Molecular Evolution and Genetic Defects of Teeth - Cells, Tissues, Organs Vol 186 No 1

Molecular Evolution and Genetic Defects of Teeth

Guest Editor

James P. Simmer, Ann Arbor, Mich.

38 figures, 6 in color, and 4 tables, 2007

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Vol. 186, No. 1, 2007

Contents

Introduction 4 Evolution and Genetics of Teeth Simmer, J.P. (Ann Arbor, Mich.) 7 Gene Duplication and the Evolution of Vertebrate Skeletal Mineralization Kawasaki, K.; Buchanan, A.V.; Weiss, K.M. (University Park, Pa.) 25 The Origin and Evolution of Enamel Mineralization Genes Sire, J.-Y.; Davit-Béal, T.; Delgado, S. (Paris); Gu, X. (Ames, Iowa) 49 Evolutionary History of Sex-Linked Mammalian Amelogenin Genes Iwase, M.; Kaneko, S.; Kim, H.; Satta, Y.; Takahata, N. (Hayama) 60 Unraveling the Molecular Mechanisms That Lead to Supernumerary Teeth

in Mice and Men: Current Concepts and Novel Approaches D’Souza, R.N. (Dallas, Tex.); Klein, O.D. (San Francisco, Calif.) 70 Disorders of Human Dentin Hart, P.S.; Hart, T.C. (Bethesda, Md.) 78 Enamel Formation and Amelogenesis Imperfecta Hu, J.C.-C.; Chun, Y.-H.P.; Al Hazzazzi, T.; Simmer, J.P. (Ann Arbor, Mich.) 86 The Molecular Control of and Clinical Variations in Root Formation Wright, T. (Chapel Hill, N.C.)

94 Author Index Vol. 186, No. 1, 2007 94 Subject Index Vol. 186, No. 1, 2007 95 Patent Watch 96 Conference Calendar

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Introduction Cells Tissues Organs 2007;186:4–6 DOI: 10.1159/000102677

Evolution and Genetics of Teeth

Completion of the human genome project vastly improved our ability to discern the genetic etiologies of inherited diseases. Besides the human genome, partial and complete characterization of other genomes is progressing at a steady pace and already huge sequence databases are available from numerous organisms. Clever analyses of these databases have yielded surprising and interesting insights into the evolution of vertebrate mineralized tissues. This special issue of Cells Tissue and Organs contains symposia proceedings highlighting recent advances in our understanding of the genes and mutations involved in inherited dental malformations, and in the expansion of a particular gene family that brought biomineralization under genetic control in vertebrates. The major mineralized tissues in vertebrates are cartilage, bone, and teeth. The teeth are comprised of three mineralized tissues: dentin, enamel, and cementum. There is growing evidence that genes encoding extracellular matrix proteins involved in the biomineralization of bone, dentin, and enamel diverged from a common ancestor gene. The initial event was the generation of SPARCL1 (Sparc-like 1) from SPARC (secreted protein, acidic and rich in cysteine; BM-40/osteonectin) in a chromosome-wide large segmental duplication that spawned the chromosomal region ancestral to the long arm of human chromosome 4. The new gene (SPARCL1) gave rise to the secretory calcium-binding phosphoprotein (SCPP) gene family. In the first paper, Kawasaki et al. (Pennsylvania State University) advance their theory of the evolution of the SCPP gene family in Gene Duplication and the Evolution of Vertebrate Skeletal Mineralization.

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Mineralized tissues form in extracellular spaces, the contents of which are determined by specialized cells that line the space. Collagen-based extracellular matrices are extremely ancient, and are associated with the emergence of multicellularity. Sponges, for instance, which diverged from the line leading to vertebrates perhaps a billion years ago express fibrillar collagen. SPARC, the ancestor of the SCPP gene family, encodes a basement membrane protein. Collagen-based extracellular matrices must have evolved for many millions of years prior to the onset of biomineralization. Interestingly, the SCPPs share only a small region of similarity with SPARC. This region includes the signal peptide, the signal peptide cleavage site, and a Golgi casein kinase phosphorylation site (SXE) near the amino-terminus of the secreted protein. Therefore the fundamental features shared by the SCPP family are the targeting system for secretion into the extracellular environment where mineralization occurs and a near N-terminal phosphoserine that might be involved in calcium binding. The first evidence in the fossil record of mineralization in vertebrates is pharyngeal tooth-like structures (conodonts) from jawless fish that appear in the fossil record
540 million years ago. It remains a point of controversy whether or not the vertebrates that first formed conodonts had already diverged from the line leading to tetrapods. Kawasaki et al. argue that the origin of SPARCL1 coincided with the innovation of mineralized skeleton and occurred in a genome-wide duplication in the stem jawed vertebrates (after the divergence of jawless fish). In contrast, Sire (University Paris, France) et al. in

the second paper, The Origin and Evolution of Enamel Mineralization Genes, argue for a much more ancient origin for SPARCL1. During the evolution of jawed vertebrates, SPARCL1 expanded through a series of gene duplications to create the SCPP gene family. The repertoire of SCPP genes varies greatly among vertebrates. In humans, there are 22 SCPP genes arranged in two clusters on chromosome 4, in addition to the amelogenin genes (AMELX and AMELY) on the X and Y chromosomes. The two chromosome 4 SCPP clusters separated early in vertebrate evolution and diverged into two classes of extracellular matrix proteins. The dentin/bone SCPPs form the smaller, older cluster, and include DSPP, DMP1, IBSP, MEPE, and SPP1, which arose from a common ancestor by tandem gene duplication and have been designated the small integrinbinding ligand, N-linked glycoproteins, or SIBLINGs. The larger, more recent cluster of SCPPs contains the genes encoding the enamel proteins, caseins and some salivary proteins. Translocation of the amelogenin alleles to what became the sex chromosomes is explored by Iwase et al. (Graduate University for Advanced Studies, Sokendai, Japan) in the third paper, Evolutionary History of SexLinked Mammalian Amelogenin Genes. Amelogenin represents about 90% of the protein in developing dental enamel. The amelogenin gene translocated out of the SCPP cluster and into the first intron of ARHGAP6 (Rho GTPase-activating protein 6 gene) on another autosomal chromosome. In the ancestral lineage of eutherian (placental) mammals, the pair of homologous autosomes encoding AMEL translocated and became the short arms of the sex chromosomes. The amelogenin genes first resided in the autosomal-like (pseudoautosomal) region of the sex chromosomes, and then came to straddle the pseudoautosomal boundary. For some time, only the AMEL region downstream of intron 2 could undergo homologous recombination. The AMEL 5
region (upstream from transposon MER5 in intron 2) differentiated before the eutherian radiation, while the 3
region (downstream from MER5) differentiated independently within individual eutherian orders. Because of its recent history, the amelogenin genes on the X and Y chromosomes are used in PCR analyses to determine gender in forensics and archeology. The application of genomic analyses to uncover the evolution of vertebrate biomineralization is in its infancy, but it has already provided insights that could not be imagined a few years ago. Drs. Kawasaki, Sire, and Takahata have been leaders in this pioneering work, which

