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. Author manuscript; available in PMC: 2015 Jul 24.
Published in final edited form as: Cell Rep. 2014 Jul 17;8(2):596–609. doi: 10.1016/j.celrep.2014.06.026

The Extreme Anterior Domain Is an Essential Craniofacial Organizer Acting through Kinin-Kallikrein Signaling

Laura Jacox 1,2,3,4,5,*, Radek Sindelka 1,6,*, Justin Chen 1,2, Alyssa Rothman 1, Amanda Dickinson 1,7, Hazel Sive 1,2,a
PMCID: PMC4135435  NIHMSID: NIHMS607720  PMID: 25043181

SUMMARY

The extreme anterior domain (EAD) is a conserved embryonic region that includes the presumptive mouth. We show that the Kinin-Kallikrein pathway is active in the EAD and necessary for craniofacial development in Xenopus and zebrafish. The mouth failed to form and neural crest (NC) development and migration was abnormal after loss of function (LOF) in the pathway genes kng, encoding Bradykinin (xBdk), carboxypeptidase-N (cpn) that cleaves Bradykinin and neuronal nitric oxide synthase. Consistent with a role for nitric oxide (NO) in face formation, endogenous NO levels declined after LOF in pathway genes but these were restored and a normal face formed after medial implantation of xBdk-beads into LOF embryos. Facial transplants demonstrated that Cpn function from within the EAD is necessary for migration of first arch cranial NC into the face and to promote mouth opening. The study identifies the EAD as an essential craniofacial organizer acting through Kinin-Kallikrein signaling.

INTRODUCTION

The face derives from both neural crest and non-neural crest derivatives. The presumptive mouth arises from a conserved extreme anterior domain (EAD) where ectoderm and endoderm are juxtaposed (Dickinson and Sive 2006). The cranial neural crest (NC) migrates into the future facial region to abut the EAD (Dickinson and Sive 2007, Spokony 2002) during tailbud stages in Xenopus. At mouth opening, the cranial NC has begun differentiating into cranial nerves, melanocytes, connective tissue, and chondrocytes that contribute to the jaws and other facial bones (Fabio and Filippo 2003). The EAD expresses signaling regulators (Dickinson and Sive 2009), which suggested that the region might act as a facial organizer. We addressed this possibility using transplant assays where EAD lacking the secreted Wnt regulators, Frzb1 and Crescent replaced the EAD of a control embryo. Not only did the mouth fail to form but surrounding facial regions appeared abnormal, suggesting more global activity of the EAD. However, this putative organizer activity was not extensively explored for other factors impacting mouth formation and cranial NC migration.

Molecular rules for NC movement have been extensively described, and include contact inhibition of locomotion, co-attraction, chase-and-run strategies (Theveneau et al. 2013), and guidance through interaction with extracellular matrix, semaphorins, and Eph/Ephrin signals (Mayor and Theveneau 2013). Despite these elegant conclusions, the mechanisms that direct the cranial neural crest into the face primordium, and the identity of localized guidance signals that facilitate this migration are not known.

In a microarray screen to identify regulatory genes expressed in the EAD that may regulate mouth and other aspects of face formation, we isolated carboxypeptidase N (cpn), kininogen (kng), and neural nitric oxide synthase (nNOS). These genes are members of the Kinin-Kallikrein pathway (Kakoki and Smithies 2009) a regulator of blood pressure (Sharma 2009) that also participates in inflammation (Bryant 2009) and renal function. This pathway had not been described as necessary for craniofacial development in any animal. In the adult mammalian Kinin-Kallikrein pathway (Fig. 1A), Kallikrein, a protease, cleaves Kng to yield Bradykinin, a 9 amino acid (9AA) peptide. Bradykinin is a vasodilator that binds the Bradykinin B2 (BKB2) G-protein coupled receptor. BKB2 receptor activates NOS, which converts L-Arginine (Arg) to nitric oxide (NO) and citrulline. Bradykinin can also be cleaved by CPN, yielding 8AA desArg-Bradykinin and Arg that can be converted to NO (Moncada and Higgs 1995). The BKB2 receptor is constitutively expressed in adult mammals and binds Bradykinin, but not desArg-Bradykinin, to activate NOS (Kakoki and Smithies 2009). A BKB1 receptor is conditionally expressed during inflammation and binds desArg-Bradykinin but not Bradykinin. Angiotensin Converting Enzyme (ACE) degrades both Bradykinin and desArg-Bradykinin.

Figure 1. Mammalian Kinin-Kallikrein pathway and putative pathway genes are expressed in the developing face.

Figure 1

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(A) Adult mammalian Kinin-Kallikrein pathway (Kakoki and Smithies 2009). In situ hybridization for kininogen (kng) (B, B′, E, E′), cpn(C, C′, F, F′), and nNOSRNA (D, D′, G, G′) (RNA is purple). Cement gland marker (xcg) is red. Arrow: presumptive mouth. cg, cement gland. B–G frontal views; B′–G′sagittal sections. Scale bars: 200μm.

In addition to its role in the Kinin-Kallikrein pathway, NO participates in multiple processes including wound healing, tissue regeneration (Filippin 2011), angiogenesis (Cook 2003), neurotransmission (Contestabile, 2004) and possibly malignancy (Olson 2008). NO has been implicated in developmental contexts including neuronal development (Bradley et al. 2010), bone growth regulation, (Yan et al. 2010), cardiac endothelial-to-mesenchymal transition (Chang et al. 2011), control of organ size and developmental timing (Kuzin et al. 1996). Elevated NO production has been found in developing epithelial tissues, ganglia, and the notochord (Lepiller et al., 2007). In Xenopus, NO is a potent parthenogenetic activator of Xenopus eggs (Jeseta 2012), and is correlated with movement in tadpoles (Mclean 2000).

The strong expression of kng, cpn, and nNOS in the EAD led us to hypothesize that the Kinin-Kallikrein pathway is active during embryogenesis and required for facial development. We present data that support this hypothesis, and additionally show that Kinin-Kallikrein signaling localized to the EAD is necessary for movement of the first arch cranial NC into the face, and for mouth formation. The study identifies the EAD as an essential craniofacial organizing center acting through Kinin-Kallikrein signaling.

