HEMATOPOIESIS

Negative regulation of primitive hematopoiesis by the FGF signaling pathway
Fumie Nakazawa, Hiroki Nagai, Masahiro Shin, and Guojun Sheng

Hematopoiesis is controlled by multiple
signaling molecules during embryonicand postnatal development. The functionof the fibroblast growth factor (FGF) pathway in this process is unclear. Here weshow that FGF plays a key role in theregulation of primitive hematopoiesis inchicks. Using hemoglobin mRNA expression as a sensitive marker, we demonstrate that timing of blood differentiation

can be separated from that of initial mesoderm patterning and subsequent migration. High FGF activity inhibits primitive
blood differentiation and promotes endothelial cell fate. Conversely, inhibition of
FGFR activity leads to ectopic blood formation and down-regulation of endothelial markers. Expression and functional
analyses indicate that FGFR2 is the key
receptor mediating these effects. The FGF

pathway regulates primitive hematopoiesis by modulating Gata1 expression level
and activity. We propose that the FGF
pathway mediates repression of globin
gene expression and that its removal isessential before terminal differentiation
can occur. (Blood. 2006;108:3335-3343)© 2006 by The American Society of Hematology

Introduction
During the early development of amniotes, blood cells are generated exclusively from the extraembryonic yolk sac mesoderm.1-3
Intraembryonic hematopoietic niches such as aorta-gonad-mesonephrosand liver mature later and initiate the second wave of hematopoiesis.4-6 Located in the extreme lateral regions of the developing
embryo, primitive blood cells are derived from ventral mesoderm
precursors ingressing through the posterior part of the primitive
streak during gastrulation. In birds, postingression precursor cells
undergo extensive migration before the establishment of thecirculation to populate the entire hemogenic region called areavasculosa.3,7-9 Endothelial cells have a similar origin and gastrulate
through the posterior streak. Molecular, genetic, and cell biologic
analyses have demonstrated that precursor cells in the streak andduring early migration still have the potential to differentiate intoeither the blood or the endothelial lineage.10-15 Extraembryonicmesoderm (EEM) contains at least 2 additional tissue types,
somatic and splanchnic EEM, which differentiate into smoothmuscle and connective tissue and constitute dorsal and ventral
linings of the extraembryonic coelom.16,17 Blood and endothelial
cells separate early from the splanchnic mesoderm as aggregates on
its ventral side, variously called blood islands, angioblasts, orhemangioblasts.18-20 These aggregates are referred to as blood
islands in this work. In lateral EEM, blood islands (BIs) give rise toboth blood and endothelial cells, whereas in medial EEM and thelateral plate, only endothelial cells are generated.18,21,22 It is notwell understood why some BIs give rise to both, whereas others
give rise to only endothelial cells.
Chicken hemoglobins were used for initial descriptive andbiochemical studies on primitive hematopoiesis21,23 and for later
investigations on transcriptional control.24-26 More recent studies
using different model organisms revealed a complex genetic

network controlling primitive hematopoiesis.3,10,13,15,27-29 Analyses
focusing on signaling molecules indicated the involvement of the
BMP,30-32 VEGF,33-36 WNT,37,38 and Notch39,40 pathways in various
aspects of primitive hematopoiesis. The role of the fibroblast
growth factor (FGF) pathway in this process is unclear. It is known
to be important for mesoderm induction during gastrulation,41-43 fordorsoventral patterning of the mesoderm,44,45 and for mesoderm
cell movement during and after gastrulation.46,47 In addition, FGF
can act as a potent inducer of neovascularization and neoangiogenesis48,49 and plays a role in promoting proliferation and maintenance of hematopoietic progenitors and stem cells.50 Its precise
function in primitive hematopoiesis has been debated, with someevidence suggesting a positive regulatory role51-53 and otherevidence indicating a negative one.54,55
In this study, we investigate the function of the FGF pathway
in chick primitive hematopoiesis. Globin mRNA expression isused as a sensitive readout for terminal differentiation. We focus
our study on the differentiation step only, separating it frompotential roles of the FGF pathway in earlier (induction,
patterning, and migration) or later (proliferation and maturation)
steps. We demonstrate that FGF signals can elicit strong
inhibition of hemoglobin expression and up-regulate endothelial
markers. Inhibition of the FGF signaling pathway results in
ectopic blood formation and completely inhibits endothelial
marker expression. Among 4 known FGFRs, we show that
FGFR2 is the key receptor functioning in this process. These
effects are achieved through crosstalk with the VEGF pathway
yet can be attributed primarily to the FGF pathway. We also
show that it acts in primitive hematopoiesis through Gata1 bymodulating Gata1 expression level and activity.

From the Laboratory for Early Embryogenesis, RIKEN Center for
Developmental Biology, Kobe, Hyogo, Japan.

F.N. and H.N. contributed equally to this study.

Submitted May 5, 2006; accepted July 10, 2006. Prepublished online as BloodFirst Edition Paper, August 3, 2006; DOI 10.1182/blood-2006-05-021386.
The authors declare no competing financial interests.
G.S. designed the experiments; F.N., H.N., M.S., and G.S. performed theexperiments; G.S., F.N., H.N., and M.S. analyzed the data; and G.S. wrote thepaper.

BLOOD, 15 NOVEMBER 2006 䡠 VOLUME 108, NUMBER 10

Reprints: Guojun Sheng, Laboratory for Early Embryogenesis, RIKEN Center
for Developmental Biology, Kobe, Hyogo 650-0047, Japan; e-mail:sheng@cdb.riken.jp.
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 USC section 1734.
© 2006 by The American Society of Hematology

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NAKAZAWA et al

Materials and methods
Chick embryology
Fertilized hens’ eggs were purchased from Shiroyama Farm (Kanagawa,Japan) and incubated at 38.5°C to the desired stages. For tissue graft, beadgraft, chemical treatment, and electroporation experiments, we used themodified New culture method.68 Differentiation of isolated streak tissue
pieces was performed on vitelline membrane as in New culture but withouta host embryo. For in situ hybridization and immunohistochemistry,
standard published protocols were followed.56 All in situ analyses werecarried out with either digoxigenin- or fluorescein-labeled RNA probes anddeveloped with alkaline phosphatase staining. All tissue sections shown
were 10-␮m paraffin sections. Electroporation was performed with Intracel
electroporator (Intracel, Hertfordshire, United Kingdom) at the following
setting: 5 volts, 3 pulses (50 ms with 250-ms interval). All imageacquisition equipment was purchased from Olympus (Tokyo, Japan).
Whole-mount embryos were visualized in PBS solution with an Olympus
SZX12 microscope using a DF PLAPO 1.2⫻/0.133 PF2 objective lens.
Embryo sections were mounted with canada balsam and visualized with an
Olympus BX51 microscope using Olympus UPlanSApo objective lenses(20⫻/0.75; 40⫻/0.9; 100⫻/1.40 oil-immersion). For both whole-mountembryos and embryo sections, an Olympus DP70 camera was used tocapture images, and DP Controller 2.1 image software was used to acquirethem. Final images were assembled with Photoshop 8.0 (Adobe Systems,
San Jose, CA).
ChemicalsProteins and chemical inhibitors were purchased from commercial sources.
The following concentrations were used for bead soaking prior to graft: BMP2
(20 ␮g/mL, 355-BEC); BMP4 (20 ␮g/mL, 314-BP); Chordin (100 ␮g/mL,
758-CN); Noggin (12 ␮g/mL, 1967-NG); FGF4 (50 ␮g/mL, 235-F4); Nodal(12 ␮g/mL, 1315-ND); Dkk1 (25 ␮g/mL, 1096-DK); VEGF165 (10 ␮g/mL,
93-VE); Wnt3a (10 ␮g/mL, 1324-WN); Epo (25 ␮g/mL, 959-ME) (all fromR&D Systems, Minneapolis, MN); FGF8 (50 ␮g/mL; Sigma, St Louis,
MO; no. F6926); SU5402 (42 mM; Calbiochem, San Diego, CA; no. 572630).
For chemical treatment, embryos were cultured with SU5402 (85 ␮M, 6 ⫻ 50%
inhibitory concentration [IC50]) or SU5416 (10 ␮M, 10 ⫻ IC50, Calbiochem;no. 676487) dissolved in albumen, with less than 0.2% of final DMSO
presence.
Molecular biology
DNA fragments for gene-specific in situ probes were generated bypolymerase chain reaction (PCR), cloned in pGEM-T vectors, and confirmed by sequencing (␳: nucleotides [nt’s] 31-465 of CHEST324a16; ⑀ nt’s
55-475 of CHEST26i6; ␣␲: nt’s 39-488 of CHEST23l23; ␣A nt’s 40-485 of
CHEST973g24; gata1: nt’s 577-1061 of NM_205464 with full-length

