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First published online 18 January 2006
doi: 10.1242/dev.02242

Development 133, 663-673 (2006)
Published by The Company of Biologists 2006
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Context-specific requirements for Fgfr1 signaling through Frs2 and Frs3 during mouse development

Renée V. Hoch and Philippe Soriano*

Program in Developmental Biology, Division of Basic Sciences, Fred Hutchinson Cancer Research Center, 1100 Fairview Avenue North, Seattle, WA 98109, USA.

* Author for correspondence (e-mail: psoriano{at}fhcrc.org)

Accepted 8 December 2005


   SUMMARY
TOP
 SUMMARY
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Fibroblast growth factor receptor 1 (Fgfr1) plays pleiotropic roles during embryonic development, but the mechanisms by which this receptor signals in vivo have not previously been elucidated. Biochemical studies have implicated Fgf receptor-specific substrates (Frs2, Frs3) as the principal mediators of Fgfr1 signal transduction to the MAPK and PI3K pathways. To determine the developmental requirements for Fgfr1-Frs signaling, we generated mice (Fgfr1{Delta}Frs/{Delta}Frs) in which the Frs2/3-binding site on Fgfr1 is deleted. Fgfr1{Delta}Frs/{Delta}Frs embryos die during late embryogenesis, and exhibit defects in neural tube closure and in the development of the tail bud and pharyngeal arches. However, the mutant receptor is able to drive Fgfr1 functions during gastrulation and somitogenesis, and drives normal MAPK responses to Fgf. These findings indicate that Fgfr1 uses distinct signal transduction mechanisms in different developmental contexts, and that some essential functions of this receptor are mediated by Frs-independent signaling.

Key words: Fgfr1, Frs2, Frs3, Signaling, Gastrulation, Neural tube, Tail bud, Pharyngeal arches


   INTRODUCTION
TOP
 SUMMARY
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Members of the receptor tyrosine kinase (RTK) superfamily elicit similar cellular, transcriptional and biochemical responses when stimulated in cultured cells (Fambrough et al., 1999Go; Schlessinger, 2000Go), yet direct specific, non-redundant cellular responses in developing organisms. It has been proposed that qualitative differences in signal transduction underlie RTK functional specialization. Such differences could be in ligand sensitivity, effector pathway use or the strength or duration of signaling. This model has gained support from in vivo domain swap experiments, which have demonstrated biologically significant differences in RTK signal transduction (Klinghoffer et al., 2001Go; Madhani, 2001Go; Hamilton et al., 2003Go). Analyses of mice harboring signaling domain mutations have also shown that effector pathways are differentially required within the RTK superfamily, and that many signaling events observed in biochemical studies are dispensable or required only for a subset of the in vivo functions of receptors (Partanen et al., 1998Go; Tallquist et al., 2000Go; Maina et al., 2001Go; Klinghoffer et al., 2002Go; Tallquist et al., 2003Go).

The mammalian fibroblast growth factor (Fgf) signaling network consists of four high affinity RTKs and at least 22 ligands (Ornitz, 2000Go; Böttcher and Niehrs, 2005Go). Mouse knockout studies have identified numerous roles of Fgfs, and demonstrated that all essential embryonic roles are mediated by two receptors, Fgfr1 and Fgfr2 (Deng et al., 1994Go; Yamaguchi et al., 1994Go; Deng et al., 1996Go; Arman et al., 1998Go; Yu et al., 2000Go; Böttcher and Niehrs, 2005Go). We have focused our studies on Fgfr1, which is required for postimplantation growth, mesodermal migration and patterning during gastrulation, and for somitogenesis. Fgfr1null mouse embryos also exhibit posterior truncations and neural tube closure defects (Deng et al., 1994Go; Yamaguchi et al., 1994Go). Studies of hypomorphic and conditional alleles have identified additional roles of this receptor in node regression, neural stem cells, olfactory bulb development, and patterning of the anteroposterior axis, central nervous system and pharyngeal arches (Partanen et al., 1998Go; Tropepe et al., 1999Go; Xu et al., 1999Go; Hébert et al., 2003Go; Trokovic et al., 2003aGo; Trokovic et al., 2003bGo).

Though many roles of Fgfr1 are known, the mechanisms by which this receptor signals in vivo have not yet been elucidated. Active Fgfrs can directly engage relatively few proteins, namely Crk, Grb14, Shb, PLC{gamma} and Frs2,3 (hereafter referred to collectively as `Frs') (Mohammadi et al., 1991Go; Wang et al., 1996Go; Kouhara et al., 1997Go; Larrson et al., 1999Go; Reilly et al., 2000Go; Cross et al., 2002Go). Biochemical studies have implicated Frs adaptors as the key mediators of Fgfr signal transduction. These proteins interact constitutively with the intracellular juxtamembrane regions of Fgfr1 and Fgfr2, unlike signaling proteins downstream of other RTKs that are recruited after receptor activation. Fgfr activation leads to phosphorylation of Frs tyrosine residues to which Grb2 and SHP2 are subsequently recruited, initiating PI3K and MAPK signaling (Wang et al., 1996Go; Kouhara et al., 1997Go; Xu et al., 1998Go; Ong et al., 2000Go; Hadari et al., 2001Go). This adaptor-mediated mechanism may distinguish the kinetics, amplitude or subcellular localization of Fgfr signal transduction from other RTK signaling events.

Frs adaptors are conserved among vertebrates, and expression patterns suggest that the two mammalian isoforms perform non-redundant functions in vivo (McDougall et al., 2001Go; Gotoh et al., 2004Go). Loss-of-function studies have not been reported for Frs3, but Frs2-/- mouse embryos die ~E7.5 with defects in extra-embryonic development (Hadari et al., 2001Go; Gotoh et al., 2005Go). Mouse chimera and Xenopus knockdown studies have further implicated Frs2 in mesoderm development and convergent extension, respectively (Akagi et al., 2002Go; Gotoh et al., 2005Go). These studies are informative with respect to Frs2 roles, but they are difficult to interpret in terms of specific growth factor pathways because of the promiscuity of Frs adaptors. In addition to Fgfrs, Frs can interact with active neurotrophin receptors and are phosphorylated downstream of other RTKs (Rabin et al., 1993Go; Ong et al., 2000Go). Furthermore, both adaptors associate strongly with the cell cycle regulatory protein p13suc1, and may thus play a role in cell cycle regulation or progression (Rabin et al., 1993Go; Ong et al., 1996Go).