RESULTS

kininogen (kng), carboxypeptidase N (cpn) and neural nitric oxide synthase (nNOS) are expressed in the EAD during initial stages of craniofacial development

kng, cpn, and nNOS expression was identified in the Xenopus EAD region (Dickinson and Sive 2009; and Fig. S1A), suggesting activity of an embryonic Kinin-Kallikrein pathway (Fig. 1A). Protein alignment showed high conservation of Cpn and nNOS (Fig. S1B–D). Gene expression was examined by in situ hybridization and quantitative RT-PCR (qPCR) (Fig. 1 B–G; Fig. S1E–G). At tailbud (st. 20 and 26) when the EAD is present and cranial NC is migrating, kng is expressed in the prechordal plate with anterior expression adjacent to the EAD (Fig. 1 B, B′, E, E′). At st. 20, cpn was expressed in deep EAD layers (Fig. 1 C, C′, F, F′) and by st. 26 at low intensity in the first branchial arch (Fig. 1F). nNOS RNA is present in outer ectoderm of the face, excluding hatching and cement glands (Fig. 1D, D′, G, G′). Later, nNOS is expressed in the head and notochord (Peunova et al. 2007). These data show that putative Kinin-Kallikrein pathway genes are simultaneously expressed in adjacent regions of the presumptive face.

Putative Kinin-Kallikrein pathway genes are required for mouth formation and neural crest development

A requirement for kng, cpn, and nNOS during craniofacial development would be consistent with activity of the Kinin-Kallikrein pathway. This was tested by loss of function (LOF) using injection of morpholino-modified antisense oligonucleotides (morpholinos, MOs) at the one cell stage. Specificity of MO targeting was demonstrated by using two MOs, or more importantly, by “rescue” assays where a normal phenotype was observed when MO was co-injected with cognate mRNA lacking the MO target site (Fig. S2A–D′, B″). For kng and nNOS MOs targeting splice sites, qPCR showed a strong decrease in endogenous RNA levels (Fig. S2E, F). At late hatching stage (st. 40), LOF animals (“morphants”) displayed abnormal body morphology and no open mouth, with a small stomodeal invagination (Fig. 2A–D′, bracket). Nostrils were absent, eyes were small, pigment was reduced and the face was narrow (Fig. 2A′–D′). Morphant phenotypes were apparent at early tailbud (st. 22, Fig. S3A–L′), and were accompanied by elevated cell death but normal proliferation (Fig. S3M–V). Despite abnormal mouth phenotypes, the EAD was correctly specified as shown by expression of frzb1 and xanf1 (Fig. 2E–H‴).

Figure 2. kng, cpn and nNOS are required for mouth opening and face formation.

Figure 2

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(A–D, A′–D′) kng, cpn, and nNOS loss of function (LOF) using antisense morpholinos. Embryos assayed at stage 40, four experiments. Arrow: mouth region. Bracket: Unopened mouth. cg, cement gland. Scale bar (A–D): 2000μm. Scale bar (A′–D′): 200μm. (A, A′) Control morphants (100% normal, n=97). (B–D, B′–D′) kng, cpn, and nNOS morphants (kng (B′) 0% normal, n=102; cpn(C′) 2% normal, n=105; nNOS(D′) 0% normal, n=129). (E –H, E′–H′, E″–H″, E‴–H‴) Kinin-Kallikrein pathway morphants at stage 22 express presumptive mouth markers, frzb1 and xanf1. Scale bar: 200μm. (I–P; I′–L′) Histology of kng, cpn, and nNOS LOF. Coronal sections (I–L, control morphant 100% normal, n=5; each Kinin-Kallikrein morphant, 0% normal, n=9) assayed at stage 26 in 2 independent experiments with β-catenin immunolabeling. Parasagittal sections with anterior to the left (I′–L′, control morpholino 100% normal, n=5; each Kinin-Kallikrein pathway morpholino, 0% normal, n=12). Parasagittal sections with Laminin immunolabeling (M–P) assayed at stage 26 in 2 independent experiments (control morphant 100% normal, n=10; each Kinin-Kallikrein morphant 8% normal, n=12). β-catenin: green. Laminin: green. Nuclear propidium iodide: red. Bracket: presumptive mouth region. cg, cement gland. Scale bars: 170 μm. (Q–X, Q′–X′) kng, cpn, and nNOS morphants showed reduced expression of neural crest markers sox10 and sox9. (Q–T, Q′–T′) sox10 in situ hybridization at stage 22. (U–X, U′–X′) sox9 in situ hybridization at stage 26. Bracket: cranial NC-free midline region. Arrow: normal extent of 1st arch cranial NC. Scale bars (Q–X): 200μm. Scale bar (Q′–T′): 800μm. Scale bar (U′–X′): 400μm. (Y-d) kng morphants showed reduced expression of slug at stage 22, while cpn and nNOS morphants and kng morphants coinjected with kng mRNA showed control slug levels. Arrow: specified neural crest. Dorsal view. Scale bars: 800μm. (e–l) Cell proliferation and death in cranial NC cells. (e–g) sox10 in situ hybridization at stage 22 in axial section. Control and cpn morpholino plus cpn mRNA embryos showed normal expression, while cpn morphants had reduced expression. Scale bar: 200μm. (h) Schematic demonstrating axial section. (i–k) Ph3 staining of axial sections show increased positive cells in control (i) relative to cpn morphants (j). Embryos injected with cpn morpholino plus cpn mRNA (k) had more Ph3 positive cells than cpn morphants. Scale bar: 170μm. (l–m) Quantification of Ph3 and TUNEL staining, with standard deviation included. P value: one-way anova with multiple comparisons.

To understand LOF defects, we analyzed tailbud embryos (st. 26) for β-catenin indicating adherens junctions, and Laminin indicating basement membrane using immunostaining. In coronal (frontal) sections, controls displayed a narrow midline strip of β-catenin positive cells running from brain to cement gland, 2–4 cells wide (Fig. 2I). However, in morphants this strip was 6–8 cells wide, indicating abnormal epithelial organization (Fig. 2J–L), also apparent in parasagittal sections (Fig. 2I′, bracket) where morphants showed a deep region of β-catenin-positive tissue (Fig. 2J′–L′). Laminin expression was largely absent in morphants in the basement membrane extending from brain to cement gland and separating epidermis and deep ectoderm (Fig. 2M–P, arrows). These data demonstrate epithelial and basement membrane abnormalities after kng, cpn and nNOS LOF at tailbud.

