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University of Ghent, University Hospital, Department of Clinical Chemistry, Microbiology and Immunology, Ghent, Belgium
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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and 
ß
T lymphocytes. These findings support the notion that the T/NK split
occurs downstream of the NK/dendritic split. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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ß
high, CD4+CD8-, or
CD4-CD8+ (single positive, SP) cells, serve as
indicators of the degree of maturation of the donor population. Using
these techniques, a very small population of thymocytes has been
identified that seems to include early progenitors. In mice, these
cells are identified by the expression of CD44, high levels of class I
MHC, low levels of CD4 and heat stable Ag, and absence of CD3
and CD8 (4, 5). In adoptive transfer experiments, these cells give rise
to B cells as well as T cells, but not to myeloid cells.
The study of intrathymic developmental pathways in humans relies on
phenotypic analysis. Particular seminal has been the discovery of CD34
as a stage-specific marker identifying immature hematopoietic stem
cells (6, 7). The expression level of CD34 and its coexpression with
differentiation markers as well as the capacity of these subsets to
differentiate in FTOC allowed the delineation of different stages in
human intrathymic development (8). More recently, on the basis of their
ability to reconstitute thymopoiesis in ectopic human fetal thymus
implants in immunodeficient C.B17 scid/scid (SCID) mice, the early
stages of lymphoid cell formation of phenotypically defined subsets of
CD34+ bone marrow cells have been defined (9). However,
although bone marrow lymphoid precursors have been carefully studied,
little is known about the sequential appearance of phenotypic markers
during the first 2 wk after the entrance of the human precursor cell in
the thymus. Phenotypic changes during the differentiation process have
been proposed on the basis of multiparametric flow cytometric studies
of the thymus (7), or more recently by assessing the differentiation of
sorted thymus subpopulations in human FTOC (8, 10) or in murine FTOC
(11), but data on the kinetics of the different lineages are lacking.
Using a hybrid human/scid mouse FTOC system (12), we address in the
present study the kinetics of the very early steps of differentiation
of CD34+CD38-Lin- precursor cells
derived from fetal liver by analysis of the sequential appearance of
surface Ags and the expression of recombination activating gene 1
(RAG-1) and pre-TCR -chain (pT). We report the existence of
different phenotypic intermediates that produce lymphocytes, NK cells,
and dendritic cells and demonstrate that the kinetics of these distinct
pathways differ.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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CB-17 SCID mice were obtained from our own specific pathogen-free breeding facility. For timed pregnancies, females were housed in separate cages from the males until mating. The appearance of vaginal plug after overnight mating was labeled as day 0 of pregnancy. The 14- to 15-day pregnant mice were sacrificed to obtain the embryos for preparation of the thymic lobes.
Abs and reagents
The mAbs used were rat anti-mouse CD45 cytochrome
(CD45, 30F1 1.1; PharMingen, San Diego, CA), biotinylated rat
anti-mouse H-2 class I MHC H-2Dd (PharMingen), and the
following mouse anti-human mAbs: anti-glycophorin-A (10F7 MN; a
kind gift of Dr. L. L. Lanier, DNAX, Palo Alto, CA), CD1a (B-B5;
Serotec, Oxford, U.K.), CD38 (OKT10 FITC; Ortho, Raritan, NJ), CD2
(Leu-5b FITC), CD3 (anti CD3
, Leu-4 FITC, phycoerythrin (PE)), CD4
(Leu-3a FITC or PE), CD7 (Leu-9 FITC), CD8 (Leu2a FITC), CD14 (LeuM3
PE), CD16 (Leu-11a FITC), CD19 (Leu-12 PE), CD34 (HPCA-2 FITC or PE),
CD45 (HLe-1 FITC), CD56 (clone NCAM16 PE), anti-TCR-ß (TCR-1
FITC), and anti-TCR- (anti-TCR-/l FITC) (all from
Becton Dickinson Immunocytometry Systems, Mountain View, CA). Human
recombinant IL-15 (CHO-derived) was kindly provided by Immunex
(Seattle, WA).
