A periodic pattern generator for dental diversity
© Fraser et al; licensee BioMed Central Ltd. 2008
Received: 25 January 2008
Accepted: 14 July 2008
Published: 14 July 2008
Periodic patterning of iterative structures is a fundamental process during embryonic organization and development. Studies have shown how gene networks are employed to pattern butterfly eyespots, fly bristles and vertebrate epithelial appendages such as teeth, feathers, hair and mammary glands. Despite knowledge of how these features are organized, little is known about how diversity in periodic patterning is generated in nature. We address this problem through the molecular analysis of oral jaw dental diversity in Lake Malawi cichlids, where closely related species exhibit from 1 to 20 rows of teeth, with total teeth counts ranging from around 10 to 700.
We investigate the expression of conserved gene networks (involving bmp2, bmp4, eda, edar, fgf8, pax9, pitx2, runx2, shh and wnt7b) known to pattern iterative structures and teeth in other vertebrates. We show that spatiotemporal variation in expression pattern reflects adult morphological diversity among three closely related Malawi cichlid species. Combinatorial epithelial expression of pitx2 and shh appears to govern the competence both of initial tooth sites and future tooth rows. Epithelial wnt7b and mesenchymal eda are expressed in the inter-germ and inter-row regions, and likely regulate the spacing of these shh-positive units. Finally, we used chemical knockdown to demonstrate the fundamental role of hedgehog signalling and initial placode formation in the organization of the periodically patterned cichlid dental programme.
Coordinated patterns of gene expression differ among Malawi species and prefigure the future-ordered distribution of functional teeth of specific size and spacing. This variation in gene expression among species occurs early in the developmental programme for dental patterning. These data show how a complex multi-rowed vertebrate dentition is organized and how developmental tinkering of conserved gene networks during iterative pattern formation can impact upon the evolution of trophic novelty.
Biology is replete with periodically patterned elements, from the sensory bristles of a fruit fly to the hair and teeth of mammals. Models of periodic patterning seek to explain the developmental origin of boundaries separating adjacent repetitive structures and the maintenance of cellular compartments once formed [1–8]. For example, the formation of feather tracts on the dorsal surface of chick embryos  and mammary (milk) lines on the ventral surface of embryonic mammals  serve to pre-pattern regions competent for the initiation of these structures. Similarly, a functionally equivalent field is established along the axis of the oral jaws in most vertebrates, competent to form tooth bud primordia [11–15]. In teleost fish this initial field is known as the primary odontogenic band (OB) [12, 13, 15] and in mammals it is termed the dental lamina [4, 14]. This band or lamina sets the regionally restricted 'field' along the jaw axis from which tooth induction is triggered.
As with all periodically patterned systems, an initial 'field of competence' is set from a once-homogeneous cellular region, followed by the establishment of positional information throughout the restricted 'field' [16, 17]. The initial field may be set up by a number of diffusing molecules such as morphogens that allow regionalization to occur, from which cellular differentiation responds along a gradient [5, 18], probably by means of a reaction-diffusion-type mechanism [19, 20]. Positional information determines cell differentiation, cellular compartmentalization and subsequent unit placode initiation, the first of which is imperative for iterative initiation of adjacent placodes via activator-inhibitor mechanisms . Placode initiation is thought to be triggered by cellular accumulation (self-organization) over a given threshold that reacts to a number of positional cues within the competent field . Within the placode itself, additional activators and inhibitors determine the boundaries of the placode unit and the spacing between units. Studies of periodically patterned systems such as the developing vertebrate dentition and developing chick feathers have led to the identification of a number of molecules that have been modelled as activators or inhibitors within the specific developing system [2, 21, 22]. In feather placode patterning, Shh and members of the Eda pathway have activator roles, while Bmp2 and Bmp4 are thought to act as inhibitors [2, 23, 24]. During mouse odontogenesis the same molecules are involved in patterning the molar cusps. Attempts have been made to model cusps according to activator-inhibitor patterning mechanisms; however, whether individual candidates can be classed as activators or inhibitors during tooth development is largely stage dependent [21, 25–27].