To assess the developmental requirements for Frs-mediated signaling downstream of Fgfr1, we have generated mice lacking the Frs-binding site on this receptor (Fgfr1{Delta}Frs). We found that Frs-mediated signaling is dispensable for Fgfr1 functions during gastrulation and somitogenesis, but is required for Fgfr1 roles in neurulation, tail bud and pharyngeal arch development. Furthermore, we found that primary embryonic cells do not require this signaling interaction to elicit strong and sustained MAPK responses to Fgf, and that MAPK activation in vivo is not grossly affected by the Fgfr1{Delta}Frsmutation. These results indicate that Frs adaptors are not the exclusive effectors of Fgfr1 signal transduction in vivo.


   MATERIALS AND METHODS
TOP
 SUMMARY
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Targeting vector construction
Targeting vectors were generated in pPGKneoF2L2DTA, a plasmid containing a DTA selection cassette and a neomycin cassette flanked by nested loxP and FRT sites. For knock-in (KI) vectors (Fgfr1wtKI, Fgfr1{Delta}Frs), the short arm of homology, an XbaI/HindIII fragment spanning introns 6-7 of Fgfr1, was inserted upstream of the 5' loxP site in the same orientation as the neomycin cassette. Intron 7-partial cDNA cassettes were generated through two rounds of PCR SOEing (splicing by overhang extension, oligos are given in Table S1 in the supplementary material) (Pogulis et al., 2000Go). In the first SOEing reaction, nucleotides 1276-1356 (amino acids 407-433) were deleted from a mouse Fgfr1 cDNA construct to generate an Fgfr1{Delta}FrscDNA. In the second round, partial Fgfr1wt, Fgfr1{Delta}Frs cDNAs (exons 8-17) were spliced to the 3' end of intron 7, preserving the endogenous intron 7/exon 8 junction. All PCR products incorporated into targeting vectors were generated using Pfu (Stratagene) and were verified by sequencing. Intron 7-partial cDNA cassettes were digested with Tth111I and ligated to the long arm of homology, a ~4.1 kb Tth111I/PstI genomic fragment. Resultant fragments were cloned between the 3' loxP site and DTA (Fig. 1B).

For the Fgfr1floxex4 vector, the short arm of homology, exon 4 fragment, and part of the 3' long arm of homology were PCR amplified from wild type 129Sv genomic DNA (oligos are given in Table S1 in the supplementary material). The exon 4 fragment was cloned between loxP sites, upstream of the FRT-flanked neomycin cassette in pPGKneoF2L2DTA. The short arm was inserted upstream of the 5' loxP site; the intron 5 PCR fragment was ligated to a genomic HindIII/EcoRV fragment to complete the long arm, which was inserted between the second loxP site and DTA (Fig. 1C).

Generation of mouse lines
AK7 ES cells were electroporated with linearized targeting vectors, G418 selected and PCR screened for integration at the Fgfr1 locus (data not shown). Targeting was verified by Southern analysis with the following probes: Fgfr1KI-5' external (NcoI/XmaI), internal (SpeI/NciI), 3' external (PstI/HindIII); Fgfr1{Delta}ex4-5' external (PCR product from 129Sv genomic DNA, oligos are given in Table S1 in the supplementary material); internal (NotI/NcoI).

Correctly targeted ES cells were injected into wild type C57BL/6J blastocysts to generate chimeras. The neomycin cassette caused early embryonic lethality of KI homozygotes and was excised through breeding to Meox2-Cre or FlpeR mice (Farley et al., 2000Go; Tallquist and Soriano, 2000Go). Complete excision of neomycin (Fgfr1KI, Fgfr1floxex4) and exon 4 (Fgfr1floxex4) was verified by Southern analysis (data not shown). Tail biopsies were routinely PCR genotyped (oligos are given in Table S1 in the supplementary material). In KI homozygotes, embryonic RT-PCR confirmed that targeting did not disrupt alternative splicing of the extracellular domain or splicing of upstream exons into the cDNA (data not shown).

Whole-mount in situ hybridization
Whole-mount in situ hybridization was performed using standard procedures.

RNA preparation and semi-quantitative RT-PCR
Embryonic RNA was prepared using Trizol (GibcoBRL) and reverse transcribed as described previously (Chen and Soriano, 2003Go). Diluted cDNA pools were amplified using the following primer pairs: FGFR1.52/FGFR1.36, FGFR2ex11.51/FGFR2ex13.31, Timm_cDNA+/Timm_cDNA- (see Table S1 in the supplementary material). Triplicate PCR reactions were cycled within a linear range of amplification and products were detected and quantified using Sybr Gold (Molecular Probes) and NIH ImageQuant software. Fgfr1 and Fgfr2 RT-PCR products were normalized to Timm (mitochondrial membrane transport protein) levels.

Immunohistochemistry
Whole-mount phospho-MAPK IHC was performed as described previously (Corson et al., 2003Go), with minor modifications.

For thin section immunohistochemistry, embryos were fixed overnight (4% PFA), embedded in paraffin wax and sectioned. After rehydration, slides were pretreated in (3% H2O2, 10% methanol, 1xPBS). Antigen retrieval was performed by microwaving slides in 10 mM citrate buffer (pH 6.0) (phosphohistone H3, Upstate Biotechnology #06-570) or 10 mM EDTA (pH 8.0) (pMAPK, Cell Signaling #9101S). Samples were blocked in 5% serum/PBS and incubated with antibody overnight. Immunocomplexes were detected using biotinylated secondary antibodies with ABC Elite and DAB substrate kits (Vectastain).

Nile Blue and TUNEL analysis
Freshly isolated embryos were stained in Nile Blue (0.003% in DMEM, 0.5% FBS) for 30-60 minutes (37°C) and washed in PBS. For TUNEL analysis, embryos were fixed, embedded and sectioned as for immunohistochemistry. Rehydrated sections were incubated in (0.3% Triton-X-100, 1x PBS), digested with proteinase K, and incubated in TdT reaction mix for 1 hour, 37°C [30 mM Tris pH 7.5, 140 mM cacodylate, 1 mM CoCl2, 10 µM dig-dUTP, 0.3 U/µL TdT (Roche)]. Reactions were stopped in PBS and TUNEL-positive nuclei were visualized by IHC with anti-Dig-AP Fab (Roche) and NBT/BCIP detection.