Preparation of human CD34+CD38-Lin- fetal liver cells
Human fetal liver tissues were obtained and used following the guidelines of the medical ethical commission of the University Hospital of Ghent. Human fetal liver cells were isolated by gentle disruption of the tissue in complete medium (Iscove’s modified Dulbecco’s medium/10% FCS, Life Technologies, Paisley, Scotland) followed by centrifugation over Lymphoprep (Nycomed Pharma, Oslo, Norway). Cells were washed and resuspended in 90% FCS/10% DMSO and frozen in liquid N2. After thawing, fetal liver cells were washed and labeled with glycophorin A, CD19, and FITC-labeled mAbs CD1, CD3, CD4, CD7, CD8, and CD38. Ab-labeled cells were depleted by immunomagnetic beads. For this purpose, cells were resuspended in 0.5 ml cold PBS/2% FCS and mixed with 0.5 ml prewashed (to remove the preservative) sheep anti-mouse Ig-coated Dynabeads (Dynal AS, Oslo, Norway) to obtain a ratio of cells to Dynabeads of 1:5. After 30 min at 4°C, the suspension was diluted by carefully adding 5 ml PBS/2% FCS, and the rosettes of cells with Dynabeads were removed by placing the tube on a magnetic particle concentrator (Dynal AS). The supernatant that contained the unlabeled and weakly stained cells was removed and centrifuged (500 x g for 6 min). These cells were labeled with CD34-PE and CD34+CD38-Lin- sorted. The sorted cells were transferred to murine thymic lobes by the hanging drop method (13).
FTOC
Thymic lobes were prepared from fetal day 14–15 SCID mice. Hanging drops were prepared in Terasaki plates by adding in each well 25 µl of complete medium containing 10,000 or less sorted human CD34+CD38-Lin- fetal liver cells and one thymic lobe. The plates were immediately inverted to form hanging drops and incubated in an humidified incubator for 48 h (7.5% CO2 in air, 37°C).
After incubation, the lobes were removed from the hanging drop, washed, and placed on the surface of a nuclepore filter in organ culture in an incubator. At different time points, thymocytes were recovered by mechanical disruption of the thymic lobes with a small tissue grinder. Thymocytes were stained with eosin and counted with a hematocytometer. At least 80% of the cells were viable.
Flow cytometry
Before labeling, cells were suspended in PBS/1% BSA/0.1%
NaN3, and the Fc receptor of the mouse thymocytes was
blocked by preincubation for 15 min with anti-FcRII/III mAb
(clone 2.4.G2) (14) to avoid aspecific binding of Abs by the murine
thymocytes. Subsequently, the cells were stained with a panel of mAb,
as indicated. The biotinylated Abs were revealed with second step
streptavidin Tri-Color (Caltag, Burlingame, CA).
All mAb against human Ags and second step reagents were checked for negative staining on SCID thymocytes after blocking with 2.4G2 mAb. Isotype controls were also included in most staining series and were found to be negative.
For intracellular staining of CD3 (cytoplasmatic (cy) CD3), cells were first labeled with mAbs against mouse and human surface Ags, and after two washes the cells were permeabilized with Ortho Permeafix (Ortho, Raritan, NJ) according to the guidelines of the manufacturer. After two washes, cells were labeled with CD3-FITC. The cells were analyzed on a FACScan (Becton Dickinson Immunocytometry Systems) with an argon-ion laser tuned at 488 nm. Forward light scattering, orthogonal scattering, and three fluorescence signals were determined on 10,000 cells and stored in listmode data files. Data acquisition and analysis was done with Lysis 2.0 software (Becton Dickinson Immunocytometry Systems). Dead cells were gated out by propidium iodide (PI) exclusion, except for permeabilized cells, where the dead cells were gated out according to different scatter properties. In most cases, viable human cells were gated by exclusion of mouse CD45+, mouse class I MHC+ cells, and PI-positive cells.
Allogeneic MLC assays
PBL were obtained by centrifugation of blood from healthy adult volunteers over Lymphoprep (Nycomed Pharma). PBL were resuspended in RPMI 1640 with 10% filtered human AB serum and 5 x 10-5 M 2-ME (complete MLC medium). MLC were set up in 0.2 ml complete MLC medium in round-bottom 96-well trays (Falcon, Becton Dickinson, Lincoln Park, NJ). The stimulator cells were either sorted human thymocytes from FTOC after 11 days of culture (2 x 103 - 2 x 104) or sorted CD4+HLA-DR+ (102–103) putative dendritic cells thereof or PBL (104 - 5 x 104), each given 30 Gy irradiation. The responder cells (5 x 104/well) were PBL from a different donor. The cultures were incubated in a humidified incubator (5% CO2 in air) for 6 days. Each culture was pulsed overnight with 1 µCi [3H]TdR (Amersham, Amersham, England) per well, before harvesting of the cultures onto fiberglass filters. Tritiated thymidine incorporation was determined by liquid scintillation counter fluid counting (Microbeta, Wallac, Turku, Finland). Results are expressed as cpm.