Reduction of pigment and narrow faces in morphants suggested cranial NC may be abnormal, and consistently, RNA expression of cranial NC markers sox9 and sox10 (Mori-Akiyama et al. 2003) was reduced at early tailbud (st. 22) and at late tailbud (st. 26) (Fig. 2Q–X′) as assayed by in situ hybridization. This was confirmed by qPCR, with > 50% reduction in RNA levels (data not shown). Frontal views of control embryos at st. 26 showed a midline strip negative for NC markers (Fig. 2U, bracket) that was not apparent or wider in morphants (Fig. 2V–X). These data suggest cranial NC induction, survival, proliferation, or migration is abnormal.

To assay NC induction in morphants, expression of slug (LaBonne and Bronner 1998) was examined at early neurula (st. 15) (Fig. 2Y-b). While nNOS and cpn morphants displayed normal slug expression (Fig. 2Y–Z, c–d), kng morphants showed a decrease that was prevented by co-injection of kng mRNA (Fig. 2a–b). Since cpn morphants show normal NC induction but a later deficit in NC marker expression, morphants were analyzed for alterations in proliferation and cell death. Axial sections of sox10 in situ embryos confirmed NC identity (Fig 2e–h). PH3 labeling demonstrated 50% reduction in mitotic cells (Fig. 2i–l) and TUNEL demonstrated a 100% increase in dying cells in cpn morphants relative to controls (Fig. 2m) that was partially prevented by co-injecting cognate mRNA (Fig. 2k–m). The data show a requirement for kng, cpn and nNOS during craniofacial development, including mouth opening. After LOF, multiple changes are observed, in epithelial organization and NC induction, proliferation or survival, consistent with an active embryonic Kinin-Kallikrein pathway.

kng and cpn LOF phenotypes are prevented by Xenopus Bradykinin peptides

In the adult, the Kng precursor is processed to release a 9AA peptide, Bradykinin (Bdk) and desArg-xBdk, an 8AA peptide, after cleavage by Cpn. Xenopus Bdk (xBdk) peptide was predicted by aligning Kng protein sequence across species and identifying putative Kallikrein cleavage sites (Fig. 3A) (Borgoño 2004). Considering the adult mammalian pathway, we predicted that both the 9AA and 8AA peptides should prevent the kng LOF phenotype, whereas only the 8AA peptide should prevent the cpn LOF phenotype (Fig. 1A). Beads soaked in peptides were implanted medially in the future facial region of kng or cpn LOF embryos at tailbud (st. 22), which were scored at tadpole (st. 40) for mouth and facial phenotypes (Fig. 3B). Relative to a scrambled xBdk peptide (Fig. 3C, F), 9AA and 8AA peptides prevented the kng morphant phenotype (Fig. 3D, E, I), as predicted. In cpn morphants, mouth opening was restored by 8AA but not by 9AA peptide, consistent with the adult model (Fig. 3G–I). However, both peptides restored normal pigment, overall facial symmetry and head size to cpn morphants (Fig. 3I).

Figure 3. Bradykinin-like peptides prevent cpn and kng loss of function phenotypes.

Figure 3

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(A) amino-acid sequence alignment of region around Bdk-l peptide. Grey highlights: Bdk-l peptide sequence. Red: conserved amino acids. Black arrows: Kallikrein and Cpn cleavage sites. Bdk-l (9AA) and Des-Arg Bdk-l peptides (8AA) used. (B) Experimental design. (C–H) Abnormal mouth phenotype after kng LOF prevented by 9AA and 8AA peptides, while in cpn morphants was prevented only by the 8AA peptide. P values: one-tailed Fisher Exact test. (C) kng morphants implanted with 9AA scrambled (9AAscr) peptide bead (28% normal, n=60). Embryos scored as abnormal if mouth failed to open, was tiny or asymmetric, nostrils failed to form, pigment was absent, or face was abnormally narrow. (D, E) kng morphant implanted with 9AA (D, 60% normal, n=105) or 8AA bead (E, 57% normal, n=75). (F) cpn morphants implanted with 9AAscr bead (mouth- 43% normal, n=67; face- 27% normal, n=67). (G, H) cpn morphants implanted with 9AA bead (mouth- 41% normal, n=54; face- 44% normal, n=54) or 8AA bead (mouth- 65% normal, n=79; face- 51% normal, n=79). Scale bar: 200μm. (I) Graph depicting percent of morphants implanted with beads, displaying normal mouth and face formation. (J–L) Expression of neural crest marker sox9 in kng morphants implanted with 9AA bead. Arrow: normal extent of 1st arch cranial NC. (J) Wild-type expression of sox9 (100% normal, n=13). kng morphant with a 9AAscr bead (K, 8% normal, n=12) and 9AA bead (L, 39% normal, n=13). Scale bar: 200μm.

To investigate whether xBdk peptide could restore NC development after kng LOF, 9AA scrambled or xBdk soaked-beads were implanted medially (Fig. 3J–L) or anterolaterally below the eye (Fig. S4A–C′) at stage 22, and sox9 expression was later assayed. Normal sox9 expression was observed with 9AA xBdk beads (Fig. 3J–L). Consistent data were obtained with lateral implants (Fig. S4B–C′), however these failed to rescue mouth formation at stage 40 (Fig. S4D–F). These data support the hypothesis that xBdk peptides derived from Kng direct mouth and NC formation.

Nitric oxide (NO) prevents kng, cpn and nNOS LOF phenotypes and endogenous NO production is regulated by xBdk

In mammals, the Kinin-Kallikrein pathway leads to production of the signaling molecule NO. We therefore hypothesized that LOF phenotypes would be prevented by application of the NO donor S-Nitroso-N-Acetyl-D, L-Penicillamine (SNAP). SNAP was co-injected with MO at the one cell stage or injected into the face at late neurula (st. 20). When applied at the one cell stage, SNAP prevented craniofacial and whole-body phenotypes (Fig. 4A–D′; Fig. S5A–G, B′–D′, J), and corrected β-catenin and Laminin localization and sox9 expression (Fig. 4E–P′). When injected into the presumptive facial region, SNAP improved facial development (Fig. S5A–D′) indicating NO can act at later stages. This rescue was not due to a general effect on all MOs, since the par1 phenotype (Ossipova 2005) was not prevented by SNAP (Fig. S5H–J). Consistent data were obtained using the NO antagonist TRIM applied at st. 20, resulting in abnormal mouth, face and sox9 expression (Fig. 4Q–R′). Although the Kinin-Kallikrein pathway has a role in angiogenesis (Westermann 2008), craniofacial phenotypes did not result from altered blood flow, shown by a head extirpation assay. Thus, an open mouth developed in isolated heads lacking a heart and cultured from pre-hatching st. 31–32, before heartbeat until st. 41 (swimming tadpole) (Fig. 4S–T′).

