription factors are not individually required for T cell commitment. All, however, appear to be required for efficient T cell development. This suggests an incomplete functional redundancy among the ets family transcription factors expressed during T cell development. The production of double and triple mutants for transcription factors within the T lineage will be required to address this issue.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Lisa
M. Spain, Wistar Institute, 3601 Spruce Street, Philadelphia, PA
19104-4268 (E-mail address:
3 A. Guerriero, L. M. Spain, and E. W. Scott. PU.1 is required for myeloid-derived but not lymphoid-derived dendritic cells. Submitted for publication.
4 Abbreviations used in this paper: FTOC, fetal thymic organ culture; HPRT, hypoxanthine phosphoribosyltransferase.
Received for publication April 27, 1999. Accepted for publication June 16, 1999.
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ß T lymphocyte lineage commitment. Immunity 9:187.[Medline]
ß T cell differentiation in mouse fetal thymic ontogeny. J. Immunol. 160:744.ß or - lineages can occur just prior to the onset of CD4 and CD8 expression among immature thymocytes. Eur. J. Immunol. 22:2185.[Medline]
RII/III and CD2 expression mark distinct subpopulations of immature CD4-CD8- murine thymocytes: in vivo developmental kinetics and T cell receptor ß chain rearrangement status. J. Exp. Med. 177:1079. and IL-1 in fetal thymocyte commitment and differentiation. Science 268:1906.This article has been cited by other articles:
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H.-G. Kim, C. G. de Guzman, C. S. Swindle, C. V. Cotta, L. Gartland, E. W. Scott, and C. A. Klug The ETS family transcription factor PU.1 is necessary for the maintenance of fetal liver hematopoietic stem cells Blood, December 15, 2004; 104(13): 3894 - 3900. [Abstract] [Full Text] [PDF]
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 E. Tiacci, S. Pileri, A. Orleth, R. Pacini, A. Tabarrini, F. Frenguelli, A. Liso, D. Diverio, F. Lo-Coco, and B. Falini PAX5 Expression in Acute Leukemias: Higher B-Lineage Specificity Than CD79a and Selective Association with t(8;21)-Acute Myelogenous Leukemia Cancer Res., October 15, 2004; 64(20): 7399 - 7404. [Abstract] [Full Text] [PDF]
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M. K. Anderson, R. Pant, A. L. Miracle, X. Sun, C. A. Luer, C. J. Walsh, J. C. Telfer, G. W. Litman, and E. V. Rothenberg Evolutionary Origins of Lymphocytes: Ensembles of T Cell and B Cell Transcriptional Regulators in a Cartilaginous Fish J. Immunol., May 15, 2004; 172(10): 5851 - 5860. [Abstract] [Full Text] [PDF]
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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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R. Schotte, M.-C. Rissoan, N. Bendriss-Vermare, J.-M. Bridon, T. Duhen, K. Weijer, F. Briere, and H. Spits The transcription factor Spi-B is expressed in plasmacytoid DC precursors and inhibits T-, B-, and NK-cell development Blood, February 1, 2003; 101(3): 1015 - 1023. [Abstract] [Full Text] [PDF]
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R. N. Laribee and M. J. Klemsz Loss of PU.1 Expression Following Inhibition of Histone Deacetylases J. Immunol., November 1, 2001; 167(9): 5160 - 5166. [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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 R. P. DeKoter and H. Singh Regulation of B Lymphocyte and Macrophage Development by Graded Expression of PU.1 Science, May 26, 2000; 288(5470): 1439 - 1441. [Abstract] [Full Text]
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K. L. Anderson, H. Perkin, C. D. Surh, S. Venturini, R. A. Maki, and B. E. Torbett Transcription Factor PU.1 Is Necessary for Development of Thymic and Myeloid Progenitor-Derived Dendritic Cells J. Immunol., February 15, 2000; 164(4): 1855 - 1861. [Abstract] [Full Text] [PDF]
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A. Guerriero, P. B. Langmuir, L. M. Spain, and E. W. Scott PU.1 is required for myeloid-derived but not lymphoid-derived dendritic cells Blood, February 1, 2000; 95(3): 879 - 885. [Abstract] [Full Text] [PDF]
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