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Gene Expression during Palate Fusion in vivo and in vitro

¹ Developmental Biology Unit, Institute of Child Health, University College London, 30 Guilford Street, London WC1N 1EH, UK;
² Unit of Paediatric Dentistry, Eastman Dental Institute and Hospital, University College London, UK; and
³ Faculty of Dentistry, Sous Section Odontologie Pédiatrique, Hôpital Civil, Louis Pasteur University, 1 place de l’Hôpital, F-67000 Strasbourg Cedex, France;

^* corresponding authors, Agnes.Bloch-Zupan{at}dentaire-ulp.u-strasbg.fr and ferretti{at}ich.ucl.ac.uk

	ABSTRACT

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Failure of secondary palate fusion during embryogenesis is a cause of cleft palate. Disappearance of the medial epithelial seam (MES) is required to allow merging of the mesenchyme from both palatal shelves. This involves complex changes of the medial edge epithelial (MEE) cells and surrounding structures that are controlled by several genes whose spatio-temporal expression is tightly regulated. We have carried out morphological analyses and used a semi-quantitative RT-PCR technique to evaluate whether morphological changes and modulation in the expression of putative key genes, such as twist, snail, and E-cadherin, during the fusion process in palate organ culture parallel those observed in vivo, and show that this is indeed the case. We also show, using the organotypic model of palate fusion, that the down-regulation of the transcription factor snail that occurs with the progression of palate development is not dependent on fusion of the palatal shelves. Abbreviations: dsg1, desmoglein1; EMT, epithelial-mesenchymal transition; MEE, medial edge epithelium; MES, medial epithelial seam; RT-PCR, reverse-transcriptase polymerase chain-reaction.

KEY WORDS: palate development • mouse • gene expression • RT-PCR • epithelial-mesenchymal transition

	INTRODUCTION

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Cleft palate, one of the most common congenital malformations, can occur as an isolated defect or may be an associated feature in various syndromes. It has been reported that 53% of infants with cleft palate have other congenital anomalies (Murray, 2002). Any interference with the cellular and molecular processes underlying palate formation, either as a consequence of teratogenic or genetic factors, can be responsible for clefting of the palate. Development of the secondary palate entails downgrowth of the palatal shelves from the maxillary processes, shelf elevation above the tongue, and their juxtaposition and fusion at the midline. When the shelves approximate, the medial edge epithelial (MEE) cells covering the tip of each shelf intercalate and form the medial epithelial seam (MES). This requires the formation of desmosomes to achieve strong cell-cell adhesion (Mogass et al., 2000). The MES gradually breaks down and disappears to allow for the complete confluence of palatal mesenchyme from both shelves. Several lines of evidence have suggested that programmed cell death (apoptosis), migration of the cells toward the nasal and oral epithelium, and epithelial-mesenchymal transition (EMT) are involved in MES disappearance (Fitchett and Hay, 1989; Clarke, 1990; Carette and Ferguson, 1992). This transformation involves modulations of cytoskeletal intermediate filaments (e.g., cytokeratin 5 and vimentin), cell adhesion molecules (e.g., E-cadherin), and extracellular matrix (ECM) (Gibbins et al., 1999; Tudela et al., 2002).

Signaling molecules such as transforming growth factor beta-3 (tgfß₃) and fibroblast growth factors (fgfs), and their receptors (tgfßRs, fgfr1 and 2), are expressed in the MEE and surrounding mesenchyme prior to and during palate fusion in rodent and human embryos (Fitzpatrick et al., 1990; Sharpe et al., 1993; Cui and Shuler, 2000; Lee et al., 2001; Britto et al., 2002). TGFß₃ has been identified as one of the candidate genes involved in palatogenesis in patients with non-syndromic cleft palate (Murray, 2002). The only craniofacial anomaly present in homozygous tgfß₃-deficient mice is cleft of the secondary palate (Proetzel et al., 1995). In palate cultures from these mutant embryos, the fusion defect is rescued by exogenous tgfß₃ (Taya et al., 1999). FGFR2 mutations are known to underlie various craniofacial defects in humans and mice, including cleft palate (De Moerlooze et al., 2000; Murray, 2002). Transcription factors such as R-twist, which is expressed during rat palatogenesis (Bloch-Zupan et al., 2001), may control FGF signaling. The heterozygous twist mutant mice show several craniofacial defects, including craniosynostosis and narrow palate, which resemble those presented by patients with Saethre-Chotzen’s syndrome (Bourgeois et al., 1998). Another transcription factor, snail, has been implicated in several developmental processes involving EMT, including secondary palate development (Carver et al., 2001; Martínez-Álvarez et al., 2004).