Figure 4. kng, cpn and nNOS loss of function phenotypes are prevented by the NO-donor, SNAP, and Kinin-Kallikrein morphants show reduced nitric oxide production that is increased by xBdk.

Figure 4

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(A–D, A′–D′) Facial morphology of kng, cpn, and nNOS loss of function (A–D) and with SNAP (A′–D′). Embryos assayed at stage 40 in 3 independent experiments, and scored as abnormal if mouth failed to open, was tiny or asymmetric, nostrils failed to form, pigment was absent, or face was abnormally narrow. Arrow: mouth region. Bracket: unopened mouth. cg, cement gland. (A) Control MO injected (98% normal, n=427) (B–D) kng, cpn or nNOS MO injected. (A′) SNAP plus control MO. (B′–D′) kng, cpn or nNOS MO plus SNAP coinjection (kng (B′) 85% normal, n= 105; cpn (C′) 86% normal, n=98; nNOS (D′) 90% normal, n=87). Scale bar: 100μm. (E–L, E′–L′) Histology of kng, cpn, and nNOS LOF embryos after SNAP treatment. Parasagittal sections with anterior to left assayed at stage 26 with β-catenin (E–H′) and Laminin immunolabeling (I–L′). β-catenin: green. Laminin: green, with nuclear propidium iodide, red. cg, cement gland. (E–H, E′–H′) β-catenin in control embryos (E, E′), LOF embryos (F–H), and LOF embryos co-injected with SNAP (F′–H′) (kng (F′) 100% normal, n= 5; cpn (G′) 100% normal, n=5; nNOS (H′) 100% normal, n=5). (I–L, I′–L′) Laminin staining in control embryos (I, I′), LOF embryos (J–L), and LOF embryos co-injected with SNAP (kng (J′) 75% normal, n= 4; cpn (K′) 80% normal, n=5; nNOS (L′) 100% normal, n=4). Scale bars: 170μm. (M–P, M′–P′). Expression of sox9 RNA (in situ hybridization) after SNAP injection into kng (N), cpn (O), and nNOS (P) LOF embryos. Lateral view. Scale bar: 100μm. (Q–R, Q′–R′) NOS inhibitor TRIM prevents mouth formation and reduces sox9 expression. (Q, Q′) Wild-type embryos (100% normal, n=6). (R, R′) TRIM treated embryos (17% normal, n=6). (Q–R) Frontal view at stage 40. (Q′–R′) Lateral view of sox9 in situ hybridization at stage 26. Scale bar (Q–R): 100μm. Scale bar (Q′–R′): 400μm. (S–T, S′–T′) Extirpated heads show open mouth and normal pigmentation at swimming tadpole (st. 41). (S, S′) Control heads (96% normal, n=27). (T, T′) Isolated heads (92% normal, n=26). (S–T) frontal view. (S′–T′) side view. Scale bar: 100μm. (U–X) Fluorescence after incubation with NO sensor DAF-FM in control embryos (U), kng (V), cpn (W) and nNOS (X) LOF embryos. cg, cement gland. Sagittal view. Scale bar: 170μm. (Y-c) Control morphant with no bead (Y). kng morphant with no bead (Z), with 9AA xBdk scrambled bead (a) or 9AA xBdk bead (b). Images collected with same exposure, gain, and fluorescent illumination. kng morphants implanted with 9AA xBdk bead displayed 50% of control florescence compared with 23% of control fluorescence in morphants treated with 9AAscr xBdk peptide. Frontal view. Scale bar: 100μm. (c) Graph showing morphant fluorescence as percentage of control fluorescence; cpn morphants: 49%, kng morphants: 24%, and nNOS morphants: 64%. Each dot represents average of 3 biological replicates from independent experiments. P values: one-tailed T test.

If NO mediates craniofacial development, it should be detectable in developing facial regions and decrease after kng, cpn and nNOS LOF. NO was measured by incubating late neurula (st. 20) embryos with DAF-FM diacetate, which emits green fluorescence after reacting with NO. Tailbud (st. 26) control embryos showed fluorescence in the outer epidermis (Fig. 4U), where nNOSis strongly expressed. Diminished fluorescence was seen and quantified in kng, cpn and nNOS LOF embryos (Fig. 4V–X, c). nNOS LOF was associated with the smallest reduction in NO production, perhaps due to other NOS isoforms. We predicted that xBdk peptides would increase NO production (Fig. 1A) and this was confirmed by implanting xBdk-beads into the presumptive mouth region of kng morphants (Fig. 4Y-c). These data demonstrate production of NO in the EAD is dependent on Kinin-Kallikrein gene function, occurs during facial development and is responsive to xBdk.

cpn is expressed in the EAD, is required locally for mouth opening and modulates Arginine levels

Based on LOF phenotypes, we hypothesized that kng, cpn and nNOS function in the EAD is locally required in the presumptive mouth and globally required for cranial NC development. This was tested by transplanting the EAD from kng, cpn and nNOS LOF embryos at early tailbud (st. 22) into sibling controls (Fig. 5A) (Jacox et al. 2014). Control transplants led to normal mouth opening, nostril formation and pigmentation (Fig. 5B, B′, and quantified in Fig. 5F). Strikingly, when cpn LOF EAD was transplanted into control embryos, open mouths or nostrils failed to form, heads were narrow and lacked pigment, similar to global cpn LOF (Fig. 5C, C′). In contrast, transplant of nNOS and kng LOFEAD into control embryos led to milder phenotypes (Fig. 5D, D′, E, E′), consistent with the highly preferential expression of cpn in the EAD, and more widespread expression of kng and nNOS. We further showed that cpn expression in the EAD is required for cranial NC formation since sox9 expression at late tailbud is abnormal and reduced after EAD cpn LOF transplants (st. 28, Fig. 5G–H′).

Figure 5. Local cpn expression is required for mouth opening.