Molecules involved in the establishment of vertebrate dentition have been well characterized from studies of the mouse [11, 21, 28]. A number of these molecules are known to have detrimental effects on the murine dentition when removed/inhibited from the dental network early in tooth development; Shh [29, 30], Pitx2  and Pax9  are among those with severe dental phenotypes . For example, inhibition of Shh in mandibular explants during the transition of dental competence to initiation (E10.5) leads to tooth arrest at the bud stage [29, 30]. Thus, it is clear that this gene is essential for the correct establishment of the global dental programme. However, these studies are specific to the mouse experimental model, which develops a single set of teeth with no replacements. We therefore know nothing of the resulting phenotypes when modifications occur to these networks, for example the hedgehog pathway, in vertebrates with numerous functional tooth rows and continuous replacement cycles.
Results and Discussion
Variation in developmental gene networks prefigures differences in adult cichlid dentitions
We cloned cichlid orthologues of genes required during oral epithelial organization and tooth germ initiation (bmp2, fgf8, pitx2, shh) [11, 38] as well as mesenchymal markers (bmp2, bmp4 , pax9, runx2) involved in reciprocal signalling to the epithelium . Teleost tooth development has been well characterized in the zebrafish and thus our nomenclature for the early stages of tooth development will follow that model. Two stages of early odontogenesis are relevant: the thickened epithelium stage and the bell-shaped epithelium stage [39–41]. Developing teeth beyond this point will be referred to as tooth germs, spanning the progression of the tooth from a bell-shaped unit to various stages of functional maturity, characterized by cytodifferentiation.
pax9, one of the earliest mesenchymal markers of odontogenesis in the mouse, is either absent from or weakly expressed in the dentitions of zebrafish and Mexican tetra . By contrast, in Malawi cichlids pax9 is expressed initially in oral mesenchymal cells as a dental field along the mesiodistal jaw axis (Figure 2b, OB stage, and Figure 3D), then it is strongly up-regulated in the underlying mesenchyme at the epithelial thickening stage of the first tooth (Figure 2b, first-tooth stage, and Figure 3D). Expression of pax9 is then restricted to cells of the dental mesenchyme enveloping the tooth during morphogenesis and is absent from the cells of the dental papilla (Figure 2b, 3–4-teeth stage, and Figure 3E and 3F). The expression of runx2 essentially replicates that of pax9 for the stages examined (data not shown).
Size and spacing of tooth germs in three species of Malawi cichlids
Diameter of first tooth
ZOI including first tooth
ZOI medial to first tooth
ZOI distal to first tooth
ZOI from First tooth to second OB
MD length of OB
Width of OB at level of first tooth
Area (μm2) of first tooth
22.91 ± 0.839
51.80 ± 1.298
18.67 ± 0.344
17.99 ± 0.482
8.80 ± 0.56
110.72 ± 7.337
22.21 ± 1.135
528.08 ± 31.348
17.62 ± 0.149
44.05 ± 0.898
16.50 ± 0.213
12.87 ± 0.322
6.84 ± 0.084
116.62 ± 5.013
23.02 ± 0.382
384.06 ± 4.754
14.87 ± 0.349
39.9 ± 1.697
11.95 ± 0.285
15.24 ± 0.392
7.28 ± 0.545
120.91 ± 3.834
19.41 ± 0.782
268.47 ± 8.084
Diameter of first tooth
Inter-germ space between first and next medial tooth
Inter-germ space between first and next distal tooth
ZOI from First tooth to second OB
MD length of second row OB
Area (μm 2 ) of first tooth
25.68 ± 0.359
16.64 ± 0.128
14.02 ± 0.096
8.29 ± 0.266
140.02 ± 3.669
684.57 ± 19.248
18.19 ± 0.231
19.63 ± 0.222
11.96 ± 0.318
10.18 ± 0.434
138.47 ± 2.897
435.09 ± 19.98
18.37 ± 0.437
13.06 ± 1.112
20.33 ± 1.065
8.02 ± 0.295
168.39 ± 5.432
404.15 ± 23.182
Organizing the periodic pattern with molecular 'spacers'
Remarkably, these genes seem also to be employed in the initiation and spacing of future tooth rows, an iterated expression pattern similar to tooth germ organization within each row. shh labels each OB for subsequent tooth rows (Figures 2c and 4); eda and wnt7b are expressed between the first tooth row and the OB of the second (Figures 5 and 6). Specifically, eda expression partly overlaps that of shh in the lingual OB, while wnt7b is expressed either side of shh. Thus, eda from the enveloping mesenchyme (Figure 6A–C) may induce and maintain shh expression in tooth germs as well as in future tooth rows, and planar epithelial wnt7b (Figure 6D–F) may inhibit dental competence in these regions, similar to the role of these molecules in other systems [9, 30, 44–48].