Skeletal preparations
Skeletons were prepared and stained with Alcian Blue and Alizarin Red using a standard protocol.

Mouse embryonic cell (MEC) isolation and culture
E12.5-14.5 embryos were trypsinized (5', 37°C), gently disaggregated with a Pasteur pipette, and plated on gelatin in DMEM, 15% FBS. To generate immortalized lines, p1 MECs were infected with an E6LTTNLoxP retrovirus transducing the SV40 Large T antigen (Berghella et al., 1999Go). Infected cells were selected with 500 µg/ml G418 and resistant colonies were pooled.

Immunoprecipitation (IPs) and western blotting
MECs were seeded at 1.5-1.8x104 cells/cm2, serum starved (24-36 hours, 0.5% FBS) and stimulated with heparin and Fgf (aFgf and bFgf gave similar results; Research Diagnostics). Cells were then washed with cold PBS and lysed in HNTG (20 mM HEPES pH 7.9, 150 mM NaCl, 1% Triton X-100, 10% glycerol, 1.5 mM MgCl2, 1 mM EGTA, 1 µg/ml aprotinin, 1 mM PMSF, 10 mM NaPPi, 0.2 mM activated Na2VO3, 50 mM NaF). IPs were performed overnight in HNTG; immunocomplexes were pulled down using protein A- or G-PLUS agarose (Santa Cruz) and washed in HNTG or (500 mM NaCl, 10 mM Tris pH 7.5, 2 mM EDTA, 1% NP40). SDS-PAGE and western blotting were performed according to standard protocols, with phospho-specific and -nonspecific antibody blots blocked in 1% BSA and 3% milk, respectively.

Antibodies: pMAPK (Cell Signaling); Frs2 (Santa Cruz H-91); Fgfr2 (mouse monoclonal, Research Diagnostics); RasGAP (70.3; gift of Andrius Kazlauskas; Valius et al., 1993Go); phosphotyrosine (clone 4G10, Upstate Biotechnology); pAkt (Cell Signaling); Crk (BD Transduction Labs); and actin (clone AC-15, Sigma).


   RESULTS
TOP
 SUMMARY
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Generation of Fgfr1{Delta}Frs and Fgfr1wtKI mice
We used a partial cDNA knock-in approach to generate mice lacking the Frs binding site on Fgfr1 without disrupting alternative splicing of exons encoding the extracellular domain (Fig. 1) (Werner et al., 1992Go; Yan et al., 1992Go; Johnson and Williams, 1993Go). This deletion mutation has previously been shown to abrogate Fgfr1 binding to both Frs2 and Frs3 without disrupting other receptor functions and interactions (Xu et al., 1998Go). As a control, we used the same approach to generate wild-type knock-in mice (Fgfr1wtKI; the untargeted allele is designated Fgfr1+).


Figure 1
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  		Fig. 1. Fgfr1 targeting. (A) Alleles introduced by gene targeting. (B) Partial cDNA knock-in approach: Fgfr1 exons 8-17, encoding the transmembrane and cytoplasmic domains, were replaced with a pseudoexon encoding the corresponding region of Fgfr1wtKI or Fgfr1{Delta}Frs. Partial cDNAs were spliced at the 5' end into exon 8 and at the 3' end into exon 17, upstream of the endogenous polyadenylation sequence. (C) To generate the null allele, the first exon common to all reported Fgfr1 splice variants (exon 4) was flanked with loxP sites and excised in vivo by Meox2-Cre (Tallquist and Soriano, 2000Go). RT-PCR from embryonic RNA confirmed the generation of a stop codon soon after the exon3-exon5 splice junction (data not shown). (D) Southern blots verifying correct targeting of all alleles in ES cell clones. Abbreviations: A, ApaI; H, HindIII; KI, knock-in; Nd, NdeI; Nh, NheI; P, PstI; Rv, EcoRV; T, Tth111I; X, XbaI; ext, external; int, internal. (E) Relative Fgfr1 mRNA levels in homozygous embryos, assessed by semi-quantitative RT-PCR. Each point is the mean of triplicate reactions for a single embryo (±s.d.).

 
Fgfr1wtKI/wtKI and Fgfr1{Delta}ex4/{Delta}ex4 phenotypes on a high percentage C57BL/6J background
Following two or more backcross generations to the C57BL/6J background, we recovered nearly the expected ratio of Fgfr1wtKI/wtKI animals at E16.5, and a number of homozygotes survived to adulthood with normal fertility (Table 1). We therefore selected this background (>75% contribution from C57BL/6J) for our studies. On this genetic background, Fgfr1wtKI/wtKI embryos are often smaller than littermates and exhibit digit 3/4 fusions (data not shown). Importantly, however, Fgfr1{Delta}Frs/{Delta}Frs phenotypes discussed below are either absent or notably less penetrant/severe in Fgfr1wtKI/wtKI embryos (see Fig. S1 in the supplementary material). Phenotypic differences between the knock in lines were not due to differences in Fgfr1 expression: Fgfr1 mRNA levels are similar in untargeted, Fgfr1wtKI/wtKI and Fgfr1{Delta}Frs/{Delta}Frs embryos on the high percentage C57 background (Fig. 1E).


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  		Table 1. Genotypes of offspring from heterozygote knock-in (KI) intercrosses at late embryonic and postnatal stages

 
Because the Fgfr1wtKI/wtKI phenotype is background dependent (data not shown), we revisited the null phenotype on the (>75%) C57BL/6J background. We generated mice harboring a null allele (Fgfr1{Delta}ex4, Fig. 1A,C) and backcrossed them more than two generations to C57BL/6J. Compared with published Fgfr1null embryos, which were analyzed on different mixed backgrounds,Fgfr1{Delta}ex4/{Delta}ex4 embryos on the (>75%) C57BL/6J background survive slightly later in embryogenesis (E11.5 vs. E7.5-9.5). However, they are always developmentally arrested prior to E9.5 and exhibit phenotypes similar to published Fgfr1null embryos, including a developmental delay, mesodermal migration and patterning defects, craniorachischisis and posterior truncations (Fig. 2; Fig. 3B) (Deng et al., 1994Go; Yamaguchi et al., 1994Go). Fgfr1{Delta}ex4/{Delta}ex4 embryos also have enlarged hearts with disrupted looping (data not shown).