Figure 5

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Local requirement of kng, cpn and nNOS expression tested with EAD transplants. (A) Experimental design: donor morphant tissue was transplanted to uninjected sibling recipients. (B–E, B′–E′) EAD transplant outcome from control, cpn, kng, or nNOS morphant donor tissue (control (B) 100% normal, n=11; cpn (C) 28% normal, n=14; kng (D) 83% normal, n=24; nNOS (E) 61% normal mouth phenotype, 72% normal facial phenotype, n=18). (B′–E′) Overlay of (B–E) with GFP fluorescence indicating donor tissue. Dots surround open mouths. Bracket: unopened mouth. Frontal view. Scale bar: 100μm. (F) Quantification of structure depending on morphant background of facial tissue. (G–H, G′–H′) sox9 expression in cpn morphant donor tissue transplants, compared with control morphant transplants. sox9 in situ hybridization in control morphant transplants (G, G′, 70% with normal expression, n=10) and cpn morphant transplants (H, H′, 36% with normal expression, n=11). Two representative embryos shown. Scale bar: 100μm. (I, J) (I) Summary of urea assay for analysis of Cpn activity. (J) Chart summarizing level of urea derived from free-Arg in cpn morphants or morphants co-injected with cpn RNA, as percent of urea derived from free-Arg in control morphants. Urea levels in control morphants and wild-type embryos were equivalent. Each dot represents an independent experiment. P value: one-tailed T test.

The activity of Cpn predicts it modulates levels of Arg (Fig. 1A). To examine this, we used a quantitative assay where Arg is converted into urea whose levels can be measured (Fig. 5I). As hypothesized, after cpn LOF, lower levels of urea relative to control embryos were present. Specificity was demonstrated as urea levels increased after injection of cpn mRNA into LOF embryos (Fig. 5J). Together, these data indicate a requirement for Cpn activity localized in the EAD during mouth, cranial NC and face development.

Localized cpn activity in the EAD is necessary for migration of first arch neural crest into the face

The reduction in sox9 expression with cpn LOF suggested that cpn expression is required for NC migration. To analyze migration, fluorescent cranial NC was transplanted into control or cpn morphant hosts at neurula (st.18) and scored at late tailbud (st. 28) (Fig. 6A). While control transplants displayed three or four distinct branchial arches at late tailbud (st. 28) (Fig. 6B, B′), control NC transplanted into cpn morphants failed to segregate into branchial arches and did not migrate (Fig. 6C, C′), indicating a requirement for Cpn in cranial NC migration.

Figure 6. Global and local cpn expression is required for cranial neural crest migration.

Figure 6

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Global requirement for cpn expression tested with cranial NC transplants. Embryos scored as normal if three or four distinct branchial arches formed and migrated normally. (A) Experimental design: donor wild-type cranial NC transplanted into cpn morphant sibling recipients. (B–C, B′–C′) (B, C) Cranial NC transplant outcomes in control and cpn morphant recipients with GFP fluorescence overlay, indicating location of donor transplant at stage 28 (control (B) 69% Normal, n=36; cpn (B′) 27% normal, n=29). (B′, C′) GFP fluorescence of cranial NC in control and cpn morphant recipient. Numbers indicate branchial arches. Side view. cg, cement gland. Scale bar: 200μm. (D) Experimental design: donor cpn morphant EAD transplanted into control morphant sibling recipients with fluorescent cranial NC. (E–H, E′–H′, E″–H″) (E, F, G, H) Bright field view of control and cpn morphant transplants at stages 28 and 40. (E′, F′, G′, H′) Cranial NC in control and cpn morphant EAD recipients at stages 28 and 40 with GFP fluorescence overlay, indicating location of cranial NC and mCherry fluorescence overlay, indicating location of EAD transplant. (control st. 28 (E′) 85% Normal, n= 41; cpn st. 28 (F′) 57% Normal, n= 42; control st. 40 (G′) 63% Normal, n= 38; cpn st. 40 (H′) 17% Normal, n=35). (E″, F″, G″, H″) GFP fluorescence of cranial NC in control and cpn morphant EAD recipients at stage 28 and 40. Arrow: Open mouth. Bracket: unopened mouth. Frontal view. cg, cement gland. Scale bar = 100μm. (I–J, I′–J′) (I–J) Cranial NC outcome in control and cpn morphant EAD recipients with GFP fluorescence overlay, indicating location of cranial NC at stage 28. (I′, J′) GFP fluorescence of cranial NC in control and cpn morphant EAD recipients. Numbers indicate branchial arches. Side view. cg, cement gland. Scale bar: 200μm. (K–L, K′–L′) Cartilage in control morphant EAD recipients (K, K′ 78% normal, n=14) and cpn morphant EAD recipients (L, L′ 6% normal, n=16). (K, L) Ventral view. (K′, L′) Dorsal view. M, Meckel’s cartilage. C, ceratohyal cartilage. Scale bar: 100μm.

We extended this to ask whether local cpn expression is required for cranial NC migration, using double NC and EAD transplants, where control cranial NC was first transplanted into control embryos, followed by a control or cpn morphant EAD transplant (Fig. 6D). Relative to controls, (Fig. 6E-E″, I-I′), embryos with a cpn LOF EAD showed reduced NC migration at late tailbud (st. 28) (Fig. 6F–F″, J-J′). In particular, first arch NC showed highly reduced migration anteriorly and medially (Fig. 6J-J′), demonstrating that cpn expression in the EAD is necessary to guide the cranial NC into the face. At tadpole (st. 40), control transplants developed a normal mouth and face with extensive NC-derived tissue (Fig. 6G-G″) and a normal cartilaginous skeleton (Fig. 6K-K′). However, cpn EAD LOF transplants failed to form normal mouths or faces (Fig. 6H-H′), had substantially less NC-derived tissue (Fig. 6H″) with deformed Meckel’s and ceratohyal cartilages (Fig. L-L′). These data demonstrate that local Cpn activity in the EAD is required for migration of the first branchial arch into the face, putatively through processing of Kng-derived peptides.