Much understanding of palatogenesis and the etiology of craniofacial clefting arises from epidemiological and genetic studies of patients and families affected by these defects, in combination with data from experiments conducted in animal models. Organotypic palate cultures have also been utilized for the study of palatogenesis (Brunet et al., 1995). However, it is essential that we carefully compare developmental processes in vitro and in vivo, to assess whether the results of in vitro analysis and manipulation can be safely extrapolated to the in vivo situation. Although localization of several transcripts and proteins during palate development has been reported, information on the relative levels of expression and how they are controlled is still missing. We have examined expression patterns of several genes presumably involved in EMT to show that fusion in vitro closely resembles the in vivo process, and have investigated the expression of the transcription factor snail to assess whether it is regulated by palatal shelf fusion.

	MATERIALS & METHODS

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Animals and Tissue Collection
Wild-type CD1 mice housed and handled according to UK Home Office regulations for animal experimentation were used. Mice were mated overnight, and the presence of a vaginal plug was designated as embryonic day (E) 0.5. Pregnant mice were killed at embryonic stages E13, E14, E15, and E16. From 3 to 5 whole embryo heads per time-point were either fixed in Bouin’s solution and embedded in paraffin for standard histological investigation or fixed in 4% paraformaldehyde overnight at 4°C for in situ hybridization. For RNA analysis of palatal shelves, the whole maxilla was dissected from the head and surrounding tissues, including the primary palate, and tooth germs were then removed. The clean palatal shelves were transferred to TRI REAGENT^TM (Sigma, Dorset, UK), frozen immediately, and stored at –20°C.

Palate Cultures
Palatal shelves were harvested from E13 embryos and cultured in a chemically defined medium (Brunet et al., 1995). To maintain the precise physiological distance between the shelves in the organ culture system, we retained the primary palate and the posterior tissue. In some experiments, the palatal shelves were placed at least 0.5 mm apart to prevent fusion. From 3 to 5 palates were gently placed on 0.8-µm Milipore filter paper and a metal grid in a Falcon organ culture dish containing serum-free DMEM/F12 culture medium supplemented with 1% glutamine, 1% ascorbate, and 1% penicillin/streptomycin. The palate nasal epithelium was in direct contact with the Millipore raft. The moat in the culture dish was partially filled with sterile Milli-Q water, and the dish was placed in a humidified incubator in 95% air/5% CO₂ and 37°C for approximately 30, 48, and 60 hrs. For RT-PCR, the cultured palates were re-dissected to remove the extra tissue and collected as described above.

RNA Analysis
For each stage, 5 pairs of palatal shelves were pooled, and 3 independent pools were analyzed. Total RNA was extracted with the use of TRI-REAGENT^TM, according to the manufacturer’s instructions, and quantified spectrophotometrically. cDNA was prepared with M-MLV reverse-transcriptase (Promega, Southampton, UK) from 1.5 µg RNA and oligo-dT primers, according to the manufacturer’s guidelines. PCR was performed with the paired-primers and conditions indicated in the Table. We used gapdh cDNA to normalize levels of expression (Marone et al., 2001). gapdh product and PCR products to be quantified were run in the same gel (1.5% agarose gel containing ethidium bromide), visualized under UV light, and imaged with AlphaImager^TM V5.5 software (Alpha Innotech, San Leandro, CA, USA). Whole-mount in situ hybridization was carried out in E14.5 heads, after the mandible was removed, and in palates cultured for 30, 48, and 60 hrs, as previously described (Nieto et al., 1992).