cDNA kindly given by T. Evans (Albert Einstein College of Medicine, New
York, NY); ets1: nt’s 41-782 of 052594.1; lmo2: nt’s 7-962 of NM204271;
vegfr2: nt’s 3477-5149 of AY382882; Vecad: nt’s 642-2368 of AF522067;
fgfr1-4: Shin et al57). Constitutively active FGFR2 (CA-FGFR2) was constructedbased on studies by Webster and Donoghue.58 A chick FGFR2 fragment lacking
the extracellular and transmembrane domains was created by PCR (forward:
5⬘-ATCGATATGGGGAGCAGCAAGCCCAAGGACCCCAGCCAGCGCCCTGACTTCAGCAGCCAGCCCGCTGTC-3⬘, and reverse: 5⬘-ACTAGTTCATGTTTTAACGCTCCCATTC-3⬘, with myristoylation sequence from c-Src:
MGSSKSKPKDPSQR tagged to the 5⬘ end of the forward primer). An
additional Lys661Glu mutation was introduced to create ligandindependent kinase activity. Final CA-FGFR2 was cloned into pCAGGS
vector for efficient expression in vivo.59 The activity was confirmed bydouble-phosphorylated ERK (dp-ERK) antibody staining. FGFR2-specificfluorescein-tagged morpholino (5⬘-CTAGAATGATTTACCTTCGGGTTCC-3⬘) was designed against exon1-intron1 boundary and purchasedfrom Gene Tools (Philomath, OR) together with standard control morpholino. Chick FGFR2 contains a 14.3-kb first intron, and blocking the splicing
of first exon led to severe reduction of the mature transcript as revealed byin situ analysis with c-terminal–specific probe (Figure 5R-S). Morpholinos
were electroporated at 3 mM and DNA constructs at 1 ␮g/␮L.

Results
Expression of hemoglobin genes during chickprimitive hematopoiesis

Based on pseudoperoxidase activity of the heme group, chemicals
such as benzidine and diaminofluorene have traditionally been used
for staining differentiated red blood cells.8,60,61 As an alternativeand more sensitive and specific method, we investigated mRNA
expression patterns of hemoglobin genes during early chickdevelopment. Chick hemoglobin loci contain 4 beta-type (␤H, ␤A,␳, and ⑀) and 3 alpha-type (␣␲, ␣A, and ␣D) globin genes. Five ofthem (␣␲, ␣A, ␣D, ␳, and ⑀) are expressed in primitive blood
cells.21,23 We generated probes for ␣A, ␣␲, ␳, and ⑀ and performed insitu analysis with precirculation chick embryos (before Hamburger
and Hamilton Stage 13 [HH13]). All 4 genes were strongly
expressed in area vasculosa. Figure 1 shows the expression patterns
at HH10 of ␣␲ (A), ␣A (B), ␳ (C), and ⑀ (D), respectively. As
revealed in sections (Figure 1E-H), each gene was expressed in all
differentiated blood cells. Since all 4 genes were detected at theearly somite stage, we performed detailed analysis on hemoglobin␳ (␳) expression (Figure 1I-M). The earliest expression was seen at
HH6-7. At HH7, ␳ can be readily detected in differentiating BIs(Figure 1J,N-P). Within individual BIs, a few inner cells were

Figure 1. Hemoglobin expression during early development. (A-H) Expression at HH10 of ␣␲ (A), ␣A (B), ␳(C), and ⑀ (D) globin transcripts and sections throughindicated level of ␣␲ (E), ␣A (F), ␳ (G), and ⑀ (H). (I-W)
Expression of ␳ at HH5 (I), 7 (J), 8 (K), 10 (L), and 12 (M).
Sections of HH7 (N-P) and HH8 (Q-S) embryos areshown as indicated. Arrows point to cells initiating expression within a BI. Representative BI morphology is shownfor HH9 (T), 10 (U), 11 (V), and 12 (W).

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observed to initiate ␳ expression (Figure 1O), followed by morecells within a cluster (Figure 1P-S). Mature and laterally locatedBIs from HH8 onward contained all inner cells positive for ␳ (␳⫹)
surrounded by negative cells (Figure 1S-W). Morphologicallydistinct blood cells were visible from HH10 (Figure 1L-M,U-W).
At the onset of circulation (HH12-13; Figure 1M,W), the majorityof blood cells had lost contact with each other and with theendothelial wall, which was a prominent feature for ␳⫹ cells up to
HH10-11 (Figure 1U-V). The detection sensitivity with globin
mRNA probes (presomite stage) is greater than with heme group–
reacting compounds (4-5 somite stage), in agreement with biochemical studies on chicken globin proteins23,62,63 and mouse globin
genes.64 Using ␳ as a sensitive marker, we performed further
experiments to investigate the regulation of primitive hematopoiesis.
Posterior streak mesoderm can differentiate into ␳-expressing
blood cells in the absence of cell migration

Explant culture showed streak tissue, and dissociated streak cells
are able to differentiate into blood cells in mice.14 In chicks,
dissociated posterior streak and early migratory mesoderm cells
can form blood colonies in culture.35 This was confirmed by streaktissue graft using ␳ as a marker (Figure 2A-D). We separated HH4
streak into 3 pieces (anterior, middle, and posterior; Figure 2A),
transplanted each piece to the extreme lateral region of the area
opaca of same-stage host embryos, and cultured to HH10. The
posterior streak piece can differentiate into ␳⫹ cells with developmental timing similar to that of the host embryo (Figure 2D). Boththe intensity of ␳ expression and the spatial distribution of BIs werecomparable to the host (Figure 2D). The middle piece can also
differentiate, but to a lesser degree as predicted from fate map
studies, into ␳⫹ cells (Figure 2C). The anterior piece failed togenerate any ␳⫹ cell (Figure 2B) and instead differentiated into
more axial structures with notochord and somites (data not shown).
These data suggested that posterior streak mesoderm precursors
contain intrinsic ability to undergo proper migration and differentiation. Since the host area opaca can provide a substrate for migration

