m
D
ELSEVIER
Mechanisms of Development 61 (1997) 23-36
Specific craniofacial cartilage dysmorphogenesis gene expression in retinoic acid-treated Debra
L. Ellies”,“, Marie-AndrCe
Robert
M. Langillea,
AkimenkoaTb,c,
coincides with a loss of dlx zebrafish embryos
C. Cristofre Marc
MartinaT’,
EkkeralbTc,*
“L3epartnzent of Anatomy and Neurobiology, University of Offawa, Ottawa, Ontario, Canada bDepartment of Medicine, University of Ottawa, Otfawa, Ontario, Canada ‘L.oeb Institute for Medical Research, Ottawa Civic Hospital, Ottawa, Onfario. Canada
Received 27 March 1996; revised version received 13 September 1996; accepted 13 September 1996
Abstract Treatments of zebrafish embryos with retinoic acid (RA), a substance known to cause abnormal craniofacial cartilage development in other vertebrates, result in dose- and stage-dependent losses of dlx homeobox gene expression in several regions of the embryo. Dlx expression in neural crest cells migrating from the hindbrain and in the visceral arch primordia is particularly sensitive to RA treatment.
The strongest effects are observed when RA is administered prior to or during crest cell migration but effects can also be observed if RA is applied when the cells have entered the primordia of the arches. Losses of dlx expression correlate either with the loss of cartilage elements originating from hindbrain neural crest cells or with abnormal morphology of these elements. Cartilage elements that originate from midbrain neural crest cells, which do not express dlx genes, are less affected. Taken together with the observation that the normal patterns of visceral arch dlx expression just prior to cartilage condensation resemble the morphology of the cartilage elements that are about to differentiate, our results suggest that dlx genes are an important part of a multi-step process in the development of a subset of craniofacial cartilage elements. 0 1997 Elsevier Science Ireland Ltd. All rights reserved Keywords:
Cartilage; Head skeleton; Homeobox genes; Neural crest; Retinoic acid; Zebrafish
1. Introduction Development of the head skeleton depends in part upon interactions between neural crest cells that populate the visceral arches and the epithelial cells of the arch primordia. Cranial neural crest cells originate from the hindbrain rhombomeres and from the mesencephalon. Once they have reached their terminal location, for example, within the visceral arch primordia, an interaction between crest cells and the epithelium is required for the ectomesenchyme that originates from neural crest cells to condense and differentiate into cartilage (Hall, 1987). This phase of cytodifferentiation is then followed by morphogenesis. The morphology of the cartilage elements formed in a given arch depends on the identity of the neural crest cells * Corresponding author. Loeb Institute for Medical Research, 725 Parkdale Avenue, Ottawa, On, KlY 4E9, Canada. Tel.: +l 613 7985555, ext. 6033: fax: +l 613 7615365; e-mail:
[email protected]
that migrate into that arch (Noden, 1983; Langille, 1993; Langille and Hall, 1993). Considerable attention has been paid to the genetic mechanisms that regulate this process. Cells from specific rhombomeres will populate defined arches and positional information carried by the neural crest cells seems to be transferred to the arches and will influence the morphology of the skeletal elements (Noden, 1983). Premigratory neural crest cells of the hindbrain express a rhombomere-specific combination of Hox genes (Hunt et al., 1991) which is thought to give them their unique segmental identity along the antero-posterior axis of the hindbrain (Hunt and Krumlauf, 1991). This unique combination of neural crest cell subpopulations carry their prepattemed Hox information into each visceral arch. Selective inactivation of Hox genes leads to homeotic transformations by which skeletal elements that originate from one arch will take the likeness of the elements that normally originate from another arch. Furthermore, ecto-
0 1997 Elsevier Science Ireland Ltd. All rights reserved 0925-4773/97/$17.00 PII SO925-4773(96)00616-8
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D.L. Ekes el al. /Mechanisms of Development 61 (1997) 23-36
pit Hex gene expression can lead to craniofacial abnormalities in both mouse and zebrafish (Balling et al., 1989; McLain et al., 1992; Alexandre et al., 1996). Some of these abnormal craniofacial phenotypes are similar to effects seen in mice following administration of retinoic acid (RA) (Kessel et al., 1990; Alexandre et al., 1996): Such craniofacial defects involve a failure of the frontonasal mass to enlarge and fuse with the nasal processes and maxillae (Wedden, 1987), classified as severe clefting of the primary palate, or cleft palate. Retinoic acid is known to directly regulate Hox gene expression by binding to regulatory elements found within the Hox clusters. The retinoic acid response elements (RAREs) found 3’ to the Hox clusters are known to upregulate Hox gene expression. However, RAREs have also been found in the mouse and chick Hoxbl flanking sequences (Marshall et al., 1994; Studer et al., 1994), and these RAREs were shown to downregulate the expression of Hoxbl within the hindbrain (Studer et al., 1994). Although a lot is known about the role of Hox genes in conferring positional identity for the different rhombomeres and arches, little is known about other genetic factors that determine which subset of neural crest cells will contribute to the visceral arches and which factors influence the differentiation program of these neural crest cells. We have studied the role of another family of homeobox genes, the distal-less (dlx) genes, on craniofacial development. It has previously been shown that migrating crest cells, apparently populating the visceral ‘arches, express the dlx2 gene, both in zebrafish (Akimenko et al., 1994) and in mice (Bulfone et al., 1993). Following crest cell migration, the lateral mesenchymal cells of the visceral arches express a complex combination of several dlx genes, which is temporally regulated (Akimenko et al., 1994). Expression of rodent dlx genes has been shown in arch mesenchymal cells at the time of cartilage condensation and differentiation (Robinson and Mahon, 1994; Simeone et al., 1994; Zhao et al., 1994). In the present study, we report that treatment of zebrafish embryos
