Institute of Neuroscience, 1254 University of Oregon, Eugene, OR 97403-1254, USA
*Author for correspondence (e-mail: maves{at}uoneuro.uoregon.edu).
Accepted 16 May 2002
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SUMMARY |
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Key words: Hindbrain, Rhombomere, Organizer, FGF3, FGF8, acerebellar, valentino, krox-20, Zebrafish
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INTRODUCTION |
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Segmentation is another mechanism that is used to promote patterning and regionalization along the anteroposterior axis of the CNS. This is particularly true in the vertebrate hindbrain, which becomes subdivided into segments, termed rhombomeres, each of which acquires distinct cellular and molecular characteristics (reviewed by Lumsden and Krumlauf, 1996). Rhombomeres function to order neuronal differentiation and cranial neural crest migration patterns that are crucial for the proper development and function of the vertebrate head. Several transcription factors have been identified that are expressed in conserved, rhombomere-specific patterns and are required for proper hindbrain development. These include krox-20 (egr2 Zebrafish Information Network) kreisler/mafB/valentino and several Hox genes (reviewed by Schneider-Maunoury et al., 1998
). While signals such as FGFs (Marín and Charnay, 2000a
) and retinoic acid (reviewed by Gavalas and Krumlauf, 2000
) have been proposed to play roles in activating the expression of these hindbrain segmentation genes, it is not clear when and from which tissues such signals might be acting. Evidence for interactions between rhombomeres, particularly in promoting krox-20 and val expression (Graham and Lumsden, 1996
; Helmbacher et al., 1998
; Marín and Charnay, 2000b
), suggests that local organizing signals may play roles in patterning the hindbrain.
Using time lapse analyses of zebrafish hindbrain development, we find that r4 is the first rhombomere to form. Reticulospinal and motoneuron differentiation occurs earliest in r4. Two FGF signals, FGF3 and FGF8, are expressed early in r4 and are together required for the development of r5 and r6. Transplantation of r4 cells can induce expression of r5/r6 markers, as can misexpression of either FGF3 or FGF8. Genetic mosaic analyses also support a role for FGF signaling acting from r4. Our findings thus demonstrate the existence of an FGF-mediated signaling center in r4. The early establishment of r4 as a signaling center promotes the development of adjacent rhombomeres and thus supports the propagation of hindbrain segmental patterning. Evidence from studies of other vertebrates suggests that this r4 signaling center is conserved.
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MATERIALS AND METHODS |
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Time-lapse analyses
Embryos were incubated in a 200 µM solution of BODIPY FL C5-ceramide (Molecular Probes) from sphere stage to bud stage. Embryos were mounted between two coverslips in a mixture of 0.5% agar in EM and 3% methyl cellulose in EM. Images were obtained using a Zeiss 310 confocal microscope. Four-dimensional images were collected every 6 minutes at 28.5°C from about the two-somite (s) stage to about the 16 s stage. Recordings were analyzed using NIH Image software (http://rsb.info.nih.gov/nih-image/index.html). The presence and identities of early boundaries were confirmed by analyzing recordings both retrospectively and prospectively.
RNA in situ hybridization
cDNA probes that detected the following genes were used: isl1 (Appel et al., 1995); krox-20 (Oxtoby and Jowett, 1993
); pax2a (Krauss et al., 1991
); fgf3 (this paper); no tail (Schulte-Merker et al., 1992
); fgf8 (Reifers et al., 1998
); val (Moens et al., 1998
); hoxb3a, hoxb1a, hoxb4a (Prince et al., 1998
); efnb4, efnb2a (Cooke et al., 2001
); and erm, pea3 (Münchberg et al., 1999
). Probe syntheses and whole-mount in situ hybridization were performed as previously described (Jowett and Lettice, 1994
; Hauptmann and Gerster, 1994
). Embryos were de-yolked using tungsten needles, mounted in 90% glycerol in phosphate-buffered saline (PBS) and photographed using a Zeiss Axiophot 2 microscope.