	RESULTS

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In vitro Palate Fusion
Cultured palates were monitored regularly under the dissecting microscope. Serial sections from palates cultured for 30, 48, and 60 hrs (Figs. 1D–1G) were examined and compared with those from in vivo palates at different developmental stages. Overall, our time-course of palatal shelf fusion in vivo (Figs. 1A–1C) and in vitro (Figs. 1D–1G) was consistent with previous studies (Brunet et al., 1995). The majority of cultured palatal shelves (75%) had extended and adhered at the midline after 24 hrs in culture (Fig. 1D). The beginning of the fusion process was observed at 30 hrs. This paralleled the early fusion events observed in vivo around late E14. Formation of the MES was detected in some sections from the mid-palate. At 48 hrs, the MES dispersed, resembling in vivo palate development at E15 (Fig. 1E). By 60 hrs, the shelves had fused, and the MES had disappeared, as observed in late E15 to early E16 palates (Fig. 1F). Some sections from palates cultured for 72 hrs showed initial ossification centers within the lateral part of the palatal mesenchyme (not shown). A summary of the time-course of the fusion process in vivo and in vitro is shown in Fig. 1G.

View larger version (125K): [in this window] [in a new window]   Figure 1. Palatal fusion in vivo (A,B,C) and in vitro (D,E,F) examined in transverse sections stained with hematoxylin and eosin. (A) At E13 (n = 3), palatal shelves (ps) lie vertically on both sides of the tongue (T). (B) At late E14 (n = 5), the shelves are elevated horizontally and juxtaposed. The MEE formed the MES (arrow). (C) At E15 (n = 3), the MES was disrupted, and epithelial islands were observed (arrows), which completely disappeared at E16 (unpublished observations). The mesenchyme from both shelves merged at the midline (asterisk), and lateral mesenchymal cells condensed (stars). (D) At 30 hrs in culture (n = 5), the MEE cells formed the multilayered MES (arrow). (E) At 48 hrs (n = 5), MES thinned and was disrupted (arrow), allowing mesenchyme to merge across the midline. (F) At 60 hrs (n = 3), MES disappeared, and the palate was completely fused (asterisk). (G) Diagram summarizing time-frame of the fusion processes in vivo (bold lines) and in vitro (dotted lines). Scale bars: A,B = 500 µm; C,D,E,F = 200 µm. ps = palatal shelf, t = tooth bud, N = nasal surface, O = oral surface, T = tongue.

View larger version (28K): [in this window] [in a new window]   Figure 2. Expression patterns of genes regulated during palate development in vivo (A,B,C,D,E,F) and in organ culture (G,H,I,J,K,L), assessed by RT-PCR. (A,G) tgfß3, fgfr1; (B,H) fgfr2IIIb; (C,I) twist, snail; (D,J) E-cadherin, keratin 5; (E,K) dsg1; (F,L) vimentin. Error bars represent the standard error of the means (SEM); n = 3. Asterisks indicate significant differences between 2 time-points (P < 0.05).

View larger version (85K): [in this window] [in a new window]   Figure 3. Expression of snail in vivo (A) and in vitro (B–G) detected by whole-mount in situ hybridization (positive signal is dark or reddish-blue). Palatal shelves (n = 3–7/group) were either cultured in proximity to allow fusion to occur (B-D) or placed apart (E-G). (A) Transverse section of the whole-mount in situ hybridization of an E14.5 (n = 5), showing strong snail expression in the mesenchyme of the fusing palatal shelves, but not in the epithelium of the MES. (B-C) A similar pattern of snail expression is observed in fusing palatal shelves cultured for 30 and 48 hrs, with a weaker signal restricted to the lateral aspect (asterisks) observed at 48 hrs when the MES is being disrupted. (D) snail expression decreases as fusion progresses and is hardly detectable after 60 hrs in culture. (E,F,G) snail expression in palates that are not allowed to fuse decreases over time, as in fusing palatal shelves (B-D). Scale bars: A = 300 µm, B-G = 500 µm. N = nasal aspect, O = oral aspect, t = tooth bud. Color version of this figure is available online as an Appendix.