promises additional insights as more genomes are characterized. In the second half of this special issue of Cells Tissues and Organs, four papers are presented that summarize another recent leap forward in our understanding of the genetic etiologies of diseases affecting the dentition. Teeth form in concert with a series of epithelial-mesenchymal interactions. Histologically, developing teeth pass through initiation, bud, cap, bell, crown formation, and root formation stages. Genetic defects affecting the earliest stages of tooth development lead to familial tooth agenesis. Numerous syndromes have hypodontia as a feature. Defects in AXIN2 (part of the Wnt signaling system) and the transcription factor genes MSX1 and PAX9 cause different patterns of familial tooth agenesis that most severely affect the molar regions. D’Souza (Baylor College of Dentistry) and Klein (University of California) review the developmental opposite of hypodontia (supernumerary teeth) in their paper, Unraveling the Molecular Mechanisms That Lead to Supernumerary Teeth in Mice and Men. During the crown formation stage, odontoblasts and ameloblasts act in concert to orchestrate the extracellular deposition of dentin and enamel. P. Suzanne and Thomas C. Hart (National Institutes of Health), in Disorders of Human Dentin, summarize the current classification system for inherited defects in dentin, which includes three types of dentinogenesis imperfecta (DI) and two types of dentin dysplasia (DD). It has long been known that defects in the genes encoding type I collagen are responsible for DI-I, which is the syndromic form of DI-II. The recent genetic breakthroughs concerning non-syndromic dentin defects relate to the single-handed importance of DSPP (dentin sialophosphoprotein). This SCPP gene has now been implicated in the etiologies of DD-II, DI-II, and DI-III, which appear to be successively more severe forms of DSPP defects. Defects in another SCPP gene, DMP1 (dentin matrix acidic phosphoprotein), cause autosomal recessive hypophosphatemia, and (surprisingly) are not part of the etiology of isolated dentin defects. Implication of DMP1 in the regulation of phosphate homeostasis suggests that vertebrate biomineralization evolved in tandem with mechanisms to regulate phosphate, and probably calcium homeostasis. Inherited enamel defects that occur in the absence of a generalized syndrome are collectively designated as amelogenesis imperfecta (AI). Hu et al. (University of Michigan School of Dentistry) review the major stages of enamel biomineralization and explain how defects in the genes encoding enamel proteins or proteases can induce

Introduction

Cells Tissues Organs 2007;186:4–6

5

the various phenotypic patterns (hypoplasia, hypocalcification, and hypomaturation) associated with inherited enamel defects. While most of the genes involved in the full spectrum of AI are unknown, recent advances show that defects in AMELX cause X-linked AI, ENAM defects cause autosomal dominant AI, and defects in the enamel protease genes MMP20 and KLK4 cause autosomal recessive pigmented hypomaturation AI. Defects in the transcription factor gene DLX3 were previously shown to cause the tricho-dento-osseous syndrome, and it has been suggested that some DLX3 mutations might cause AI hypoplastic-hypomaturation with taurodontism. Wright (University of North Carolina), in The Molecular Control of and Clinical Variations in Root Formation, clarifies the role of DLX3 in the etiology of

6

Cells Tissues Organs 2007;186:4–6

tooth defects as he reviews our understanding of developmental defects of root formation, the molecular mechanisms involved, and the impact of root variants on clinical dentistry. The papers in this special issue present original and interesting ideas and new evidence concerning common evolutionary relationships among vertebrate mineralized tissues, and also review recent advances in our understanding of genetic diseases affecting primarily the dentition. These advances are central to our understanding of normal and pathological biomineralization and are important steps in discerning the molecular mechanisms underlying biomineralization in vertebrates. James P. Simmer, Ann Arbor, Mich.

Introduction

Cells Tissues Organs 2007;186:7–24 DOI: 10.1159/000102678

Gene Duplication and the Evolution of Vertebrate Skeletal Mineralization Kazuhiko Kawasaki Anne V. Buchanan Kenneth M. Weiss Department of Anthropology, Pennsylvania State University, University Park, Pa., USA

Key Words Gene duplication
Genome duplication
Mineralized tissue
Skeletal mineralization
Vertebrate evolution

Abstract The mineralized skeleton is a critical innovation that evolved early in vertebrate history. The tissues found in dermal skeletons of ancient vertebrates are similar to the dental tissues of modern vertebrates; both consist of a highly mineralized surface hard tissue, enamel or enameloid, more resilient body dentin, and basal bone. Many proteins regulating mineralization of these tissues are evolutionarily related and form the secretory calcium-binding phosphoprotein (SCPP) family. We hypothesize here the duplication histories of SCPP genes and their common ancestors, SPARC and SPARCL1. At around the same time that Paleozoic jawless vertebrates first evolved mineralized skeleton, SPARCL1 arose from SPARC by whole genome duplication. Then both before and after the split of ray-finned fish and lobe-finned fish, tandem gene duplication created two types of SCPP genes, each residing on the opposite side of SPARCL1. One type was subsequently used in surface tissue and the other in body tissue. In tetrapods, these two types of SCPP genes were separated by intrachromosomal rearrangement. While new SCPP genes arose by duplication, some old genes were eliminated from the genome. As a consequence, phenogenetic drift occurred: while mineralized skeleton is maintained by natural selection, the underlying genetic basis has changed. Copyright © 2007 S. Karger AG, Basel