FGF PATHWAY IN PRIMITIVE HEMATOPOIESIS

3337

and possible inducing signals from the extraembryonic ectoderm
and endoderm, we cultured posterior pieces directly onto thevitelline membrane (Figure 2E). The posterior half of the HH4
streak was further divided into anterior (third quarter) and posterior(fourth quarter) pieces. Explants were cultured until control
embryo reached HH10 (Figure 2F). The ␳⫹ cells can be seen in boththe third and the fourth quarters (Figure 2G-H), with more ␳⫹ cells
and stronger expression in the fourth quarter (Figure 2G). In
vitelline membrane culture, the cells failed to migrate and as aconsequence the characteristic spatial distribution of BIs was lost.
Differentiation of primitive blood cells is inhibitedby FGF signaling

The previous experiment suggested that signals regulating precursor cell migration can be separated from those regulating differentiation. In chicks, these 2 processes can also be separated temporally. Differentiation of primitive blood as marked by ␳ expression
is initiated at about HH7 (Figure 1J). Migration of EEM cells starts
before HH4 and the majority of them reach their destinations by
HH7. We focused on the step of hemoglobin gene transcription as amarker for the initiation of terminal differentiation. We first lookedat the ability of molecules involved in various signaling pathways
to affect the endogenous ␳ expression. Beads soaked with secreted
molecules were grafted either within or outside the future ␳ domain(Figure 3A). HH6-7 embryos were used to minimize possible
secondary effects due to their influence on cell migration. Results
are summarized in Table 1. We found no single factor capable ofinducing ␳ ectopically, whereas only FGF beads (FGF4, 40/53,
75%; FGF8, 15/29, 52%) showed remarkable inhibition of endogenous ␳ expression. As shown in Figure 3B and D, control beads had
no effect on ␳ expression, whereas FGF4 beads exerted strong
inhibition (Figure 3C,E). A similar effect was seen with FGF8
beads (Figure 3G). The inhibitory effect of FGFs was not due totheir ability to repel mesoderm cells, as sections revealed large
numbers of ␳⫺ mesoderm cells near beads (Figure 3F). As
summarized in Table 1 and shown in Figure 3H (BMP7) and Figure
3I (Noggin), we did not observe a similar prominent effect withother factors tested. We then asked whether the cells that failed toadopt blood fate can differentiate into endothelial cells. The effectof FGF bead graft was analyzed with 2 endothelial markers, Vegfr2
and Ets1. Both have been described to specifically mark endothelial
cells at HH10.65,66 We generated RNA probes for Vegfr2 and Ets1
and extended the expression study by focusing on EEM differentiation (Figure 3J-L, Vegfr2; Figure 3O-Q, Ets1). When FGF4 beads
were grafted from HH6-7 and analyzed at HH10, strong upregulation of both transcripts was observed in cells adjacent tografted beads (Figure 3M-N,R-S).
Inhibition of the FGF signaling pathway leads to ectopic blood
cell differentiation

Figure 2. Differentiation potential of posterior streak mesoderm. (A-D) HH4
streak pieces are grafted to the lateral area opaca of host HH4 embryo and cultured
to HH10 (A). Differentiation is assayed by ␳ expression for anterior (B), middle (C),and posterior (D) pieces. Red dotted lines indicate the limit of host ␳ expression. (E-H)
HH4 streak pieces are cultured on vitelline membrane (E). Differentiation is assayedby ␳ when control reaches HH10 (F) for posterior (G) and third (H) quarters.

Our graft analysis suggested that FGF mediates the choice between
blood and endothelial fates. Activation of FGF signaling inhibits
blood differentiation and promotes endothelial formation. We thenasked whether more blood cells can be generated when thispathway is inhibited. The ␳ expression was analyzed in embryos
treated with SU5402, a small molecule inhibitor specific for FGFR
class RTK proteins.67 We applied SU5402 starting from HH6-7,
when the induction and extensive migration of mesoderm cells haslargely been accomplished. Beads soaked in control DMSO had noeffect (Figure 4A,C). Grafting of SU5402 beads resulted in large
numbers of blood cells in their vicinity (Figure 4B,D-E). In
particular, we saw blood cells being induced in medial regions

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NAKAZAWA et al(compare Figure 4Oi, Pi, and Qi with Figure 4Oii, Pii, and Qii),
indicating that endothelial development was greatly compromised
with the inhibition of FGFR activity. This was further confirmed byanalyzing the effect on Lmo2. Analysis of Lmo2 RNA expression(Figure 4Ri-iii) showed similar patterns as previously described.8
At HH10, Lmo2 was expressed in both blood and endothelial cells(Figure 4S,Ui-ii,Wi-ii) in control embryos. With SU5402 treatment, Lmo2 expression in blood cells was not affected, whereas
expression in endothelial cells was completely abolished (Figure
4T,Vi-ii,Xi-ii). In agreement with these observations, no vascular
network or blood flow was observed (data not shown) when treatedembryos were cultured until controls reached circulation stage(⬎ HH13). These data indicated that inhibition of the FGF pathway
severely blocked endothelial differentiation.
FGFR2 is a key negative regulator of primitive
blood differentiation

Figure 3. FGF inhibits ␳ and enhances endothelial marker expression. (A) HH6-7
embryos are grafted with factor-soaked beads either within (black) or outside (yellow)
the future ␳-expressing domain and cultured to HH10. (B-I) FGF4 beads inhibit ␳
expression (C,E) and control beads have no effect (B,D). Boxed areas in panels B
and C are magnified in panels D and E, respectively. Arrows point to grafted beads
and arrowheads show the repression. F (section of E) shows presence of ␳⫺ cells
around the bead. A similar effect is seen with FGF8 (G) but not BMP7 (H) or Noggin(I). (J-N) Expression of Vegfr2 at HH4 (J), 6 (K), and 10 (L). FGF4 beads enhance its
expression (M-N). Arrows point to grafted beads. (O-Q) Normal expression of Ets1 at
HH4 (O), 6 (P), and 10 (Q). FGF4 beads enhance its expression (R-S).

(Figure 4F), which was not observed in normal embryos or in graftexperiments with secreted factors (Table 1). This prompted us toinvestigate the effect of SU5402 in more detail.
To confirm this effect, we cultured whole embryos with SU5402
dissolved in the albumen in the modified New culture setting.68 As
mentioned earlier, we chose to test the effect with embryos from
HH6-7 onward. Earlier treatment resulted in malformations in theembryo proper as well as a greatly reduced area vasculosa, likely as
a consequence of mesoderm induction and migration defects (datanot shown). No obvious defect or delay in embryonic development
was seen with the concentration used (DMSO: Figure 4G,I;
SU5402: Figure 4H,J; Paraxis staining: I-J insets). Control-treatedembryos showed normal ␳ expression (Figure 4G,K). SU5402
caused dramatic medial expansion of the ␳-expressing domain(Figure 4H,L). Sections of treated embryos revealed ␳⫹ cellclusters below the splanchnic mesoderm in the lateral plate (Figure
4Ni-v), far more medial than in control embryos (Figure 4M). In
the endogenous hemogenic domain, the intensity of ␳ expression
and the pattern of BI distribution were not greatly affected (Figure
4G-H). In most cases, we saw ␳⫹ clusters still surrounded by ␳⫺cells with endothelial cell morphology (Figure 4Ni-v).
Since endothelial-like cells were still observed in treatedembryos, we next investigated whether FGF activity is required forendothelial marker expression. We compared the effect of SU5402
with DMSO control using endothelial markers Ets1, Vegfr2, andVecad. Expression of all 3 was potently inhibited by SU5402