RNA isolation, reverse transcription, and RT-PCR
A total of 2 x 104–105 cells were
harvested for determination of mRNA encoding human pT, RAG-1, or
hypoxanthin phosphoribosyltransferase (HPRT) by reverse
transcription-PCR (RT-PCR). The procedure for RNA isolation and RT-PCR
was performed with minor modifications as described (15). In brief,
total RNA was isolated using Trisol (Life Technologies), and further
processed following the instructions of the supplier. Then, 10 µg of
Escherichia coli tRNA (Boehringer, Indianapolis, IN) was
added as a carrier before chloroform addition and cDNA was prepared
using Superscript (Life Technologies) according to the specifications
of the supplier. An aliquot of each cDNA sample was subsequently
amplified in twofold serial dilutions by PCR using pT-, RAG-1-, and
HPRT-specific primers. In each PCR, water, SCID-mouse fetal thymus
cDNA, and human and mouse genomic DNA were included as negative
controls. All primer pairs amplified human cDNA only. PCR amplification
using the different primers was performed in 50 µl volumes with 1 U
of Taq polymerase (Perkin-Elmer, Emeryville, CA) using a
96-well thermocycler (Omnigene, Hybaid Teddington, U.K.) under the
following conditions: pT, 35 cycles of 30 s at 94°C, 30
s at 65°C, and 1 min at 72°C; RAG-1 and HPRT, 30 cycles of 30
s at 94°C, 30 s at 55°C, and 1 min at 72°C. The sequences of
the primers were as follows: for pT, 5'-TCC AGC CCT ACC CAC AGG
TG-3' (sense primer) and 5'-TAG AAG CCT CTC CTG ACA GAT GCA T-3'
(anti-sense primer); for RAG-1, 5'-CCA AAT TGC AGA CAT CTC AAC-3'
(sense primer) and 5'-CAA CAT CTG CCT TCA CAT CGA TCC-3'
(anti-sense primer); for HPRT, 5'-AAT TAT GGA CAG GAC TGA ACG T-3'
(sense primer) and 5'-TCA AAT CCA ACA AAG TCT GGC TTA-3'
(anti-sense primer). HPRT was visualized on ethidium bromide
stained gels. PCR products of pT and RAG-1 were Southern blotted to
Nytran paper (Schleicher & Schuell, Keene, NH). The membrane was
hybridized at 60°C using a 32P-end-labeled internal
oligonucleotide, for pT 5'-CCT TCT CTG GCC CCA CCA ATC A-3' and for
RAG-1 5'-CAT CCT GTG ACA TCT GCA AC-3'.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We have already reported that during hanging drop approximately
1000 to 2000 progenitors cells, i.e., 1/5th to 1/10th of
the input number of cells, enter the thymic lobe (12) and that
afterward the number of human cells initially decreases (<500
cells/lobe) (data not shown) before expanding. As a consequence of this
transient decrease, the number and the frequency of human cells during
the first days of the culture is low. When we electronically gated out
the dead as well as the murine cells by eliminating the cells that
stained with PI and anti-mouse CD45-Tri-Color, a population
remained that did not stain for human CD45 (Fig. 1
). When anti-mouse class I MHC
biotin-streptavidin-Tri-Color was added, we obtained a neater picture
as nearly all cells that were gated out electronically now stain for
human CD45. Most likely, this CD45- population, which can
be stained by anti-mouse class I MHC, represents a population of
mouse epithelial cells. By gating out all nonhuman cells, we could
carefully trace the human cells and study their phenotypic changes at
the early stage of differentiation, even when the total cell number is
very minute. Other systems, such as injection of mismatched HLA donor
cells into transplanted SCID-human mice, do not offer the
possibility to clearly gate out donor cells, because the Ag
density of the HLA-marker is relatively weak during the first stages of
differentiation (16). Therefore, the hybrid human/mouse FTOC system
used in our study gives us a unique opportunity to address the earliest
steps of differentiation.
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A shows the cell surface
expression of CD34, CD7, and CD38 and the cytoplasmic expression of CD3
(cyCD3) in function of duration of the culture. According the
purification strategy, the sorted precursor cells were CD7- and
CD38-negative (Fig. 2A) at the start of the culture. After
three days of culture, most of the CD34+ cells already
expressed CD7 and CD38, 70 and 85%, respectively. Very rapidly some
cells coexpress CD34 cells with cyCD3 upon culture. However, an
important fraction of the cells at that time has down-regulated CD34
and does not express cyCD3. After 4 to 5 days of culture, 
1000 to
2000 human cells can be found in one reconstituted thymic lobe. An
important fraction of the human cells expresses CD4 (30–50%, 300–900
cells/lobe) but does not express cyCD3 nor CD7 (Fig. 2B). At
that time, cyCD3+ cells are
CD4-CD7+ (Fig. 2B) and the
expression of CD4 and CD7 is mutually exclusive (data not shown). After
7 days of culture, the number of human cells has increased to
3000–4000/lobe, part of the cyCD3+ cells starts to express
CD4, but almost all the cyCD3- cells remain
CD7-. From this we conclude that there is a first wave of
two cell populations that are either
CD4-cyCD3+CD7+ or
CD4+cyCD3-CD7- followed by the
generation of a second wave of cells that are
CD4+cyCD3+CD7+. As discussed below,
further phenotypic analysis allowed us to delineate the differentiation
of four different cell lineages: dendritic cells, NK cells, TCR-,
and TCR-ß cells.