fgf3 cDNA cloning and misexpression
Based on the published zebrafish fgf3 genomic DNA sequence (Kiefer et al., 1996), we screened a zebrafish 15-19 h cDNA library (Appel and Eisen, 1998
) by PCR to isolate a full-length fgf3 cDNA. Full-length fgf3 cDNA was subcloned into pCS2+ (Rupp et al., 1994
), yielding pCS2+-fgf3. To make full-length fgf3 antisense RNA probe, pCS2+-fgf3 was linearized with BamHI and transcribed with T7 polymerase. To tag fgf3 with a Myc epitope, we subcloned full-length fgf3 cDNA into pCS2+MT (Rupp et al., 1994
), which contains a 6xMyc epitope, yielding pCS2+-fgf3-MT. To make heat shock (HS)-fgf3, the CMV promoter of pCS2+MT was removed, and a zebrafish hsp70 promoter (Halloran et al., 2000
) and full-length fgf3 cDNA were inserted into that plasmid. Sequencing confirmed proper fgf3 sequence in frame with the Myc tag at the 3' end of fgf3. The Myc tag does not affect FGF3 activity in overexpression assays.
For misexpression experiments, HS-fgf3 or HS-fgf8 (Roehl and Nüsslein-Volhard, 2001) plasmid was injected into one- to four-cell-stage embryos at about 5 ng/µl. Uptake of the transgenes by early blastomeres is very mosaic, leading to small clones of fgf-expressing cells upon heat shock. Embryos were heat shocked at 37°C for 1 hour at about bud stage, then were fixed for staining at about the 6-8 s stage. Transgene-injected/non-heat-shocked control embryos and non-injected/heat-shocked control embryos show largely normal val and krox-20 expression.
To make FGF8-soaked beads, 45 µm polystyrene beads (Polysciences) were rinsed in PBS, then were incubated in 5 mg/ml heparin for 1 hour at room temperature, then were incubated in 250 µg/ml [in 0.5% bovine serum albumin (BSA) in PBS] mouse FGF8b (R&D Systems) for 2 hours at room temperature. Control beads were incubated in 0.5% BSA in PBS. Beads were then rinsed in PBS. Bead implants were carried out similar to transplantation experiments. To test efficacy of FGF8 beads, embryos with implanted beads were stained for pea3, an FGF target gene (Roehl and Nüsslein-Volhard, 2001; Raible and Brand, 2001
). Nine out of nine embryos showed a ring of pea3 expression around the FGF8 bead.
fgf3 morpholinos and embryo injections
Based on the published sequence of zebrafish fgf3 (Kiefer et al., 1996) (we also confirmed fgf3 5'UTR sequence in our AB wild-type line), four antisense morpholino oligos (MOs) were ordered (Gene Tools): fgf3A, 5'-CATTGTGGCATGGCGGGATGTCGGC-3'; fgf3B, 5'-GGTCCCATCAAAGAAGTATCATTTG-3'; fgf3C, 5'-TCTCGCTGGAATAGAAAGAGCTGGC-3'; and fgf3MMA, 5'-CAaTGTcGCATGGCGGGtTGTgGGC-3'. Sequence complementary to the predicted start codon is underlined in fgf3A. fgf3B and fgf3C are designed to non-overlapping 5'UTR sequence upstream of fgf3A. fgf3MMA is a mis-match control for fgf3A; the mis-paired bases are in lower case. For stock solutions, MOs were dissolved in water or Danieau buffer (Nasevicius and Ekker, 2000
) at 25 mg/ml. For working solutions, MOs were dissolved in 0.2 M KCl with 0.1% phenol red.
Embryos were pressure injected, using a pulled glass micropipette and a microinjector (ASI), into the yolk at the one- to four-cell stage. We inject a 2-3 nl volume into each embryo.
To show that the MOs can block translation of FGF3, we co-injected fgf3A and fgf3C with pCS2+-fgf3-MT, which contains the target sequence for these MOs, and found that the MOs can block expression of FGF3-Myc protein; the MOs also block the effects of overexpression of an fgf3 cDNA containing the MO target sequence (not shown).
