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Vol. 19, Issue 12, 5478-5489, December 2008
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*Departments of Cell Biology, Medicine, and Neurobiology, Division of Gastroenterology, Department of Medicine, Duke University Medical Center, Durham, NC 27710; and Institute of Experimental and Clinical Pharmacology, University of Freiburg, Germany
Submitted May 2, 2008;
Revised August 29, 2008;
Accepted September 17, 2008
Monitoring Editor: Marianne Bronner-Fraser
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
In spite of the number and diversity of these proteins, 7TMR signaling is regulated by a relatively small number of proteins. These proteins include the kinases that phosphorylate activated receptor, so called G protein–coupled receptor kinases (GRKs), and the multiadaptor proteins, arrestins, which bind to the phosphorylated receptor (Claing et al., 2002). Binding of arrestin to the receptor acts not only to uncouple the receptor from G-protein activation, but also to function as scaffolds to facilitate the internalization of the receptor and to promote signaling through alternative, non-G protein–dependent cascades (Lefkowitz and Shenoy, 2005). Excluding the visual system, mammals have five GRKs (GRK2–6) and two arrestins, (also known as β-arrestin 1 and 2 [barr1 and 2]). The GRKs are divided by sequence homology into two subfamilies: GRK2 and 3 (previously called βARK1 and 2 for their ability to phosphorylate the β-adrenergic receptor) and GRK4–6. These genes display an apparent high functional redundancy in mammals, thwarting the analysis of GRK and barr function during development, as only the GRK2 knockout (KO) mice die during embryonic development (Jaber et al., 1996). More recently, experiments in our laboratory in the vertebrate Danio rerio have uncovered a role for barr2 in promoting Hedgehog (Hh) signaling in vivo (Wilbanks et al., 2004). Knockdown of barr2 resulted in a phenotype with multiple defects, including faulty somite patterning and craniofacial development, aspects of zebrafish development regulated by Hh. These findings were consistent with the simultaneous report of the activity-dependent interaction of barr2 with the Hh pathway component Smoothened (Smo) in a cellular system (Chen et al., 2004).
The Hh pathway regulates multiple aspects of embryonic development in both vertebrates and invertebrates. Reduced Hh signaling leads to the human developmental disorders of polydactyly and holoprosencephaly, as well as severe craniofacial and skeletal malformations (McMahon et al., 2003). Constitutive pathway activity is implicated in the formation of basal cell carcinoma and medulloblastoma and in the incidence of an increasing number of cancers (Beachy et al., 2004). The core components of the pathway include the ligand Hh; its receptor, the 12TM membrane protein, Patched (Ptc); the 7TM protein Smo; and the Gli transcription factors (called Cubitus interruptus, Ci in flies). Ptc catalytically inhibits the activity of Smo in the absence of Hh (Taipale et al., 2002). When Hh binds to Ptc, Smo is released from tonic repression and undergoes rapid changes in its subcellular localization (Denef et al., 2000; Corbit et al., 2005). Recent studies in mice and in mammalian cell culture implicate Smo localization to the primary cilia as critical to Hh signal transduction (Huangfu et al., 2003; Corbit et al., 2005; Huangfu and Anderson, 2005; Rohatgi et al., 2007), an organelle not found in most Hh sensing cells in flies (Huangfu and Anderson, 2006). Also, phosphorylation, as demonstrated in flies, represents one important event regulating Smo signaling. The kinases, PKA and CK1, phosphorylate Smo at multiple sites promoting full pathway activity (Jia et al., 2004; Zhang et al., 2004; Apionishev et al., 2005), but the phosphorylation sites are not conserved from Drosophila to vertebrates. Recently, it has been reported, that Smo activity in the wing disk anterior–posterior compartment boundary requires and enhances the expression of Gprk2 (Molnar et al., 2007), which represents the Drosophila homolog of mammalian GRK5 (Fan and Schneider, 2003).
Regulation of Smo signaling by direct phosphorylation has not been described in vertebrates; indeed, this is critical as the intracellular mechanisms controlling pathway activity appear to have diverged considerably between vertebrates and invertebrates (Varjosalo et al., 2006). In heterologous cell systems it has been demonstrated, that Smoothened-mediated signal transduction can be regulated by GRK2 (Chen et al., 2004). However, it has remained unanswered, whether this regulation can be applied to Hedgehog signaling in the whole organism. To determine if GRK2 functions in the facilitation of Smo signal transduction in vertebrates, we chose a trifold approach using mammalian cells, zebrafish and KO mice.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
Cloning of Zebrafish GRK2/3 and Mutagenesis
A sequence with significant homology to the mammalian G protein–coupled receptor kinases GRK2 and GRK3 was identified by BLAST search of the Sanger zebrafish genome database (http://www.ensembl.org/Danio_rerio/index.html). A partial clone (amino acids 88–688 of final sequence) was obtained by PCR with pfu polymerase (Stratagene, La Jolla, CA) from cDNA (Protoscript, New England Biolabs, Ipswich, MA) generated from 24 hpf (hours after fertilization) embryos using the following primers: zGRK2/3 forward 5'-ATTAAAGAGTACGAGAAGTTGGACTCA; zGRK2/3 reverse 5'-TCACAGGCCGTTGCTGTTGCGGTG. 5'RACE PCR was performed (first choice RLM RACE, Ambion, Austin, TX) to obtain the full-length coding sequence and 5'-untranslated region (UTR). Full-length cDNA was cloned into the pCS2+ vector for expression in HEK293T cells and generation of capped mRNA for rescue experiments. Alignment was compiled using T-coffee (Notredame et al., 2000) and illustrated using the Boxshade software. Mutagenesis of the full-length clone was performed using the QuikChange Site-Directed Mutagenesis Kit (Stratagene).