with exogenous retinoic acid results in a loss of dlx expression in several regions of the embryo. Dlx expression in the migrating cranial neural crest cells derived from rhombencephalic regions and in the visceral arches they populate is particularly sensitive to RA treatment. The loss of dlx expression in these migrating crest cells and arch primordia correlates closely with the loss or abnormal development of craniofacial cartilage components, suggesting an essential role for dlx genes in development of the head cartilage. 2. Results 2.1. Dlx expression
during visceral arch cartilage
differentiation
A specific temporal and spatial combinatorial expression of zebrafish dlx genes during development of the visceral arches up to 55-60 h has been reported (Akimenko et al., 1994). To better correlate patterns of dlx expression with the development of cartilage elements, we examined dlx expression in visceral arches at the time of arch differentiation. Craniofacial cartilage begins to condense between day 2 and day 3 (Kimmel et al., 1995; our unpublished observations). With respect to the chondrocranium, condensation begins at day 2 and by day 3 matrix deposition is evident. Cartilage differentiation in the visceral arches occurs later than the chondrocranial elements. At day 3, the visceral arch prechondrocytes are condensed and matrix production can only be seen by alcian blue staining by the end of the third day (Schilling and Kimmel, 1994; Schilling et al., 1996; our unpublished observations). Prior to and at day 2, expression patterns of dlx genes can be seen in the mandibular, hyoid, and branchial arches (Fig. lA,B). From 2.5 days (d), the visceral arch primordia (just prior to condensation) express dlx genes in areas that correlate with the future discretely patterned arch condensations. Thus, during the third day, cells in the position of
Fig. 1. Progressively restricted expression patterns of dlx genes in visceral arches prefigure the morphology of condensing cartilage elements, (A) Ventral view of a 2 d embryo showing dlx4 gene expression in the mandibular arch (arrow), extending to the opening of the oral cavity (arrowhead). (B) A sagittal view of the same embryo showing dlx4 expression in the mandibular, hyoid and in branchial arches 1, 2, and 3. (C) Sagittal view of a 2.5 d embryo showing dlx2 expression just prior to visceral arch cartilage condensation. The mandibular and hyoid arch pre-cartilaginous derivatives are becoming visibly discrete. (D) Al&m blue staining of a 3 d larva showing the cartilage of the jaw. The quadrate (Q) and Meckel’s cartilage (MC) derive from the mandibular arch. The hyosymplectic (HY), the ceratohyal (C) derive from the hyoid arch. The morphological patterns of these condensed cartilage elements resemble the patterns of dlx2 expression shown in (C). (E) Dlx4 expression in a 4 d larva in areas that correspond to the developing ceratobranchials. (F) Craniofacial cartilage elements derived from the visceral arches at 4 d. 1-3, branchial arches 1 to 3: ABC, anterior basicranial commissure; C, ceratohyal; CB, ceratobranchials; F, forebrain; H, hyoid arch; HY, hyosymplectic; M, mandibular arch; MC, Meckel’s cartilage; OP, olfactory placode; OV, otic vesicle: P, pectoral fin bud; Q. quadrate. Scale bar: 107 nrn for (A,B,E). 67 pm for (C). 32 pm for (D), and 51 pm for (F). Fig. 2. DLr expression correlates with neural crest derived chondrocyte condensations. (A,B) Transverse sections through the head of 2 d (A), and 3 d (B) zebrafish. The cells expressing dlxl transcripts (A, arrowhead) occupy a position similar to those that will condense to form Meckel’s cartilage 1 day later (B, arrow). (CD) Transverse sections through a more posterior region of the head of 2 d (C). and 3.5 d (D) zebrafish. The position of cells expressing dk3 (C, arrowhead) corresponds to that of cells that will condense to form the ceratobranchial cartilage elements (D, arrow). T, trabeculae; hollow arrowhead, oral cavity; F, forebrain. Scale bar: 2.5 pm for (A), 14 pm for (B). 18 nrn for (CD).
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D.L. Ellies et al. /Mechanisms of Development 61 (1997) 23-36
Neural Crest Migration Gastrulation
I
Segmentation Of Arch Primordio
Ilead /Jaw Cartilage BegIns Condensation
Fig. 4. Retinoic acid treatment schedule. The time is indicated as hours post-fertilization. The approximate times of relevant developmental stages are indicated.
Fig. 3. Schematic representation of neural crest derived craniofacial cartilage components. The visceral arch from which the various cartilage components derive is indicated in bold. M, mandibular arch; H, hyoid arch; B, branchial arches. The trabeculae originate from midbrain neural crest cells and the anterior basicranial commissure is the mesodermallyderived component of the posterior neurocranium with which the jaw articulates.