Hedgehog signalling is required for initiation of periodic dental patterning
In all treated individuals that were allowed to develop for a further six days (fixed at 12 dpf), we found that the first tooth continues partial development and shows signs of mineralization, although it does not complete development or attachment (Figure 7d). With the exception of a mineralized remnant of the first tooth, all other teeth, adjacent to the first and in subsequent rows, failed to develop (Figure 7d). Knockdown of the hedgehog pathway at the 3–4-teeth stage resulted in a functional, patterned and replacing dental system (data not shown). These observations demonstrate that when perturbed (via the hedgehog pathway) at the first-tooth stage, the dental programme cannot recover, despite continued cycles of periodic patterning past this stage in untreated individuals.
The periodic pattern generator for dental diversity
The comparison of gene expression across Malawi cichlid species with divergent dentitions suggests a simple model implicating pitx2, eda and wnt7b, and their interaction with shh and edar, as primary features of a periodic pattern generator for diversity in Lake Malawi cichlid dentitions (Figure 8). The model accounts for two aspects of dental patterning: how to put tooth rows in jaws, and how to put teeth in tooth rows. Our data suggest that the combination of pitx2 and shh is required for a competent field of tooth initiation (the OB, Figures 2, 3, 4 and 8). M. zebra and L. fuelleborni exhibit expanded expression of pitx2 lingually on the embryonic lower jaw; C. afra does not (Figure 2). pitx2 and shh are also co-expressed in each subsequent OB for M. zebra and L. fuelleborni (Figure 4); C. afra does not initiate a third OB. Therefore, the lack of lingual/oral co-expression of pitx2 and shh in C. afra (Figures 4, 8F and 8I) may account for the reduction in row number compared with the other species (Figures 4, 8G and 8J, H and 8K, respectively). The lack of combinatorial expression of shh and pitx2 in the oral region of zebrafish may partially explain the lack of teeth . Here we show that this mechanism likely accounts for variation in tooth row number among Malawi cichlids. Thus, molecular mechanisms used to pattern the first row of teeth (the only row of teeth in mammals and most vertebrates) are redeployed as 'triggers' of dental competence and initiation in each subsequent row. We suggest that the initiation of new tooth rows follows a 'copy and paste' mechanism wherein the dental expression network is redeployed for each new tooth row. Therefore, our model posits that preceding tooth rows are required as a source of signal to initiate the next lingual row during sequential addition.
The combination of comparative gene expression data and perturbation of the hedgehog pathway suggests that the correct initiation and maintenance of the first-tooth germ, via activation of shh, is necessary for the periodically patterned dental programme in Malawi cichlids (Figure 8). Comparison of the cyclopamine phenotype at the first-tooth to the 3–4-teeth stages shows that disturbing the development of the first-tooth germ has an effect on the entire dentition, whereas disrupting the dentition at later stages results in a mildly reduced phenotype with additional teeth forming and completing development. We do not yet understand the molecular mechanisms (for example, decreased epithelial proliferation and/or increased cell death) of this severe dental phenotype at the first-tooth stage.
Our data imply that eda and wnt7b, expressed in the ZOI, regulate initial tooth germ size and position within rows, through interactions with shh; wnt7b inhibits the germ through planar epithelial signals (Figures 5 and 6D–F) and eda maintains the tooth germ (shh and edar) from within the surrounding mesenchyme (Figures 5 and 6A–C). The ZOI may not lie solely within the layers of the epithelium and we suggest that inhibitor/activator controls signal from within the underlying mesenchyme that envelops the thickened dental epithelium . Once the periodic pattern is established, other molecules may act as inhibitors from within the developing tooth unit, for example bmp2, which is present both in the early epithelial thickening and within the dental papilla (mesenchyme) during maturation (Figure 3J–L), and bmp4, which is restricted to the dental papilla (data not shown; ).