Morpholino Design and Microinjections
All zGRK2/3 morpholino antisense oligos (MOs) were designed and synthesized by Gene Tools (Philomath, OR), based on submitted sequences. The Shh MO has been published before (Nasevicius and Ekker, 2000). The zGRK2/3 ATG MO with the sequence, AGGTCCGCCATCTTCGCCCTCTGGG, was designed to the region encompassing the 5'-UTR and sequence downstream of the start codon. A five-base mismatch control MO (5_mis CT MO) was designed with the following sequence: AGCTCCCCCATCTTCCCCGTCTCGG. The genomic sequence spanning from exon 5 to exon 9 was amplified, cloned, and sequenced. Splice-blocking MOs were created to the exon/intron junctions of exon 6 and 7 (Supplemental Figure S3). Effective dose concentrations of 0.05 mM for the ATG and 5_mis MOs were determined by dose dilution experiments where the 5_mis control MO exhibited no morphological effects to 120 hpf. One nanoliter of 0.05 mM ATG and 5_mis MOs were injected into the yolk of 1–2-cell embryos using a Femtojet microinjector (Eppendorf, Fremont, CA), and pulled needles were calibrated with a micrometer under magnification. Capped mRNAs for injections were generated using the T7 and SP6 message machine kit (Ambion) using linearized and purified cDNA as the template.
In Situ Hybridization and Immunohistochemistry
In situ hybridization was performed following standard protocols. DIG-labeled probes were generated for the following zebrafish genes as described for ptc-1(Concordet et al., 1996), shh (Krauss et al., 1993), nkx2.2 (Barth and Wilson, 1995), smo (Chen et al., 2001), and myoD (Weinberg et al., 1996), as well as for mouse patched (Adolphe et al., 2004). A 1002-base pair fragment amplified from NM_130952
[GenBank]
.1 was used to detect zebrafish dmrt2. The probe for wnt11 was ordered from ZIRC (clone ID cb748). Pictures of stained zebrafish embryos were obtained with a Zeiss Axiovert compound microscope under Nomarski optics (Thornwood, NY) or alternatively with a Nikon stereoscope (Melville, NY). Immunostainings of whole zebrafish embryos and mouse cryosections (20 µm) were performed following standard protocols. The 4d9 antibody used 1:100, which recognizes the engrailed protein in the nuclei of muscle pioneer cells, was kindly provided by Nipam Patel (University of California, Berkeley, CA; Patel et al., 1989). The Prox1 antibody (R&D Systems, Minneapolis, MN), which labels the nuclei of slow muscle fibers in zebrafish (Ochi et al., 2006) was used at 1:500 dilution. Fluorescent secondary antibodies (Molecular Probes, Eugene, OR) were used for detection. Stained embryos were photographed on a Zeiss Axiovert compound microscope utilizing the Apotome or using a Zeiss LSM510 confocal microscopy system. Antibodies used on cryosections of E11.5 mouse embryos were obtained from the following sources: HB9 (1:2000, Abcam, Cambridge, MA), Pax7 (1:100, Developmental Hybridoma Bank, University of Iowa, Iowa City, IA).
Quantification of Slow Muscle Defects
Twenty-seven hpf embryos were fixed and stained using Prox1 and 4d9 antibodies. Numbers of slow-twitch muscle nuclei and muscle pioneer cells were counted in five somites over the yolk extension per embryo. Statistical analysis was carried out using GraphPad software (San Diego, CA).
Luciferase Assay
C3H10T1/2 cells were transfected using TransIT-LT1 reagent (Mirus, Madison, WI) at a density of 105 cells/well in six-well dishes with indicated plasmids along with a Gli-luciferase reporter and CMV–β-galactosidase (β-gal) as transfection control (Meloni et al., 2006). Cells were harvested 72 h after transfection in 1x reporter buffer (Promega, Madison, WI). The Luciferase assay system (Promega) was used to measure the raw Gli-promoted signaling, which was normalized to the β-gal activity.
Smo Phosphorylation by zGRK2/3
HEK293T cells were transfected with human myc-tagged Smo and bovine GRK2 or zGRK2/3. After 48 h, the cells were washed in phosphate-free medium and labeled with 32P-orthophosphate (0.1 mCi/ml) for 1 h. Cells were washed with cold PBS and lysed with buffer A (20 mM HEPES, 0.5% Nonidet P-40, 250 mM NaCl, 10% glycerol, 2 mM EDTA, 1 mM PMSF, 2.5 µg/ml aprotinin, 2.5 µg/ml leupeptin, 100 µM sodium orthovanadate, 50 mM sodium fluoride, and 1 µM microcystin). Smo was immunopurified from clarified supernatant with the use of anti-Myc affinity gel (Covance, Princeton, NJ). Immunocomplexes were washed with buffer A and analyzed on 10% SDS-PAGE gels and detected by autoradiography.
Western Blotting
Fifteen to 20 zebrafish embryos (24 hpf) were deyolked (Link et al., 2006), and homogenized in 200 µl of lysis buffer (50 mM HEPES, 250 mM NaCl, 2 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 10% glycerol, and 0.5% NP-40) containing proteinase inhibitors (Complete Mini, Roche, Alameda, CA). The lysates were cleared by short centrifugation. Samples were loaded with an equal amount of protein as measured using Bradford reagent (Bio-Rad, Richmond, CA) on SDS 10% polyacrylamide gel, resolved by electrophoresis (Invitrogen, Carlsbad, CA) and transferred to nitrocellulose membranes. zGRK2/3 was probed using the C-14 antibody (1:500, Santa Cruz Biotechnology, Santa Cruz, CA) and detected by chemiluminescence. Recombinant zGRK2/3 was expressed in HEK293T cells by standard transfection using calcium phosphate and extracted for subsequent gels by lysing cells in lysis buffer. Anti-GAPDH antibody (1:500, Abcam) was used as loading control.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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GRKs are defined by a highly conserved kinase domain. The kinases are modular proteins with domains both C- and N-terminal to the common kinase domain that control their localization and activity (Penela et al., 2003; Supplemental Figure S1). Searching all available sequences, we did not identify another GRK with significant homology to the βARK subfamily of GRKs, suggesting that the zebrafish may have only one βARK-like kinase. Therefore, we have called this kinase the zebrafish GRK2/3 (zGRK2/3). Given the high homology in all regions between the human and fish proteins, zGRK2/3 is likely to possess the biochemical activities of the mammalian proteins.