the developing quadrate express dlx2 (compare Fig. lC,D), and dlxl (not shown); cells in the position of the differentiating Meckel’s cartilage express dlxl (Fig. 2A) and dlx2 (out of focus in Fig. 1C); cells forming the ceratohyal express dlx2 (Fig. lC), dlxl (not shown), and dlx4 (not shown); and cells of the hyosymplectic anlage express dlx2 (Fig. lC), dlxl and dlx3 (not shown). At 3 d, condensations of the branchial arches, whose shape already prefigures the ceratobranchial elements (Kimmel et al., 1995), express dlx3 (Fig. 2C). Morphogenesis and differentiation of the various visceral arch cartilage elements continue during the fourth day and the trabeculae (anterior neurocranial), basihyal, quadrate, hyosymplectic, and ceratohyal (viscerocranial) components can be seen elongating. Meckel’s cartilage now articulates with the anterior tip of the quadrate, and the hyosymplectic articulates with the neurocranium at the anterior basicranial commissure (Fig. 1F and Fig. 3). Matrix production within the basibranchial and ceratobranchial components becomes visible by day 4. During this time, dlx expression is progressively down-regulated
and transcripts are only detectable in areas that correspond to those cartilage elements in which alcian blue staining is weak or undetectable. Thus, dlx2 expression is no longer detectable after 3 d and dlx4 expression is then restricted to the ceratobranchials (Fig. 1E). At day 5, dlx3 transcripts are detected in the condensations of the operculum, as well as the circumoral area (not shown). The above results suggest that dlx expression pattern within the visceral arches is progressively restricted to areas whose morphology prefigures the cartilage elements that will form during visceral arch differentiation and persists until a stage corresponding to matrix production. 2.2. Dlx gene expression in visceral arch prirnordia particularly sensitive to exogenous retinoic acid
is
In addition to the primordia of the visceral arches, cells in the ventral forebrain, olfactory placodes, otic placode and vesicle, pectoral fin buds, and median fin fold express multiple dlx genes (Akimenko et al., 1994). When embryos are treated with a dose of 1 PM RA for 2 h starting either during or immediately after gastrulation, (6 h, 8 h, or 10 h after fertilization; Fig. 4), we observe a complete loss of all dlx gene expression in the head region when measured at 24 h (Fig. 5A,B) while weak expression of dlx3 persists at the caudal end of the developing median fin fold (Fig. 5B). Development of the embryos is also extremely retarded (Fig. 5A,B) and the loss of dlx expression may reflect the loss of anterior head structures. When the dose of RA is reduced from 1 PM to 0.1 PM and administered at either 6 h, 8 h, or 10 h, transcripts of the dlx2, dlx3, and dlx4 are seen in structures such as the
Fig. 5. Effects of early RA treatments on dlx 3 expression. (A.0 Control embryos (DMSO-treated) that show the normal patterns of dlx3 expression at 16 h. side view (A) and 27 h. dorsal view (C). (B) Embryo treated with 1 PM RA at 6 h. D1x3 expression was determined at 27 h and is undetectable in the olfactory placode area, otic vesicle and visceral arches. Some dlx3 transcripts are detected at the caudal end of the median fin fold (arrowhead). Development of RA-treated embryos is severely retarded. Compare the embryo in (B) with those in (A) and (C). (D) Embryo treated with 0.1 PM RA at 10 h. DM expression in the pharyngeal arches is lost, whereas it is reduced in the olfactory placodes and normal in the otic vesicle. Note that the otic vesicle (OV) is located in a position closer towards the optic cup (OC), consistent with the loss of the midbrain-hindbrain border region that was previously described (Holder and Hill, 1991; Hill et al., 1995). Other symbols are as described in Fig. 1. Scale bar: 79 pm for (A), 74 pm for (B), 89 pm for (CD). Fig. 6. RA effects on dlx2 expression in migrating neural crest cells, (A-C) Control embryos. (A) Fourteen hour embryo showing dlx2 (blue; hollow arrowheads) and krx20 (red) expression; (B) dLr2 and eng2 (arrowhead) at 16 h, and (C) dorsal view of a 20 h embryo showing dln2 (blue) and krx20 (red). (D) dln2 and krx20 expression at 14 h in embryos that were treated at 12 h with 0.1 PM RA. Dlx2 expression is abolished but krx20 expression (red) is normal. F, forebrain: R3, rhombomere 3: R5, rhombomere 5. Scale bar: 100 Frn for (A,C) 68 pm for (B). 75 Frn for (D).
D.L. Ellies et al. /Mechanisms
ventral forebrain, the otic vesicle, and the olfactory placodes but not in the visceral arches, as shown in Fig. 5C,D for dlx3. Therefore, expression of dlx genes in visceral arch primordia seems to be particularly sensitive to RA treatment.
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2.3. RA treatment affects dlx2 expression neural crest cells
in migrating
Zebrafish hindbrain neural crest cells express dlx2 during their migration into the visceral arches from 12 h to 19
28
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h (Akimenko et al., 1994). To determine whether the RAdependent loss of dlx gene expression in the visceral arch primordia can be related to altered dl.x2 expression in migrating neural crest cells, we treated embryos with varying doses of RA before and during neural crest cell migration. First, we used double labeling to better localize the position of dlx2-expressing cells. We performed in situ hybridization with probes for dlx2 and for either krx20, a gene expressed in cells of rhombomeres 3 and 5 (Oxtoby and Jowett, 1993) or for erzg2, which is expressed in cells at the midbrain-hindbrain border (Ekker et al., 1992b). In control embryos, neural crest cells expressing dZx2 migrate into the periphery along pathways adjacent to rhombomeres 1, 2, 4 and 6 (Fig. 6A-C) (Akimenko et al., 1994). Furthermore, double-labeling with dlx2 and eng2 shows that those cells that express dZx2 and appear to be migrating towards the mandibular arch primordium originate from the hindbrain but that their pathway seems to deviate rostrally (Fig. 6B). We never observed any dZx2expressing cells migrating from the midbrain, at 14 h nor at any other stage. We cannot totally exclude the possibility that neural crest cells originating from the midbrain express dZx2 but, contrarily to crest cells originating from the hindbrain, they would initiate expression of this gene after commencement of their migration, possibly after contact with dlx2-expressing hindbrain crest cells. Our observations would suggest that, although neural crest cells from both the hindbrain and the midbrain are thought to contribute to the mandibular arch in fish (Hall and Horstadius, 1988; Langille and Hall, 1988a; Langille and Hall, 1988b; Schilling and Kimmel, 1994), only those migrating from the hindbrain express dlx2 prior to entering the mandibular arch. In embryos treated with a dose of 0.1 PM RA, administered at either 12 h (Fig. 6D), or at earlier times (not shown), dlx2 expression is no longer detected in migrating crest cells when measured at 14 h. Krx20 expression is still detected in embryos that received this dose of RA at 12 h (Fig. 6D). When 0.1 PM or 1 PM RA was administered at either 14 h or at 16 h, dlx2 expression is reduced and transcripts are restricted to two thin stripes of cells, one anterior and one posterior to the otic vesicle (data not shown). In summary, RA administration before neural crest cell migration abolishes dlx2 expression within the migrating neural crest cells, whereas treatments during neural crest cell migration result in apparently lower levels of dlx2 transcripts in migrating neural crest cells. 