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After 3 days of culture, some human cells had down-regulated CD34
without cyCD3 expression (Fig. 2
A). We analyzed whether
these cells might represent dendritic cells. Because Sotzik et al. (17)
have shown that human thymic dendritic cells are large cells expressing
high levels of CD4 and class I and II MHC but no CD7, we analyzed these
markers at different time points of FTOC. As seen in Fig. 3A, at day 4 the
CD4+ cells express HLA-DR. Later, the intensity of both CD4
and HLA-DR increases, so that after 11 days a small but distinct
population of HLA-DR++CD4+ cells can be
discerned. The number of these cells is 200–500 cells/lobe at day
4–6 and increases only two- to threefold until day 12; afterward, the
number remains constant. As the total number of human thymocytes
increases 10- to 20-fold (20,000–40,000 human cells/lobe), the
frequency of these putative dendritic cells decreases in function of
length of culture (Fig. 3A). After 12 days of FTOC, it is
possible to define cells that highly express CD4, HLA-DR, and HLA-class
I are negative for CD7 and cyCD3 (Fig. 3B) and have a high
forward and side scatter (Fig. 3B, R6). We
further corroborated that these cells were functional dendritic cells
by assessing their ability to stimulate allogeneic T cells in an MLC.
The maximal extent of proliferation of responder lymphocytes was
similar to that seen with PBL as stimulators, but 80-fold fewer
stimulator cells were required for peak stimulation (Fig. 3C).
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, we purified these populations (which
were >96% pure upon reanalysis). It is clear that dendritic cells and
CD4+ precursor cells were generated starting from the
CD4- population, whereas the
CD4+HLA-DR- population has lost its potential
to develop into dendritic cells. Finally, the sorted CD4+
HLA-DR+ cells were end-stage cells that retained the
dendritic phenotype and were not able to give rise to T cells (data not
shown). These results demonstrate that the lymphoid/dendritic common
precursor is CD4-, whereas the CD4+
(HLA-DR-) cell populations contain no dendritic
precursors. Taken together, these results indicate that during
differentiation, dendritic cells down-regulate CD34, express HLA-DR and
CD4 and do not express cyCD3 and CD7.
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As only a small number of NK cells develop spontaneously in FTOC
(Fig. 5), we took advantage of the
property of IL-15 as a strong stimulator of NK cell development (18, 19) to study whether different thymic subsets were able to generate NK
cells. Starting with
CD34+Lin-CD38- cells, we were not
able to generate NK cells with IL-15 in cell suspension (data not
shown). However, when put into FTOC and cultured in the presence of
IL-15, large numbers of CD3-CD56+ cells were
generated. Fig. 5 shows that the addition of 20 ng/ml IL-15 to FTOC
resulted in a dramatic increase in CD56+ cells. These cells
had the appearance of large granular lymphocytes (data not shown).
Further phenotypic analysis showed that part of the cells coexpresses
CD16 (Fig. 5). In addition, we performed sorting experiments to obtain
CD4+HLA-DR- and CD4- subsets from
day 10 FTOC, and both subsets were able to generate NK cells when
reintroduced in FTOC in the presence of IL-15 (Fig. 6). However, the starting
CD34+CD38-Lin- population, as
well as CD4+HLA-DR- cells from day 10 FTOC,
were not able to generate NK cells in suspension cultures with IL-15.
In contrast, CD4- cells from day 10 FTOC generated NK
cells when cultured in suspension in the presence of IL-15 (data not
shown). This apparent paradox might be ascribed to a difference in
sensitivity of the assays and that a small contamination of the
CD4- cells in the CD4+ cells are able to
generate NK cells in FTOC, whereas this does not occur in suspension.
In this respect, the purity of the sorted cell populations was at least
96%, and the input of cells was 5000. Therefore, the absolute number
of cells with the NK phenotype CD56+CD3-
recovered per lobe, which is 8,000–12,000 after 10 days of culture,
does not rule out the possibility of outgrowth of contaminating cells.