Fgfr1-Frs signaling is not required during gastrulation or somitogenesis
Fgfr1{Delta}Frs/+ animals are viable and indistinguishable from littermate controls, but Fgfr1{Delta}Frs/{Delta}Frs mice were never recovered postnatally (Table 1). Timed matings indicated that Fgfr1{Delta}Frs/{Delta}Frs embryos survive much later in embryogenesis than do Fgfr1{Delta}ex4/{Delta}ex4 embryos, with a drop in viability only after E15.5 (Fig. 2A).

Fgfr1 roles in early mesodermal development were rescued in ~80% of Fgfr1{Delta}Frs/{Delta}Frs embryos. During gastrulation, Fgfr1 is required for mesodermal migration through the primitive streak. In Fgfr1{Delta}ex4/{Delta}ex4 (and published null) embryos, this migration is impaired and cells accumulate in the streak. Consequently, there is an expansion of axial mesoderm and reduction of paraxial mesoderm, which remains disorganized and fails to form somites (Fig. 2B,C) (Deng et al., 1994Go; Yamaguchi et al., 1994Go; Ciruna et al., 1997Go; Ciruna and Rossant, 2001Go). We examined the development of axial and paraxial mesoderm in Fgfr1{Delta}Frs/{Delta}Frs embryos by Shh and Meox1 in situ hybridizations, respectively. Although Fgfr1{Delta}Frs/{Delta}Frs embryos are developmentally delayed by ~1 day, most mutants exhibit normal expression of both mesodermal markers following gastrulation (Fig. 2D-G). This indicates that mesodermal migration is rescued in these embryos. Furthermore, whereas Fgfr1{Delta}ex4/{Delta}ex4 embryos never form somites, Fgfr1{Delta}Frs/{Delta}Frs embryos undergo normal somitogenesis, as evidenced by the segmented pattern of Meox1 staining.


Figure 2
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  		Fig. 2. Rescue of early Fgfr1 functions by Fgfr1{Delta}Frs. (A) Viability (percent of expected) of homozygotes recovered from Fgfr1{Delta}Frs/+ and Fgfr1{Delta}ex4/+ intercrosses. (B-G) Sonic hedgehog (Shh, E9.5) and Meox1 (E10.5) in situ hybridization. Arrows indicate width of the Shh expression domain (more than two cell diameters in Fgfr1{Delta}ex4/{Delta}ex4 compared with one cell diameter in control embryos). (C) The paraxial mesoderm population is reduced and disorganized in Fgfr1{Delta}ex4/{Delta}ex4 embryos, so Meox1 staining is faint and diffuse. (Color development time required to visualize any positive signal was notably longer in this embryo than in F,G.)

 
Fgfr1-Frs signaling is essential during neurulation and tail bud development
Thirty to 50% of Fgfr1{Delta}Frs/{Delta}Frs embryos isolated E10.5-11.5 exhibited a defect in spinal neural tube closure (Fig. 3A,C). This was never observed in Fgfr1wtKI/wtKI embryos, and differs from the neurulation phenotype observed in Fgfr1{Delta}ex4/{Delta}ex4 embryos in which the neural tube remains open along the entire rostrocaudal axis (craniorachischisis; Fig. 3B). The spinal neural tube (caudal to approximately somite 8) closes by the elevation and fusion of neural folds in a progressive rostral-to-caudal manner (Shum and Copp, 1996Go; Copp et al., 2003Go). In some Fgfr1{Delta}Frs/{Delta}Frs embryos, the spinal neural tube remained completely open at E10.5, which could indicate a mere delay in the closure process (Fig. 3C, right). In others, however, the spinal neural tube was closed at discrete points but open in the intervening regions (Fig. 3C, left). Later in development, some Fgfr1{Delta}Frs/{Delta}Frs embryos completed neural tube closure and others exhibited varying degrees of spina bifida (Fig. 3D-F, data not shown). In one embryo (E17.5), spina bifida was accompanied by loss of spinal cord marginal layer tissue and caudal meningomyelocele (extrusion of the spinal cord and meninges from the vertebral column; data not shown).


Figure 3
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  		Fig. 3. Neural tube defects. (A-C) Hoxb1 whole-mount in situ hybridization, labeling dorsal neural tube in the spinal region. Edges of open neuroepithelium are outlined in B. In C, the open cranial region (left embryo) is a procedural artifact. (D-F) Hematoxylin and Eosin stained sections through spinal neural tubes of E10.5 littermates. Embryo in E completed closure and embryo in F exhibited the intermittent neural tube closure phenotype. (G) Mitotic and apoptotic indices of neural tube sections (E10.5), assessed at similar spinal levels by phosphohistone H3 immunostaining and TUNEL. Each data point is the mean of positive/total cells on 12-16 sections per embryo, two embryos/genotype. Error bars indicate ±1 s.d.; P values were determined by Student's t-test.

 
Histological analysis revealed altered morphology of the canal through mutant neural tubes in the spinal region. This was observed in mutant embryos that completed closure, as well as in those that retained regions of open neural tube (Fig. 3E,F). During neurulation, spinal neural folds bend towards the midline through the formation of neuroepithelial hingepoints; this morphogenetic process occurs by different mechanisms at different axial levels. In the upper- and mid-spinal regions, the neuroepithelium bends at one medial hingepoint and the lateral edges fold towards the dorsal midline, generating a slit-shaped neural canal (e.g. Fig. 3D). In the lower spinal region, two dorsolateral hingepoints (DLHP) form in addition to the medial hingepoint, resulting in a diamond-shaped canal (Shum and Copp, 1996Go). In Fgfr1{Delta}Frs/{Delta}Frsembryos, the canal is diamond shaped at all spinal levels (Fig. 3E,F), suggesting that this second morphogenetic mechanism is used to close the entire spinal neural tube. Cranial neuroepithelial folding appeared normal in all Fgfr1{Delta}Frs/{Delta}Frs embryos (data not shown).