To reduce or eliminate nonspecific MO side-effects, we determined the highest dose for each MO that still yielded normal embryos and larvae (larvae that develop swim bladders) and used these doses in pair-wise combinations to achieve synergistic effects on fgf3 while minimizing nonspecific effects. The three pair-wise combinations of the fgf3 MOs all cause the same phenotype: a reduced forebrain, a slightly reduced tail and small ears. We presume this phenotype corresponds to a severe reduction of fgf3 function. The four-base mis-match control MO (fgf3MMA), when injected (at the same dose used for fgf3A) alone or in combination with the others, generates normal embryos and larvae. To generate fgf3-MO; fgf8- embryos, we injected the fgf3B (1.0 mg/ml) + fgf3C (0.25 mg/ml) combination into ace embryos. Each fgf3 MO (A, B, C) causes loss of r5 krox-20 expression when injected into ace. fgf3MMA does not.
Immunocytochemistry
Embryos were fixed in 4% paraformaldehyde in PBS for 2 hours at room temperature or, for RMO-44, in 2% trichloroacetic acid in water for 4 hours at room temperature. After fixation, embryos were rinsed with PBS, then rinsed with distilled water, permeabilized with acetone treatment at 20°C for 10 minutes, rinsed again with distilled water and then PBS, and then blocked in PBDTX (PBS with 1% bovine serum albumin, 1% DMSO, pH to 7.3, and 0.1% Triton X-100) with 2% normal goat serum (NGS) for 30 minutes. Embryos were then incubated overnight at 4°C in primary antibodies at the following dilutions in PBDTX with 2% NGS: RMO-44, 1:25 (Zymed); Islet 39.4D5, 1:100 (Ericson et al., 1992); zn-8, 1:1000 (Trevarrow et al., 1990
); anti-Myc, 1:100 (Oncogene Research Products). After PBDTX rinses, embryos were incubated in secondary antibody goat anti-mouse Alexa Fluor 488 (Molecular Probes) at 1:200 dilution in PBDTX with 2% NGS for 5 hours at room temperature. Embryos were then rinsed in PBS and analyzed using a Leica MZ FLIII fluorescence stereomicroscope. Images were obtained using a Zeiss 310 confocal microscope.
Transplantation and mosaic analysis
Transplantation techniques were adapted from Moens et al. (Moens et al., 1996) and Woo and Fraser (Woo and Fraser, 1997
). All embryos used for transplantations were raised in filter-sterilized EM supplemented with penicillin (5000 U/l)/streptomycin (100 mg/l; Sigma). Donor embryos were labeled at the one-cell stage with a spin-filtered mixture of 3% lysine-fixable fluorescein dextran (or 2% Alexa Fluor 488 dextran) and 3% lysine-fixable biotin dextran (10,000 Mr; Molecular Probes) in 0.2 M KCl. Dechorionated donor and host embryos were mounted in 2% methyl cellulose in Ringers (Westerfield, 1995
) on a glass depression slide. Working with a Nikon SMZ-U fluorescence stereomicroscope and using a pulled glass micropipette as a knife, donor tissue (about 50-100 cells) was excised and then inserted into a host embryo. The position of labeled donor tissue in the host was checked briefly with UV light. Embryos on the depression slide were then submerged in EM with pen/strep. Donor and host embryos were processed for in situ hybridization as described above. To detect donor-derived biotin-labeled cells, embryos were rinsed with water following the in situ, then were rinsed with PBDTX, then were incubated overnight at 4°C in streptavidin Alexa Fluor 488 (1:100 in PBDTX with 2% NGS; Molecular Probes). Embryos were then rinsed and analyzed as they were for immunocytochemistry.
To test the identity of r4 transplants, we fixed shield-stage host embryos within an hour after transplantation and stained for rhombomere markers: 8/10 transplants expressed hoxb1 (r4), 9/10 transplants expressed fgf3 (r4), and 5/10 transplants expressed val (r5/6). To assess whether we were transplanting any paraxial head mesoderm along with the r4 cells, we stained for follistatin (Bauer et al., 1998): six out of nine transplants were follistatin-negative; three out of nine showed only two to three follistatin-expressing cells.
In cases where fgf3-MO; fgf8-MO embryos were used as hosts, the morpholinos were co-injected at the one-cell stage. The fgf8 morpholinos E2I2 and E3I3 (Draper et al., 2001) were used at 0.5 mg/ml each.