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Introduction

Among the most important characteristics of vertebrates is a mineralized skeleton. Its evolution made possible various adaptive traits: endoskeleton for locomotion, dermal skeleton for protection, and teeth for predation. The evolution of this important trait has long been investigated in the fossil record but critical information for deciphering this process is also to be found in our genome. The vertebrate tooth consists of three principal mineralized tissues, a hard surface tissue called enamel or enameloid, more resilient body dentin, and supportive bone [Hall, 2005]. Paleozoic jawless vertebrates (Agnatha) also had a similar structure; their dermal skeleton (body armor) consisted of tubercles, containing tissues

Abbreviations used in this paper

BGLAP ECM ML NJ SCPP SIBLING UTR WGD

bone
-carboxyglutamic acid protein extracellular matrix maximum likelihood neighbor joining secretory calcium-binding phosphoprotein small integrin-binding ligand, N-linked glycoproteins untranslated region whole genome duplication

See table 1 for the names of SCPP genes.

Dr. Kazuhiko Kawasaki Department of Anthropology, Pennsylvania State University 409 Carpenter Building University Park, PA 16802 (USA) Tel. +1 814 863 2977, Fax +1 814 863 1474, E-Mail [email protected]

Tunicates Lampreys Hagfish Conodonts Anaspids Heterostracans

mineralization

Thelodonts dermal skeleton Galeaspids Osteostracans Placoderms Cartilaginous fish jaw

Acanthodians Rayfins

tooth

bichir teleosts

Lobefins coelacanth

WGD1

WGD2

mammals

SPARC SPARCL1 SCPP (Acidic) SCPP (P/Q)

Fig. 1. Vertebrate phylogeny and the evolution of skeletal miner-

alization. There are many extinct vertebrates (dashed lines) split from our ancestral lineage after the modern jawless vertebrates (lampreys and hagfish) but before cartilaginous fish. The most ancient mineralized tissue has been found in the feeding apparatus of conodonts [Donoghue and Sansom, 2002; Donoghue et al., 2006]. Subsequent jawless vertebrates evolved dermal skeleton. Cartilaginous fish have teeth but some placoderms, the first jawed vertebrates, may have independently innovated teeth [Johanson and Smith, 2005]. Two WGD are thought to have taken place, first in the stem jawless vertebrates (WGD1) and second in the stem jawed vertebrates (WGD2). SPARC descended from the common ancestor of protostomes and deuterostomes, whereas SPARCL1 arose through the WGD2. SCPP genes originated from SPARCL1 but have been found only in teleosts and tetrapods to date. Genes for acidic SCPPs and Pro/Gln-rich (P/Q) SCPPs arose before the divergence of ray-finned fish and lobe-finned fish. The vertebrate phylogeny is based on Donoghue et al. [2006]. Tunicates are shown as the outgroup.

comparable to teeth, and underlying bony plates (fig. 1) [Smith and Hall, 1990; Donoghue and Sansom, 2002; Donoghue et al., 2006]. For this reason, the investigation of dental tissues is critical for elucidating the evolution of vertebrate skeletal mineralization. 8

Cells Tissues Organs 2007;186:7–24

Mineralization of dental tissues is cooperatively regulated by extracellular matrix (ECM) proteins, secreted from the cells that develop through reciprocal interactions between epithelial cells and neural-crest-derived ectomesenchymal cells [Nanci, 2003; Veis, 2003; Hall, 2005]. Many transcription factors, signal molecules, and their receptors, involved in these processes, are all coded by old genes, i.e. their orthologs (genes that split by speciation) and paralogs (genes that arose by gene duplication) exist widely in metazoans [Jacob, 1977; Duboule and Wilkins, 1998]. In contrast, ECM proteins are more specialized to mineralized tissues. Some of these proteins are abundant only in a particular mineralized tissue, but their evolutionary relationship had been largely unknown until recent studies. We have previously reported that many of these highly specialized ECM protein genes have evolved from a common ancestor and they constitute the secretory calcium-binding phosphoprotein (SCPP) gene family [Kawasaki and Weiss, 2003]. Before their origin, the last common ancestor of this gene family, SPARCL1 (secreted protein, acidic, cysteine-rich like 1) arose from SPARC (also called osteonectin) that also codes a major ECM protein for both dentin and bone [Kawasaki et al., 2004; Sanetra et al., 2005]. Many SCPP genes, involved in tissue mineralization, have been identified in tetrapods and teleost fish. However, in these two lineages, SCPP genes arose independently by parallel gene duplication [Kawasaki et al., 2005]. Later in tetrapods, SCPPs were co-opted for milk caseins and salivary proteins in mammals, and an eggshell matrix protein in birds. All these descendants are also involved in some sort of mineralization: milk caseins help infants to grow teeth and bone, salivary SCPPs maintain enamel integrity, and the eggshell SCPP organizes calcification of the shell [Kawasaki and Weiss, 2006]. Here we describe the duplication histories of the SPARC, SPARCL1, and SCPP genes, and discuss possible scenarios linking these genes and skeletal mineralization. Dentin and Bone ECM Proteins of Mammals Dentin and bone both form on collagen fibrils. Approximately 90% of dentin and bone protein matrix consist of type-I collagen, the most abundant proteins in the body [Veis, 2003]. In teleost fish, type-I collagen also serves as the mineralization scaffold for enameloid. However, collagen fibrils alone cannot initiate mineralization and many other molecules are required for regulated mineralization. Among these molecules, DSPP (dentin sialophosphoprotein), DMP1 (dentin matrix acidic phosKawasaki/Buchanan/Weiss