Alternatively, the fact that with IL-15 CD4+ cells can
generate NK cells in FTOC and not in suspension, whereas
CD4- cells generate NK cells in FOTC and in suspension,
can be interpreted as the CD4- populations contain more
mature NK precursor cells (day 10). Taken together, these results
indicate that up-regulation of the CD4 marker on HLA-DR-
precursor cells is accompanied with a loss of differentiation potential
for dendritic cells, but that NK and T potential is still preserved.
Therefore, these results suggest that the branching point of dendritic
cells is earlier in the developmental scheme than the T/NK cell
branching point.
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We have previously shown that 28 days of FTOC started with 10,000
CD34+ fetal liver cells resulted in the appearance of
CD4+CD8-CD3+CD1+ human
thymocytes (12). Here we show that after 40 days of culture
CD1-TCR-ß+ and
CD1-TCR+ thymocytes were generated by
CD34+CD38- fetal liver cells and that the
thymocytes had partially down-regulated CD45RO and acquired CD45RA
(Fig. 7). At that time point of the
culture, 50,000–150,000 human cells are found per lobe. In addition,
part of the CD3+CD1-TCR-ß+
cells were either CD4+CD8- or
CD4-CD8+ (data not shown). We have shown
previously that CD1-CD3+CD4+ or
CD1-CD3+CD8+ cells have acquired
functional capacity and are considered as complete differentiated
thymocytes (20). In addition, we have also demonstrated that
CD1-TCR-+ cells have functional
characteristics of mature T cells (21).
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To assess the immaturity of the starting population, we analyzed
the presence of RAG-1 and pT mRNA transcripts. Expression of RAG
genes is absolutely required for rearrangement of TCR genes, but is
also present in developing B cells as it is required for the
rearrangement of Igs. pT is restricted to cells of the T cell
lineage (22, 23). The expression of RAG-1 and pT mRNA has already
been addressed in the analysis of human thymic subsets (24). In Fig. 8, it is shown that pT is already
present in the CD34+CD38-Lin-
starting population and that the expression is higher after 3 days of
culture in FTOC. In contrast, although RAG-1 expression is sometimes
weakly present in the starting population, it is not detectable in day
+3 samples. RAG-1 mRNA expression could be clearly demonstrated in
human thymocytes after 11 and 14 days of FTOC.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, and TCR-ß T cells. This system allows a detailed
kinetic analysis of the different cell populations, because a cohort of
CD34+CD38- stem cells starts to develop
resembling a synchronized culture. This is in contrast to analysis of
different subsets in a human thymus, where all different cell
populations are already present.
Moreover, in this study we are able to address the
differentiation of pluripotent hemapoietic precursor cells from fetal
liver, whereas most data on human T cell differentiation are founded on
studies with postnatal thymus. However, the human/mouse FTOC method
that we have used has potential shortcomings. First, we have introduced
human pluripotent fetal liver cells with myeloid differentiation
capacity (data not shown) in FTOC. This might not represent the default
pathway, because it remains an open question whether an uncommitted
pluripotent cell seeds the thymus or a more committed cell. Secondly,
although the differentiation capacity of the fetal liver cells in the
xenogeneic environment is impressive, we cannot exclude that critical
factors or ligands that are species-specific are lacking and might
influence the default differentiation pathway. Nevertheless, our
experience with cord blood-derived hematopoietic precursors shows that
the differentiation obtained in in vitro xenogeneic FTOC (25) is very
similar to that obtained by precursor cells that were injected in vivo
in human thymic tissue transplanted under the kidney capsule of
SCID/human reconstituted mice (our unpublished observations).
Therefore, we consider that our approach gives us an unique opportunity
to study the T developmental pathway of prenatal fetal liver precursor
cells. If the human/mouse FTOC recapitulates the normal differentiation
of the human thymus, our results imply that immature
CD34+CD38- human stem cells develop very
rapidly into dendritic cells, that the differentiation toward NK cells
takes somewhat longer in time, and, finally, that TCR- cells are
generated before TCR-ß cells.