Knockdown of zGRK2/3 in Zebrafish
To study the function of zGRK2/3 in early development, we utilize MOs directed at the ATG of the first codon and sequences in the 5'-UTR to knock down zGRK2/3 expression. To control for nonspecific effects, another MO with five base changes (5_mis CT MO) is included in all of the experiments. Furthermore, we characterize two additional splice-blocking MOs (Supplemental Figure S3). Western blot analysis using antibodies raised against the mammalian GRK3 protein reveals a single protein of the correct predicted molecular weight in 24 hpf whole embryo lysates of the noninjected and 5_mis CT MO-injected embryos (control embryos), but the signal is markedly reduced in zGRK2/3 ATG MO-injected embryos, demonstrating effective knockdown of zGRK2/3 expression (Figure 1H). Embryos with knocked down levels of zGRK2/3 develop a reproducible, distinct phenotype that is lethal between 72 and 96 hpf. In comparison, the control embryos survive and develop no apparent phenotype. The salient morphological features of the zGRK2/3 ATG MO-injected embryos include a shortened anterior–posterior axis, a smaller head, and malformed somites (Figure 1, A and B). Nomarski optics of 24 hpf embryos reveals U-shaped somites in the morphant embryos in comparison to control embryos, which display the typical "chevron" shape (Figure 1, C and D).
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; Concordet et al., 1996). During early somitogenesis, we detect a down-regulation of ptc-1 transcripts in the hypoblast adjacent to the axial mesoderm in embryos injected with MOs against zGRK2/3 (18/27 embryos; Figure 2, E and H), whereas two Hh-independent genes (Ochi et al., 2006), doublesex-related (dmrt2; Figure 2, F and I), and wnt11 (Figure 2, G and J) were unaffected. This effect persists at later stages, where ptc-1 levels in the somites and the neural tube remain decreased compared with control embryos (34/50 embryos; Figure 2, K and N). Similarly to the earlier stage analyzed, dmrt2 levels are not altered in zGRK2/3 morphants (Figure 2, L and O). However, wnt11 appears to be down-regulated in the medial somite, where muscle pioneer cells are located (Figure 2, M and P). These results suggest a specific reduction in Hh signaling in embryos with reduced zGRK2/3 expression.
zGRK2/3 Directly Phosphorylates Smo and Promotes Gli-directed Transcription
Given the morphology of the zGRK2/3 morphants and the down-regulation of ptc-1 transcript levels, we analyzed the effect of zGRK2/3 and two mutated versions on Hh signaling in a cell-based model. The assay in C3H10T1/2 cells utilizes a luciferase reporter under the control of the Gli1 promoter, providing a sensitive readout of Smo signaling (Figure 3A; Meloni et al., 2006). zGRK2/3 alone, like bovine GRK2 (bGRK2), does not activate Gli-promoted transcription in these cells. However, when cotransfected with Smo, zGRK2/3, and bGRK2 enhance Smo-mediated activity of Gli-promoted transcription to a high level, indicating a synergistic effect. This synergy is dependent on the kinase activity of zGRK2/3, as the kinase dead mutant (K220R) is unable to increase Smo-induced pathway activity (Kong et al., 1994). We also tested a previously characterized point mutant (R587Q) that was described to maintain full kinase activity but was unable to interact with Gβ
subunits (Carman et al., 2000). GRKs of the βARK-like kinase subfamily have the ability to interact with β subunits of G proteins through a binding motif in the C-terminus. This interaction has been shown to be critical for targeting of the kinase to the plasma membrane during receptor activation. Indeed, this mutant displayed a substantially reduced ability to synergize with Smo to promote Gli signaling, suggesting that zGRK2/3 may function in Hh signaling in a manner analogous to other 7TM receptor systems (Figure 3B).
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Given that zGRK2/3 can synergize with Smo to enhance Hh signaling in cells, we analyzed the ability of zGRK2/3 to directly phosphorylate Smo (Figure 3, E and F). Human Smo overexpressed in HEK293T cells is constitutively phosphorylated at low levels. However, phosphorylation of Smo is stimulated about sevenfold by cotransfection of zGRK2/3. This activity is similar to the observed activity for bGRK2 (Chen et al., 2004). In accordance with the signaling data in cells and rescue data in zebrafish embryos, the kinase dead mutant of zGRK2/3 is unable to phosphorylate Smo. Taken together, the signaling and phosphorylation data imply that zGRK2/3 stimulates Hh signaling in cells and that this activity likely relates to its ability to enhance Smo activity by directly phosphorylating Smo.
zGRK2/3 in Slow Muscle Development
Because the kinase activity of zGRK2/3 promotes Smo phosphorylation and Smo-directed Hh signaling in cellular assays and zebrafish, we sought to understand the role of this kinase in Hh signaling in vivo. Studies in the zebrafish have revealed that the Hh pathway mediates many aspects of the development of the somites and skeletal musculature (Lewis et al., 1999; Barresi et al., 2000; Wolff et al., 2003; Ingham and Kim, 2005). Mutations identified in forward genetic screens have demonstrated that a reduction of Hh signaling during development hinders the development of the two slow-muscle cell types, the superficial slow muscles and the slow muscle pioneer cells. In brief, Hh released from the notochord specifies the identity of these slow muscle precursors emerging from the surrounding presomitic mesoderm (Blagden et al., 1997). During somitogenesis, adaxial cells adjacent to the notochord as well as cells in the forming somites express the myogenic transcription factor myoD in response to Hh (Barresi et al., 2000).