phoprotein 1), IBSP (integrin-binding sialoprotein), MEPE (matrix extracellular, phosphoglycoprotein), SPP1 (secreted phosphoprotein 1; also called osteopontin), SPARC, and BGLAP (bone
-carboxyglutamic acid protein; also called osteocalcin) are especially important in mammals [Nanci, 2003; Veis, 2003; Qin et al., 2004]. These proteins are present in both dentin and bone but their composition in each tissue is significantly different. For example, DSPP is abundant in dentin, and mutations in this gene cause dentin hypoplasia and dentinogenesis imperfecta, whereas the expression of DSPP is marginal in bone and its known mutations have no effect on bone development [Rajpar et al., 2002; Kim et al., 2005]. By contrast, IBSP constitutes 8–12% of the total non-collagenous proteins in human bone, with substantially lower amounts (
1%) in dentin [Ganss et al., 1999]. Among these dentin/bone ECM proteins, DSPP, DMP1, IBSP, MEPE, and SPP1 have been shown to have arisen from a common ancestor by tandem gene duplication and have consequently been named small integrin-binding ligand, N-linked glycoproteins (SIBLINGs) [Fisher et al., 2001]. SIBLING genes are primarily expressed in mineralized tissues, but their expression has also been detected in many soft tissues [Giachelli and Steitz, 2000; Ogbureke and Fisher, 2004, 2005]. Here we refer to this group of proteins as dentin/bone SCPPs, because they are part of the larger SCPP family. Enamel ECM Proteins of Mammals Tetrapod enamel is highly specialized among vertebrate mineralized tissues in that this tissue does not contain collagen but forms on a distinctive protein scaffold of epithelial (ameloblast) origin [Fincham et al., 1999; Veis, 2003; Bartlett et al., 2006]. In mammals, ameloblasts secrete three major enamel ECM proteins: AMEL (amelogenin), AMBN (ameloblastin), and ENAM (enamelin) [Hu et al., 2005; Paine and Snead, 2005]. The genes for these proteins are all primarily expressed in ameloblasts, although their marginal or transient expression has also been detected in odontoblasts of mesenchymal origin [Nagano et al., 2003; Papagerakis et al., 2003]. Mutations in human AMEL or ENAM result in enamel hypoplasia or amelogenesis imperfecta, and mice with disrupted AMBN develop severe enamel hypoplasia [Hu and Yamakoshi, 2003; Wright et al., 2003; Fukumoto et al., 2004]. These facts show the essential role of these proteins in enamel mineralization. AMEL constitutes more than 90% of enamel ECM protein and serves as the transient scaffold for mineralizing enamel [Moradian-Oldak and Goldberg, 2005; Margolis et al., 2006], whereas the Gene Duplication and Skeletal Mineralization

functions of ENAM and AMBN are less clear [Bouropoulos and Moradian-Oldak, 2004; Fukumoto et al., 2004]. During maturation, these enamel proteins are actively degraded by proteinases secreted by ameloblasts and subsequently removed from the mineralizing enamel [Zeichner-David et al., 1995; Simmer and Hu, 2002; Hu et al., 2005]. As a result, the mature enamel becomes highly mineralized, and grows into the hardest tissue in the vertebrate body. In addition to these three enamel genes, another, AMTN (amelotin), was recently found to be preferentially expressed in ameloblasts. AMTN localizes in the basal lamina between ameloblasts and maturing enamel, suggesting an important role in enamel mineralization [Iwasaki et al., 2005; Moffatt et al., 2006b]. Characteristics of SCPP Genes We have previously shown that a series of gene duplications created the SCPP gene family, which includes ECM protein genes for dentin/bone (DSPP, DMP1, IBSP, MEPE, and SPP1) and enamel (AMEL, ENAM, AMBN, and AMTN), as well as milk casein, and some salivary protein genes [Kawasaki and Weiss, 2006]. Table 1 lists 22 members of this gene family found in the human genome. Although, with just a few exceptions, these proteins have virtually no sequence homology, the following three lines of evidence show their common evolutionary origin. First, their protein products share some basic biochemical characteristics: (i) all these proteins have a signal peptide, and hence are secretory proteins; (ii) most of these proteins are rich in charged, particularly acidic amino acids (Glu and Asp); (iii) most have at least one, but usually many Ser-Xaa-Glu motifs (Xaa represents any amino acid; Asp or phospho-Ser may replace Glu), in which the Ser residue is phosphorylated, and (iv) these proteins bind to calcium ions through these acidic amino acids (Glu, Asp, and phospho-Ser). We thus call these proteins ‘secretory calcium-binding phosphoproteins’ [Kawasaki and Weiss, 2003]. Second, SCPP genes share the common genomic structure: (i) the entire exon 1 and the 5-end of exon 2 together comprise the 5-untranslated region (5-UTR); (ii) exon 2 additionally codes the entire signal peptide and the Nterminus of the mature protein, and (iii) all the introns are phase 0, which means that they are located between two adjacent codons instead of disrupting a codon [Kawasaki and Weiss, 2006]. Some of these structural elements may be found in other genes. However, only SCPP genes (and the 5 portion of their ancestral SPARC and SPARCL1 as described below) have been found to possess all these characteristics. Cells Tissues Organs 2007;186:7–24

9

Table 1. SCPP genes and ancestors

Gene symbol (alias)

Protein name (alias)

Protein distribution

Ancestor SPARC (OSN) SPARCL1 (HEVIN)

secreted protein, acidic, cysteine rich (osteonectin) secreted protein, acidic, cysteine-rich like 1 protein (high endothelial venule protein)

skeleton brain

dentin sialophosphoprotein dentin matrix acidic phosphoprotein 1 integrin-binding sialoprotein (bone sialoprotein) matrix extracellular phosphoglycoprotein secreted phosphoprotein 1 (osteopontin) amelogenin enamelin ameloblastin (sheathlin, amelin) amelotin (UNQ689) odontogenic, ameloblast associated (APin protein) follicular dendritic cell secreted peptide mucin 7 (MG2) proline-rich 1 (basic proline-rich lacrimal protein 1) proline rich 3 [submaxillary gland androgen-regulated protein 3 homolog B (mouse)] proline rich 5 [submaxillary gland androgen-regulated protein 3 homolog A (mouse)] LOC401137 histatin 1 histatin 3 statherin -casein -casein S1-casein

dentin, bone dentin, bone dentin, bone dentin, bone dentin, bone enamel enamel enamel enamel milk, saliva, enamel milk, saliva, PDL salivaa saliva saliva

SCPP DSPP DMP1 IBSP (BSP) MEPE SPP1 (OPN) AMEL ENAM AMBN AMTN (UNQ689) ODAM (APIN) FDCSP (C4ORF7) MUC7 PROL1 (BPLP) PROL3 (SMR3B) PROL5 (SMR3A) LOC401137 HTN1 HTN3 (HTN2) STATH CSN3 (CSN10) CSN2 CSN1S1

saliva saliva saliva saliva saliva milk milk milk

PDL = Periodontal ligament; LOC401137 = the locus symbol given in the genome sequence database. a Many salivary SCPPs are also present in tears.