Normally, notochordal Shh inhibits DLHP formation at mid- and upper spinal levels, while signals from the dorsal ectoderm are thought to play an inductive role (Copp et al., 2003Go). In Fgfr1{Delta}Frs/{Delta}Frsembryos, the notochord appears normal, extends through the spinal region, and expresses Shh (Fig. 2E, Fig. 3D-F). Although ectodermal-inducing signals have not yet been identified, it has been proposed that DLHP formation involves localized proliferation or apoptosis in the dorsolateral neuroepithelium (Copp et al., 2003Go). We analyzed proliferation and apoptosis in Fgfr1{Delta}Frs/{Delta}Frs neural tubes and found that although the mitotic index is not significantly altered in these embryos, they do exhibit an increase in cell death within the spinal neural tube (Fig. 3G). However, as apoptosis was not specifically localized to dorsolateral regions (data not shown), this probably reflects a role of Fgfr1-Frs signaling separate from its role in neurulation.

Neural tube closure defects are often associated with posterior truncations, and it has been postulated that the curvature of the body axis generated during tail bud outgrowth provides tension in the neural folds that facilitates closure (Copp et al., 1982Go; Copp, 1985Go; Gofflot et al., 1997Go; Peeters et al., 1998Go; Finnell et al., 2003Go). Over 80% of Fgfr1{Delta}Frs/{Delta}Frs embryos exhibit posterior truncations of varying severity (Fig. 4A-C, see Fig. S1A in the supplementary material). Truncations persist through embryogenesis and in severe cases are accompanied by a truncation of the notochord at the level of the hindlimb and failure to bifurcate caudal limb fields and organs (Fig. 4D,E, data not shown). In previous Fgfr1 reports, posterior truncations were associated with ectopic posterior neuroectoderm and suggested to be secondary to the mesodermal defects (Ciruna et al., 1997Go; Ciruna and Rossant, 2001Go). However, in Fgfr1{Delta}Frs/{Delta}Frs embryos, we observed truncations in the absence of these other phenotypes (Fig. 2, data not shown), indicating that Fgfr1-Frs signaling directly impacts tail bud development.

All tissue layers of the tail bud form during secondary gastrulation from remnants of the primitive streak and node. In most Fgfr1{Delta}Frs/{Delta}Frs embryos, Shh in situ hybridization and Hematoxylin and Eosin staining of embryo sections indicated full extension of the notochord, and hence, correct positioning of tail bud progenitors, at the onset of secondary gastrulation (Fig. 2E, data not shown). We investigated whether the Frs site deletion directly affected patterning or outgrowth of the tail bud. Mutant analysis and expression studies have identified genes expressed and required in the tail bud during secondary gastrulation (Roelink and Nusse, 1991Go; Takada et al., 1994Go; Greco et al., 1996Go; Gofflot et al., 1997Go; Copp et al., 2003Go). Except in the most severely truncated embryos, tail bud gene expression (T, Wnt3a, Fgf8, Hoxb1) and proliferation appeared normal in Fgfr1{Delta}Frs/{Delta}Frs tail buds (Fig. 4F-I, data not shown). However, TUNEL analysis revealed an increase in cell death throughout caudal tissues of Fgfr1{Delta}Frs/{Delta}Frs embryos (Fig. 4I). Cell death may hinder cell migrations or reduce the number of tail bud progenitors during secondary gastrulation.


Figure 4
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  		Fig. 4. Posterior truncations and tail bud development. (A-C) Bright-field photographs of E13.5 littermates (caudal region). (D,E) Hematoxylin and Eosin stained sections through kidneys of wild-type and severely truncated Fgfr1{Delta}Frs/{Delta}Frs embryos; bar indicates relative magnification. (F) T (Brachyury) and (G,H) Wnt3a whole-mount in situ hybridization. (I) Quantitation of proliferation (pH3, P=0.33) and apoptosis (TUNEL, P=0.09) in E10.5 tail bud sections. Each data point represents the mean of three to eight sections through each of at least three embryos. Error bars indicate ±1 s.d.; P values were determined by Student's t-test.

 


Figure 5
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  		Fig. 5. Pharyngeal arch (PA) development. (A-C) Phosphohistone H3 immunostained PA sections (E10.5). PA1 is labeled, arrowheads indicate PA2. (D,E) Fgf8 whole-mount in situ hybridization (PA region); arrows indicate PA2 Fgf8 expression domain lost in mutant embryos. (F-H) Migrating NCCs in the pharyngeal region of stage-matched embryos, visualized by Sox10 in situ hybridization. Arrows indicate limits of NCC migration into PA1, 2. (I-L) Nile Blue staining of cell death in the pharyngeal region (E10.5). I/J and K/L are stage-matched control/mutant embryo pairs.

 
Fgfr1-Frs signaling in pharyngeal arch (PA) development
The second PA is hypoplastic in Fgfr1{Delta}Frs/{Delta}Frs embryos by E10.5, and histological analysis revealed a dramatic reduction in the number of mesenchymal cells populating this arch. This population normally includes both mesoderm and neural crest cells. Unlike cells within wild-type PAs, mesenchymal cells within mutant PA2 are neither proliferating nor closely apposed to the surrounding epithelia (Fig. 5A-C). We also observed a loss of Fgf8 expression in PA2, which has been shown to promote survival of PA mesenchyme (Fig. 5D,E) (Frank et al., 2002Go; Macatee et al., 2003Go). Although TUNEL analysis on sections did not reveal ectopic apoptosis of cells within Fgfr1{Delta}Frs/{Delta}Frs arches (data not shown), the loss of Fgf8 signals may contribute to the lack of PA2 mesenchymal proliferation or affect migration of neural crest cells into the arches of mutant embryos. In a previous study of Fgfr1 hypomorphs, PA2 was found to be hypoplastic because of a neural crest migration defect (Trokovic et al., 2003aGo). Likewise, we observed impaired neural crest cell migration into Fgfr1{Delta}Frs/{Delta}Frs PAs by Sox10 in situ hybridization (Fig. 5F-H) (Kuhlbrodt et al., 1998Go). Nile Blue staining revealed ectopic cell death along the path of neural crest migration (prior to PA entry; Fig. 5I-L).

At later stages, we noted some recovery of PA2 size and did not observe prominent craniofacial defects that would be expected if PA neural crest development failed entirely. We therefore analyzed the formation of PA crest derivatives to determine whether neural crest migration recovered later in development. Analysis of craniofacial skeletal elements at E15.5 indicated that PA2 NCC derivatives are selectively affected in Fgfr1{Delta}Frs/{Delta}Frs embryos. We did not observe major hypoplasia or malformation of the jaws (PA1-derived; data not shown) or PA1 derivatives in the middle ear (Fig. 6A-D). By contrast, cartilages derived from PA2, including the stapes and styloid process of the middle ear and the lesser horns of the hyoid, were missing or hypoplastic in mutant embryos (Fig. 6).