	DISCUSSION

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To gain further knowledge of molecular mechanisms underlying the normal and abnormal palate fusion, it is imperative that we identify not only spatial expression of relevant genes, but also their levels of expression and how they change with time. Various novel technologies have allowed researchers to gain a better insight into the regulatory genetic networks governing embryogenesis. Some methods, however, involve complicated and time-consuming procedures or require large amounts of starting material, which may be difficult to obtain. In this study, we have used two relatively simple techniques, organ culture and semi-quantitative RT-PCR, to study palate fusion. Semi-quantitative RT-PCR is a sensitive technique, and changes in expression levels as low as 1.2-fold can be detected (Marone et al., 2001). The gene expression profiles we have obtained using this technique provide a useful baseline to study the effect of disrupting key regulatory molecules either by using teratogen or via gene-targeting technology.

Our results have shown that fusion of palatal shelves in vivo is rather accurately reproduced in culture at both the morphological and molecular levels, though with a small time shift, as also observed by others (Cui and Shuler, 2000). Therefore, careful monitoring of the time of fusion is crucial when in vitro results are interpreted. In our case, gene expression after 30 hrs in culture is approximately comparable with that observed in E14.5 palates. Gene expression patterns in developing palates appear to follow very similar trends, both in vivo and in vitro, and the few small temporal discrepancies observed are likely due to slight developmental variations within the litter from which the palates are pooled.

Overall, the changes in gene expression we have detected by RT-PCR during secondary palate fusion are in agreement and complement those reported in previous studies in which qualitative localization techniques were used. Co-expression of TGFß₃, FGFR1, and FGFR2 IIIb has been shown in fusing the embryonic human palate (Britto et al., 2002). The expression of tgfß₃ and fgfr1 we have reported is consistent with this observation and with a crucial role for these molecules in the fusion process. At later developmental stages (E15), tgfß₃ and fgfr2 IIIb expression coincides with the condensation of the lateral mesenchyme, suggesting that these genes play a role in palatal bone development. Changes in expression profile of the cell adhesion, cytoskeletal, and desmosomal transcripts reflect the phenotypic changes occurring in the MEE, toward mesenchymal characteristics, prior to its disappearance.

Snail is a regulator of EMT during gastrulation, organogenesis, and in carcinoma cells when their invasiveness increases (Carver et al., 2001), and its expression in the palate is developmentally regulated. The down-regulation of snail we have observed following palate fusion, by both RT-PCR and in situ hybridization, and its localization in vivo and in vitro are consistent with the in vivo distribution of this transcript recently reported in the palate of another mouse strain (Martínez-Álvarez et al., 2004). These results suggest that snail is likely to be involved in the early stages of palate fusion. When fusion of palatal shelves is prevented in culture, snail down-regulation still occurs. This indicates that the decrease in the snail transcripts with palate development is not regulated by changes in gene expression occurring during the fusion process. Our finding that changes in snail expression with fusion progression parallel those of twist suggests a close interaction between these two transcription factors during the fusion process. Although the expression patterns of the genes we have studied may point to EMT as the mechanism underlying palate fusion, other mechanisms, such as apoptosis and migration of the MEE cells, may play a role. Indeed, these occur simultaneously during fusion and may be initiated by the same regulatory genes activating different pathways (Martínez-Álvarez et al., 2000). It is known that snail and twist have anti-apoptotic effects and can, therefore, play a role in cell survival (Maestro et al., 1999; Nieto, 2002). Further studies will be required to determine the significance of different pathways in inducing changes within the MEE cell population.

	ACKNOWLEDGMENTS

 
P. Pungchanchaikul is supported by a Royal Thai Government Scholarship for Civil Officers. This work was supported by research grants from the Unit of Paediatric Dentistry, Eastman Dental Institute and Hospital and Developmental Biology Unit, Institute of Child Health, London, UK. Part of the work presented here is based on the dissertation submitted to the Eastman Dental Institute and Hospital, University College London, in partial fulfillment of the requirements for the Master in Clinical Dentistry (Paediatric Dentistry), year 2003. A preliminary report was awarded the President’s prize for presentation at the President’s Evening, Royal Society of Medicine (Odontology Section), London, June 2, 2003, and was awarded the J. Morita prize for poster presentation at the 19th Congress of the International Association of Paediatric Dentistry, New Orleans, LA, USA, on the October 18, 2003.

	FOOTNOTES

 
A supplemental appendix to this article is published electronically only at http://www.dentalresearch.org.

Received April 20, 2004; Last revision February 17, 2005; Accepted March 20, 2005

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