Third, with one exception, SCPP genes all reside on a single chromosome, clustered in two regions. In the human genome, these clusters are separated 17 Mb apart on chromosome 4 (fig. 2). The only exception is AMEL, which is located on the sex chromosomes in placental mammals; some species, including humans, retain this gene both on the X- and Y-chromosomes but others, such as the mouse, have it only on the X [Iwase et al., 2001, 2003; Sire et al., 2005]. And the X-copy, AMELX, resides within the first intron of ARHGAP6 (Rho GTPase-activating protein 6 gene). SCPP Gene Clusters in the Human Genome These two SCPP gene clusters contain different classes of genes. One cluster consists of genes for dentin/bone ECM proteins, whereas the other contains genes for 10

Cells Tissues Organs 2007;186:7–24

enamel ECM proteins as well as caseins and salivary proteins. In this study, we will show evidence that suggests an early divergence of these two classes of SCPP genes. By contrast, we have described the relatively recent history of the current milk/saliva/enamel SCPP gene cluster [Kawasaki and Weiss, 2006]. Among these SCPPs, only AMBN has been found in both frogs and crocodilians [Shintani et al., 2002, 2003, 2006], while no milk or salivary SCPP genes have been identified in non-mammalian tetrapods. We thus assume that the milk/saliva/enamel SCPP gene cluster initially arose from an ancient enamel SCPP gene, and many milk and salivary SCPP genes first evolved in the stem group of mammals [Kawasaki and Weiss, 2006]. ODAM (odontogenic, ameloblast associated) and FDCSP (follicular dendritic cell secreted peptide) both Kawasaki/Buchanan/Weiss

SCPP Gene Cluster in the Fugu Genome Among the SCPP family, only orthologs of SPP1 have been found in both tetrapods and teleosts [Kawasaki et al., 2004]. However, this dentin/bone SCPP gene, identified from zebrafish and trout, has not as yet been found in the genomes of medaka (Oryzias latipes) or two puffer fish species (Takifugu rubripes and Tetraodon nigroviridis). Similar to the mammalian and chicken genomes, in which dentin/bone SCPP genes reside adjacent to the ancestral SPARCL1, the fugu genome also has seven distinct SCPP genes adjacent to SPARCL1: SCPP1–SCPP5 with three closely related SCPP3 genes, SCPP3A, SCPP3B, and SCPP3C (fig. 2; their names reflect the history of gene discovery) [Kawasaki et al., 2005]. With the exception of SCPP3C, the expression of all these fugu SCPP genes has been detected in dental tissues. However, none of these genes show significant sequence homology to any known tetrapod SCPPs. Furthermore, none of the well-conserved short sequence elements, present in tetrapod SCPPs, exist in any of these fugu proteins. These facts suggest that, in the teleost and tetrapod lineages, SCPP genes arose independently by parallel gene duplications [Kawasaki et al., 2005]. The Origin of the SCPP Gene Family SPARCL1 is closely related to SPARC; both are multifunctional ECM proteins and associate with various other ECM proteins, cations, growth factors, and cells [Yan and Sage, 1999; Sullivan and Sage, 2004]. In mouse embryos, SPARCL1 is primarily expressed in brain, whereas SPARC is preferentially expressed in skeleton [Mothe and Brown, 2001]. In adults, SPARC is one of the most abunGene Duplication and Skeletal Mineralization

a Human Milk / Saliva

Enamel

Dentin / Bone

SPA

RC L1 DS P DM P P IBS 1 P ME PE SP P1

CS N CS 1S1 N STA 2 T HT H N HT 3 N LO 1 C OD 4011 3 A FD M 7 CS CS P N3 PR OL PR 5 O PR L3 O MU L1 C7 AM TN AM EN BN AM

DM P IBS 1 P OC 1 SP 16 P1

SPA

RC

L1

17 Mb

DM P IBS 1 P

AM

BN

b Chicken

c Frog SC PP SC 5 P SC P4 P SC P3B PP SC 3C P SC P3A PP 2 SPA RC L1 SC PP 1

belong to the SCPP gene family and are adjacent to each other within the cluster of milk and saliva class SCPP genes (fig. 2). The expression of these two genes had been detected in both salivary glands and lactating mammary glands [Rijnkels et al., 2003]. More recently, however, their expression was also detected in developing teeth, ODAM in the enamel organ (including ameloblasts) and FDCSP in fibroblasts producing periodontal ligament (a soft connective tissue surrounding the roots of teeth) [Nakamura et al., 2005; Moffatt et al., 2006a]. The last exon of these two genes has no protein-coding region and consists only of the 3-UTR. This untranslated last exon is present in all milk and salivary SCPP genes located between CSN1S1 (S1-casein) and CSN3 (-casein), suggesting their close evolutionary relationship (fig. 2).

d Fugu P/Q rich

Acidic

Fig. 2. SCPP gene clusters in the human (a), chicken (b), frog (c), and fugu (d) genomes. Location and transcriptional orientation of SPARCL1 (open box) and SCPP (closed box) genes are schematically illustrated. a Two large SCPP gene clusters are separated by 17 Mb. The distribution of SCPPs is indicated above (see table 1 for more details). Location of genes for Pro/Gln-rich (P/Q rich) or acidic SCPPs is shown at the bottom.

dant non-collagenous ECM proteins in both dentin and bone [Termine et al., 1981]. These two proteins both consist of three functional modules, acidic N-terminal domain I, follistatin-like domain II, and extracellular calcium-binding domain III [Girard and Springer, 1995; Sullivan and Sage, 2004]. Among these, domains II and III are both globular, tightly folded through many disulfide bonds between cysteine residues abundant in these two domains [Hohenester et al., 1996; Sullivan and Sage, 2004; Chun et al., 2006]. By contrast, domain I is free from cysteine and consists largely of an intrinsically disordered structure, which does not fold into a fixed globular structure [Hambrock et al., 2003; Sullivan and Sage, 2004]. The critical difference between SPARC and SPARCL1 is the size and amino acid composition of this domain [Kawasaki et al., 2004]. In humans, domain I of SPARC is 52 amino acids in length, whereas that of SPARCL1 consists of 414 amino acids, in which 339 residues are coded by a large exon, Cells Tissues Organs 2007;186:7–24