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RESULTS |
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fgf3 and fgf8 are both expressed in early r4
We hypothesized that the early development of r4 may indicate a possible function for r4 in promoting subsequent development of neighboring rhombomeres by acting as an early signaling center. In support of this hypothesis, we find that an FGF signaling gene, fgf3, is expressed early in presumptive r4. Zebrafish fgf3 (Kiefer et al., 1996) is expressed in the presumptive hindbrain beginning at 9 h/90% epiboly (Fig. 2A) in a transverse stripe, which, relative to the midbrain-hindbrain boundary (MHB) marker pax2a (Krauss et al., 1991
), is in the center of the early hindbrain (arrow in Fig. 2A). At 10 h/bud stage, when krox-20 expression first appears in presumptive r3 (Oxtoby and Jowett, 1993
), the fgf3 stripe is just caudal to r3 (and possibly some expression in r3, Fig. 2B). From 10.3-12 h/1-6 s, we find strong fgf3 expression in presumptive r4, as well as some weaker expression in r3, r5 and r6 (Fig. 2C,D). Beginning at about 13 h/8 s, fgf3 expression in the hindbrain is restricted to r4 (Fig. 2E), and fgf3 continues to be expressed in r4 until about 18 h/18 s, at which time hindbrain expression of fgf3 becomes undetectable (not shown). During gastrulation and somitogenesis stages, fgf3 also has other domains of expression, some of which have been described by Fürthauer et al. (Fürthauer et al., 2001
) and Phillips et al. (Phillips et al., 2001
), including the margin during gastrulation (not shown), the forebrain (Fig. 2C), the MHB (Fig. 2E), the tail bud (Fig. 2C) and the head periphery (Fig. 2E). Zebrafish fgf3 is thus expressed in several domains that are similar to those observed for fgf3 expression in other vertebrates (Wilkinson et al., 1988
; Tannahill et al., 1992
; Mahmood et al., 1995
; Mahmood et al., 1996
) (see Discussion).
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fgf3 and fgf8 are two of the earliest genes known to be expressed in the zebrafish hindbrain primordium, and their overlapping expression in presumptive r4, beginning with the initiation of fgf3 expression at 9 h/90% epiboly, precedes that of krox-20 in r3 and r5 (late bud stage) (Oxtoby and Jowett, 1993), val in r5 and r6 (bud stage) (Moens et al., 1998
) and rhombomere-specific expression of the Hox genes (Prince et al., 1998
). fgf3 and fgf8 are thus present at the proper time and place to mediate a potential early signaling activity of r4 and regulate the expression of hindbrain segmentation genes.
FGF3 and FGF8 are required for r5 and r6 development
We next addressed the requirements for FGF3 and FGF8 in hindbrain patterning. Although a mutant line exists [acerebellarti282a (ace)] that is a strong hypomorph for fgf8 (Reifers et al., 1998; Draper et al., 2001
), an fgf3 mutant line has not yet been isolated. To knock down the function of FGF3, we used antisense morpholino oligos (MOs), which have been shown to block translation and function of target genes when injected into zebrafish embryos (Nasevicius and Ekker, 2000
). We performed several controls to demonstrate that the fgf3 MOs can indeed block translation of FGF3 and also act specifically on fgf3 (see Materials and Methods). As we will document in the following sections, we do not see severe defects in hindbrain patterning in fgf3 morphant (fgf3-MO) embryos. Similarly, fgf8 mutant, or fgf8-, embryos, aside from loss of the cerebellum (Reifers et al., 1998
), show only subtle defects in hindbrain patterning (Roehl and Nüsslein-Volhard, 2001
) (this paper). Thus, severe reduction of either FGF3 or FGF8 has little effect on hindbrain patterning.