SU5402 can inhibit tyrosine kinase activity of all FGF receptors.
We performed further experiments to investigate which FGF
receptor(s) may be mediating our observed effects. As in mammals,
chicks have 4 FGFRs. The expression patterns of FGFR1, 2, and 3
have been described in chicks69 but not in the context of EEM
differentiation. We first carried out detailed in situ analyses for
FGFR1-4 from HH4 to HH12, with special attention to their
possible involvement in blood and endothelial differentiation.
Among them, we found FGFR2 (Figure 5Bi-vi) to be the likely
candidate in mediating these observed effects. FGFR2 was expressed in the endothelial cells in the extraembryonic region(Figure 5Bvi,F), whereas primitive blood cells were devoid of its
transcripts (Figure 5F inset). The splanchnic mesoderm in thelateral plate and medial EEM was also positive for FGFR2 (Figure5F). FGFR1 transcripts were not detected in EEM in either blood or
endothelial cells (Figure 5Avi,E), although they were present inboth somitic and lateral plate mesoderm (Figure 5E). FGFR3(Figure 5Ci) was expressed at HH4 in all ectoderm cells except
those adjacent to the streak. At HH10, nascent somites andneighboring intermediate mesoderm were strongly positive (Figure
5Ciii,G). In lateral plate and medial EEM, signals can be detectedin somatic and splanchnic mesoderm but not in endothelial cells or
blood cells at any axial level (Figure 5G). Expression patterns of
FGFR1 and 3 indicated they may also play a role in restricting
primitive blood differentiation to more lateral regions. For FGFR4(Figure 5Di-iii), we did not observe obvious expression in theextraembryonic region at any stage. FGFR4 signals were seen in
the neural plate from HH4 (Figure 5Di-iii).
Table 1. Effect of signal molecules on ␳ expression
FactorInduction (%)

Inhibition, no. beads with effect/
no. beads tested (%)

BMP2

0/13 (0)0/9 (0)BMP7

0/16 (0)0/12 (0)Chordin

0/37 (0)1/12 (8)Noggin0/28 (0)0/20 (0)FGF4

0/62 (0)40/53 (75)FGF80/40 (0)15/29 (52)Nodal

0/24 (0)0/18 (0)Dkk1

0/29 (0)0/22 (0)VEGF165

0/27 (0)0/15 (0)Wnt3A

1/52 (2)0/28 (0)Epo

0/6 (0)0/4 (0)Beads are grafted as indicated (Figure 3A). The effect of each factor is assayedby its ability to either induce (in ectopic or endogenous domain) or inhibit (endogenous) ␳ expression in adjacent cells.

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Figure 4. Inhibition of FGFR causes ectopic blood formationand down-regulation of endothelial markers. (A-F) Control
DMSO bead has no effect (A,C) and SU5402 bead has excessive
surrounding ␳⫹ clusters (B,D). Panels E and F show additional
examples of blood induction by SU5402 beads. Arrows point tografted beads. (G-N) Whole embryo culture in the presence ofDMSO (G,I) or SU5402 (H,J) does not affect general growth (I-J,insets: Paraxis in situ). SU5402 drastically expands medial limit ofblood formation (H,L) compared with DMSO (G,K), as outlined byyellow dotted line. Sections of panels L (Ni-v) and K (M) show
clear induction of ectopic ␳⫹ clusters (arrowhead). Panels Nii-v
show presence of endothelial-like cells (arrows) surrounding
induced (Niii-v) and endogenous (Nii) clusters. (O-X) Downregulation of Ets1 (Oii), Vegfr2 (Pii), and Vecad (Qii) by SU5402
compared with respective controls (Oi, Pi, and Qi). Lmo2 expression is shown for HH4 (Ri), 5 (Rii), and 8 (Riii). At HH10 (S), it isexpressed in both endothelial and blood cells laterally (Ui andsection in Uii) and only in endothelial cells medially (Wi-ii).
SU5402 abolishes endothelial expression of Lmo2 while maintaining blood expression laterally (Vi-ii). Ectopic blood clusters in the
medial region also express Lmo2 (Xi-ii) but only in inner cells like ␳
globin (Niii-v). Arrows indicate blood expression; arrowheads,
endothelial expression.

Since FGFR2 was expressed in endothelial cells and absent inprimitive blood cells, we next tested whether activation of the FGF
pathway mediated through FGFR2 can influence cell fate. For thispurpose, we constructed an expression vector for the constitutive
active form of FGFR2 (CA-FGFR2; see “Molecular biology”). A
DNA construct expressing CA-FGFR2 together with a reporter
GFP gene was cointroduced into posterior streak mesoderm at HH4
by electroporation. Control embryos were electroporated with the
GFP construct alone. We scored the distribution of CA-FGFR2–
positive cells at HH10 in comparison with control embryos. To
exclude a possible secondary effect due to migratory defect, only
cells that had reached the endogenous ␳⫹ domain were scored. In
control embryos (Figure 5H-I,L; electroporated cells, brown; blood
cells, blue), GFP-positive cells were distributed among different
lineages. For our analysis, we grouped the location of positive cells
into 3 categories: primitive blood (␳⫹), endothelial (cells withflattened morphology surrounding ␳⫹ cluster), and others (all otherpositive cells). Among 1571 cells scored (Table 2), 36.0% control
GFP-positive cells (565) became blood cells, whereas 29.9% (470)went to endothelial lineage. The remaining 34.1% (536) contributed to other cell types. Among the CA-FGFR2–positive cells,
distribution ratios were greatly altered (Figure 5J-K,M). In 1703
total cells scored (Table 2), we found fewer than 50 (2.8%)
CA-FGFR2–positive cells differentiated into primitive blood,
whereas the vast majority (1366, 80.2%) became endothelial cells.
To confirm these effects, we performed a knockdown experiment using FGFR2-specific splice-block morpholino (see “Molecular biology”). Antimorpholino staining did not yield sharp
cell boundaries, making statistical analysis difficult. Nevertheless, the general distribution patterns of morpholino-positive
cells were easily recognizable. Cells with control morpholino
were distributed among different lineages (Figure 5N,P; morpholino cells, blue; blood cells, brown), similar to control GFP cells.
FGFR2 morpholino cells showed a distribution pattern opposite

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Figure 5. FGFR2 is the key mediator. (A) FGFR1 at HH4 (Ai), 5(Aii), 6 (Aiii), 8 (Aiv), and 10 (Av). (B) FGFR2 at HH4 (Bi), 6 (Bii), 7(Biii), 9 (Biv), and 10 (Bv). (C) FGFR3 at HH4 (Ci), 6 (Cii), and 10(Ciii). (D) FGFR4 at HH4 (Di), 7 (Dii), and 10 (Diii). (E-G) Sectionsof FGFR1 (E), FGFR2 (F), and FGFR3 (G). Inset in panel F shows
FGFR2 in endothelial but not blood cells. (H-M) Brown indicateselectroporated cells; blue, blood cells stained with ␳; arrowheads,
endothelial distribution; arrows, blood distribution. CA-FGFR2
cells contribute mainly to endothelium (J,K,M), whereas control
cells are seen among different lineages (H-I,L). See Table 2. (N-S)
Blue indicates morpholino electroporated cells; brown, blood cells
stained with ␳. Knockdown of FGFR2 with a splice block morpholino leads to blood localization (O,Q), and control morpholino cells
are distributed among different lineages (N,P). Arrowheads indicate endothelial distribution; arrows, blood distribution. FGFR2
splice morpholino blocks FGFR2 mature transcript accumulationin neural ectoderm at HH4 (compare R with Bi). (S) Brown
indicates morpholino-positive cells and blue FGFR2 expression(R shows embryo before morpholino staining).