11

present only in SPARCL1. Furthermore, domain I of SPARC has 18 acidic but no basic amino acids, in contrast with the same domain of SPARCL1 that contains 104 acidic plus 55 basic amino acids. These differences are common to all teleost and tetrapod SPARC and SPARCL1 found to date, and hence appear to reflect an important functional division that took place before the split of these two vertebrate lineages. Due to the acidic nature, domain I of both SPARC and SPARCL1 weakly binds to calcium ions, although the amino acid sequences are highly variable across species [Engel et al., 1987; Oritani et al., 1997; Brekken and Sage, 2000; Chun et al., 2006]. Many SCPPs appear to retain an intrinsically disordered or unfolded nature as discussed later, and dentin/bone SCPPs in particular have the amino acid composition similar to SPARCL1 [Huq et al., 2005]. In addition to these chemical characteristics, SPARC and SPARCL1 both show striking structural similarities to SCPP genes: the location of the signal peptide coding region within exon 2 and all phase-0 introns between the exons coding domain I [Kawasaki and Weiss, 2003]. Furthermore, SPARCL1 is located on 4q22.1 in the human genome adjacent to the dentin/bone SCPP gene cluster (fig. 2), whereas SPARC resides on 5q31.3-q32. For these reasons, we concluded that gene duplication created SPARCL1 from SPARC first, then the initial SCPP gene from SPARCL1, which was accompanied by simultaneous or subsequent loss of two globular domains [Kawasaki et al., 2004]. SPARC and SPARCL1 Gene Duplication We previously estimated that SPARCL1 arose from SPARC not so long before the divergence of ray-finned fish (Actinopterygii) and lobe-finned fish (Sarcopterygii), based on the molecular clock hypothesis [Kawasaki et al., 2004]. However, we used no SPARC sequences from cartilaginous fish (Chondrichthyes) or jawless vertebrates in the previous analysis. Mineralized skeleton initially evolved in extinct jawless vertebrates, which split from our ancestral lineage after modern jawless vertebrates but before cartilaginous fish (fig. 1) [Donoghue and Sansom, 2002; Donoghue et al., 2006]. Thus, the relative timing between the origin of SPARCL1 and the divergence of these two vertebrate clades is critical to resolve questions about the early evolution of skeletal mineralization. In this study, we added new SPARC sequences from the shark and lamprey, and show that the origin of SPARCL1 coincided with the innovation of mineralized skeleton.

12

Cells Tissues Organs 2007;186:7–24

Methods Identification of SPARC Genes Nurse shark (Ginglymostoma cirratum) and lamprey (Petromyzon marinus) SPARC genes were isolated from dental tissues and eyes, respectively. mRNA molecules were purified using the RNeasy mini kit (Qiagen), and cDNA libraries were constructed using the SMART cDNA library construction kit (Clontech). SPARC genes were amplified from these cDNA libraries by PCR. Three degenerate PCR primers were designed within regions conserved in teleost and tetrapod SPARC and SPARCL1 genes: dSPARCup2 (5-TTCCCYCTGCGYATGMGNGACTGGCT-3), dSPARCdwn1 (5-TGRTCCAGCTGNCCRAAYTGCCAGTG3), and dSPARCdwn2 (5-AARAAGCGNGTGGTGCARTGYTCCAT-3). dSPARCup2 and dSPARCdwn1 were used to amplify shark SPARC cDNA by the rapid amplification of cDNA ends according to specifications of the SMART library kit. dSPARCup2 was also used with dSPARCdwn2 to obtain a middle portion of lamprey SPARC. These products were cloned into plasmid vector and their nucleotide sequences were determined as previously described [Kawasaki et al., 2005]. The sequence analysis revealed two distinct lamprey SPARC genes, and hence three PCR primers specific to each of these genes were additionally designed and used for rapid amplification of cDNA ends: dSPARCup2 and pSPARCAdwn2 (5-ACGACGACGTTCTTGAGCCAGTCC-3) for one gene, and pSPARCBup2 (5-CATCTACCCGGTGCACTGGCAGTT-3) and pSPARCBdwn1 (5-CCGCTCGTCGCTCACGATCTTCC-3) for the other. Bioinformatic Analysis The amino acid sequences of well-conserved domains II and III of SPARC and SPARCL1 were aligned using CLUSTALW (http://www.ddbj.nig.ac.jp/) and manually corrected. Gaps in the multiple sequence alignment were not used for further analysis. Phylogenetic trees were constructed by the neighbor-joining (NJ) method and the maximum-likelihood (ML) method using MEGA3.1 and TREE-PUZZLE 5.2, respectively [Schmidt et al., 2002; Kumar et al., 2004]. The NJ trees were based on the
-distance using the Johns-Taylor-Thornton substitution model [Kumar and Nei, 2000]. The
-parameter was estimated using GAMMA [Gu and Zhang, 1997]. Genome sequences of various vertebrate species were examined through the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/), Ensembl (http://www.ensenbl. org/), and University of California, Santa Cruz, genome bioinformatics (http://genome.ucsc.edu/) websites using the BLAST or BLAT program. Duplicate genes in the human genome were sought for functional gene sequences (annotated as best RefSeq by the NCBI in the Build36.1 genome sequence assembly; nonfunctional pseudogenes among these entries were not used in this analysis) located on chromosome 5q31-qter (the region between HINT1 and TRIM52) around SPARC using the BLASTP program against the human RefSeq database. Nucleotide sequences of tunicate (Ciona intestinalis) orthologs of human genes were reconstructed from many entries in the expressed sequence tag database (http://ghost.zool.kyoto-u.ac.jp/).