To assess the role of zGRK2/3 in the development of slow muscle, we analyzed the affect of zGRK2/3 knock-down on the expression of slow muscle markers. Accordingly, the zGRK2/3 morphant embryos display a marked reduction in myoD expression in the adaxial cells as well as a slight reduction in the somites compared with control embryos (53/79 embryos in three injections; Figure 4, A and D). Moreover, differentiation of slow muscle cells is perturbed in the morphant embryos. Both, superficial slow fibers as visualized by Prox1 staining of their nuclei (Figure 4, B and E; green; 22–28 embryos per condition in two separate injections) and engrailed+ muscle pioneers were significantly reduced in numbers in zGRK2/3 ATG MO-injected embryos (Figure 4, B and E, as well as H and J; 4d9, red; 67/79 embryos from three injections). This phenotype could be successfully rescued by coinjection of zGRK2/3 mRNA-bearing silent mutations for the MO-binding site (Figure 4, C and F; n = 24–25 embryos per condition in two separate injections). The incomplete penetrance of the slow muscle phenotype may be a result of a functionally redundant kinase partially compensating for the loss of zGRK2/3 or alternatively, that low levels of zGRK2/3 activity persist in these tissues. Indeed, similar varying levels of slow muscle developmental defects have been described in the different Hh pathway genetic mutants (Wolff et al., 2004), potentially indicating compensatory effects by other genes or pathways in the development of slow muscle. Nevertheless, these data demonstrate that embryos with reduced zGRK2/3 expression are unable to respond effectively to Hh signals during early segmentation, resulting in significant impairment in the development of slow muscle.
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; Schauerte et al., 1998; Barresi et al., 2000), as visualized by an increase in the number of engrailed+ muscle pioneer cells (Figure 4, I compared with H). In the absence of normal levels of zGRK2/3, the expansion of the number of muscle pioneer cells still occurs, suggesting that zGRK2/3 acts upon the pathway upstream of Gli (12/12 embryos; Figure 4, K compared with J). To further investigate the point at which zGRK2/3 exerts its activity on the Smo signaling cascade, we examined the expression of the Hh target gene nkx2.2 in the developing brain (Barth and Wilson, 1995). As shown in Figure 5, A and B, zGRK2/3 ATG MO causes a notable reduction of nkx2.2 in the brain. As previously reported, overexpression of Shh leads to the expansion of ventral cell fates in the nervous system as can be seen by the ectopic induction of nkx2.2 (27/34 embryos; Figure 5C; Wilbanks et al., 2004). Consistent with the notion that zGRK2/3 functions downstream of Smo, coinjection of the zGRK2/3 ATG MO with Shh mRNA normalizes the ectopic expression of nkx2.2 in the brain toward control levels (16/24 embryos; Figure 5D, compare with 5A).
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In our studies we see a similar, moderate decrease of Hh signaling if zGRK2/3 levels are reduced. Therefore we tested for synergistic effects with zGRK2/3 by titrating Shh levels. Ptc-1 has been reported to be a transcriptional target associated with high Hh-signaling levels in flies and fish (Sekimizu et al., 2004; Molnar et al., 2007). As demonstrated earlier in here, knockdown of zGRK2/3 decreases ptc-1 (Figure 6, A and B). Likewise, embryos injected with a low dose of Shh MO express lower levels of ptc-1 (Figure 6C). Simultaneous injection of zGRK2/3 ATG MO and Shh MO synergistically abrogated Smo-mediated Hh signal transduction, as ptc-1 was barely detectable (Figure 6D; n = 35–63 embryos per condition).
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), whereas slow muscle fibers require moderate to low Hh. Accordingly, zGRK2/3 morphants display a stronger defect in muscle pioneer development, although slow muscle fiber formation is affected, too. Gradual depletion of Shh by injection of increasing doses of MO reduces more strongly the number of muscle pioneer cells than those of slow muscle fibers (Figure 6, G and I). Consistently with our gene expression result above, coinjection of Shh MO and zGRK2/ATG MO synergistically decreased both muscle subtypes (Figure 6, H and I; n = 18–35 embryos per condition in two separate injections). Thus, similarly to Gprk2 in flies, low levels of Hh can augment the phenotype of zGRK2/3 in zebrafish.
zGRK2/3 Is Involved in Patterning of the Neural Tube
Developmental studies in the fish, mouse, and chick have identified Hh as a key morphogen in the patterning of the ventral neural tube (Ruiz i Altaba et al., 2003). To assess the contribution of zGRK2/3 to Hh mediated signaling in the neural tube, we analyzed again the expression of nkx2.2. In noninjected and control injected embryos, nkx2.2 is expressed in the middiencephalic boundary regions of the brain and in lateral floor plate of the neural tube at 24 hpf (Figures 5A and 7A). In the zGRK2/3 morphant embryos, tissues express significantly lower levels of nkx2.2, consistent with the idea that zGRK2/3 is important in transducing the Hh signal (36/45 embryos in two injections; Figure 7B). These data indicate a requirement of zGRK2/3 in the developing neural tube for the appropriate cellular response to the Hh signals from the floor plate and notochord. Indeed, in the morphant embryos, the floor plate and notochord produce the shh transcript at normal levels, suggesting that the defect is in the response to Hh not in its production (Figure 7, C and D). Moreover, smo transcript levels are comparables in the control and zGRK2/3 ATG MO-treated embryos suggesting that the Hh signaling defects are not a result of reduced levels of Smo (Figure 7, E and F).
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; Lewis and Eisen, 2001). Accordingly injection of islet1-GFP transgenic embryos with zGRK2/3 ATG MO causes a marked reduction in both the number of isletl1-positive cells in the neural tube and in motor axon outgrowth (Figure 7, G and H, arrows and arrowhead; 23/31 embryos observed from two injections).
Similarity of GRK2-deficient Mice with Hh Mutants
In mice, deletion of GRK2 causes embryonic lethality, which has been attributed to a defect in heart development (Jaber et al., 1996). However, the embryonic lethality of the constitutive KO presumably cannot solely be explained by a heart defect (Matkovich et al., 2006). Apart from the apparent failure of cardiac development, GRK2 KO embryos display multiple aberrations from normal embryonic development. Approximately half of the dissected GRK2 KO embryos at embryonic day (E)11.5 are dramatically retarded in growth and embryonic development stalls around E9.5. They usually lack visible external eye structures and limbs (Figure 8, A and B). Similar phenotypes have been described in mouse models of impaired Hh signaling (Chiang et al., 1996; Zhang et al., 2001; Kawakami et al., 2002). Consistent with the hypothesis that zGRK2/3 is required for Hh signal transduction, we observe a reduction in the number of motoneurons in the lumbosacral region of the spinal cords of the GRK2 KO animals, when compared with sections of wild-type (wt) littermates (GRK2 KO: n = 4, wt: n = 4). Also, the remaining motoneurons are not concentrated in the ventrolateral region of the spinal cord; instead they appear scattered along the dorsal–ventral axis of the neural tube (Figure 8, C and D).