to that of CA-FGFR2 and contributed primarily to blood cells(Figure 5O,Q). The efficacy of FGFR2 morpholino was confirmed by down-regulation of the mature FGFR2 transcript inneural ectoderm at HH4 (Figure 5R-S). These observations
demonstrated that FGFR2 plays a critical role in preventing
primitive blood differentiation and promoting endothelial formation, consistent with phenotypical analysis of fgfr2 null mice.70
Relationship with the VEGFR signaling pathway

Some of our observed effects may be explained by proposed roles
of VEGFR signaling during primitive hematopoiesis. VEGFR2 isinitially expressed in blood and endothelial precursors and vegfr2
null embryos die lacking both lineages.34,35 In our analyses, FGF
can induce the VEGFR2 expression and SU5402 resulted in its
suppression, suggesting some effects were mediated via VEGFR.
In bead graft with VEGF165 protein, however, we did not see

FGFR activity modulates expression level of hematopoietic
marker Gata1

Table 2. Statistical distributions

Total

inhibition of ␳ (Table 1). Since SU5402 can block kinase activity ofboth FGFRs and VEGFRs, we used a VEGFR-specific inhibitor
SU541671 to assess its contribution. As a control, we incubated
15–hours after fertilization (hpf) zebrafish embryos with SU5416
until 30 hpf and observed enlarged pericardial cavity and failure of
the establishment of circulation (data not shown), similar to what
has been previously described.72 We then cultured chick embryosfrom HH6-7 in the presence of SU5416. Unlike in treatment with
SU5402, no obvious effect on ␳ expression was observed with
SU5416 (Figure 6A,E), nor did we observe any remarkabledifference with Lmo2 (Figure 6D,H). Interestingly, we observed aslight up-regulation of endothelial marker Ets1 and Vegfr2 with
SU5416 treatment (Figure 6B-C, F-G), contrary to strong inhibition seen with SU5402. This evidence suggested that FGFR plays
important roles during hematopoietic and endothelial differentiation, distinct from those mediated via VEGFR.

Control, no. cells (%)

CA-FGFR2, no. cells (%)

1571 (100)1703 (100)Blood

565 (36.0)

47 (2.8)

Endothelial

470 (29.9)

1366 (80.2)

Others536 (34.1)

290 (17.0)

Primitive blood progenitors express earlier lineage markers Gata2,
Scl, and Gata1 before hemoglobin expression, indicating thatevents to separate the endothelial and hematopoietic lineages takeplace prior to terminal differentiation. Gata1 has been suggested asa definitive marker for the hematopoietic lineage, before theBLOOD, 15 NOVEMBER 2006 䡠 VOLUME 108, NUMBER 10

FGF PATHWAY IN PRIMITIVE HEMATOPOIESIS

3341

gram. Supporting this hypothesis, we observed up-regulated Gata1
expression and expanded Gata1-positive blood island clusters inmedial EEM and lateral plate mesoderm in SU5402-treated embryos (compare Figure 6T,V with Figure 6S,U), similar to the effecton ␳ expression (Figure 4L).

Discussion

Figure 6. Contributions of VEGFR and hematopoietic marker Gata1 in FGFRmediated effects. (A-H) Specific inhibition of VEGFR pathway by SU5416 does notaffect ␳ (E) or Lmo2 (H) expression and slightly up-regulates Vegfr2 (F) and Ets1 (G)compared with their respective controls (A,D,B,C). (I-M) Gata1 expression at HH4(Ii-ii), 6 (Ji-ii), 7 (K), 8 (L), and 10 (Mi-ii). Arrowhead points to weak but broad
expression in EEM. Sections of panels Ii, Ji, and Mi at indicated levels are shown in Iii,
Jii, and Mii, respectively. (N-R) Control bead does not affect Gata1 expression (N-O),
which at HH10 is up-regulated in blood and absent in nonblood cells (Mi-ii). FGF4
bead maintains low-level expression of Gata1 in adjacent cells (P-Q and section in
R). These cells are completely negative for ␳ (Figure 3F). Dotted line marks bead andarrow points to weak expression. (S-V) SU5402 treatment of whole embryo results in
up-regulated and expanded Gata1 expression (T,V) compared with control (S,U),
similar to the effect on ␳ globin (Figure 4L).

expression of terminal differentiation genes. To investigate whether
Gata1 plays a role in FGF-mediated inhibition of terminal differentiation, we first analyzed Gata1 expression carefully during early
development (Figure 6Ii-ii,Ji-ii,K-L,Mi-ii). The earliest expressionof Gata1 in ventral mesoderm was detected at HH5-6 (Figure
6Ji-ii), which covered the entire EEM area. Compared with globin
genes, Gata1 was expressed both earlier and in a wider region.
Up-regulation of Gata1 expression was observed from HH6-7 infuture globin-positive cells (Figure 6Ji-ii,K). This suggested thatthe expression or activity level of Gata1 is critical for its role asterminal activator. We then investigated the effect of FGF beadgraft on Gata1 expression. Control beads did not affect Gata1
expression (0/40; Figure 6N-O), which at HH10 is up-regulated inblood cells and down-regulated in nonblood cells. In cells adjacent
to FGF4 beads, Gata1 can be detected at low levels (Figure 6P-Q),
which were weaker than in blood cells (Figure 6R) but comparable
to the level seen at HH5-6 in the EEM (48/60). These Gata1expressing cells adjacent to the graft beads were completely devoidof globin expression (Figure 3E-F). This suggested that FGF
activity can block the up-regulation of Gata1, which is critical indetermining whether to initiate the terminal differentiation pro-