Kawasaki/Buchanan/Weiss

Results and Discussion

SCPP Genes of Placentals, Marsupials, and Monotremes Different lineages of placental mammals have distinct duplication and deletion histories of milk and saliva class SCPP genes. For instance, humans have three functional casein genes (CSN1S1, CSN2, and CSN3), whereas the mouse has five (Csn1s1, Csn2, Csn1s2a, Csn1s2b, and Csn3) [Rijnkels et al., 2003]. CSN1S2 became a pseudogene in the human, chimpanzee, and rhesus monkey genomes. Exon duplication and deletion also occur frequently in casein genes. In particular, human CSN1S1 consists of 11 exons, while the mouse ortholog has 33 with many small duplicate exons. The evolution of salivary SCPP genes is similar to casein genes. Three closely related salivary SCPP genes, STATH, HTN1, and HTN3, are clustered in the human genome, whereas only two pseudogenes are present in the orthologous mouse genomic region [Kawasaki and Weiss, 2003]. The substitution rates are extremely high in these salivary and casein genes [Sabatini et al., 1993]. In contrast, dentin/bone and enamel SCPP genes evolve more slowly; significant sequence homology is detectable among the orthologous dentin/bone and enamel genes of mammals, birds, and amphibians [Toyosawa et al., 1998; Shintani et al., 2003; Kawasaki and Weiss, 2006; Sire et al., 2006]. Within marsupials, the opossum (Monodelphis domestica) has at least 14 SCPP genes: all five dentin/bone SCPP genes (DSPP, DMP1, IBSP, MEPE, and SPP1), six SCPP genes expressed in teeth (ODAM, FDCSP, AMTN, AMBN, ENAM, and AMEL), and three milk casein genes (CSN1S1, CSN2, and CSN3). Although no other SCPP genes have been identified, there may be more within unsequenced gaps that still remain in the current version of the genome sequence (monDom4 released in January 2006). Similar to placental mammals, the opossum also has two SCPP gene clusters, one for dentin/bone and the other for milk/enamel genes, and AMEL is located within the first intron of ARHGAP6. The order and orientation of these SCPP genes are all conserved in the opossum and human genomes. We also confirmed a similar configuration of these SCPP genes in the preliminary duck-billed platypus (Ornithorhynchus anatinus) genome sequence (version OANA5 released in December 2005): the dentin/bone gene cluster (DMP1, IBSP, and SPP1, identified so far), the enamel gene cluster (AMTN, AMBN, and ENAM), and isolated AMEL within ARHGAP6. These conserved arGene Duplication and Skeletal Mineralization

rangements suggest that the two large SCPP gene clusters had been established before the origin of monotremes. Evolution of Tetrapod SCPP Genes We have previously reported that the chicken has three dentin and bone class SCPP genes (DMP1, IBSP, and SPP1) and the eggshell SCPP gene, ovocleidin-116 (OC116), all forming a cluster adjacent to ancestral SPARCL1 [Kawasaki et al., 2004]. OC116 has been thought to be orthologous to mammalian MEPE based on their equivalent chromosomal location within the SCPP gene cluster and slight sequence homology [International Chicken Genome Sequencing Consortium, 2004; Kawasaki and Weiss, 2006]. No enamel/milk/saliva SCPP genes have been found in the chicken genome [Kawasaki and Weiss, 2006]. In the frog (Xenopus tropicalis) genome, we have identified two dentin/bone SCPP genes (DMP1 and IBSP) and two enamel SCPP genes (AMBN and AMEL) common to mammals [Kawasaki and Weiss, 2006]. No frog SPP1 has been found in the genome, although some teleosts have this gene [Kawasaki et al., 2004]. The differences between SCPP genes found in the frog and chicken genomes suggest that two enamel SCPP genes, AMEL and AMBN, were secondarily lost from the chicken genome after modern birds became toothless in the Late Cretaceous [Kawasaki and Weiss, 2006]. We also assume that DSPP, primarily expressed in odontoblasts, and three SCPP genes expressed in ameloblasts (or enamel organ), ODAM, AMTN, and ENAM, all evolved in the lineage leading to mammals after the divergence from ancestral birds. Alternatively, some of these genes may have been secondarily lost from the chicken genome. Milk casein genes are likely to have arisen in mammal-like reptiles early in the evolution of lactation [Kawasaki and Weiss, 2006]. Ancient Tetrapod SCPP Gene Cluster No SPARCL1 has been found in the frog genome, probably because this gene was also secondarily lost. Instead, AMBN is located approximately 170 kb apart from the DMP1-IBSP cluster, much closer than their distances in mammalian genomes [Huq et al., 2005; Kawasaki and Weiss, 2006]. Our previous analysis showed that intrachromosomal rearrangement is common around the SCPP gene clusters both in the human and chicken genomes; hence we speculated that intrachromosomal rearrangements split the primordial tandem array of SCPP genes, in which dentin/bone and enamel genes had been co-localized (no milk or salivary genes may have existed at the time) [Kawasaki and Weiss, 2006]. Cells Tissues Organs 2007;186:7–24

13

HERC6

PKD2

PPM1K

IBSP

ABCG2

DMP1

SPARCL1

G3BP2

VDP

CDKL2

ABCG2

RCHY1

PKD2

AMBN

IBSP

Human

Original

Frog HERC6

PPM1K

DMP1

CDKL2

G3BP2

VDP

AMBN

RCHY1

Fig. 3. Reconstruction of ancient tetrapod SCPP gene cluster. Lo-

cation and orientation of human and frog genes around SCPP genes are illustrated. Three SCPP genes, AMBN, DMP1, and IBSP, found both in the frog and human genomes, are depicted here. Many genes present within two gaps between human AMBN and RCHY1, and between VDP and SPARCL1 are not shown here. Different gray scales represent conserved gene blocks or SCPP genes. See text for possible inversions that occurred in the lineages leading to humans and the frog.