2.4. RA affects dlx expression in the visceral arches
and cartilage development
To determine whether the loss of dlx gene expression in visceral arch primordia in response to exogenous RA can be related to abnormal development of the craniofacial
cartilage that will develop subsequently, we performed a more detailed analysis of the effects of RA. Various doses of RA were administered immediately prior to, during and after neural crest cell migration. Dlx gene expression was then determined in 27 h embryos and cartilage development was analyzed in 5 day larvae. The data are compiled in Table 1 for all dlx genes and representative examples of dlx2 expression and altered cartilage development are shown in Fig. 7. Treatments with RA starting during gastrulation (6 h, 8 h) or at the end of gastrulation (10 h) always led to the total loss of dlx expression in the arches and to the absence of visceral arch-derived cartilage elements as well as of other head structures (Fig. 5B,D). We therefore chose to administer RA at later stages for the more detailed analysis presented in Fig. 7 and Table 1. There was little variability in the effects of RA treatments on dlx expression as identical expression patterns were observed in all embryos of a given group of at least five embryos. We did, however, observe some variability in the resultant cartilage phenotypes as indicated in Table 1. 2.5. Effects of 0.1 PM RA: mandibular
arch
Zebrafish embryos treated with 0.1 PM RA at 12 h, 14 h, 16 h, 19 h, and 24 h, all display reduced levels of dlx2 and dlx3 expression in the mandibular arch regardless of the schedule of RA administration (Table 1, Fig. 7B,C), whereas mandibular dlx4 expression is either absent or much reduced (Table 1). Development of the two cartilage elements derived from the mandibular arch, the quadrate and Meckel’s cartilage, as measured in 5 day larvae, shows defects which are dependent upon the schedule of RA administration. Thus, embryos treated at 12 h, 14 h, 16 h and 19 h show either a loss, or altered morphology, of the mandibular cartilage components (Table 1, Fig. 7F,G), whereas embryos treated at 24 h display no loss of cartilage (Table 1) except for a modest ventral deviation of Meckel’s cartilage. 2.6. Effects of 0.1 PM RA: hyoid arch Embryos treated with 0.1 PM RA at various stages show strong reductions in the intensity of dlx3 and dlx4 gene expression in the hyoid arch (Table 1). Dlx2 expression is generally less affected in intensity but the patterns of expression are sometimes abnormal and sometimes fused with the patterns of expression of this gene in the first branchial arch (Fig. 75). A dose of 0.1 PM, given at 12 h, results in the absence or abnormal morphology of the hyoid arch cartilage elements in a significant proportion of the embryos. Treatment at either 14 h, 16 h or 19 h has generally weaker effects on the ceratohyal, the basihyal, or the hyosymplectic cartilage elements (Table 1) and hyoid cartilage elements are normal in embryos that received RA at 24 h.
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Table 1 Visceral
arch dlx expression
and cartilage
development
Wildtype dlx expression
dlx expression
Mandibular
Hyoid dlx expression
Branchial
Branchial
arch I dlx expression
arch 2 dlx expression
dlx2 dlx3 dlx4 dlx2 dlx3 dlx4 dlxl dlx2 dlx3 dlx4 dlx2 dlx4
Number of embryos assessed Mandibular arch cartilage Meckel’s cartilage
Quadrate
Hyoid arch cartilage Basihyal Ceratohyal
Branchial arch cartilage Basibranchial and ceratobranchials
Other cartilage elements Trabeculae/ethmoid Posterior
neurocranial
RA administration
base
retinoic acid treatment dose and stage
lo-‘M
1O-7 M
10m7 M
IO-‘M
lo-‘M
10-6M
10m6M
12 h
14 h
16 h
19 h
24 h
12 h
14 h
10-6M 16 h
10-6M 19 h
10-6M 24 h
RiR+ A R A A na. R A A A A
R+ n.a. A R n.a. A n.a. AP, R n.a. A R A
R+ R A AP, AP, AP, A AP, AP, AP, AP, AP,
R R+ R+ N R+ R+ n.a. N R+ R+ N A
R R+ A N R+ A A R R+ R+ R A
A n.a. A R+ na. A na. Ri n.a. A A A
A n.a. A R+ n.a. A n.a. R+ n.a. A A A
A A A AP, A A A AP, AP, A AP, A
R+ A A R+ A AP, A AP, AP, AP, AP, A
A A A AP, R+ A A A AP, R+ A A AP, R+ A
16
11
30
24
18
17
7
40
22
13
VD+4 A5 VD,F7
VD+9 Al Nl
VD+28 Al Nl
VD+21 VD3
VD9 N9
A 17
A7
A 40
A 12 AM 1
VD+lOVD+6 A5 Al VD,Fl N4
VD+28 Al Nl
VD24
N18
A 17
A7
A 40
A 15 AM3 Fl N3 A 18 AM 1 VD+l N2
A 16
N6 A 24 VD21 N6 A3 VD21 N6 A3
N4 A 20 VD20 N4
N 18
A 17
A7
A 40
N18
A 17
A7
A 40
AM20 N4
N18
A 17
A7
A 40
VD+4 VD,Fl A 11 VD+4 VD,FI A 11
Hyosymplectic
following
VD+9 N2 VD+6 N5 VD+6 N5
R R+ R+ R+ R+ Ri R+ R+
R+
R+ R+ R+
R+ R+ R+ R+ R+
A 20 N2 A 18 VD+l N3 A 18 VD+l N3
A 13
A 13 A 12 AM1 A 10 AM3
VD+4 Al2
VD+4 N7
AM21 N6 A3
AM20 N4
Nl8
A 17
A7
A 40
A 19 N3
A 13
N6 s4 N 16
N7 s4 N 11
N7 s3 N 30
N4 so N 24
N7 so N 18
NO s 17 N 17
NO s7 N7
NO s 20 N 40
N4 S7 N 22
N4 S6 N 13
Dlx expression was analyzed at 27 h. At this time, only dlx2, dlx3 and dlx4 are expressed in the mandibular and hyoid arch, dlxl expression is only found in branchial arch 1, dlx2 expression is found in all of the branchial arches, dlx3 is only found in branchial arch 1, and dLx4 expression is found in branchial arches 1 and 2. Dlx expression is shown as N, normal; R, reduced; R+, severely reduced; AP. abnormal pattern; A, absent; or na., not assessed. Gene expression was determined on a minimum of five embryos per gene and was comparable for all embryos in a given group. The craniofacial cartilage was analyzed at 5 d and was either N. normal; A, absent; showed VD+, severe; VD, modest ventral deviation; AM, of abnormal morphology, F, fused with pair of adjacent elements; S, shortened trabeculae (see legend to Fig. 7). The number of larvae in each group showing the phenotype is indicated at the right of the symbol.