Fgf signaling responses in Fgfr1{Delta}Frs/{Delta}Frs primary cells
Phenotypic data demonstrated that Fgfr1{Delta}Frs is expressed and functional in vivo: we observed neither recapitulation of the null phenotype, as would be expected if the mutant receptor were non-functional, nor gain-of-function phenotypes indicative of ligand-independent activation. However, phenotypic data also indicated that Fgfr1 transduces Frs-independent signals in some developmental contexts. To investigate the nature of these signals, we derived mouse embryonic cells (MECs) from wild-type and mutant embryos, and used them in biochemical studies.

We first used immortalized MECs to verify that the mutant receptor signaled as expected based on previous reports. In these cells, we observed rapid and robust phosphorylation of Frs2 in wild type, but not Fgfr1{Delta}Frs/{Delta}Frs, cells stimulated with Fgf (see Fig. S2A, part i in the supplementary material). We also observed an Fgf-stimulated decrease in Frs2 protein level in wild-type but not mutant cells, concomitant with Frs2 phosphorylation (see Fig. 2A, part ii in the supplementary material). In MAPK response assays, mutant immortalized MECs exhibited reduced sensitivity to Fgf and drove weaker signaling responses as assessed by phosphorylation of MAPK (see Fig. S2B in the supplementary material). These data recapitulate findings of previous studies performed in immortalized Frs2-/- MECs (Hadari et al., 2001Go).


Figure 6
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  		Fig. 6. PA2 neural crest derivatives. Skeleton preparations of E15.5 middle ears (A,B,D) and hyoid cartilages (E,F). PA1, 2 derivatives are traced in C,G,H. PA1 neural crest derivatives, traced in blue: i, incus, m, malleus, ty, tympanic ring, mc, Meckel's cartilage. PA2 derivatives, traced in red, are hypoplastic or missing in mutants: (C) s, stapes, sp, styloid process; (G,H) lesser horns of hyoid. Ectopic cartilages were observed in mutant ears (arrows in B,D; yellow in C).

 
Cellular transformation may affect signal transduction and thus confound results of biochemical studies. Therefore, to approximate the in vivo situation, we re-examined Fgf responses in low passage (p2) non-immortalized MECs. Surprisingly, Fgfr1{Delta}Frs/{Delta}Frs and wild-type p2 MECs responded with indistinguishable MAPK activation profiles, both in terms of response duration and Fgf dose sensitivity (Fig. 7A). We did not observe a notable PI3K pathway response, assayed by Akt phosphorylation, in cells of either genotype (Fig. 7A). In addition, neither Crk nor PLC{gamma} were tyrosine phosphorylated (i.e. activated) in p2 MECs following Fgf stimulation (Fig. 7B, data not shown). Thus, these pathways are probably not mediating the observed MAPK response in Fgfr1{Delta}Frs/{Delta}Frs cells. Furthermore, compared with wild-type cells, the Frs2 phosphorylation response was severely diminished in mutant cells despite comparable levels of total Frs2 (Fig. 7C). We expect that residual Frs activation in Fgfr1{Delta}Frs/{Delta}Frs cells occurs indirectly, as it has previously been shown by yeast two hybrid that the amino acid 407-433 deletion completely abolishes the interaction of the Fgfr1 cytoplasmic domain with Frs2 (Xu et al., 1998Go).

We next investigated whether signaling through Fgfr2 could compensate for the lack of Fgfr1-Frs signaling by activating Frs2 (low level) and MAPK in Fgfr1{Delta}Frs/{Delta}Frs MECs. Fgfr2 expression levels are not altered in Fgfr1{Delta}Frs/{Delta}Frs (p2) MECs or embryos, as might be expected if this receptor was upregulated to compensate for reduced Fgfr1 signaling (Fig. 7D,E). Furthermore, we analyzed the activation state (tyrosine phosphorylation) of Fgfr2 in p2 MECs and found that although both wild-type and mutant MECs had basal levels of tyrosine phosphorylated Fgfr2, the amount of active Fgfr2 was not increased in cells of either genotype in response to Fgf. Surprisingly, the basal (-Fgf) level of activated Fgfr2 was elevated in Fgfr1{Delta}Frs/{Delta}Frs MECs, and these cells responded to Fgf with a decrease in Fgfr2 phosphorylation (Fig. 7D). These data are not consistent with a role of Fgfr2 in compensatory activation of Frs and MAPK, and instead suggest that Fgfr1-Frs signaling transregulates Fgfr2 activity.

The MAPK signaling responses in p2 MECs did not directly translate into Fgf responsiveness in a proliferation assay: only two out of four mutant MEC cultures proliferated in response to Fgf, whereas all four responded with comparable MAPK and PI3K responses (Fig. 7A,F, data not shown). The growth curves did, however, reflect the relative phenotypic severity of embryos from which mutant cells were derived (data not shown). Variation in mutant MEC proliferative responses is probably due to differences in cell physiology or developmental staging between severely and mildly affected embryos. Nonetheless, the ability of two mutant lines to proliferate in response to Fgf indicates that Fgfr1-Frs signaling is not essential in directing this cellular response.

Impact of the Fgfr1{Delta}Frs mutation on MAPK signaling in vivo
Previous reports have implicated the MAPK pathway as an effector of Fgf signaling in vivo (Corson et al., 2003Go). Our primary MEC data support this model and suggest that the Fgfr1-Frs signaling is not required for Fgf-induced MAPK activation. To determine the physiological relevance of the MEC results, we examined MAPK activation in Fgfr1-dependent developmental contexts by whole-mount immunohistochemistry. First, we analyzed stage-matched embryos at E8.5-9.5, when the caudal region of the embryo is still undergoing gastrulation and more rostral axial levels are undergoing somitogenesis and neurulation. We found that phospho-MAPK staining in gastrulating mesoderm and somites was not altered in mutant embryos (Fig. 8A-F). In addition, we observed a thin line of phospho-MAPK staining along the medial edges of the neural folds, where neuroepithelial fusion takes place. This domain of MAPK activation was present in both mutant and control embryos (arrows, Fig. 8C,F), despite the disruption of neural tube closure in Fgfr1{Delta}Frs/{Delta}Frs embryos. Just after neurulation, when we observed an increase in cell death in mutant neural tubes, MAPK is not activated in wild-type or mutant neural tubes (data not shown). Thus, the cell survival role of Fgfr1-Frs signaling in the neural tube is probably facilitated by a different downstream pathway.