To address whether fgf3 and fgf8 interact or function redundantly in hindbrain patterning, we injected fgf3 morpholinos into fgf8- embryos, thus generating fgf3-MO; fgf8- embryos. We examined the expression of several hindbrain patterning genes at the 18-19 h/18-20 s stage, a period when these genes show well-defined rhombomere-specific expression. We find that, although krox-20 (Oxtoby and Jowett, 1993) expression appears fairly normal in fgf3-MO embryos and fgf8- embryos (although rhombomeres often appear reduced in width; Fig. 3A, parts a and c), the r5 stripe of krox-20 expression is completely lost in fgf3-MO; fgf8- embryos and the r3 stripe appears reduced (Fig. 3A, part d). fgf3-MO; fgf8- embryos also show complete loss of val (Moens et al., 1998
) expression in r5 and r6 (Fig. 3A, part h) and loss of high levels of hoxb3 (Prince et al., 1998
) in r5 and r6 (Fig. 3A, part l). However, r4 expression of hoxb1a (Prince et al., 1998
) is maintained in fgf3-MO; fgf8- embryos (Fig. 3A, part p). Furthermore, we find that expression of hoxb4, which normally has a rostral boundary at the r6/7 boundary (Prince et al., 1998
) (Fig. 3A, part q), is contiguous with hoxb1a in fgf3-MO; fgf8- embryos (Fig. 3A, part t). Staining of fgf3-MO; fgf8- embryos with hoxb4 (without hoxb1a) reveals a space corresponding to the width of r4 between the anterior hoxb4 expression and krox-20 in r3 (not shown). These results show that while loss of either FGF3 or FGF8 has only subtle effect on hindbrain patterning, rhombomeres adjacent to r4 are lost (r5, r6) or reduced (r3) in fgf3-MO; fgf8- embryos; however, r4 is maintained.
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To further address the timing and extent of FGF signaling activity in the early hindbrain, we analyzed the expression of two known targets of FGF signaling, erm and pea3 (Roehl and Nüsslein-Volhard, 2001; Raible and Brand, 2001
), which encode ETS-domain transcription factors (Münchberg et al., 1999
; Brown et al., 1998
). Although erm and pea3 are broadly expressed during late epiboly stages (Münchberg et al., 1999
; Roehl and Nüsslein-Volhard, 2001
), both erm and pea3 show a more restricted hindbrain expression domain beginning at about 10-10.3 h/bud stage-1 s (Münchberg et al., 1999
) (not shown), just after the time when fgf3 and fgf8 begin to be co-expressed in r4. These hindbrain domains extend from the MHB to r6 (Fig. 3B, parts m,q), and r6 is where we find the caudal extent of requirement for FGF signaling, as shown above. erm and pea3 expression is slightly upregulated in fgf3-MO embryos (Fig. 3B, parts n,r) and reduced in fgf8- embryos (Fig. 3B, parts o,s) (Roehl and Nüsslein-Volhard, 2001
; Raible and Brand, 2001
). However, in fgf3-MO; fgf8- embryos, erm and pea3 expression are completely absent from the hindbrain (Fig. 3B, parts p,t), yet are maintained, at reduced levels, in other domains, such as the forebrain (Fig. 3B, part p). These results support the timing of FGF signaling in the hindbrain beginning at about bud stage-early somite stages, support the caudal extent of FGF hindbrain signaling at the level of r6, and show that we have identified the critical FGFs involved in hindbrain patterning.
FGF3 and FGF8 are required for neuronal development in r5 and r6
We next asked whether the loss of r5 and r6 in fgf3-MO; fgf8- embryos was evident at the level of neuronal development. We analyzed the reticulospinal interneurons of the hindbrain using the anti-neurofilament antibody RMO-44 (Pleasure et al., 1989). The reticulospinal neurons show rhombomere-specific cell body shapes and axonal projection patterns (Metcalfe et al., 1986
) (Fig. 4A). For example, at 48 h, RMO-44-labeled reticulospinal cells in r2, r4 and r6 have axons that cross the midline (Fig. 4A). In addition, the T interneurons, which are present in r7 and extend into the rostral spinal cord, have characteristically large, round cell bodies and T-shaped axonal branching patterns (Kimmel et al., 1985
) (Fig. 4A). The reticulospinal pattern appears largely normal in fgf3-MO embryos and fgf8- embryos (Fig. 4B,C). However, fgf3-MO; fgf8- embryos have T cells just caudal to the pair of Mauthner cells in r4 (Fig. 4D) and no r5 or r6 cells can be identified, providing further support for the loss of r5 and r6.