Conservation of kng function during craniofacial development in zebrafish

To investigate whether function of kng in face formation is conserved, we used antisense MOs to target zebrafish (Danio) kng and assayed facial cartilages in 5-day post fertilization embryos by alcian blue staining (Fig. 7A–E′, F; Fig. S6A–C′, D). The MO used targets the kng1 isoform, the only transcript that includes the 9AA Bdk-l peptide. Zebrafish kng is expressed during NC development and mouth opening (Fig. S6E–F). kng LOF led to abnormally shaped Meckel’s and ceratohyal cartilages, or abnormal spacing between Meckel’s cartilage and the ethmoid plate. As in Xenopus, LOF led to absence of an open mouth (Fig. 7G–I). The LOF phenotype was prevented by co-injection of zebrafish kng that does not bind the MO(Fig. 7D, D ′) or by human KNG RNA indicating specificity (data not shown). Morphants injected with Xenopus laeviskng RNA showed no rescue (Fig. 7E, E′) consistent with the greater identity between human and zebrafish Bradykinin than with Xenopus (Fig. 3A).

Figure 7. Function of kng in craniofacial development is conserved in zebrafish.

Figure 7

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(A-A′) Camera lucida of facial cartilages. E: Ethmoid plate. C: Ceratohyal cartilage. M: Meckel’s cartilage. (B–E, B′–E′) kng loss of function using splice morpholinos and rescue with zebrafish (zf) kng mRNA. Embryonic cartilage scored at 5dpf after alcian blue staining in three independent experiments. Scale bar: 250μm.(B, B′) Control morphants coinjected with mRNA were normal (88% normal, n=50). (C-C′, E-E′) kng morphants and kng morphants coinjected with 200ng Xenopus kng mRNA showed abnormal facial cartilage. Meckel’s cartilage was truncated, boxy, and pointed at an abnormal angle. The ceratohyal cartilage was positioned at an abnormal angle, perpendicular to the midline. (kng (C, C′) 3% normal, n=61; kng mo plus frog mRNA (E, E′) 0% normal, n=65). (D, D′) kng morphants coinjected with 200ng zebrafish (zf) mRNA showed partial rescue. Embryos scored as partially rescued if Meckel’s cartilage was longer, more rounded, and pointed dorsally and if ceratohyal cartilage pointed more anteriorly, compared to kng morphants (54% partial rescue, n=89). (F) Quantification of phenotypes. P-values: one-tailed Fisher Exact test. N, Normal or partially rescued phenotype. A, Abnormal phenotype. (G–I) Ventral views of mApple-injected embryos at 48hpf. White arrow: open mouth. White bracket: closed mouth. Scale bar: 100 μm. (G) Control morphants (100% normal, n=5). (H) kng splice morphants failed to form open mouths (0% normal, n=6). (I) kng splice morphants coinjected with 200ng zf mRNA had open mouths (67% normal, n=6). (J–Q, J′–Q′) Confocal images of Sox10::GFP zebrafish coninjected with 75pg mApple and 4ng control morpholino (100% normal, n=5) or 4ng kng splice morpholino (0% normal, n=5). Paired images of the same embryo show GFP signal alone and GFP with mApple. Numbers indicate pharyngeal arches (PA). Bracket: uncondensed/disorganized cartilage. Lateral view. M, Meckel’s cartilage. Scale bar: 100μm. (J–K, J′–K′) At 36hpf, NC has migrated into the face of both morphant and control embryos to form 1st and 2nd PA. (L–M, L′–M′) At 48hpf, the 1stPA has begun to extend under eye to form the lower jaw in both morphant and control embryos. (N, N′) At 60hpf, 1stPA has condensed into Meckel’s cartilage in control embryos. (O, O′) At 60hpf, 1stPA remains disorganized in morphants and does not condense. (P, P′) At 72 hpf, Meckel’s cartilage is prominent in control embryos. (Q, Q′) At 72hpf, cartilage of the lower jaw remains disorganized and uncondensed in morphants.

Sox10::GFP transgenic fish were used to observe NC specification and migration after kng LOF. In both controls and morphants, NC was properly specified at the 10-somite stage (not shown) and migration to form the 1st and 2nd pharyngeal arches was normal until 48 hpf (Fig. 7J–Q′). However, by 60 hpf, Meckel’s cartilage, derived from the 1st pharyngeal arch, fails to condense in morphants (Knight and Schilling, 2006). We conclude that zebrafish kng is necessary for NC and mouth development, demonstrating a conserved requirement for Kinin-Kallikrein signaling. The phenotypes observed in zebrafish are apparent at a later stage than observed in Xenopus, indicating temporal control of facial development by Kinin-Kallikrein signaling may differ between species.

DISCUSSION

This study demonstrates activity of the Kinin-Kallikrein pathway during embryogenesis and localized control of craniofacial development through this pathway for the first time in any species. Three major conclusions are reached. First, the embryonic pathway in Xenopus functions through a signaling sequence similar to that described for the adult mammalian pathway, and conservation is present in zebrafish. Second, nitric oxide (NO) production is an outcome of the pathway and is necessary for mouth and neural crest development. Third, the extreme anterior domain (EAD) functions as a craniofacial organizer and facilitates migration of first arch cranial neural crest (NC) into the face via Kinin-Kallikrein signaling. These findings add insight into localized signaling essential for craniofacial development.

Epistatic relationships demonstrated for the adult pathway appear to be conserved in the embryo, such that loss of function in kng, cpn and nNOSis overcome by application of the predicted peptide xBdk or by the downstream effector NO. Further, cpn activity and xBdk modulate levels of endogenous NO, connecting NO and Kinin-Kallikrein signaling. Consistent with a role in craniofacial signaling, pathway genes are expressed at the front of the embryo, however their non-overlapping expression domains suggest that initial processing of Kng to yield xBdk occurs distal to the site of xBdk processing and NO production. We did detect different sensitivity of the embryo for intact xBdk and xBdk after C-terminal Arg removal. Thus, with reduced cpn activity, an open mouth is formed in response only to the 8AA peptide, whereas overall face morphology is corrected by both peptides, suggesting that different downstream receptors or alternate forms of peptide processing may be available to the NC.

NO has not previously been appreciated as critical for craniofacial development. In Xenopus, it was proposed that NO suppresses cell proliferation and promotes convergent extension, but a facial phenotype was not explored (Peunova et al. 2007). The requirement for kng in zebrafish facial development implies involvement of NO, and this is in accord with effects of treating zebrafish embryos with a NO inhibitor (Kong et al, 2014). In Zebrafish, NOS isoforms are expressed in the developing face, specifically in the mandibular primordium and surrounding the oral cavity, consistent with this role (Poon et al. 2003, 2008). Another route to NO production is the endothelin pathway and consistent with our results, mice deficient in endothelin-1 have craniofacial abnormalities (Yurlhara et al. 1994).