The early phase of primitive hematopoiesis is dedicated to generating erythrocytes, providing the sole source for circulating erythrocytes until embryonic day 12 (E12) in mice73 and day 10 inchicks.74 Hemoglobin serves as a critical marker for its terminal
differentiation. In this study, we report that 4 of the globin genes are
expressed from early stages of primitive hematopoiesis. Eachglobin is expressed in all blood cells, suggesting no allelicexclusion of embryonic globins within single blood cells. The
initiation of ␳ expression always takes place in a couple of cells
followed by expression in surrounding cells, suggesting a stepwise
terminal differentiation from precursor cells.
At HH7, ␳⫹ cells are limited to the posterior and outer edge of areavasculosa. The majority of EEM cells, having migrated extensively
from their origin in the streak, are still ␳⫺ at this stage. Our data suggestthat migration and differentiation events are distinctly regulated. Nevertheless, early steps of differentiation marked by Gata2, Scl, and Gata1
for hematopoiesis and Vegfr2, Est1, and Vecad for vasculogenesis takeplace during migration, indicating differentiation itself is a multistep
process. Aside from Gata1, however, these markers are initially expressed in both lineages and only later resolve to a specific one. Other
described terminal transcriptional regulators for hematopoiesis, such as
EKLF, have their onset of expression prior to the up-regulation of Gata1.
The delayed expression of hemoglobin indicates a tight regulation of theonset of terminal differentiation. Further investigation will be needed tounderstand the precise mechanism of this delay.
We found that none of the factors tested can ectopically induce
hemoglobin expression and only FGFs cause inhibition. Since we havenot exhausted the list or tested combinations, we do not rule out the
involvement of pathways that did not elicit an effect. The effects weobserve with FGF are achieved through crosstalk with the VEGFR
pathway, as Vegfr2 is up-regulated by high FGFR activity and downregulated by its inhibition. However, our data suggest the FGF pathway
plays a distinct role and is the primary mediator in this process, as
specific inhibition of VEGFR did not cause similar phenotypes.
Crosstalk between these 2 pathways as well as their distinct roles havebeen well documented in other systems.48,75-77
Our evidence suggests FGF acts by modulating Gata1 expression
level or activity. Gata1 expression is considered the last step before
terminal differentiation. However, it is expressed both earlier and more
widely than globins, suggesting the requirement of intermediate steps. A
recent study on the global gene regulation profile of gata1 nullerythroblasts demonstrated a significant delay of globin gene transcription after reintroduction of Gata1, suggesting the involvement ofadditional signals.78 Up-regulation of Gata1 expression, however, coincides with its purported role as a terminal activator, suggesting theimportance of Gata1 level in terminal differentiation. A reduction of 4to 5-fold in Gata1 expression was shown to markedly impair erythroid
cell maturation in mice.79 In our analyses, FGF strongly inhibits globin
gene expression, although the cells surrounding FGF beads still express
weak but detectable levels of Gata1. The up-regulation of Gata1
expression, however, is inhibited by high FGF activity, and inhibition of
FGFR activity prominently up-regulates Gata1 expression in ectopicallyinduced blood cells. Whether this is achieved by a direct or indirect
effect of the FGF signaling pathway will require further investigation.

3342

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NAKAZAWA et al

In chicks, prestreak-stage marginal zone tissue can be induced
to form blood by bFGF.51 This is likely due to FGF’s role in early
mesoderm induction and reflects different FGF functions at different stages of early development. A similar report with dissociatedquail prestreak-stage epiblast cells showed that FGF can induce
both blood and endothelial cells in culture.80 In Xenopus, midblastula animal cap cells can be induced by a combination of FGF and
BMP to form Gata1-positive cells.52 Another Xenopus study
reported that FGF inhibits blood island formation and dominantnegative FGFR expands blood islands to the lateral plate mesoderm.54 Although our analyses tend to agree with the latter
observation, and beads soaked with both BMP and FGF caused
similar inhibition of ␳ expression as with FGF alone (F.N. and G.S.,
unpublished data, 2006), these effects in Xenopus cannot be easily
compared with ours. The FGF pathway is well documented to playpleiotropic roles in multiple early developmental processes and forthis reason we have chosen to study only the terminal differentiation step within a defined developmental window. Our analyses,
however, support studies on cultured progenitor cells. bFGF wasreported to maintain the immature state of primitive blood progenitors while blocking final differentiation.55 Removal of FGF signaling effectively promoted final maturation.55 Similarly, FGF activity
was recently shown to block the final maturation of presomitic
mesoderm (PSM) cells during somitogenesis.81 In addition, ourstudy provides strong evidence that FGFR2 is the main mediator inthis process. It is premature, however, to hypothesize which
ligand/receptor pair may be involved, as there are at least 22 FGF
ligands and 2 FGFR2 isoforms.82 FGF4 and 8 used in this study
were described to be expressed medially in mesoderm precursor
cells83,84 and possibly create an FGFR2 activity gradient along themedio-lateral axis to control the timing of differentiation. Progressive decay of FGF8 mRNA can serve as a key mechanism forcreating a morphogenetic gradient of FGF8 protein.83 Both FGFs
were shown to bind and activate FGFR2 effectively.70 In addition tothese 2 FGFs, other FGF transcripts (3, 10, 12, 13, 14, and 19) werereported to be expressed in ventral mesoderm precursors.84,85 None
has been studied in detail in the context of EEM differentiation.

Through gene array analysis, we have identified 5 additional FGFs
expressed medially and at least 3 FGFs expressed more laterally in
differentiating EEM cells (M.S., F.N., and G.S.; unpublished data,
2006). Additional complexity was highlighted by specific roles
played by different isoforms.86 Comprehensive analysis of dynamic expression patterns of different FGF molecules and theirisoforms during EEM differentiation is currently being carried out.
Blood and endothelial differentiation are developmentally linked.
The existence of bipotential precursor cells has been shown with invitro differentiation assays. Whether bipotential precursor cells
exist at the single-cell level remains to be shown in vivo. Within apopulation of undifferentiated blood island cells, the bias toward aparticular lineage, the restriction of differentiation potentials, andthe terminal differentiation happen progressively and may require adelicate balance of inputs from transcriptional regulators andsignaling molecules. A surprising find in our studies is the tightnessof this link. In our SU5402-treated embryos, we found strong
inhibition of all endothelial markers tested, yet endothelial-likecells still surround blood cells. However, they are not functional
endothelial cells and fail to establish a proper vascular network
later in development. In medial regions where ectopic blood cells
are induced, globin-negative cells with endothelial morphology arestill observed within the same BI, no matter how few cells the BI
contains. These observations suggest that primitive hematopoiesis
is tightly regulated to occur in conjunction with endothelial
differentiation, possibly controlled by yet unknown mechanisms toensure that all blood cells generated are properly connected to thecirculatory system.

Acknowledgments
We thank Erike Sukowati for help in embryology; Masahiko Hibi
for help with fish experiments; and Shinichi Nishikawa, Leonard
Zon, Claudio Stern, Fred Wilt, and Brendan McIntyre for criticalcomments. This work was initiated in Claudio Stern’s laboratory,
and we thank him for continuous support.

References
1. Baron MH. Early patterning of the mouse embryo:
implications for hematopoietic commitment anddifferentiation. Exp Hematol. 2005;33:1015-1020.
2. McGrath KE, Palis J. Hematopoiesis in the yolk
sac: more than meets the eye. Exp Hematol.
2005;33:1021-1028.
3. Dieterlen-Lievre F, Le Douarin NM. From the hemangioblast to self-tolerance: a series of innovations gained from studies on the avian embryo.
Mech Dev. 2004;121:1117-1128.
4. Orkin SH, Zon LI. Hematopoiesis and stem cells:
plasticity versus developmental heterogeneity.
Nat Immunol. 2002;3:323-328.
5. Kondo M, Wagers AJ, Manz MG, et al. Biology ofhematopoietic stem cells and progenitors: implications for clinical application. Annu Rev Immunol. 2003;21:759-806.
6. Dzierzak E. The emergence of definitive hematopoietic stem cells in the mammal. Curr Opin Hematol. 2005;12:197-202.
7. Psychoyos D, Stern CD. Fates and migratoryroutes of primitive streak cells in the chick embryo. Development. 1996;122:1523-1534.
8. Minko K, Bollerot K, Drevon C, Hallais MF, Jaffredo T. From mesoderm to blood islands: patterns of key molecules during yolk sac erythropoiesis. Gene Expr Patterns. 2003;3:261-272.
9. Garcia-Martinez V, Schoenwolf GC. Positional
control of mesoderm movement and fate during
avian gastrulation and neurulation. Dev Dyn.
1992;193:249-256.