2.7. Effects of 0.1 pM RA: branchial
arches
Expression of dlx genes in branchial arches 1 and 2 seems to be sensitive to RA treatments in a manner that is dependent both upon the gene and the schedule of administration. Thus, as noted for dlx expression in the other arches, the earlier treatments (12 h, 14 h) result in stronger reductions or in the
absence of dlx expression in branchial arches 1 and 2 (Table 1, Fig. 7B). Furthermore, dlx2 is consistently less affected than the other dlx genes by the RA treatments (Table 1). The two branchial arch-derived cartilage components which have developed sufficiently at 5 d, the time at which embryos were fixed for cartilage staining, are the basibranchials and the ceratobranchials. With a dose of 0.1
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Fig. 7. RA affects pharyngeal arch dlx expression and cartilage development, (A-D,I,J) dlx2 expression in embryos at 27 h; (E-H,K,L) craniofacial cartilage from 5 d embryos. (A,E) Control embryos. (B,F) Embryos treated at 12 h with 0.1 PM RA. Dlx gene expression in pharyngeal arches is reduced and many cartilage components are lost. (C,G) Embryos treated at 19 h with 0.1 PM RA. The reduction in d1.r expression in the pharyngeal arches is not as severe as in embryos that received the same dose at 12 h (B). Cartilage elements are present but their morphology is abnormal, resembling the embryo shown in (L). Cartilage derived from the branchial arches is also shifted posteriorly. (D,H) Embryos treated at 16 h with 1 PM RA. This treatment results in the loss of d/x expression in the mandibular arch and in a total loss of pharyngeal arch-derived cartilage components. (I,J) Sagittal views of a control embryo (I) at 27 h, and of an embryo treated at 14 h with 0.1 PM RA (J). The patterns of dlx2 expression in the positions corresponding to the hyoid and first branchial arch appear to be fused and non-uniform compared to the control embryo. (K,L) Sagittal views of a control larva (K) and of a larva which received 0.1 FM RA at 14 h (L). Both first arch elements show abnormal position within the face. The quadrate (Q) is shortened and curves ventrally away from rather than antero-dorsally towards the front of the mouth. The positioning of Meckel’s cartilage (MC) is even more aberrant. The ceratohyal (C) is positioned more ventrally than normal. The trabeculae (T) appears shortened but in proper position and probably reflects the overall shortening of the rostra1 region of the face. EP, ethmoid plate; BP. basilar plate. Other symbols as in Fig. 1. Scale bar: 147 pm for (A), 109 pm for (B), 90 pm for (C), 119 pm for (D), 159 pm for (E), 145 ym for (F). 175 pm for (G,L), 133 pm for (H), 32 PM for (1.J). and 114 nm for (K).
Fig. 8. Increased number of apoptotic cells in the hindbrain and visceral arch areas of zebrafish embryos following RA treatment. Dorsal views of 18 h control embryos (A) or of embryos that received 1 pM (B) or 0.1 PM (C) RA at 12 h or 1 PM RA at 16 h (D). An increased number of apoptotic cells in the hindbrain region (arrows), as revealed by TUNEL staining. is seen only in those embryos that received RA at 12 h. Scale bar: 100 pm.