The expression of eda in the mesenchyme surrounding the developing dental germs of cichlids (Figures 5 and 6A–C) is more similar to that deployed during the patterning of feather placodes and salivary primordia  than that observed in mammalian dentitions, where it is restricted to epithelium [44, 46]. In our model, a large initial tooth germ in C. afra results from sustained local and intense eda expression on a comparatively similar inhibitory background of wnt7b (Figure 5a). The size of this tooth germ is reduced in M. zebra (Figure 5b) and L. fuelleborni (Figure 5c) because the eda expression broadens earlier (especially for L. fuelleborni), a heterochronic imbalance setting the stage for more, closely packed shh-positive tooth germs (Table 1 and Figure 8). Consistent with our results, transgenic mice (K14-eda) with increased levels of Ectodysplasin expression exhibit larger tooth germs [48, 52]. Furthermore, Eda null mutant mice have reduced tooth germs [48, 53–55]. However, in the mouse, effects of Eda on tooth size correlate positively with effects on tooth number; for example, higher levels of Eda lead to a single extra molar [49, 52]. Our data and model point to an important distinction between overall levels of eda and its spatial expression over time. An earlier dispersion of eda expression after initiation of the first tooth (as in L. fuelleborni), rather than continued localized expression around that first-tooth germ (as in C. afra), may in fact lead to the production of more, smaller tooth germs (Figure 8).
The position of subsequent tooth rows is also specified in part by the expression of wnt7b and eda in our model. Mesenchymal eda plays a permissive role in the positioning of the lingual OB (Figures 5 and 6A). In C. afra, its expression is strongest medial to the first tooth, while in M. zebra and L. fuelleborni it appears more as a band along the mesiodistal axis (Figures 5 and 6A–C; also see the second row tooth positions in Figure 1). wnt7b also appears to demarcate the location of the second row, as its expression is either side of the shh-positive second OB (Figures 5 and 6D–F) and, in a similar iterative manner to the patterning of individual tooth units, wnt7b is restricted to the inter-row space (Figure 5a–c). Once the initiation of the primary dental pattern for each row is established, the essential nature of shh and genes that occupy the ZOI is lost; although they likely continue to be expressed during further morphogenesis (Figures 3 and 6), these molecules are probably no longer required for initiation of the secondary, replacement dentition .
Periodically patterned phenotypes such as the dentitions of Lake Malawi cichlids present important exemplars for evolutionary developmental biology. The discipline has heretofore focused on the molecular basis of evolutionary novelty among distantly related organisms [35, 56] or the genetic/transcriptional basis of discrete trait loss among closely related groups [51, 57]. Trait elaboration (for example, bigger, longer, stronger [58, 59]) is more difficult to study because phenotypes are subtler, but this remains the more common type of evolutionary change . Dental diversity is an intermediate case; quantitative elaboration takes the form of gain or loss of discrete units. Our results support the general model that old genes, and entire developmental modules, are deployed anew to generate micro-evolutionary novelty in iterative structures.
Embryos and fry of three species of Lake Malawi cichlids (C. afra, M. zebra and L. fuelleborni) were raised to the required stage in a recirculating aquarium system at 28°C. Embryo ages (in dpf) were set after the identification of mouth brooding females (day 0). Embryos were then removed from the mouths of brooding females and, if required, were maintained for further development in separate culture tanks at 28°C.
Cloned sequences used to generate digoxigenin-labelled antisense riboprobes from Malawi cichlid species have been deposited in GenBank (accession numbers: EU867210 – EU867219). Many of the genes were identified through partial genome assemblies of L. fuelleborni and M. zebra  and cloned from M. zebra and L. fuelleborni cDNA libraries. Sequences of cDNA used to generate the probes are identical across the three species. Overall, these species exhibit almost no sequence divergence; the average nucleotide diversity for comparisons across the Malawi assemblage is 0.2%, less than among laboratory strains of the zebrafish .