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; Eggenschwiler et al., 2006), it has been shown, that loss of Smo or Shh itself caused the complete absence of ventral neural cell types such as motoneurons (Liu et al., 2005; Eggenschwiler et al., 2006). In contrast to Shh KOs, more dorsal populations of interneurons appear less affected in GRK2 KO embryos, as Pax7-positive cells are still present although potentially less abundant in the mutant mice (Figure 8, E and F). Similar findings have been reported in mice bearing a hypomorphic mutation for IFT88, a protein involved in ciliary function, which has been shown to act as an important modulator of Smo function (Liu et al., 2005). In our zebrafish studies, we find, that zGRK2/3 is necessary for Smo signaling. By in situ hybridization for ptc-1 we reproducibly detect a decrease in ptc-1 in commissural neurons and motoneurons of spinal cords of GRK2 KO embryos (KO: n = 5; wt: n = 4), consistent with a reduction in Hh signaling (Figure 8, G and H). Together, these data reveal a significantly blunted Hh transcriptional response in the neural tube with defects in the differentiation of motor neurons of teleost and mammalian embryos.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Jia et al., 2004; Zhang et al., 2004; Apionishev et al., 2005). However, in vertebrates, strong evidence suggests that a prerequisite of Smo activity is its translocation to the primary cilia (Corbit et al., 2005). In fact, Kovacs et al. (2008) recently demonstrated, that in NIH3T3 cells Smo is driven into cilia by an interaction between β-arrestin and the ciliary motor protein Kif3a. Phosphorylation of Smo by GRK2 may be the prerequisite for a stable Smo/β-arrestin interaction, in turn regulating the trafficking of Smo to and from the cilia. Interestingly, flies without cilia develop morphologically normally, most probably because most cells in flies do not form ciliary structures with the exception of sperm and sensory neurons (Basto et al., 2006).
Another possible mode for GRK regulation of Hh signaling is through its promotion of a signaling complex. The scaffolding activity of β-arrestin 2 in the signal transduction of other 7TMR systems has been described extensively in cell culture and more recently in mice (DeFea et al., 2000; McDonald et al., 2000; Luttrell et al., 2001; Beaulieu et al., 2005; Lefkowitz and Shenoy, 2005). Hh signaling in Drosophila, involves the kinesin-like protein, Costal-2, which regulates signaling by orchestrating a series of interactions between Ci, Fused, Suppressor of Fused, and Smo (Lum et al., 2003). Morpholino studies in zebrafish have suggested that the Costal-2 homolog can regulate Hh signaling similarly to its role in flies (Tay et al., 2005). So far, Kif7/Kif27, the mammalian costal-2 homologues have not been genetically ablated in mice. However, the studies in zebrafish suggest that deletion of Kif7/Kif27 in mice may result in ectopic activation of the Hh pathway. These studies and others intimate that there may be significant differences in the spatial regulation of the fundamental Hh pathway components between vertebrates and invertebrates, but that there may also be variation in the fundamental players of the pathway between vertebrates (i.e., fish and mice).
Indeed, it appears, that a single GRK may be sufficient to facilitate Smo signaling in some tissues in mice, as the deletion of GRK2 only, already causes embryonic lethality. GRK2 KO embryos display many of the defects that have been reported in mice with impaired Hh signaling (Chiang et al., 1996; Zhang et al., 2001; Kawakami et al., 2002; Liu et al., 2005; Eggenschwiler et al., 2006). Conversely, the GRK3 KO mice develop normally. However, these KO mice express detectable levels of GRK3 mRNA, which may explain why developmental abnormalities are not observed (Peppel et al., 1997). β-arrestin 1 and 2 KO mice, on the other hand, are viable as single KOs and are anatomically indistinguishable from their wt littermates (Conner et al., 1997; Bohn et al., 1999; Gainetdinov et al., 2004). However, a double deletion of barr1 and 2 is embryonic lethal and possibly mimics mice defective in Hh signaling (DeWire et al., 2007). GRK2 KO embryos display a mild Hh phenotype, more similar to mutants of intraflagellar transport (IFT) proteins than to mice lacking Smo. As with the GRK2 KOs, motoneurons in IFT mutant mice are scattered throughout the spinal cord, whereas dorsal neuronal populations are less affected (Liu et al., 2005). Consistent with the diminished expression of ptc-1 in GRK2 KO embryos, the embryonic lethal phenotype of GRK2 KO may indeed be due to a pleiotropic, extracardiac function of GRK2 and not simply be borne from insufficiencies in heart function (Matkovich et al., 2006).
At present, we cannot exclude the importance of any other GRKs for Hh signal transduction. The zebrafish genome contains three more nonvisual GRKs with significant homology to mammalian GRKs 4–6. It remains to be uncovered, if any of those may correlate to Drosophila Gprk2 in function. However, considering, that deletion of any of the GRK4–6 family kinases does not interfere with embryonic development, strongly suggests that GRK2 is the vertebrate G protein–coupled receptor kinase enabling Smo signaling.
In summary, herein we provide evidence for a potential physiological role for a component of the GPCR desensitization machinery to facilitate the signaling of a GPCR-like protein. We demonstrate in zebrafish embryos that a βARK-like GRK is critical to Hh-mediated patterning. Our data indicate that by direct phosphorylation of Smo zGRK2/3 acts as a permissive factor in high level Hh signaling. As in the zebrafish, interfering with the function of this kinase in mice leads to embryonic phenotypes that resemble loss-of-function mutants of the Hh-signaling pathway. Although further studies will be necessary to fully elucidate the mechanism(s) of GRK2 action in Hh signaling, our observations in multiple experimental systems, strongly suggest a model where this kinase actively participates in positively mediating cellular signaling and tissue patterning through the direct regulation of Smo in vertebrates.