The demonstration that the EAD is necessary for migration of the first arch NC into the facial region addresses the long-standing question of what region might guide the migratory cranial NC into the face. Our findings not only underscore the organizer capacity of the EAD, but identify cpn locally expressed in the EAD as required for NC ingress, possibly through processing of Kng-derived peptides. Consistent with a guidance function for xBdk, midline or lateral placement (into the EAD) of xBdk-impregnated beads was sufficient to overcome the NC migration defect after Kinin-Kallikrein LOF. Bradykinin is pro-migratory in other settings, for malignant cells and trophoblasts, while NO is involved in inflammation-induced cell migration (Chen et al. 2000; Cuddapah et al. 2013; Erices et al. 2011; Yu et al. 2013). Interestingly, another substrate for CPN is C3a, a small complement peptide required for more local aspects of cranial NC migration (Carmona-Fontaine et al., 2011; Matthews K., 2004).

In addition to a role for Kinin-Kallikrein signaling in NC migration, kng is necessary for NC induction, whereas cpn is needed later for NC proliferation and survival, highlighting complex spatio-temporal requirements for Kinin-Kallikrein signaling during NC development. Unlike NC specification, mouth specification does not depend on Kinin-Kallikrein signaling. However, mouth opening is tightly linked to NC that abuts the EAD, suggesting that the Kinin-Kallikrein pathway may indirectly regulate mouth opening through the NC. Consistently, application of a xBdk peptide or NO donor after mouth specification and neural tube closure restored a normal NC, normal face morphology and concomitantly, an open mouth to LOF embryos.

Genes that encode Kinin-Kallikrein pathway factors are found in all vertebrates, raising the question of whether activity of this pathway during craniofacial development is conserved. The requirement for kng function during zebrafish NC development and mouth formation supports broad conservation. Additionally, ACE inhibitors that stabilize Bradykinin, used to treat high blood pressure, are teratogens associated with human craniofacial defects (Barr and Cohen 1991). In mammals, single gene LOF in Kinin-Kallikrein pathway proteins do not obviously result in craniofacial defects (Cheung et al. 1993; Mashimo and Goyal 1999; Merkulov et al. 2008; Mueller-Ortiz et al. 2009), however, certain double mutants or compound heterozygotes have not been examined. Humans heterozygous for CPN function suffer from angioedema without developmental manifestation, however no reported patients have complete CPN deficiency, indicating an essential function for this protein (Matthews and Rijli 2004). A screen for mouse genes involved in craniofacial development identified a Glutamate Carboxypeptidase and a Protein Inhibitor of Nitric Oxide (PIN), suggesting that NO activity is involved in mammalian facial development (Fowles et al. 2003). It is also possible that redundant genes or another pathway such as endothelin signaling work together with Kinin-Kallikrein signaling.

Our study newly defines the Kinin-Kallikrein pathway and nitric oxide as key for craniofacial development in Xenopus and zebrafish, and addresses the long-standing question of how the NC specifically moves into the face. The observations suggest important future directions, including mechanistic studies addressing a putative NC guidance function for xBdk and other EAD-derived activities, and the relationship between NC migration and mouth formation.

EXPERIMENTAL PROCEDURES

Embryo preparation

Xenopus laevis and zebrafish, Danio rerio embryos were cultured using standard methods (Sive et al. 2000; Westerfield et al. 2001). Xenopus embryos were staged according to Nieuwkoop and Faber, 1994, Danio embryos according to Kimmel et al., 1995. Lines used: Sox10::GFP (Curtin et al. 2011).

RNA and qPCR

RNA extraction, cDNA preparation, and qPCR measurements were conducted according to Dickinson and Sive, 2009. Primer sequences available on request. Three sets of five heads at stage 22 for sox10 and at stage 26 for sox9 were collected for each of four conditions, including control MO, cpn MO, kininogen (kng) MO, and nNOS MO to provide biological replicates. Equal amounts of RNA were used for reverse transcription (RT) and qPCR to measure sox9 or sox10 RNA. qPCR data from three readings for each of four conditions were averaged and their distribution was plotted to determine standard deviation. Average morphant qPCR value divided by control morphant qPCR value gave expression level relative to control.

In situ hybridization

cDNA sequences used to transcribe in situ hybridization probes including cpn (BC059995), kng (BC083002), nNOS (Peunova 2007), sox9 (AY035397), sox10 (Saint-Jeannet 2003), xanf1 (Ermakova 2007), frzb1 (BC108885) and XCG (Sive 1989). In situ hybridization was performed as described by Sive et al. 2000, without proteinase K treatment. Double-staining protocol adapted from Wiellette and Sive 2003.

Morpholinos and RNA rescues

Xenopus antisense morpholino-modified oligonucleotides (“morpholinos; MOs”) included one start site MO targeting cpn, two splice site MOs against kng and nNOS, and a standardcontrol MO. Sequences are: cpn MO 5′-ACCACAATCCCAGTGCCATTCTCCC- 3′, kng MO 5′-TTTTACCCATTGTCTCTTACCTGTC- 3′, nNOSMO 5′ - TGGCTAAAAGAACACAGGACATCAA-3′. nNOS MO resulted in an intron inclusion with an early stop codon at AA313, while kng MO resulted in an aberrant transcript that could not be amplified by RT-PCR, suggesting it was too large to be amplified or the primer binding sites spanning the MO sequence were missing. qPCR in Figure S2, Panel E confirms a reduction in normal kng mRNA transcript following MO treatment. Danio morpholinos include a start site and a splice-site MO targeting Kng1. Sequences are: Kng1 MO (5′-CAAGCTCTTGTCCAGCGCCATTGTC-3′) and Kng1 MO (5′-AGCCTGAGGAAACACAAACGCACGT-3′). The splice site Kng1 morpholino binds the terminal 22bp of intron 2 and the first 3bp of exon 3.

kng cDNA, nNOS cDNA, and cpn cDNA without 5′ UTRs were cloned into the CS2+ vector. RNA was generated in vitro using the mMESSAGE mMACHINE kit (Ambion). RNA (~1ng) and morpholino (14–18 ng) were co-injected at the one cell stage to test morpholino specificity via RNA rescue.