Dendritic cells comprise a family of professional APC responsible for the activation of primary T cell responses. Large numbers of functional dendritic/Langerhans cells can be generated by treating dendritic cell precursors from several sources with granulocyte-macrophage-CSF alone or in combination with other growth factors. More recently, it was also shown that ligation of CD40 induces the proliferation and differentiation of CD34 hematopoietic stem cells into dendritic cells (26). Sotzik et al. (17) have shown that dendritic cells in the human thymus can be identified as CD4highCD44+CD1-CD2-CD3-CD7-CD8-CD34- class I MHChigh class II MHChigh cells with efficient stimulator capacity for allogeneic T cells. We show here that dendritic cells are generated spontaneously within the thymus microenvironment from CD34+CD38- progenitor cells within 8–10 days. These cells are the earliest cells to be generated and to be functionally mature. The functional relevance of this population within the thymus remains to be established. There is no doubt from mouse studies that dendritic cells are essential for the negative selection of T cells, whereas epithelial cells mediate positive selection (27). However, at present it is not clear whether dendritic cells in the thymus originate from precursor cells that differentiate in the thymus or whether mature dendritic cells migrate from the periphery to the thymus. This is of interest because in the latter situation, Ags that are loaded in the periphery can participate in the negative selection process, whereas otherwise these Ags have to be produced locally or have to reach the thymus. Our findings show that dendritic cells are able to differentiate in the thymus from precursor cells and that the kinetics are faster than for the other cell types. This allows that negative selection can be mediated very rapidly when the T cells are at a very immature stage and supports the assumption that a linked development of T cells and dendritic cells may ensure that developing T cells are negatively selected predominantly by self-Ags presented on newly formed thymic dendritic cells. This idea was already proposed for the murine thymus, as it was shown that after irradiation bone marrow cells injected intrathymically restore simultaneously T cells and dendritic cells (28, 29).
Another cell type that develops spontaneously albeit in low
numbers in human/mouse FTOC are NK cells (Fig. 5). It is obvious that
most NK cells are generated outside the thymus as nude mice have a
normal NK activity. However, the thymus provides a microenvironment
that allows stem cells to differentiate toward the NK cell pathway.
This is shown by the fact that the CD34+Lin-
population from fetal liver did not differentiate toward NK cells in
cell suspension when IL-15 was added, even in the presence of stem cell
factor and IL-7 (data not shown), whereas in FTOC IL-15 resulted
in a massive differentiation toward NK cells. Similarly,
CD4+ and CD4- precursor cells from day 10 FTOC
also had the capacity to develop into NK cells in FTOC in presence of
IL-15. However, only day 10 FTOC CD4- precursor cells, in
contrast to the CD34+CD38- start population,
were able to develop into NK cells in the presence of IL-15 in
suspension cultures. This means that the stromal environment and/or
cytokines present in the thymus can allow the differentiation of the
IL-15 refractory CD34 precursor cell to a cell population that
responds vigorously to IL-15 by cell proliferation and
differentiation toward NK cells. It is clear from our studies that
IL-15 is a potent promoter for the growth and differentiation of human
NK cells. We also found that part of the NK cells, generated in
IL-15-supplemented FTOC, coexpress CD16 (Fig. 5) and CD94 (data not
shown). We have already shown previously the role of IL-15 as NK cell
promoter in mouse fetal thymus (18). Our present observations point to
an essential role of IL-15 in human NK development and are in agreement
with the observation of Mingari et al. (30), who have been able to show
that IL-15 provides an efficient stimulus for differentiation of thymic
precursors toward phenotypically and functionally mature NK cells.
The precise role of NK development in the thymus remains unknown. Because the main physiological maturation of NK cells occurs in the bone marrow, it is possible that NK maturation in the thymus is not relevant. In particular, the fact that only CD94 is up-regulated and other HLA class-I specific inhibitory receptors are not expressed (data not shown, and 30 may support the hypothesis that this maturation is incomplete and not physiological.
The first T cells that develop in the mouse thymus are TCR-
cells. Indirect evidence that this is also the case for the human
thymus is based on studies on T cell acute lymphoblastic leukemia. As
CD3- T cell acute lymphoblastic leukemias have been
found with rearranged TCR- genes and germline TCR-ß and TCR-
genes (31), it is probable that TCR- genes rearrange early during T
cell development before other TCR genes. In addition, RT-PCR analysis
performed by Ktorza et al. (32) showed expression of full-length
TCR- transcripts in the CD34+CD1- TN
thymocyte subset, before expression of full length TCR-ß transcripts.
In the earliest phase of fetal thymus colonization at 8.2 wk of
gestation, by immunohistochemistry 5% of the thymocytes stained
intracellularly with ß-F1 mAb, suggesting expression of
TCR-ß-chain, whereas only a few TCR- cells were found. The time
of first appearance of TCR- is at 9.5 wk (33). In our study we do
not have direct proof that differentiating thymocytes start earlier
in their attempt to rearrange the TCR- genes than
TCR-ß genes. We consistently found that cells with cell surface
expression of TCR- are present before we can detect T cells with
TCR-ß. However, as it is known that TCR-ß cells first have to
express TCR-ß-chain with pT before completion of the
differentiation and expression of TCR-ß, the difference in time of
appearance of TCR- and TCR-ß cells might rather reflect a
different kinetic in maturation of the two cell types.