Figure 7
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  		Fig. 7. p2 MEC responses to Fgf. (A) pMAPK and pAkt responses of Fgfr1{Delta}Frs/{Delta}Frs and control MECs stimulated with Fgf, assayed over a dose titration (Ai, 5 minute stimulations) and a 1 hour timecourse (Aii). RasGAP, loading control. (B,C) Crk and Frs2 activation responses to Fgf (50 ng/ml aFgf, 5 µg/ml heparin, 5'), assayed by IP-phosphotyrosine (4G10) western blot. Blots were stripped and reprobed with anti-Crk or anti-Frs2 (lower panels). *No antibody IP control. (D) Fgfr2 responses to Fgf (50 ng/ml aFgf, 5 µg/ml heparin, 5'), assayed by 4G10 IP-Fgfr2 western blot (upper panel). Total cell lysate Fgfr2 (middle panel) and actin (loading control, lower panel) western blots show relative protein levels. Matching arrowheads indicate bands of the same size recognized by different antibodies within B-D. (E) Relative Fgfr2 mRNA levels in homozygous embryos, determined by semi-quantitative RT-PCR. (F) Growth curves of p2 MECs derived from four mutant and two wild-type embryos cultured in DMEM, 4% FBS, 50 ng/ml aFgf.

 
Later, during early PA2 development, phospho-MAPK immunostained dispersed cells throughout the pharyngeal arch region in both mutant and control embryos (Fig. 8G,J). As development progressed, phospho-MAPK became concentrated in control embryos at the rostral edge of PA1 and the caudal edge of PA2 (Fig. 8H). In mutant embryos at this stage, although PA1 immunostaining appeared normal, intensification of phospho-MAPK staining was not observed in caudal PA2 (Fig. 8K). This domain corresponds to the Fgf8 expression domain that is lost in mutants; however, the difference in MAPK activation observed in whole mount may be secondary to the failure of NCCs to populate this arch. Premigratory NCCs and NCCs within PA3 stained strongly with the phosphoMAPK antibody (Fig. 8I,L, data not shown).

Together, these results demonstrate that MAPK activity is not globally disrupted in Fgfr1-dependent contexts in Fgfr1{Delta}Frs/{Delta}Frsembryos. Thus, either Fgfr1 does not contribute significantly to cellular phosphoMAPK levels in these contexts, or Fgfr1 signaling to MAPK is not affected by the Frs site deletion.


   DISCUSSION
TOP
 SUMMARY
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Developmental roles of Fgfr1-Frs signaling
Our results demonstrate that Fgfr1-Frs signaling is essential during embryogenesis but is required only in a subset of Fgfr1-dependent contexts. Abrogation of the Fgfr1-Frs interaction does not disrupt Fgfr1 functions during gastrulation or somitogenesis, but does result in late embryonic lethality and defects in the development of the tail bud, neural tube, and pharyngeal arches. The specific array of phenotypes observed in Fgfr1{Delta}Frs/{Delta}Frs embryos is distinct from that observed in Fgfr1wtKI homo- or hemizygotes, previously described hypomorphs with reduced Fgfr1 levels, or embryos harboring an Fgfr1PLC{gamma} binding site mutation (Table 2, see Fig. S1 in the supplementary material) (Partanen et al., 1998Go; Xu et al., 1999Go; Trokovic et al., 2003aGo). This suggests that Fgfr1{Delta}Frs/{Delta}Frs phenotypes are not general consequences of reduced Fgfr1 activity, but are indeed due to disrupted signaling through the Frs interaction site. Interestingly, activating mutations in Fgfr1 that cause human craniosynostosis syndromes also have context-specific effects restricted mainly to craniofacial and limb development (Passos-Bueno et al., 1999Go). Thus, developmental contexts differ in their requirements for specific signaling pathways downstream of Fgfr1, as well as for regulation of Fgfr1 activity.


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  		Table 2. Comparison of Fgfr1 mutant phenotypes

 


Figure 8
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  		Fig. 8. Phospho-MAPK immunohistochemistry. Whole-mount phospho-MAPK immunohistochemistry of stage-matched embryos: (A-C) E8.5, (D-F) E9.5. High magnification views of (B,E) caudal gastrulating regions and (C,F) more rostral areas undergoing somitogenesis and neurulation. Arrowheads and arrows indicate phospho-MAPK-positive somites and neural folds, respectively. (G-L) G/J, H/K and I/L are stage-matched embryo pairs (E10.5). (G,H,J,K) Pharyngeal arch 1, 2 region; (I) dorsal and (L) lateral views of premigratory neural crest cells (arrows). Lateral view is shown in L because this embryo has an open neural tube.

 
Because Fgfr1{Delta}Frs/{Delta}Frs embryos exhibited rescue of Fgfr1null mesodermal phenotypes and early lethality, analysis of these signaling mutants enabled us to clarify later roles of Fgfr1 in neural tube and tail bud development. We thus identified a novel role of Fgfr1 in spinal neurulation. In contrast to craniorachischisis, which in Fgfr1{Delta}ex4/{Delta}ex4 embryos is probably secondary to mesodermal defects, Fgfr1{Delta}Frs/{Delta}Frs neural tube closure defects affect only the spinal region and are not accompanied by notochord or paraxial mesoderm defects. Our histological data demonstrate that Fgfr1-Frs signaling impacts the morphogenetic folding of the spinal neuroepithelium. Frs2 was previously implicated in convergent extension movements (Akagi et al., 2002Go), disruption of which can alter the size, shape and signaling properties of the notochord and neural plate, and thus result in neural tube closure defects. However, whereas previously described convergent extension mutants lack medial hinge points and exhibit notochord defects, Fgfr1{Delta}Frs/{Delta}Frsembryos form ectopic DHLPs and do not exhibit defects in mesoderm or medial bending, (Copp et al., 2003Go). Our results are therefore more consistent with a role of Fgfr1-Frs signaling in modulating neuroepithelial responsiveness to DLHP-inducing or -repressing signals.