Peptide and NO donor rescues

Peptides (Thermo Scientific) were designed according to predicted sequences including 9 amino acid (AA) Xenopus Bradykinin (xBdk) (SYKGLSPFR) and 8AA Des-Arg xBdk (SYKGLSPF), and diluted to 0.1 mg/ml or 0.2mg/ml. Affi-gel blue agarose beads (50–100 mesh, Bio-Rad) loaded with peptides were prepared according to Carmona-Fontaine, Thesis, 2011. For rescues, beads resuspended in 0.1mg/ml peptide solution were implanted in the presumptive mouth region at stage 22 and scored at stage 40. For NC assays, beads resuspended in 0.2mg/ml peptide solution were implanted in side of head or presumptive mouth at stage 20–22. Embryos were fixed at tailbud (st. 26) for in situ hybridization analysis. For peptide-rescue assays, partial LOF morphants were employed to maximize viability.

NO donor, S-Nitroso-N-acetyl-DL-penicillamine (SNAP) (Sigma) was diluted to 100 mM in a 50% DMSO solution. For early rescues, 1nl of SNAP was coinjected with 17ng of morpholino into one-cell stage embryos. For late rescues (st. 20), 2–3nl of SNAP was injected into the presumptive mouth region. The nNOS inhibitor, TRIM (Sigma- T7313), was diluted to 1M concentration in DMSO, and applied to late neurula (st. 20) embryos. Embryos were collected at tailbud (st. 26) for sox9 in situ hybridization and at swimming tadpole (st. 40) for craniofacial morphology.

Nitric oxide staining and quantification

Embryos were incubated in NO-indicator 4-Amino-5-Methylamino-2′,7′-Difluorofluorescein diacetate (1:150), (DAF-FM diacetate – Invitrogen, Lepiller 2007) for 2–3 hours at 26°C. Embryos were fixed in 4% PFA overnight, embedded in 4% agarose, vibrotome sectioned (100 um), counterstained with DAPI and imaged on a Zeiss LSM 700 Laser Scanning Confocal. For NO quantification, 120 embryos per condition were decapitated, washed, dounced, and spun (10min, 1300rpm). The clear fraction was divided in triplicate, loaded on a microplate (Corning 3993- half-area, flat bottom, black), and fluorescence was measured using a Teican microplate reader. Untreated head solution was used to measure background fluorescence

Urea assay

A bovine Arginase solution (2mg/ml lyophylized bovine Arginase [Sigma # A3233] in 50mM MnCl2) was incubated for 1 hour at 37°C. Stage 28–29 embryos were anesthetized and decapitated, with 180 heads per condition. Heads were dounced in 90μl of water, spun for 10 minutes at 1100rpm at 4°C, and 100μl of clear, cytoplasmic fraction was mixed with 75μl of Arginase solution for a 2 hour incubation at 37°C. Urea content was detected using the Abcam Urea Assay Kit (Abcam #AB83362). Absorbance was read on a Teican “Infinite Pro” microplate reader and calculated as a percent of wild-type or control morphant level.

Immunohistochemistry

Immunohistochemistry was performed as described (Dickinson and Sive 2006). Primary antibodies included polyclonal anti-Laminin antibody (Sigma L-9393) diluted 1:150 and polyclonal anti-β-catenin (Invitrogen) diluted 1:100. Secondary antibody was Alexa 488 goat anti-rabbit (Molecular Probes) diluted 1:500 with 0.1% propidium iodide as a counterstain. Sections were imaged on Zeiss LSM 700 and 710 Laser Scanning Confocal microscopes. Images were analyzed using Imaris (Bitplane) and Photoshop (Adobe).

Whole mount TUNEL, PH3, and alcian blue labeling

TUNEL and PH3 labeling were performed according to Dickinson and Sive, 2006 and Dickinson and Sive, 2009. Alcian blue staining was performed according to Kennedy and Dickinson, 2012.

Transplants and head extirpation

EAD transplants were performed according to Jacox et al., 2014, NC transplants according to Mancilla and Mayor, 1996. For head extirpation, morphant and wild-type embryos were grown to stage 31–32, when the stomodeum forms. Embryos were anesthetized in Tricaine, and heads removed below the cement gland excluding the developing heart. Heads were moved to 0.5XMBS for healing and growth. Whole embryos and heads were scored for facial and mouth development at stage 40.

Highlights.

  • The Kinin-Kallikrein pathway regulates craniofacial development

  • Nitric oxide regulates mouth and neural crest development

  • Bradykinin stimulates nitric oxide production in the embryo

  • The extreme anterior domain (EAD) facilitates neural crest migration through Cpn

Acknowledgments

We thank Cas Bresilla for frog husbandry, our colleagues for discussion and critical input, especially Jasmine McCammon and Ryann Fame. Thanks to George Bell for help with bioinformatics, Nicki Watson and Wendy Salmon for imaging support and Tom diCesare for assistance with graphics. We thank Eric Liao for his gift of Sox10::GFP fish, for collegial discussion and for communicating results prior to publication. Thanks to Natalia Peunova, Andrey Zaraisky, and Jean-Pierre Saint-Jeannet for gifts of plasmids. We are grateful to the NIDCR for support (1R01 DE021109-01 to HLS and F30DE022989 to LJ), Harvard University for the Herschel Smith Graduate Fellowship (to LJ) and the American Association of Anatomists (for a postdoctoral fellowship to RS).

Footnotes

AUTHOR CONTRIBUTIONS

LJ designed and conducted all bead, extirpation, transplant, and migration LOF and rescue assays (Fig. 3, 4S–T, 5, 6), NO and Urea quantification assays (Fig 4Y-c, Fig 5I–J) and in situ hybridization experiments (Fig. 2Q-g, 3J–L, 5G–H′). LJ wrote and revised the manuscript drafts. RS designed and tested morpholinos, executed LOF rescues with cognate RNA and SNAP, and conducted immunohistochemistry and NO staining (Fig. 2A–P, 4A–R′, Fig. 4U–X), except for Ph3 and TUNEL experiments, conducted by LJ (Fig. 2h–l). RS contributed in situ hybridization data (Fig. 4M–P′, 2E–H‴), and obtained or cloned all plasmids. RS and LJ assembled and modified figures, and contributed in situ hybridization data shown in Fig. 1.

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