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ACKNOWLEDGMENTS |
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Footnotes |
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These authors contributed equally to this work. 
|| Present address: Hubrecht Institute, 3584 CT Utrecht, The Netherlands.
Address correspondence to: Marc G. Caron (caron002{at}mc.duke.edu).
Abbreviations used: barr, β-arrestin; GPCR, G protein–coupled receptor; GRK, G protein–coupled receptor kinase; Hh, Hedgehog; Ptc-1, Patched-1; Smo, Smoothened; 7TMR, 7-transmembrane spanning receptor.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Alexandre, C., Jacinto, A., and Ingham, P. W. (1996). Transcriptional activation of hedgehog target genes in Drosophila is mediated directly by the cubitus interruptus protein, a member of the GLI family of zinc finger DNA-binding proteins. Genes Dev 10, 2003–2013.
Apionishev, S., Katanayeva, N. M., Marks, S. A., Kalderon, D., and Tomlinson, A. (2005). Drosophila Smoothened phosphorylation sites essential for Hedgehog signal transduction. Nat. Cell Biol 7, 86–92.[CrossRef][Medline]
Barresi, M. J., Stickney, H. L., and Devoto, S. H. (2000). The zebrafish slow-muscle-omitted gene product is required for Hedgehog signal transduction and the development of slow muscle identity. Development 127, 2189–2199.[Abstract]
Barth, K. A., and Wilson, S. W. (1995). Expression of zebrafish nk2.2 is influenced by sonic hedgehog/vertebrate hedgehog-1 and demarcates a zone of neuronal differentiation in the embryonic forebrain. Development 121, 1755–1768.[Abstract]
Basto, R., Lau, J., Vinogradova, T., Gardiol, A., Woods, C. G., Khodjakov, A., and Raff, J. W. (2006). Flies without centrioles. Cell 125, 1375–1386.[CrossRef][Medline]
Beachy, P. A., Karhadkar, S. S., and Berman, D. M. (2004). Tissue repair and stem cell renewal in carcinogenesis. Nature 432, 324–331.[CrossRef][Medline]
Beaulieu, J. M., Sotnikova, T. D., Marion, S., Lefkowitz, R. J., Gainetdinov, R. R., and Caron, M. G. (2005). An Akt/beta-arrestin 2/PP2A signaling complex mediates dopaminergic neurotransmission and behavior. Cell 122, 261–273.[CrossRef][Medline]
Blagden, C. S., Currie, P. D., Ingham, P. W., and Hughes, S. M. (1997). Notochord induction of zebrafish slow muscle mediated by Sonic hedgehog. Genes Dev 11, 2163–2175.
Bohn, L. M., Lefkowitz, R. J., Gainetdinov, R. R., Peppel, K., Caron, M. G., and Lin, F. T. (1999). Enhanced morphine analgesia in mice lacking beta-arrestin 2. Science 286, 2495–2498.
Bulgakov, O. V., Eggenschwiler, J. T., Hong, D. H., Anderson, K. V., and Li, T. (2004). FKBP8 is a negative regulator of mouse sonic hedgehog signaling in neural tissues. Development 131, 2149–2159.
Carman, C. V., Barak, L. S., Chen, C., Liu-Chen, L. Y., Onorato, J. J., Kennedy, S. P., Caron, M. G., and Benovic, J. L. (2000). Mutational analysis of Gbetagamma and phospholipid interaction with G protein-coupled receptor kinase 2. J. Biol. Chem 275, 10443–10452.
Chen, W., Burgess, S., and Hopkins, N. (2001). Analysis of the zebrafish smoothened mutant reveals conserved and divergent functions of hedgehog activity. Development 128, 2385–2396.
Chen, W., Ren, X. R., Nelson, C. D., Barak, L. S., Chen, J. K., Beachy, P. A., de Sauvage, F., and Lefkowitz, R. J. (2004). Activity-dependent internalization of smoothened mediated by beta-arrestin 2 and GRK2. Science 306, 2257–2260.
Chiang, C., Litingtung, Y., Lee, E., Young, K. E., Corden, J. L., Westphal, H., and Beachy, P. A. (1996). Cyclopia and defective axial patterning in mice lacking Sonic hedgehog gene function. Nature 383, 407–413.[CrossRef][Medline]
Claing, A., Laporte, S. A., Caron, M. G., and Lefkowitz, R. J. (2002). Endocytosis of G protein-coupled receptors: roles of G protein-coupled receptor kinases and beta-arrestin proteins. Prog. Neurobiol 66, 61–79.[CrossRef][Medline]
Concordet, J. P., Lewis, K. E., Moore, J. W., Goodrich, L. V., Johnson, R. L., Scott, M. P., and Ingham, P. W. (1996). Spatial regulation of a zebrafish patched homologue reflects the roles of sonic hedgehog and protein kinase A in neural tube and somite patterning. Development 122, 2835–2846.[Abstract]
Conner, D. A., Mathier, M. A., Mortensen, R. M., Christe, M., Vatner, S. F., Seidman, C. E., and Seidman, J. G. (1997). beta-Arrestin1 knockout mice appear normal but demonstrate altered cardiac responses to beta-adrenergic stimulation. Circ. Res 81, 1021–1026.
Corbit, K. C., Aanstad, P., Singla, V., Norman, A. R., Stainier, D. Y., and Reiter, J. F. (2005). Vertebrate Smoothened functions at the primary cilium. Nature 437, 1018–1021.[CrossRef][Medline]
DeFea, K. A., Zalevsky, J., Thoma, M. S., Dery, O., Mullins, R. D., and Bunnett, N. W. (2000). beta-arrestin-dependent endocytosis of proteinase-activated receptor 2 is required for intracellular targeting of activated ERK1/2. J. Cell Biol 148, 1267–1281.