To ensure the embryos of the three species were of equivalent stages (especially during gene expression comparisons), specimens were stage-matched based on external features, including pectoral and caudal fin development and eye development and maturity. Specimens for in situ hybridization were anaesthetized in tricaine methanesulfonate (MS222, Argent) and fixed overnight in 4% paraformaldehyde (PFA) in 0.1% phosphate-buffered saline (PBS) at 4°C. Whole-mount in situ hybridization experiments were based on protocols from  and modified as follows: embryos were transferred to methanol for dehydration and stored at -20°C. Specimens were rehydrated through to PBS with Tween-20 and digested with 4–10 μg/ml proteinase K (PK); the final concentration was based on the specific stage of embryo/fry (for example, embryos at approximately 5 dpf were digested with 5 μg/ml PK). Following hybridization, embryos were washed in TST (10 mM NaCl, 10 mM Tris-HCl, Tween-20 in depc-H2O). During the colour reaction stage of the protocol, all embryos were allowed to fully develop the colour. Thus, embryos were continuously transferred into fresh NBT/BCIP solution (Roche) in NTMT until full staining had ensued; this was determined after multiple regions of known expression became positive. Specimens were stage-matched based on external features, including pectoral and caudal fin development and eye development and maturity. All in situ hybridization experiments were performed with multiple specimens (multiple individuals were fixed at regular intervals, within single broods, then repeated at least twice with alternative broods) to fully characterize the expression patterns within and across the three species. After colour reaction (NBT/BCIP, Roche) embryos were washed in PBS and fixed again in 4% PFA, before whole-mount imaging using a Leica Microsystems stereomicroscope (MZ16). Embryos were embedded in gelatin and chick albumin with 2.5% gluteraldehyde. The gelatin-albumin blocks were post-fixed in 4% PFA before sectioning. Thin sections were cut at 15–25 μm using a Leica Microsystems VT1000 vibratome.
Cyclopamine manipulation of the hedgehog pathway
From a single brood of 24 individuals, 14 C. afra embryos were treated with cyclopamine (LC Laboratories) compound (50 μM) from a stock (5 mM cyclopamine in DMSO) to make up a final 1% DMSO solution in fish water. Five C. afra individuals were used as a 1% DMSO control, under the same incubation conditions as the treated embryos (Figure 7a and 7c). A further five individuals were kept as standard controls (wild-type), developing in the Georgia Institute of Technology aquarium. Treatment and control experiments were performed in ventilated Petri dishes spinning at 28°C in an oscillating platform culture incubator (Barnstead Lab-Line Max 4000). Following the treatment experiments and for the controls with DMSO, fishes were washed 10 times in fresh fish water to remove any remnant of cyclopamine compound or DMSO before transferring to culture vessels containing at least 300 ml of fish water, changed daily until ready for analysis. Although initial experiments with 50 μM cyclopamine using 1% (of 95%) ethanol as the solvent (suggested by the manufacturer, LC Laboratories and previous reports [51, 61]) showed differential expression patterns of shh to the 1% ethanol control experiments, alizarin red preparation of embryos raised to 12 dpf showed gross phenotypic effects on the ethanol-administered controls. Therefore, we substituted 1% DMSO for ethanol solvent, after which controls could not be distinguished from standard controls (untreated). While DMSO is not the best solvent for cyclopamine because of limited solubility above concentrations of 4 mg/ml, at the low concentrations used for enhanced viability of treated embryos, DMSO proved to be a better solvent than ethanol because of lower solvation temperatures and faster solvation times from -20°C storage temperatures.
We thank Craig Albertson, Marty Cohn, Anthony Graham, Darrin Hulsey, Moya Smith, and three anonymous reviewers for comments on previous drafts of the manuscript. Alizarin red-prepared fish (Figure 1) were collected from Lake Malawi and stained by Darrin Hulsey. Cichlid pitx2 was cloned by Keen Wilson, University of Georgia, Athens, GA, USA. Research is supported by grants, from the Petit Institute for Bioengineering and Bioscience (IBB 1241318), the NIH (DE 017182), and the Alfred P Sloan Foundation (BR-4499), to JTS. RFB was a GIT Presidential Undergraduate Research Fellow.
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