Our finding that RAG-1 and pT mRNA is present in the starting
CD34+CD38- fetal liver cell population differs
from the failure to detect expression of pT and RAG-1 in the
CD34+CD38- cell population of cord blood, as
reported by Blom et al. (34), and in the
CD34+CD38- population of fetal liver, as
reported by Jaleco et al. (19). In contrast, Ramiro et al. were able to
demonstrate pT mRNA in CD34+ progenitors in fetal liver,
cord blood, and adult bone marrow (24). This could be due to
differences in the sensitivity of the test or in the strategies used to
purify the progenitor cells. In this regard we used FITC-labeled Abs
directed against CD38 instead of PE-labeled CD38 reagents. As PE stains
the cells brighter, it is possible that cells with a weak expression of
CD38 were present in our starting population. It is still a matter of
debate whether an uncommitted pluripotent precursor cell seeds the
thymus and becomes T cell committed afterward or whether commitment
occurs extrathymically and committed cells enter the thymus to complete
their differentiation. Definitive proof awaits study of cells at the
clonal level, but our findings suggest that
CD34+CD38- fetal liver express RAG-1 and pT
mRNA reminiscent of committed T cells or that uncommitted cells could
start the transcription of a variety of genes. Single-cell PCR will
inform us on the frequency of these cells.
Proposed model for development of different lineages in human thymus
The kinetic study examining the acquisition of different lineage
markers in our in vitro system, with particular emphasis on the
earliest steps in differentiation, revealed the sequential expression
of different surface Ags, shown in Fig. 9.
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in the starting population is consistent with the concept of
T-commitment of stem cells before the entry in the thymus or could
indicate the presence of cells engaged in the extrathymic T cell
pathway. At present, there is no direct evidence to conclude that one
single progenitor cell in the thymus can differentiate into T, NK, and
dendritic cells or whether there are different progenitor cells. Our
observation that RAG-1 mRNA expression is absent after 3 days of FTOC
is compatible with the view that the precursor cells with RAG-1 mRNA
transcripts are engaged in B cell differentiation and are not able to
develop in FTOC.
It is clear that the first Ags that appear on the cell membrane
of differentiating CD34+CD38-
Lin- are CD38 and CD7 (Fig. 2). These markers are not
lineage specific and are considered as activation markers. However, CD7
is not expressed by all cells, and we find evidence that strong
expression of CD7 never occurs during differentiation of dendritic
cells. This assumption is based on the fact that cyCD3 is very rapidly
expressed within 3 days of culture on part of the CD34+
cells and that this cyCD3 expression is almost exclusively found on
cells with a strong CD7 expression. Cells with a lower CD7 expression
might be precursor cells of cyCD3-positive cells. However, we cannot
exclude that part of these cells express CD7 transiently and will never
express cyCD3 and are cells at the early stages of the dendritic cell
development. In this respect, we have been able to generate dendritic
cells in FTOC starting from sorted CD34+CD7low+
cells from fetal liver (data not shown). It is generally considered
that expression of cyCD3 is an early marker for both NK and T cell
commitment. As CD7 expression correlates with cyCD3 expression, CD7, in
absence of membrane CD3 and CD4 expression, is an early T/NK marker in
the thymus. The next Ag that is expressed is CD4. It is obvious, as
shown in Fig. 3, that CD4 expression occurs in the absence of cyCD3 in
these cells that will become dendritic cells. HLA-DR, which is already
present from the start of the culture, is up-regulated during
differentiation toward dendritic cells. We could never demonstrate CD5
on these cells, whereas a small fraction of these cells expresses CD1a
(data not shown). Therefore, we can conclude that during
differentiation toward dendritic cells, progenitor cells down-regulate
CD34, possibly transiently express low levels of CD7, up-regulate
HLA-DR and CD4, and never express cyCD3.
The next CD7+ population that can be detected expresses cyCD3, which is indicative for cells engaged in the T/NK cell pathway. At that time, CD4 is not expressed. When CD4 is expressed, HLA-DR is down-regulated (data not shown). Resort experiments have shown that these cells have lost the potential to generate dendritic cells, whereas they can generate both NK and T cells.