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Transplantation and mosaic analyses reveal r4 signaling activity
The loss of r5 and r6 upon reduction of FGF3 and FGF8 suggests that FGF3 and FGF8, from their overlapping domain in r4, normally promote the development of r5 and r6. To test for signaling activity from r4, we heterotopically transplanted r4 cells at bud/1-s stage, when we propose FGF signaling from r4 is promoting r5/6 development, to see if they could non-autonomously induce r5/6 fates. Seven out of 10 control transplants, from r4 to r4, incorporated into r4 (Fig. 5A,B-B''); however, donor cells could additionally be found in other rhombomeres (from r3-r6). The remaining three out of ten transplants incorporated into r5/6. The presence of donor cells outside of r4 may in part be due to dispersal; when such transplants are fixed at an earlier stage (4 s), 16/17 were in r4 (and overlapped with r3 or r5 in most cases; not shown). These controls, with the expression of r4 markers in donor cells (see Materials and Methods), demonstrate that we are largely targeting r4 in our transplants. Such isochronic transplants showed no non-autonomous induction of krox-20 expression either within the host hindbrain or if transplants were placed in other regions (not shown), so we heterochronically transplanted to the naïve ventral ectoderm of shield-stage hosts (Woo and Fraser, 1998) (Fig. 5A). r4 transplants in these hosts remain clumped together, reside in the ventral or lateral surface ectoderm on the yolk, and most express r4/5 markers (Fig. 5C-G''). Five out of 21 r4 transplants induced non-autonomous expression of krox-20 and val (Fig. 5D-G''). When hosts carrying r4 transplants are allowed to develop further, we find (in five out of 20 cases) non-autonomously-induced differentiated tissue that is normally associated with the r5/6 region, including pigment and ears (not shown). These transplants reveal that r4 cells have signaling activity outside of the hindbrain. To show that this signaling activity is restricted to the r4 region and is not a general activity of early hindbrain tissue, we transplanted cells from the r1 region to shield-stage hosts and found no expression of val either in the transplanted cells or induced by the transplant (n=8). We also transplanted cells from the r6 region and found that even though many of these transplants expressed val (5/8; Fig. 5H-H''), none showed non-autonomous induction of val. Taken together, these transplant experiments reveal a signaling activity that is strongly associated with, or centered on, r4.
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Misexpression of either fgf3 or fgf8 can induce ectopic expression of val and krox-20
To test whether FGF3 and FGF8 are sufficient to promote the development of r5 and r6, we asked whether misexpression of fgf3 or fgf8 is sufficient to induce expression of val and krox-20. To mimic the r4 transplantation experiments, we implanted beads coated with FGF8 protein in the ventral ectoderm of shield-stage embryos. At a low frequency (three out of 26 embryos), ectopic val and krox-20 expression was induced in non-neural ectoderm next to the FGF8 bead (Fig. 7A,B). When placed near the presumptive hindbrain in shield-stage embryos, FGF8 beads show strong induction (20/28 embryos) of val and krox-20 expression in the caudal hindbrain, anterior spinal cord and associated neural crest (Fig. 7C). To show that FGFs can signal in the neuroepithelium, we used a heat shock promoter to activate either fgf3 or fgf8 at 10 h/bud stage, a stage when we propose these FGFs are normally signaling from r4. We find that only fgf-expressing cells in the caudal hindbrain (Fig. 7D,E) or in surface ectoderm above the caudal hindbrain (Fig. 7E,G-G'') are able to induce ectopic val and krox-20 expression in neighboring cells within the caudal hindbrain; fgf-expressing cells elsewhere in the embryo, including the rostral hindbrain (Fig. 7F), do not induce val or krox-20. These results show that FGF signaling is sufficient to promote val and krox-20 expression, but it appears that only the caudal hindbrain is competent to activate val and krox-20 expression in response to FGF signaling at bud stage.