Denef, N., Neubuser, D., Perez, L., and Cohen, S. M. (2000). Hedgehog induces opposite changes in turnover and subcellular localization of patched and smoothened. Cell 102, 521–531.[CrossRef][Medline]
DeWire, S. M., Ahn, S., Lefkowitz, R. J., and Shenoy, S. K. (2007). Beta-arrestins and cell signaling. Annu. Rev. Physiol 69, 483–510.[CrossRef][Medline]
Eggenschwiler, J. T., Bulgakov, O. V., Qin, J., Li, T., and Anderson, K. V. (2006). Mouse Rab23 regulates hedgehog signaling from smoothened to Gli proteins. Dev. Biol 290, 1–12.[CrossRef][Medline]
Fan, S., and Schneider, L. E. (2003). The role of maternal and zygotic Gprk2 expression in Drosophila development. Biochem. Biophys. Res. Commun 301, 127–135.[CrossRef][Medline]
Gainetdinov, R. R., Premont, R. T., Bohn, L. M., Lefkowitz, R. J., and Caron, M. G. (2004). Desensitization of G protein-coupled receptors and neuronal functions. Annu. Rev. Neurosci 27, 107–144.[Medline]
Hammerschmidt, M., Bitgood, M. J., and McMahon, A. P. (1996). Protein kinase A is a common negative regulator of Hedgehog signaling in the vertebrate embryo. Genes Dev 10, 647–658.
Higashijima, S., Hotta, Y., and Okamoto, H. (2000). Visualization of cranial motor neurons in live transgenic zebrafish expressing green fluorescent protein under the control of the islet-1 promoter/enhancer. J. Neurosci 20, 206–218.
Huangfu, D., and Anderson, K. V. (2005). Cilia and Hedgehog responsiveness in the mouse. Proc. Natl. Acad. Sci. USA 102, 11325–11330.
Huangfu, D., and Anderson, K. V. (2006). Signaling from Smo to Ci/Gli: conservation and divergence of Hedgehog pathways from Drosophila to vertebrates. Development 133, 3–14.
Huangfu, D., Liu, A., Rakeman, A. S., Murcia, N. S., Niswander, L., and Anderson, K. V. (2003). Hedgehog signalling in the mouse requires intraflagellar transport proteins. Nature 426, 83–87.[CrossRef][Medline]
Ingham, P. W., and Kim, H. R. (2005). Hedgehog signalling and the specification of muscle cell identity in the zebrafish embryo. Exp. Cell Res 306, 336–342.[CrossRef][Medline]
Jaber, M., Koch, W. J., Rockman, H., Smith, B., Bond, R. A., Sulik, K. K., Ross, J., Jr, Lefkowitz, R. J., Caron, M. G., and Giros, B. (1996). Essential role of beta-adrenergic receptor kinase 1 in cardiac development and function. Proc. Natl. Acad. Sci. USA 93, 12974–12979.
Jia, J., Tong, C., Wang, B., Luo, L., and Jiang, J. (2004). Hedgehog signalling activity of Smoothened requires phosphorylation by protein kinase A and casein kinase I. Nature 432, 1045–1050.[CrossRef][Medline]
Kawakami, T., Kawcak, T., Li, Y. J., Zhang, W., Hu, Y., and Chuang, P. T. (2002). Mouse dispatched mutants fail to distribute hedgehog proteins and are defective in hedgehog signaling. Development 129, 5753–5765.[CrossRef][Medline]
Kong, G., Penn, R., and Benovic, J. L. (1994). A beta-adrenergic receptor kinase dominant negative mutant attenuates desensitization of the beta 2-adrenergic receptor. J. Biol. Chem 269, 13084–13087.
Kovacs, J. J., Whalen, E. J., Liu, R., Xiao, K., Kim, J., Chen, M., Wang, J., Chen, W., and Lefkowitz, R. J. (2008). Beta-arrestin-mediated localization of smoothened to the primary cilium. Science 320, 1777–1781.
Krauss, S., Concordet, J. P., and Ingham, P. W. (1993). A functionally conserved homolog of the Drosophila segment polarity gene hh is expressed in tissues with polarizing activity in zebrafish embryos. Cell 75, 1431–1444.[CrossRef][Medline]
Lefkowitz, R. J. (2007). Seven transmembrane receptors: something old, something new. Acta Physiol 190, 9–19.[CrossRef]
Lefkowitz, R. J., and Shenoy, S. K. (2005). Transduction of receptor signals by beta-arrestins. Science 308, 512–517.
Lewis, K. E., Currie, P. D., Roy, S., Schauerte, H., Haffter, P., and Ingham, P. W. (1999). Control of muscle cell-type specification in the zebrafish embryo by Hedgehog signalling. Dev. Biol 216, 469–480.[CrossRef][Medline]
Lewis, K. E., and Eisen, J. S. (2001). Hedgehog signaling is required for primary motoneuron induction in zebrafish. Development 128, 3485–3495.
Link, V., Shevchenko, A., and Heisenberg, C. P. (2006). Proteomics of early zebrafish embryos. BMC Dev. Biol 6, 1–9.[CrossRef][Medline]
Liu, A., Wang, B., and Niswander, L. A. (2005). Mouse intraflagellar transport proteins regulate both the activator and repressor functions of Gli transcription factors. Development 132, 3103–3111.
Lum, L., Zhang, C., Oh, S., Mann, R. K., von Kessler, D. P., Taipale, J., Weis-Garcia, F., Gong, R., Wang, B., and Beachy, P. A. (2003). Hedgehog signal transduction via Smoothened association with a cytoplasmic complex scaffolded by the atypical kinesin, Costal-2. Mol. Cell 12, 1261–1274.[CrossRef][Medline]
Luttrell, L. M., Roudabush, F. L., Choy, E. W., Miller, W. E., Field, M. E., Pierce, K. L., and Lefkowitz, R. J. (2001). Activation and targeting of extracellular signal-regulated kinases by beta-arrestin scaffolds. Proc. Natl. Acad. Sci. USA 98, 2449–2454.
Matkovich, S. J., Diwan, A., Klanke, J. L., Hammer, D. J., Marreez, Y., Odley, A. M., Brunskill, E. W., Koch, W. J., Schwartz, R. J., and Dorn, G. W., 2nd. (2006). Cardiac-specific ablation of G-protein receptor kinase 2 redefines its roles in heart development and beta-adrenergic signaling. Circ. Res 99, 996–1003.