10. Hsia N, Zon LI. Transcriptional regulation of hematopoietic stem cell development in zebrafish.
Exp Hematol. 2005;33:1007-1014.
11. Park C, Ma YD, Choi K. Evidence for the hemangioblast. Exp Hematol. 2005;33:965-970.
12. Jaffredo T, Nottingham W, Liddiard K, Bollerot K,
Pouget C, de Bruijn M. From hemangioblast to
hematopoietic stem cell: an endothelial connection? Exp Hematol. 2005;33:1029-1040.
13. Lacaud G, Robertson S, Palis J, Kennedy M,
Keller G. Regulation of hemangioblast development. Ann N Y Acad Sci. 2001;938:96-107; discussion 108.
14. Huber TL, Kouskoff V, Fehling HJ, Palis J, Keller
G. Haemangioblast commitment is initiated in theprimitive streak of the mouse embryo. Nature.
2004;432:625-630.
15. Nishikawa SI. A complex linkage in the developmental pathway of endothelial and hematopoietic
cells. Curr Opin Cell Biol. 2001;13:673-678.
16. Kuo Z. Ontogeny of embryonic behavior in Aves,
I: the chronology and general nature of the behavior of the chick embryo. J Exp Zool. 1932;61:395-430.
17. Pierce M. The amnion of the chick as an independent effector. J Exp Zool. 1933;65:443-473.
18. Sabin FR. Studies on the origin of blood-vessels
and of red blood-corpuscles as seen in the living
blastderm of chicks during the second day of incubation. Carnegie Inst Wash Publ Contribs Embryol. 1920;9:213-262.

19. Sabin FR. Origin and development of the primitive vessels of the chick and of the pig. Carnegie
Inst Wash Publ Contribs Embryol. 1917;6:61-124.
20. Murray PDF. The development in vitro of theblood of the early chick embryo. Proc R Soc Lond
B Biol Sci. 1932;11:497-521.
21. Wilt FH. The control of embryonic hemoglobin
synthesis. Adv Morphog. 1967;6:89-125.
22. Ferkowicz MJ, Yoder MC. Blood island formation:
longstanding observations and modern interpretations. Exp Hematol. 2005;33:1041-1047.
23. Bruns GA, Ingram VM. The erythroid cells andhaemoglobins of the chick embryo. Philos TransR Soc Lond B Biol Sci. 1973;266:225-305.
24. Ginder GD, Singal R, Little JA, Dempsey N, Ferris R, Wang SZ. Silencing and activation of embryonic globin gene expression. Ann N Y Acad
Sci. 1998;850:70-79.
25. Evans T, Felsenfeld G, Reitman M. Control of globin gene transcription. Annu Rev Cell Biol. 1990;
6:95-124.
26. Minie M, Clark D, Trainor C, et al. Developmental
regulation of globin gene expression. J Cell Sci
Suppl. 1992;16:15-20.
27. Sieweke MH, Graf T. A transcription factor party
during blood cell differentiation. Curr Opin Genet
Dev. 1998;8:545-551.
28. Engel I, Murre C. Transcription factors in hematopoiesis. Curr Opin Genet Dev. 1999;9:575-579.

BLOOD, 15 NOVEMBER 2006 䡠 VOLUME 108, NUMBER 10

29. Cantor AB, Orkin SH. Hematopoietic development: a balancing act. Curr Opin Genet Dev.
2001;11:513-519.
30. Walmsley M, Ciau-Uitz A, Patient R. Adult andembryonic blood and endothelium derive fromdistinct precursor populations which are differentially programmed by BMP in Xenopus.
Development. 2002;129:5683-5695.
31. Snyder A, Fraser ST, Baron MH. Bone morphogenetic proteins in vertebrate hematopoietic development. J Cell Biochem. 2004;93:224-232.
32. Schmerer M, Evans T. Primitive erythropoiesis isregulated by Smad-dependent signaling in postgastrulation mesoderm. Blood. 2003;102:31963205.
33. Drake CJ, LaRue A, Ferrara N, Little CD. VEGF
regulates cell behavior during vasculogenesis.
Dev Biol. 2000;224:178-188.
34. Shalaby F, Ho J, Stanford WL, et al. A requirement for Flk1 in primitive and definitive hematopoiesis and vasculogenesis. Cell. 1997;89:981990.
35. Eichmann A, Corbel C, Nataf V, Vaigot P, BreantC, Le Douarin NM. Ligand-dependent development of the endothelial and hemopoietic lineages
from embryonic mesodermal cells expressing
vascular endothelial growth factor receptor 2.
Proc Natl Acad Sci U S A. 1997;94:5141-5146.
36. Gerber HP, Malik AK, Solar GP, et al. VEGF regulates haematopoietic stem cell survival by an internal autocrine loop mechanism. Nature. 2002;
417:954-958.
37. Van Den Berg DJ, Sharma AK, Bruno E, Hoffman
R. Role of members of the Wnt gene family in human hematopoiesis. Blood. 1998;92:3189-3202.
38. Austin TW, Solar GP, Ziegler FC, Liem L, Matthews W. A role for the Wnt gene family in hematopoiesis: expansion of multilineage progenitor
cells. Blood. 1997;89:3624-3635.

FGF PATHWAY IN PRIMITIVE HEMATOPOIESIS

Rusnati M. Fibroblast growth factor/fibroblast
growth factor receptor system in angiogenesis.
Cytokine Growth Factor Rev. 2005;16:159-178.
49. Auguste P, Javerzat S, Bikfalvi A. Regulation ofvascular development by fibroblast growth factors. Cell Tissue Res. 2003;314:157-166.
50. Moroni E, Dell’Era P, Rusnati M, Presta M. Fibroblast growth factors and their receptors in hematopoiesis and hematological tumors. J Hematother Stem Cell Res. 2002;11:19-32.
51. Gordon-Thomson C, Fabian BC. Hypoblastic
tissue and fibroblast growth factor induce blood
tissue (haemoglobin) in the early chick embryo.
Development. 1994;120:3571-3579.
52. Huber TL, Zhou Y, Mead PE, Zon LI. Cooperative
effects of growth factors involved in the inductionof hematopoietic mesoderm. Blood. 1998;92:4128-4137.
53. Allouche M, Bikfalvi A. The role of fibroblast
growth factor-2 (FGF-2) in hematopoiesis. Prog
Growth Factor Res. 1995;6:35-48.
54. Xu RH, Ault KT, Kim J, et al. Opposite effects of
FGF and BMP-4 on embryonic blood formation:
roles of PV.1 and GATA-2. Dev Biol. 1999;208:352-361.
55. Bartunek P, Pajer P, Karafiat V, Blendinger G,
Dvorak M, Zenke M. bFGF signaling and v-Myb
cooperate in sustained growth of primitive erythroid progenitors. Oncogene. 2002;21:400-410.
56. Stern CD. Detection of multiple gene products
simultaneously by in situ hybridization and immunohistochemistry in whole mounts of avian embryos. Curr Top Dev Biol. 1998;36:223-243.
57. Shin M, Watanuki K, Yasugi S. Expression of
Fgf10 and Fgf receptors during development of
the embryonic chicken stomach. Gene Expr Patterns. 2005;5:511-516.

39. Shawber CJ, Kitajewski J. Notch function in thevasculature: insights from zebrafish, mouse andman. Bioessays. 2004;26:225-234.

58. Webster MK, Donoghue DJ. Enhanced signaling
and morphological transformation by a membrane-localized derivative of the fibroblast growth
factor receptor 3 kinase domain. Mol Cell Biol.
1997;17:5739-5747.