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DISCUSSION |
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r4 as an early-differentiating rhombomere
The results of our time-lapse imaging, in combination with the finding that isl1-positive cranial motoneurons (this work) and reticulospinal neurons (Mendelson, 1986; Hatta, 1992
) differentiate earliest in r4, support r4 as an early-differentiating rhombomere in zebrafish. Studies of rhombomere boundary development in chick and alligator (Vaage, 1969
; Pritz, 1999
), including time-lapse analysis in chick (Kulesa and Fraser, 1998
), show that the r5/6 and r3/4 boundaries develop earliest, suggesting that an r4-5 pro-rhombomere is established earliest in these animals. However, differentiation of hindbrain motor and reticular neurons occurs earliest in r4 in chick and mouse (Lumsden and Keynes, 1989
; Layer and Alber, 1990
; Sechrist and Bronner-Fraser, 1991
; Nardelli et al., 1999
; Pata et al., 1999
). Thus, there is support for the conserved early differentiation of r4, and further analyses of rhombomere formation and hindbrain neuronal differentiation in different vertebrates should reveal the extent of the conservation.
Redundant functions of FGFs in hindbrain patterning
We find that reduction of either fgf3 or fgf8 alone has little effect on hindbrain segmental patterning. However, severe reduction of both signals leads to loss of the entire r5/6 domain. Expression of either fgf3 or fgf8 does not depend on the other, and both transcripts are still expressed in the hindbrain of fgf3-MO; fgf8- embryos (L. M., unpublished). We therefore suspect that these FGFs function redundantly in the hindbrain. Although some functional FGF3 and FGF8 may remain in fgf3-MO; fgf8- embryos, increasing doses of fgf3 and fgf8 MOs together show no heightened hindbrain effects (L. M., unpublished). The complete loss of hindbrain expression of two FGF target genes, erm and pea3, shows that fgf3 and fgf8 are the critical FGFs used in zebrafish hindbrain patterning. We are not aware of another FGF expressed in the zebrafish hindbrain, nor is there any evidence for duplicate fgf3 or fgf8 genes in zebrafish (B. Draper, personal communication).
The expression of fgf3 in other vertebrates is consistent with a conserved role for fgf3 in r4 signaling. In frogs (Tannahill et al., 1992; Lombardo et al., 1998
), chicks (Mahmood et al., 1995
) and mice (Mahmood et al., 1996
; McKay et al., 1996
), fgf3 is expressed broadly in the middle of the early hindbrain, including strong expression in r4, and this expression precedes that of mafB in r5/6 (Cordes and Barsh, 1994
; Eichmann et al., 1997
) and krox-20 in r5 (Nieto et al., 1991
). In chicks and mice, fgf3 expression resolves into its more recognized r5/6 expression domain, which was initially observed in mice (Wilkinson et al., 1988
). However, in frogs, fgf3 becomes restricted to r4 (Lombardo et al., 1998
). If the r4-restricted expression of zebrafish fgf3 (although we do observe some weak r5/6 and possibly r3 expression during late gastrulation and early somitogenesis) is indicative of fgf3 expression in more primitive vertebrates, then this would provide even more support for a conserved role for r4 FGF signaling in the vertebrate hindbrain. The fgf3 knockout in mice has little effect on hindbrain patterning (Mansour et al., 1993
), similar to fgf3-MO zebrafish embryos (this work). fgf8 has not been found to be expressed in r4 in other vertebrates as in zebrafish; however, fgf4 is expressed in the chick hindbrain before initiation of mafB expression in r5/6 (Shamim and Mason, 1999
). We expect that in other vertebrates, additional FGFs to fgf3 will show early hindbrain expression, and we predict that loss of two or more of these FGFs will lead to dramatic defects in hindbrain patterning.