McDonald, P. H., Chow, C. W., Miller, W. E., Laporte, S. A., Field, M. E., Lin, F. T., Davis, R. J., and Lefkowitz, R. J. (2000). Beta-arrestin2, a receptor-regulated MAPK scaffold for the activation of JNK3. Science 290, 1574–1577.
McMahon, A. P., Ingham, P. W., and Tabin, C. J. (2003). Developmental roles and clinical significance of hedgehog signaling. Curr. Top Dev. Biol 53, 1–114.[Medline]
Meloni, A. R., Fralish, G. B., Kelly, P., Salahpour, A., Chen, J. K., Wechsler-Reya, R. J., Lefkowitz, R. J., and Caron, M. G. (2006). Smoothened signal transduction is promoted by G protein-coupled receptor kinase 2. Mol. Cell. Biol 26, 7550–7560.
Molnar, C., Holguin, H., Mayor, F., Jr, Ruiz-Gomez, A., and de Celis, J. F. (2007). The G protein-coupled receptor regulatory kinase GPRK2 participates in Hedgehog signaling in Drosophila. Proc. Natl. Acad. Sci. USA 104, 7963–7968.
Nasevicius, A., and Ekker, S. C. (2000). Effective targeted gene ‘knockdown’ in zebrafish. Nat. Genet 26, 216–220.[CrossRef][Medline]
Notredame, C., Higgins, D. G., and Heringa, J. (2000). T-Coffee: a novel method for fast and accurate multiple sequence alignment. J. Mol. Biol 302, 205–217.[CrossRef][Medline]
Ochi, H., Pearson, B. J., Chuang, P. T., Hammerschmidt, M., and Westerfield, M. (2006). Hhip regulates zebrafish muscle development by both sequestering Hedgehog and modulating localization of Smoothened. Dev. Biol 297, 127–140.[CrossRef][Medline]
Patel, N. H., Martin-Blanco, E., Coleman, K. G., Poole, S. J., Ellis, M. C., Kornberg, T. B., and Goodman, C. S. (1989). Expression of engrailed proteins in arthropods, annelids, and chordates. Cell 58, 955–968.[CrossRef][Medline]
Penela, P., Ribas, C., and Mayor, F., Jr. (2003). Mechanisms of regulation of the expression and function of G protein-coupled receptor kinases. Cell Signal 15, 973–981.[CrossRef][Medline]
Peppel, K., Boekhoff, I., McDonald, P., Breer, H., Caron, M. G., and Lefkowitz, R. J. (1997). G protein-coupled receptor kinase 3 (GRK3) gene disruption leads to loss of odorant receptor desensitization. J. Biol. Chem 272, 25425–25428.
Rohatgi, R., Milenkovic, L., and Scott, M. P. (2007). Patched1 regulates hedgehog signaling at the primary cilium. Science 317, 372–376.
Ruiz i Altaba, A., Nguyen, V., and Palma, V. (2003). The emergent design of the neural tube: prepattern, SHH morphogen and GLI code. Curr. Opin. Genet. Dev 13, 513–521.[CrossRef][Medline]
Schauerte, H. E., van Eeden, F. J., Fricke, C., Odenthal, J., Strahle, U., and Haffter, P. (1998). Sonic hedgehog is not required for the induction of medial floor plate cells in the zebrafish. Development 125, 2983–2993.[Abstract]
Sekimizu, K., Nishioka, N., Sasaki, H., Takeda, H., Karlstrom, R. O., and Kawakami, A. (2004). The zebrafish iguana locus encodes Dzip1, a novel zinc-finger protein required for proper regulation of Hedgehog signaling. Development 131, 2521–2532.
Taipale, J., Cooper, M. K., Maiti, T., and Beachy, P. A. (2002). Patched acts catalytically to suppress the activity of Smoothened. Nature 418, 892–897.[CrossRef][Medline]
Tay, S. Y., Ingham, P. W., and Roy, S. (2005). A homologue of the Drosophila kinesin-like protein Costal2 regulates Hedgehog signal transduction in the vertebrate embryo. Development 132, 625–634.
Varjosalo, M., Li, S. P., and Taipale, J. (2006). Divergence of Hedgehog signal transduction mechanism between Drosophila and mammals. Dev. Cell 10, 177–186.[CrossRef][Medline]
Weinberg, E. S., Allende, M. L., Kelly, C. S., Abdelhamid, A., Murakami, T., Andermann, P., Doerre, O. G., Grunwald, D. J., and Riggleman, B. (1996). Developmental regulation of zebrafish MyoD in wild-type, no tail and spadetail embryos. Development 122, 271–280.[Abstract]
Wilbanks, A. M., Fralish, G. B., Kirby, M. L., Barak, L. S., Li, Y. X., and Caron, M. G. (2004). Beta-arrestin 2 regulates zebrafish development through the hedgehog signaling pathway. Science 306, 2264–2267.
Wolff, C., Roy, S., and Ingham, P. W. (2003). Multiple muscle cell identities induced by distinct levels and timing of hedgehog activity in the zebrafish embryo. Curr. Biol 13, 1169–1181.[CrossRef][Medline]
Wolff, C., Roy, S., Lewis, K. E., Schauerte, H., Joerg-Rauch, G., Kirn, A., Weiler, C., Geisler, R., Haffter, P., and Ingham, P. W. (2004). iguana encodes a novel zinc-finger protein with coiled-coil domains essential for Hedgehog signal transduction in the zebrafish embryo. Genes Dev 18, 1565–1576.
Zhang, C., Williams, E. H., Guo, Y., Lum, L., and Beachy, P. A. (2004). Extensive phosphorylation of Smoothened in Hedgehog pathway activation. Proc. Natl. Acad. Sci. USA 101, 17900–17907.
Zhang, X. M., Ramalho-Santos, M., and McMahon, A. P. (2001). Smoothened mutants reveal redundant roles for Shh and Ihh signaling including regulation of L/R symmetry by the mouse node. Cell 106, 781–792.[Medline]
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