40. Siatskas C, Boyd R. Regulation of chicken haemopoiesis by cytokines. Dev Comp Immunol.
2000;24:37-59.

59. Niwa H, Yamamura K, Miyazaki J. Efficient selection for high-expression transfectants with a novel
eukaryotic vector. Gene. 1991;108:193-199.

41. Slack JM, Darlington BG, Heath JK, Godsave SF.
Mesoderm induction in early Xenopus embryosby heparin-binding growth factors. Nature. 1987;
326:197-200.

60. Palis J, McGrath KE, Kingsley PD. Initiation ofhematopoiesis and vasculogenesis in murine yolksac explants. Blood. 1995;86:156-163.

42. Yamaguchi TP, Harpal K, Henkemeyer M, Rossant J. fgfr-1 is required for embryonic growth andmesodermal patterning during mouse gastrulation. Genes Dev. 1994;8:3032-3044.
43. Sun X, Meyers EN, Lewandoski M, Martin GR.
Targeted disruption of Fgf8 causes failure of cell
migration in the gastrulating mouse embryo.
Genes Dev. 1999;13:1834-1846.
44. Furthauer M, Van Celst J, Thisse C, Thisse B. Fgf
signalling controls the dorsoventral patterning of
the zebrafish embryo. Development. 2004;131:2853-2864.
45. Griffin K, Patient R, Holder N. Analysis of FGF
function in normal and no tail zebrafish embryos
reveals separate mechanisms for formation of thetrunk and the tail. Development. 1995;121:29832994.
46. Nutt SL, Dingwell KS, Holt CE, Amaya E. Xenopus Sprouty2 inhibits FGF-mediated gastrulation
movements but does not affect mesoderm induction and patterning. Genes Dev. 2001;15:11521166.
47. Yang X, Dormann D, Munsterberg AE, Weijer CJ.
Cell movement patterns during gastrulation in thechick are controlled by positive and negative chemotaxis mediated by FGF4 and FGF8. Dev Cell.
2002;3:425-437.
48. Presta M, Dell’Era P, Mitola S, Moroni E, Ronca R,

61. Worthington RE, Bossie-Codreanu J, Van Zant G.
Quantitation of erythroid differentiation in vitro
using a sensitive colorimetric assay for hemoglobin. Exp Hematol. 1987;15:85-92.
62. Wilt FH. The beginnings of erythropoiesis in theyolk sac of the chick embryo. Ann N Y Acad Sci.
1974;241:99-112.
63. Wainwright SD, Wainwright LK. A kinetic study ofthe effects of delta-aminolevulinic acid upon thesynthesis of embryonic and fetal hemoglobins in
the blood islands of the developing chick blastodisc. Can J Biochem. 1970;48:400-406.

3343

tal study of endoderm formation in avian embryos. Anat Embryol (Berl). 1981;163:245-263.
69. Walshe J, Mason I. Expression of FGFR1, FGFR2and FGFR3 during early neural development in thechick embryo. Mech Dev. 2000;90:103-110.
70. Xu X, Weinstein M, Li C, et al. Fibroblast growth factor receptor 2 (FGFR2)-mediated reciprocal regulation loop between FGF8 and FGF10 is essential for
limb induction. Development. 1998;125:753-765.
71. Fong TA, Shawver LK, Sun L, et al. SU5416 is apotent and selective inhibitor of the vascular endothelial growth factor receptor (Flk-1/KDR) that
inhibits tyrosine kinase catalysis, tumor vascularization, and growth of multiple tumor types. Cancer Res. 1999;59:99-106.
72. Cross LM, Cook MA, Lin S, Chen JN, Rubinstein
AL. Rapid analysis of angiogenesis drugs in a live
fluorescent zebrafish assay. Arterioscler Thromb
Vasc Biol. 2003;23:911-912.
73. Palis J, Robertson S, Kennedy M, Wall C, Keller
G. Development of erythroid and myeloid progenitors in the yolk sac and embryo proper of themouse. Development. 1999;126:5073-5084.
74. Dieterlen-Lievre F, Beaupain D, Martin C. Originof erythropoietic stem cells in avian development:
shift from the yolk sac to an intraembryonic site.
Ann Immunol (Paris). 1976;127:857-863.
75. Poole TJ, Finkelstein EB, Cox CM. The role of
FGF and VEGF in angioblast induction and migration during vascular development. Dev Dyn.
2001;220:1-17.
76. Jih YJ, Lien WH, Tsai WC, Yang GW, Li C, Wu
LW. Distinct regulation of genes by bFGF and
VEGF-A in endothelial cells. Angiogenesis. 2001;
4:313-321.
77. Nico B, de Falco G, Vacca A, Roncali L, Ribatti D.
In vivo absence of synergism between fibroblast
growth factor-2 and vascular endothelial growth
factor. J Hematother Stem Cell Res. 2001;10:
905-912.
78. Welch JJ, Watts JA, Vakoc CR, et al. Global regulation of erythroid gene expression by transcription factor GATA-1. Blood. 2004;104:3136-3147.
79. McDevitt MA, Shivdasani RA, Fujiwara Y, Yang H,
Orkin SH. A “knockdown” mutation created bycis-element gene targeting reveals the dependence of erythroid cell maturation on the level oftranscription factor GATA-1. Proc Natl Acad Sci
U S A. 1997;94:6781-6785.
80. Flamme I, Risau W. Induction of vasculogenesis
and hematopoiesis in vitro. Development. 1992;
116:435-439.
81. Delfini MC, Dubrulle J, Malapert P, Chal J, Pourquie
O. Control of the segmentation process by graded
MAPK/ERK activation in the chick embryo. Proc Natl
Acad Sci U S A. 2005;102:11343-11348.
82. Eswarakumar VP, Lax I, Schlessinger J. Cellularsignaling by fibroblast growth factor receptors.
Cytokine Growth Factor Rev. 2005;16:139-149.

64. Silver L, Palis J. Initiation of murine embryonicerythropoiesis: a spatial analysis. Blood. 1997;89:1154-1164.

83. Dubrulle J, Pourquie O. fgf8 mRNA decay establishes a gradient that couples axial elongation topatterning in the vertebrate embryo. Nature.
2004;427:419-422.

65. Eichmann A, Marcelle C, Breant C, Le Douarin
NM. Two molecules related to the VEGF receptor
are expressed in early endothelial cells during
avian embryonic development. Mech Dev. 1993;
42:33-48.

84. Karabagli H, Karabagli P, Ladher RK, Schoenwolf
GC. Comparison of the expression patterns ofseveral fibroblast growth factors during chick gastrulation and neurulation. Anat Embryol (Berl).
2002;205:365-370.

66. Tahtakran SA, Selleck MA. Ets-1 expression isassociated with cranial neural crest migration andvasculogenesis in the chick embryo. Gene Expr
Patterns. 2003;3:455-458.

85. Wright TJ, Ladher R, McWhirter J, Murre C,
Schoenwolf GC, Mansour SL. Mouse FGF15 is
the ortholog of human and chick FGF19, but is
not uniquely required for otic induction. Dev Biol.
2004;269:264-275.

67. Mohammadi M, McMahon G, Sun L, et al. Structures of the tyrosine kinase domain of fibroblast
growth factor receptor in complex with inhibitors.
Science. 1997;276:955-960.
68. Stern CD, Ireland GW. An integrated experimen-

86. Olsen SK, Li JY, Bromleigh C, et al. Structural
basis by which alternative splicing modulates theorganizer activity of FGF8 in the brain. Genes
Dev. 2006;20:185-198.