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PM RA, we observe a loss of cartilage in most of the embryos that were treated at 12 h (Fig. 7F), whereas embryos treated at later stages with the same dose display normal cartilage morphology but often show an apparent posterior shift in the position of the elements at the time of differentiation (Fig. 7G). 2.8. EfSects of I pM RA Zebrafish embryos treated with 1 PM RA at 12 h, 14 h, 16 h, 19 h, and 24 h, all display a loss of dlx2, dlx3, and dZx4 expression in the mandibular arch (Table 1, Fig. 7D), except for dZx2 in embryos treated at 19 h, whose expression is much reduced. Hyoid arch expression of dlx3 and dlx4 is also abolished or strongly reduced by these treatments, whereas the dlx2 patterns are less affected in intensity but show abnormal morphology (Table 1). The cartilage components derived from the mandibular arch (the quadrate and Meckel’s cartilage) and hyoid arch (the ceratohyal, basihyal, and hyosymplectic) are always lost in embryos treated with 1 PM RA at 12 h, 14 h, or 16 h (Table 1, Fig. 7H). These cartilage elements are generally absent or affected in their morphology in embryos that received 1 PM RA at either 19 h or 24 h. Expression of dlx genes in branchial arch primordia seems to be less affected by RA treatment than it is in the mandibular and hyoid arches. Cells in the branchial arch primordia of embryos that received 1 PM RA express at least a subset of dlx genes, with dlxl and dlx4 being more sensitive to RA treatment (Table 1). At a dose of 1 PM RA, the branchial arch cartilage derivatives are completely absent regardless of the time of administration, except for a small number of fish that received RA at 19 h (Table 1, Fig. 7H). Because RA treatments at a dose of 1 PM produce much stronger effects on dlx gene expression and cartilage formation than treatments with a dose of 0.1 FM, we tested intermediate doses ranging from 0.2 PM to 0.8 PM, administered at either 10 h or 16 h. These treatments result in a loss of expression for all dlx genes in the mandibular arch, and abnormal patterns of dlx gene expression in the hyoid arch. Intermediate doses of RA also result in a loss of both the mandibular and hyoid arch cartilage derivatives (data not shown). In all of the above experiments, the trabeculae and the posterior neurocranial base were much less affected, if at all, by the RA treatments (Table 1, Fig. 7). It is interesting to note that these two regions never show any dlx expression (Fig. 1). To determine whether the loss of dlx expression and the abnormal cartilage development following RA treatment could be attributed to a general loss of cranial neural crest cells, we examined zebrafish embryos for the presence of the trigeminal ganglia, another cranial neural crest cell derivative, by staining with the HNK-I monoclonal antibody (Holder and Hill, 1991). Treatments with 0.1 PM RA
at 12 h resulted in a small reduction in the number of neurons in the trigeminal ganglia and in abnormal extension of axons over the surface of the head when embryos are stained at 27 h (not shown), whereas treatment with 1 PM RA at 19 h resulted in normal or nearly normal number of neurons in the trigeminal ganglia but in the loss of some of the projections (not shown). There is a significant contribution of the neural crest to trigeminal ganglia neurons, based on studies in other vertebrates, but some cells are also of placodal origin (Noden, 1978). In our experiments, we cannot assess the origin of the few cells affected by RA, as other placodal derivatives such as lateral line ganglia can also be affected by RA treatments (Holder and Hill, 1991). However, the very small reductions in numbers of cells suggest that the loss of dlx expression in migrating neural crest cells and in visceral arch primordia following post-gasttulation RA treatments does not result from a general loss of hindbrain neural crest. By contrast, when embryos received RA at earlier stages (e.g. during gastrulation), we observed larger decreases in the number of neurons in the trigeminal ganglia (not shown), as previously reported by Holder and Hill (1991), which is also in accordance with the other profound defects which we observed (Fig. 5) following RA treatments at these early stages. We also performed the TUNEL procedure to look for apoptotic cells. Embryos that received either a dose of 1 PM or 0.1 PM RA at 12 h showed an increased number of apoptotic cells in the hindbrain and visceral arch area when compared to control at 18 h (Fig. 8A-C) or at 24 h (not shown). However, the same doses of RA given at 16 h did not lead to an increase in the number of apoptotic cells at either time (Fig. 8D and data not shown). 3. Discussion Treatments of zebrafish embryos with exogenous retinoic acid are known to modify the development of the midbrain and hindbrain, affect cranial ganglion formation (Holder and Hill, 1991) and respecify the position of the Mauthner neuron (Hill et al., 1995). RA treatments also cause duplication of the retina (Hyatt et al., 1992), result in the loss of anterior heart structures (Stainier and Fishman, 1992) and affect the expression of retinoic acid receptors (RAR), notably that of RAR-?/ in the visceral arches (Joore et al., 1994). We have now shown that treatment of zebrafish embryos with retinoic acid at various stages of their development affects both the expression of dlx genes in the primordia of the visceral arches and the morphology of the differentiating cartilage that originates from these arches. We observe a strong correlation between the effects of a particular RA treatment on dlx gene expression and cartilage development. When zebrafish embryos are treated with doses of RA ranging from 0.1 PM to 1 PM before neural crest cell migration, expression of d/x2 is no longer observed in the migrating crest cells which will populate
D.L. Ellies et al. /Mechanisms