The neural tube closure defect in Fgfr1{Delta}Frs/{Delta}Frs embryos may be exacerbated by the accompanying defect in tail bud development. Previously, posterior truncations in Fgfr1null animals were postulated to be secondary to mesoderm defects. However, the high penetrance of posterior truncations, together with the low incidence of mesodermal defects in Fgfr1{Delta}Frs/{Delta}Frs embryos suggest that Fgfr1 instead plays a direct, Frs-dependent role in tail bud development. The Frs site deletion did not affect patterning or proliferation in the tail bud during secondary gastrulation, but resulted in ectopic apoptosis throughout the caudal region of mutant embryos. Cell death could contribute to posterior truncations by depleting tail bud progenitors or hindering cell migration during secondary gastrulation.

In Fgfr1{Delta}Frs/{Delta}Frs embryos, Sox10-positive neural crest cells are hindered in their migration into the first and second pharyngeal arches, though at later stages, craniofacial defects in these embryos are restricted predominantly to PA2 NCC derivatives. This implies that the Sox10-positive population migrating towards PA1 recovers or is not essential for the development of some PA1 NCC derivatives. Specific disruption of PA2 and/or PA2 NCC development in Fgfr1{Delta}Frs/{Delta}Frs embryos contrasts with general Fgfr1 hypomorph phenotypes that affect NCC derivatives of both PA1 and PA2 (Trokovic et al., 2003aGo). Fgfr1-Frs signaling may be primarily required for PA2 patterning, or may participate in reciprocal signaling between the epithelium and NCCs that facilitates PA2 entry or survival of NCCs. In Fgfr1{Delta}Frs/{Delta}Frs embryos, an epithelial Fgf8 expression domain is lost in PA2, and the NCC migration defect is accompanied by an increase in cell death along the migration pathway. Preliminary results of conditional mutagenesis experiments indicate that ectopic cell death in the pharyngeal region reflects a NCC-nonautonomous effect of the Frs site deletion (R.V.H. and P.S., unpublished).

Potential mechanisms of Fgfr1 signaling during development
It has long been presumed that Fgfrs signal primarily through the MAPK pathway, and previous biochemical studies implicated Frs adaptors as important mediators of MAPK activation downstream of Fgfr1. However, we have demonstrated that the Fgfr1-Frs interaction is dispensable for early developmental roles of Fgfr1 and for activation of the MAPK pathway, both in primary embryonic cells and in vivo. Wild-type and Fgfr1{Delta}Frs/{Delta}Frs primary cell responses to Fgf stimulation were indistinguishable in terms of both dose sensitivity and signaling kinetics, and MAPK phosphorylation appeared normal in several Fgfr1-dependent contexts in Fgfr1{Delta}Frs/{Delta}Frs embryos. Furthermore, proliferative responses of non-immortalized Fgfr1{Delta}Frs/{Delta}Frs cells suggest that this pathway is not essential in driving Fgf-induced proliferation, although cells from embryos with more severe phenotypes were growth impaired. The discrepancy between our biochemical data and those published previously probably reflects cell type-specific signaling mechanisms, effects of Frs2 disruption on receptors other than Fgfr1, or effects of cellular transformation on the ability of other pathways to signal downstream of Fgfr1.

Our work, as well as previous studies of Fgfr1-PLC{gamma} signaling, demonstrates that individual signaling events downstream of Fgfr1 are required in a context-specific manner during development. In contexts where Fgfr1 functions are not compromised by the Frs site deletion, other pathways may drive cellular responses either independently of or additively with Frs adaptors. Aside from Frs2 and Frs3, only PLC{gamma}, Crk, Grb14 and Sef have been shown to interact directly with activated Fgfr1, although Src, STAT and Shc are also activated downstream of Fgfr1 in some cell types (Mohammadi et al., 1991Go; Klint and Claesson-Welsh, 1999Go; Larrson et al., 1999Go; Reilly et al., 2000Go; Tsang et al., 2002Go; Kovalenko et al., 2003Go). Sensitivity to the Frs site deletion could reflect the availability of alternate signaling pathways or the level of overall Fgfr1 signal required in a given context, rather than selective use of the Frs pathway. Thus, Fgfr1-Frs signaling may be used more broadly during development than is revealed by Fgfr1{Delta}Frs/{Delta}Frs phenotypes.

Fgfr1 and Fgfr2 can form heterodimers in vitro (Bellot et al., 1991Go). If such heterodimers were to form in vivo, Fgfr1{Delta}Frs signals could get shunted to the Frs pathway downstream of the Fgfr2 subunit in Fgfr1{Delta}Frs/{Delta}Frs embryos. This predicts that heterodimers would rescue Fgfr1{Delta}Frs function in contexts co-expressing Fgfr1 and Fgfr2, and that Fgfr2 would be activated by Fgf stimulation in primary cells. Our data do not support this model: Fgfr1{Delta}Frs/{Delta}Frsembryos exhibit defects in the tail bud and pharyngeal arches, where Fgfr1 and Fgfr2 are co-expressed, but not during gastrulation or somitogenesis when the receptors have distinct expression patterns (Orr-Urtreger et al., 1991Go; Yamaguchi et al., 1992Go; Walshe and Mason, 2000Go). Furthermore, we did not observe compensatory upregulation of Fgfr2 mRNA in vivo, or ligand-dependent activation of Fgfr2 in primary cells. Instead, basal levels of active Fgfr2 are notably elevated in Fgfr1{Delta}Frs/{Delta}Frs cells, suggesting that Fgfr1-Frs signaling transregulates Fgfr2. This may coordinate Fgfr responses and/or serve a homeostatic role in vivo. Deregulation of Fgfr2 activity may contribute in a gain-of-function manner to Fgfr1{Delta}Frs/{Delta}Frs phenotypes. Resolution of this issue will require further biochemical analysis and crosses between Fgfr1{Delta}Frs and Fgfr2 mutant mouse lines.

Supplementary material
Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/133/4/663/DC1


   ACKNOWLEDGMENTS
 
We thank Janet Rossant for providing Fgfr1 genomic clones; Mitch Goldfarb for advice on the Frs site deletion; Philip Corrin, Jason Frazier and Marc Grenley for excellent technical assistance; and Raj Kapur, Henk Roelink and our laboratory colleagues for critical reading of the manuscript. This work was supported by an NSF predoctoral fellowship and the CMB Training Grant (NIH #GM07270) to R.V.H., and by grants HD 24875 and HD 25326 to P.S.


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