r4 as a conserved signaling center
Signaling from r4 has previously been implicated in hindbrain patterning. Transplantation studies in chick have shown that signaling from r4 regulates neural crest cell death in r3 and r5 (Graham et al., 1993), and krox-20 and follistatin expression in r3 (Graham and Lumsden, 1996
). Mice deficient in Hoxa1, which is not known to be expressed rostral to r4, show defects in the development of r3, including patchy loss of krox-20 expression (Helmbacher et al., 1998
). However, in these studies, the identity of the r4 signal(s) is unknown. Chick embryo transplantations have also implicated signaling from the r2-r6 region in promoting expression of krox-20 in r5 and mafB in r5 and r6 (Marín and Charnay, 2000b
). FGFs have been proposed to mediate this signaling because FGF-soaked beads can induce ectopic expression of krox-20 and mafB, and application of an FGF receptor-inhibitor drug inhibits krox-20 and mafB expression (Marín and Charnay, 2000a
). However, in these studies, it was not determined when and from which tissue (neuroectoderm or adjacent mesoderm) these signals acted and the FGFs required were not identified. In light of our findings, these studies all support a conserved role for FGF signaling from r4 in promoting the development of adjacent rhombomeres.
If r4 is truly a signaling center in the hindbrain, then r4 tissue should be both necessary and sufficient to promote development of r5 and r6. Our transplantation and mosaic analyses demonstrate that such signaling activity is centered in r4. Although our transplants express r4 markers such as hoxb1a, the transplants are not exclusively r4 tissue and we can not rule out the possibility that cells from other rhombomeres contribute to the signaling activity. Our gain- and loss-of-function studies with fgf3 and fgf8 show that these FGF signals mediate the signaling activity of r4. It is possible that other signals participate as well, as FGF8 beads promote val and krox-20 expression with low frequency outside of the hindbrain. Another factor that may contribute to the ability of both r4 transplants and FGF misexpression to induce r5/6 markers is that competence to respond to r4/FGF signaling may be tightly spatially and temporally regulated. Indeed, misexpression of fgf3 or fgf8 at bud stage is only able to induce val and krox-20 in the caudal hindbrain (this work). This apparent localized competence to respond to FGFs may be necessary because FGFs are used in many tissues for diverse functions. It is not clear how such a localized response is achieved, although the transcription factor POU2 (Reim and Brand, 2002) or the different FGF receptors (Thisse et al., 1995
; Sleptsova-Friedrich et al., 2001
) (S. Solinsky and L. M., unpublished) may play a role. The fact that the r4 transplants and the FGF8 beads can induce hindbrain markers in ventral ectoderm shows that these signals have neural inducing activity, which has previously been attributed to FGFs (Lamb and Harland, 1995
; Streit et al., 2000
; Wilson et al., 2000
). Intriguingly, FGFs themselves have been proposed to impart a differential competence to neural inducers in the zebrafish ectoderm (Koshida et al., 1998
).
Another conserved signaling center, the MHB, promotes patterning of the rostral hindbrain via FGFs (Irving and Mason, 2000). FGF signaling from the MHB has been proposed to interact with, or antagonize, signaling by retinoic acid (RA) from more posterior tissues to define the r1 territory (Irving and Mason, 2000
; Gavalas and Krumlauf, 2000
). We propose that a second, conserved, FGF-mediated signaling center, r4, be incorporated into this model of signals involved in early hindbrain patterning (Fig. 8). FGF3 and FGF8 signaling from r4 promote the development of r5 and r6, at least in part by promoting expression of val and krox-20. FGF3 and FGF8 from r4 may also promote the development of rostral rhombomeres 1-3, possibly acting with FGFs from the MHB. RA has been implicated in the regulation of val and krox-20 expression (reviewed by Gavalas and Krumlauf, 2000
). It is likely that FGF signaling from r4 and RA signaling interact in regulating r5/6 development, and val may be a key target on which these two signals converge.
|
Recently, fgf3 and fgf8 have been shown to be redundantly required for otic placode induction in zebrafish (Phillips et al., 2001; Maroon et al., 2002
). Otic placodes develop adjacent to r5, and fgf3 has previously been implicated in vertebrate otic placode induction (Represa et al., 1991
; Vendrell et al., 2000
). We have found that fgf3 and fgf8 are also required for other head periphery structures associated with the r5/6 region, including pharyngeal cartilages and cranial ganglia (L. M. and C. B. K., unpublished). Further studies should reveal whether FGF signaling from r4 acts directly on structures in the head periphery.
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ACKNOWLEDGMENTS |
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