the visceral arches (Fig. 6) and subsequently, expression of all dlx genes is lost or largely reduced in the primordia of all visceral arches at these doses (Table 1, Fig. 7B). These treatments also result in the loss of cartilage components or severe morphological abnormalities (Table 1, Fig. 7F). When the same doses of RA are administered during or immediately after neural crest cell migration, dlx2 expression within the migrating crest cells is reduced. Expression of dlx genes is reduced or abolished in the mandibular and hyoid arches (Table 1) with a subsequent loss of cartilage components in embryos treated with 1 PM RA (Table 1, Fig. 7H) or abnormal morphology in embryos that received 0.1 PM RA (Table 1, Fig. 7G). However, expression of dlx genes in the branchial (gill) arches is generally less severely affected by RA treatments and this can be correlated with more moderate effects on cartilage formation. Nevertheless, the effects of RA on gene expression and on cartilage formation remain coupled over the dose range and with the different schedules of administration. It is not clear how treatment with RA impairs expression of dlx genes. Direct regulation of dlx genes by RA has yet to be demonstrated. There are presently no cell culture systems that are known to express dlx genes. The immediate 5’-flanking region of the Xenopus Xdll-2 gene is the only regulatory region of a vertebrate dlx gene that has been characterized (Morass0 et al., 1994; Morass0 et al., 1995) and this region does not contain sequences that match consensus RAREs (Umesono et al., 1988; Studer et al., 1994). Sequence analysis of about 1.5 kb of 5’-flanking region of both zebrafish dlx3 and dlx4 (Ellies, 1995) revealed elements that resemble RAREs but none that match perfectly. We cannot at this time exclude the possibility that the loss of gene expression in RA-treated zebrafish embryos is simply the result of the absence of cells that normally express dlx genes. However, only the early RA treatments result in increases in cells undergoing apoptosis in the hindbrain and not a large number of cells were dying (Fig. 8). RA treatments at later stages did not result in any increase in apoptotic cells. Furthermore, the presence of normal or nearly normal numbers of trigeminal ganglia neurons, which also derive in part from hindbrain neural crest, indicates that the loss of dZx expression and the abnormal head cartilage morphogenesis after RA treatments, especially the later ones, is unlikely to be due to a general loss of hindbrain neural crest cells. It is still possible that the different progressions of differentiation of trigeminal neurons and neural crest-derived cartilage would confer to them a different sensitivity to RA treatments. An intriguing finding is that expression of dlx genes and cartilage formation are affected by RA treatment that occurred before the onset of neural crest cell migration and before the onset of dlx2 expression, the first dlx gene to be expressed in migrating crest cells and arch primordia. One possible explanation is that RA treatments during or at
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the end of gastrulation (6 h, 8 h, or 10 h) result in severe perturbations in neural tube patterning leading to the respecification of neural crest cells. This would be consistent with studies carried out in other vertebrate models (Morris-Kay et al., 1991; Conlon and Rossant, 1992; Marshall et al., 1992). RA treatments could also affect the subsequent migration of hindbrain crest cells as suggested for other vertebrates (Lee et al., 1995; Gale et al., 1996). The respecification or impaired migration of crest cells would severely perturb visceral arch development and result in the concomitant abnormal expression of all dlx genes. Interestingly, misexpression of the zebrafish HoxuI gene, which acts as a regulator of hindbrain patterning, results in down-regulation of dlx2 and cartilage defects that resemble some of those reported in the present study (Alexandre et al., 1996). However, we have further shown that RA treatments at developmental stages that coincide with neural crest cell migration or with the end of migration also lead to a loss of dlx gene expression and of cartilage formation. This last observation further supports a direct effect of RA on the expression of dlx genes and a requirement for normal expression of dlx genes for proper cartilage formation. It is thus possible that RA may exert two effects leading to a loss of dlx expression and perturbed craniofacial cartilage development: an effect through perturbations in neural tube patterning when given at early stages and a direct effect on dlx genes when given during or after crest cell migration. The lack of dlx2 expression in migrating crest cells that we have shown (Fig. 5) could in turn prevent induction of the other dlx genes in the arch primordia. It has previously been suggested that expression of genes like dlx3 and dlx4 in the visceral arch primordia is activated, following migration, in the same crest cells that express dlx2 and/ or in mesodermally derived cells as a result of interactions with these dlx2-expressing crest cells (Akimenko et al,, 1994). This could also reconcile our observations that dZx2-expressing neural crest cells seem to migrate only from the hindbrain, whereas expression of dlx genes is uniform across the mandibular arch, which is thought to receive neural crest contributions from both the midbrain and the hindbrain (Hall and Horstadius, 1988; Langille and Hall, 1988a; Langille and Hall, 1988b; Langille and Hall, 1993; Schilling and Kimmel, 1994). Cross-regulation of dlx gene expression would be consistent with the observation that dlx genes are, in general, equally affected in the visceral arches following treatment with RA. In addition to the possible roles of dlx genes during crest cell migration and patterning in the early primordia as described above, the patterns of dlx expression at later stages coincide with cartilage formation. In the present study we have shown a strong correlation between the expression patterns of the zebrafish dlxl, dlx2, dlx3, and dZx4 genes and subsequent visceral arch cartilage differentiation. Moreover, when dlx expression in a specific facial region is abolished by exogenous RA, there is a
34
D.L. Elks
et al. /Mechanisms
direct one to one loss, or abnormal modification, of the visceral arch cartilage elements in that area alone. Tellingly, two regions of the head skeleton that did not ever display dlx expression, the posterior neurocranial base (the basilar plate) and the trabeculae, were largely unaffected by exogenous RA (Table 1). Based on studies of medaka, the posterior neurocranial base and the trabeculae are thought to be derived from the prechordal plate mesoderm and the more anterior mesencephalic crest, respectively (Langille and Hall, 1987; Miyake and Hall, 1994). Studies on other vertebrates also point to a role for dlx genes in craniofacial cartilage differentiation although the evidence mostly comes from the description of patterns of expression: the mouse ortholog of the zebrafish dlx2 is expressed in migrating neural crest cells that will populate the visceral arches (Bulfone et al., 1993; Akimenko et al., 1994); dlx2 and other dlx genes are expressed in the primordia of visceral arches (Doll6 et al., 1992; Papalopulu and Kintner, 1993; Akimenko et al., 1994); mammalian dlx genes are expressed during chondrocyte condensation (Robinson and Mahon, 1994; Simeone et al., 1994; Zhao et al., 1994); and mouse Dlx5 and Dlx6 genes are known to be expressed in all tissues