These data indicate that in humans the T/NK split occurs downstream of the NK/dendritic cell split. These data are compatible with the finding in human bipotential T/NK cells that have been demonstrated by Sanchez et al. (11). Others have shown that the human thymus contain CD34+ intrathymic precursors that develop into T and dendritic cells (35, 36). In our study, there is no evidence that dendritic T cells are tightly linked to T cell development beyond a common precursor that is multipotent. This is in contrast to the observation in the mouse system, where a common T/dendritic cell progenitor was found (28). This could be due to species differences, difference between adult and fetal precursors, or to our model where an early precursor with myeloid potential is introduced. However, postnatal thymocytes sorted according to phenotypic differentiation markers and tested for their differentiation capacity in FTOC are consistent with the notion that dendritic cells branch off earlier that the T/NK bipotential cell progenitors. The CD34+CD1- subset, which is considered as the most immature thymic progenitor, gives rise to HLA-DR+CD4+CyCD3-, putative dendritic cells, T cells, and in the presence of IL-15 to cells with the CD56+ NK cell phenotype (data not shown). On the contrary, CD34+CD1+ thymic progenitors, considered to be at a later step of differentiation than the CD34+CD1- cells, develop into T cells but not into cells with the dendritic phenotype. In the presence of IL-15, the CD34+CD1+ cells develop into CD56+ cells (data not shown).
From day 6–7 of FTOC, CD5 expression appears on human thymocytes (data not shown). On day 6, besides the dendritic cell population that expresses CD4 and lacks cyCD3, we find a cyCD3-positive population that is still CD4-negative but already expresses CD5. This indicates that the expression of CD5 either precedes CD4 expression or that CD4-negative cells acquire CD5 and could be precursors of NK cells, whereas T-committed cells first express CD4 and later acquire CD5. However, as NK cells can be generated from CD4-positive cells, we assume that at least part of the NK cells are derived from a precursor with dim CD4 expression. Based on the findings of Sanchez et al. (11), who showed that CD5low cells are able to generate NK cells, we propose also an intermediate CD5-positive stage.
Finally, for the development of TCR-ß and TCR- cells,
our model is based on our ability to show the intracellular presence of
the ß-chain of the TCR after 10 days of culture. At that time, the
cells did not stain for TCR-ß on the cell surface. The first TCR
that could be demonstrated on the cell surface was TCR- on day
11–14, whereas on day 17–20 TCR-ß cells appeared (data not shown
and 12 . Finally, we find mature TCR- CD1- and
TCR-ß CD1-
CD4+CD45RA+CD45RO- cells after
14–16 and 35–40 days, respectively. This indicates that it takes
longer for TCR-ß cells to achieve full maturation end-stages as
compared with TCR- cells. This is could be due to a more
elaborated selection process. These results show that our experimental
model is able to support full maturation of the different cell
populations. Our model was based on kinetic studies compiling at least
25 different cultures, which gave a consistent picture. Moreover,
several experiments starting from CD34+CD38-
populations from postnatal thymus gave the same kinetics and sequential
order of phenotypes (data not shown). These findings strongly support
that the hybrid mouse-human FTOC is a reliable model system for human
thymocyte differentiation.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Prof. Jean Plum, University of Ghent, University Hospital, Department of Clinical Chemistry, Microbiology and Immunology, 4 Blok A, De Pintelaan 185, B-9000 Ghent, Belgium. E-mail address:
3 Abbreviations used in this paper: cy, cytoplasmatic; DP, double positive; FTOC, fetal thymus organ culture; PT, pre-TCR -chain; RAG, recombination activating gene; SP, single positive; PI, propidium iodide; HPRT, hypoxanthin phosphoribosyltransferase.
Received for publication May 22, 1998. Accepted for publication September 1, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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cells in the human thymus. J. Immunol. 158:4634.[Abstract]
gene in development of ß but not T cells. Nature 375:795.[Medline]
-TCRß) gene expression during human thymic development. J. Exp. Med. 184:519.This article has been cited by other articles:
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  M. Garcia-Peydro, V. G. de Yebenes, and M. L. Toribio Sustained Notch1 signaling instructs the earliest human intrathymic precursors to adopt a {gamma}{delta} T-cell fate in fetal thymus organ culture Blood, October 1, 2003; 102(7): 2444 - 2451. [Abstract] [Full Text] [PDF]
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 M. De Smedt, K. Reynvoet, T. Kerre, T. Taghon, B. Verhasselt, B. Vandekerckhove, G. Leclercq, and J. Plum Active Form of Notch Imposes T Cell Fate in Human Progenitor Cells J. Immunol., September 15, 2002; 169(6): 3021 - 3029. [Abstract] [Full Text] [PDF]
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M. O. Muench and A. Barcena Broad Distribution of Colony-Forming Cells with Erythroid, Myeloid, Dendritic Cell, and NK Cell Potential Among CD34++ Fetal Liver Cells J. Immunol., November 1, 2001; 167(9): 4902 - 4909. [Abstract] [Full Text] [PDF]
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Q.-L. Hao, J. Zhu, M. A. Price, K. J. Payne, L. W. Barsky, and G. M. Crooks Identification of a novel, human multilymphoid progenitor in cord blood Blood, June 15, 2001; 97(12): 3683 - 3690. [Abstract] [Full Text] [PDF]
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