undergoing cartilage condensation (Simeone et al., 1994). Analysis of the rat rdlx gene during cartilage formation was performed by Zhao et al. (1994), who reported the expression of rdlx transcripts in the primordium of Meckel’s cartilage, in which the rdlx transcripts were more abundant in cells on the periphery than in the center. It was suggested, based on expression patterns, that rdlx plays a role in either the morphogenesis and pattern formation of the skeleton or in determining the fate and phenotype of cells that are constituents of the skeleton. Recently, the mouse Dlx3 gene was also shown to be expressed in a group of condensed cells of the mandibular arch, in the proximity of Meckel’s cartilage (Robinson and Mahon, 1994). Orthologous zebrafish distal-less genes (dlx3, dlx4) are also found to be expressed in regions which correspond to condensing visceral arch cartilage. Finally, Qiu and collaborators showed that a null mutation of the murine Dlx-2 gene results in abnormal morphogenesis of proximal first and second branchial arch derivatives (Qiu et al., 1995). Here we have shown that impaired expression of all zebrafish dlx genes correlates with more severe perturbations in craniofacial morphogenesis such as the loss of several cartilage elements. Thus, dlx genes may be an integral part of visceral arch differentiation and/or morphogenesis; the abolition of the expression of these genes by factors such as exogenous RA, as reported here, would then impede skeletogenesis as we have described. 4. Experimental
procedures
4. I. Animals Adult
zebrafish
embryos
were obtained
from a local
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supplier and were maintained according to standard procedures (Westerfield, 199.5). Embryos were staged at 285°C according to hours and days (d) post fertilization. 4.2. Retinoic
acid treatment
A 1 mM stock solution of all-trans retinoic acid (Sigma) was prepared in DMSO. For treatment of zebrafish embryos, RA was adjusted to final concentrations ranging from 0.1 PM to 1 PM in 0.5% DMSO. Embryos were transferred to the retinoic acid solution for 2 h after which they were washed twice with tank water. The schedules of RA administration, which extended from late gastrulation to arch segmentation, are shown in Fig. 4. Control embryos were taken from the same spawning as those treated with RA and left to develop for 2 h in 0.5% DMSO. RA-treated embryos and controls were fixed at 27 h for whole mount in situ hybridizations, or at 5 d for cartilage staining. All groups contained a minimum of five embryos. 4.3. Whole-mount
in situ hybridizations
Embryos were fixed in a phosphate buffer saline solution containing 4% paraformaldehyde. In situ hybridizations were performed on whole mount embryos using antisense riboprobes as previously described (Ekker et al., 1992a). The templates used for synthesizing the antisense riboprobes for dlx2, dlx3, dlx4, eng2, and krx20 have been previously described (Ekker et al., 1992b; Oxtoby and Jowett, 1993; Akimenko et al., 1994). The template for the dlxl probe consisted of a 900 bp cDNA fragment which will be described in details elsewhere (Ekker et al., unpublished observations). Double labeling was carried out on whole mount embryos using the same protocol as described above with the following modifications. A fluorescein- 12-UTP (no. 1427857, Boehringer Mannheim) probe was synthesized like the digoxygenin (DIG) probe; however, the DIG precipitation step was omitted. Instead, the fluorescein probe was filtered through a Nuctrap TM Push Column (Stratagene). Hybridization was carried out synchronously using both probes (digoxygenin and fluorescein). Once reactions with the anti-DIG antibody and alkaline phosphatase detection were complete, but prior to post-fixation, the embryos were immersed for 10 min in glycine-HCl (pH 2.2)/0.1% Tween-20 to inactivate the anti-digoxygenin antibody. The embryos were then washed four times for 5 min with 1 x phosphate buffered saline/ 0.1% Tween-20 (PBST) and incubated with the antifluorescein antibody (no. 1426338, Boehringer). The embryos were washed six times for 15 min in PBST, and twice for 5 min in 0.1 M Tris-HCl (pH 8.2)/O. 1% Tween-20. Detection of the fluorescein probe was done using two tablets of Fast Red (no. 1496549, Boehringer) dissolved in 4 ml of 0.1 M Tris-HCl (pH 8.2)/0.1% Tween-20. A 2 h post-
D.L. Ellies et al. /Mechanisms
fixation in 4% paraformaldehyde followed by a wash with PBS were carried out. The embryos were then stored at 4°C in PBS/5 mM sodium azide. 4.4. Cartilage
staining
Embryos were grown in a 0.2 mM solution of PTC (Phenylthiocarbamide, Sigma) in tank water, to inhibit pigment formation (Hyatt et al., 1992) and were stained with alcian blue. Briefly, 5 d larvae were fixed in 4% paraformaldhyde and washed as above. They were stained for 1- 12 h in a filtered 0.1% alcian blue solution in 30% acetic acid, 70% ethanol, rehydrated in a water-ethanol series, and cleared with a 0.5% KOH solution overnight at 4°C. Stained larvae were kept in Hz0 at 4°C. The general morphology of the cranial cartilage is shown schematically in Fig. 3. 4.5. TUNEL staining To detect apoptotic cells, we used a modification of the TUNEL staining procedure of Gavrieli et al. (1992). Briefly, embryos were fixed and washed as for in situ hybridization. They were then washed once in terminal transferase (TdT) buffer containing triton (100 mM cacodylate, pH 6.8, 2.5 mM CoC12, 0.1 mM dithiothreitol, 100 pg/ml BSA, 1% T&on). They were then incubated for 3 h at 37°C with 0.2 U/ml of terminal transferase, 10 PM dUTP (2: 1, dUTP:dUTP-biotin) in TdT buffer + 1% Triton. After washing 3 x 30 min in PBST, the embryos were incubated overnight with a streptavidin-horseradish peroxidase conjugate (1:500 in PBST), then washed 3 x 1 h in PBST, 2 x 30 min in PBS. Development was carried out in diaminobenzidine. After washing with PBS to stop the reaction, the stained embryos were mounted in 90% glycerol, 10% PBS. 4.6. Antibody
staining
Immunolocalization with the HNK- 1 monoclonal antibody was carried out as described (Holder and Hill, 1991; Westerfield, 1995). Acknowledgements We thank Wei Lin and Lynda Laforest for technical assistance, William Staines for the gift of the HNK-1 monoclonal antibody, Anthony Graham for advice on the whole-mount TUNEL procedure, Ann Graveson and Brian K. Hall for useful discussions and critical reading of the manuscript. This work is supported by grants to M.E. from the Medical Research Council (MRC) and the Natural Sciences and Engineering Research Council of Canada (NSERC). C.C.M is supported by studentships from NSERC, the Government of Ontario and the U. of Ottawa. M.E. is an MRC Scholar.
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