In this study, we compared genes in the frog and human genomes (fig. 3). The order and orientation within two gene blocks CDKL2-G3BP2-VDP and DMP1-IBSPPKD-ABCG2-PPM1K-HERC6 are both conserved in these two genomes, strongly suggesting that this was the ancient, original configuration of these genes. In the human genome, the CDKL2-G3BP2-VDP block is adjacent to RCHY1, whereas in the frog genome, this block is separated from RCHY1 by AMBN and is arranged in the opposite direction. These current configurations would be explained by a small number of intrachromosomal rearrangements: an inversion of the original CDKL2-G3BP2VDP-AMBN block in the lineage toward the frog and two other inversions involving AMBN and the RCHY1CDKL2-G3BP2-VDP block, together with their adjacent genes, in the lineage leading to humans. We thus assume that AMBN was originally situated adjacent to SPARCL1 with the transcriptional direction opposite to the dentin/ bone SCPP genes (fig. 3). In this model, we hypothesize that only a single small inversion occurred in the frog genome. A small number of intrachromosomal rearrangements have also been observed in the frog MHC region, supporting our model [Ohta et al., 2006]. This model also suggests that the gap between the dentin/bone and enamel SCPP gene clusters in modern mammals have generated after the split from amphibians, but before the origin of monotremes that we described above. 14

Cells Tissues Organs 2007;186:7–24

Intriguingly, this hypothetical configuration of the primordial tetrapod SCPP gene cluster is comparable with that of the fugu SCPP gene cluster; SCPP1, involved in dentin formation, and SPARCL1 are arranged in the opposite direction, whereas SCPP2 and SCPP4, expressed at the secretory stage of enameloid matrix, are located on the other side of SPARCL1 with the same orientation (fig. 2, 3) [Kawasaki et al., 2005]. Furthermore, principally, dentin SCPPs are rich in charged, particularly acidic amino acids, while enamel and enameloid SCPPs have more Pro and Gln both in teleosts and tetrapods (fig. 2) [Kawasaki and Weiss, 2007]. From these characteristics, we assume that the two classes of SCPP genes, one for the tooth body and the other for the tooth surface, had been separated before the split of ray-finned fish and lobefinned fish. This characteristic amino acid composition of enamel/enameloid SCPPs appears to persist in their more recent descendants; all milk and salivary class SCPPs also have abundant Pro and/or Gln residues, with the exception of the His-rich salivary proteins HTN1 and HTN3. Notably, Pro, Gln, and charged amino acids (Glu, Lys, and Arg), all these amino acids are thought to promote the intrinsically disordered structure, an important characteristic of SCPPs as we will describe in more detail below [Dunker et al., 2001; Radivojac et al., 2004; Huq et al., 2005; Kawasaki and Weiss, 2007]. The Origin of SPARCL1 In this study, we cloned the SPARC genes from cartilaginous fish (shark) and jawless vertebrates (lamprey). The lamprey has two distinct SPARC genes, which we call here SPARCA and SPARCB. All these new sequences are well conserved in domains II and III, whereas domain I has basic amino acids (one in SPARCA and two in shark SPARC and SPARCB) that are always found in non-vertebrate SPARC but have never been identified in any teleost or tetrapod SPARC. In addition, domain I of SPARCB contains 97 amino acids, the largest among vertebrates and comparable to tunicate SPARC (111 amino acids in Ciona), although still considerably smaller than SPARCL1 (378 amino acids in medaka is the smallest known) [Kawasaki et al., 2004]. Domain I of shark SPARC and SPARCA consist of 47 and 53 amino acids, respectively, similar to that of the other vertebrates. We used these shark and lamprey SPARC sequences for phylogenetic analysis. The results obtained by the NJ and ML methods show exactly the same topology (fig. 4). Shark SPARC is closer to bony fish (Osteichthyes) SPARC genes rather than SPARCL1 genes with high statistical support (96 and 86% by the NJ and ML methods, respecKawasaki/Buchanan/Weiss

Fig. 4. Phylogenetic tree of SPARC and

96/86 92/98

Frog Fugu Medaka Shark

90/87 100/100 100/98 65/64 94/83

0.2 98/92 NJ/ML

SPARC

72/90 100/98

Human Mouse Quail

Human Mouse Quail

Fugu Medaka Lamprey A Lamprey B Tunicate Sea urchin

SPARCL1

98/93 83/81

100/99

SPARC

SPARCL1. A phylogenetic tree obtained by the NJ method is shown here (
parameter:  = 1.19). The ML method yielded trees with the identical topology. Statistical supports obtained by the NJ (bootstrap value) and ML (quartet puzzling support value) methods are shown at the nodes. Lamprey SPARCA and SPARCB are represented by A and B, respectively. GenBank accession numbers used in this analysis are: NP_ 003109.1 (human), NP_033268.1 (mouse), O93390 (quail), NP_989347.1 (frog), NP_ 001027722.1 (fugu), ABB70180.1 (medaka), EF033075 (shark), EF033074 (lamprey A), EF033076 (lamprey B), NP_001027592 (tunicate), and AB276122 (sea urchin) for SPARC; and NP_004675.2 (human), NP_ 034227.1 (mouse), AAA49500.1 (quail), AB276121 (medaka), and NP_001027724.1 (fugu) for SPARCL1.

tively), and both lamprey SPARC genes split before the divergence of the jawed vertebrate (Gnathostomata) SPARC and SPARCL1 clades with relatively high statistical support (90 and 87% by the NJ and ML methods, respectively). These results suggest that SPARCL1 arose from SPARC before the shark-bony fish divergence but after the origin of the lamprey (fig. 4). The branch patterning of two lamprey SPARC genes is not statistically well supported (65 and 64% by the NJ and ML methods, respectively). This is due to the relative position of these two genes: the lamprey SPARCA may have branched off earlier than SPARCB, or both copies split in the lamprey lineage after the divergence from the stem jawed vertebrates. The sequences of hagfish SPARC would resolve the branch patterning of these two lamprey genes. Based on the generally accepted molecular clock hypothesis, we previously estimated that SPARCL1 arose from SPARC not so long before the divergence of rayfinned fish and lobe-finned fish [Kawasaki et al., 2004]. In contrast, a different study estimated the divergence date between SPARC and SPARCL1 to be in the Precambrian [Delgado et al., 2001; Sire et al., 2006], which is significantly earlier than our estimate. However, neither of these two studies used SPARC genes of cartilaginous fish or jawless vertebrates. Moreover, the molecular clock analysis potentially has large statistical errors. We now suggest that SPARCL1 arose from SPARC in Paleozoic jawless vertebrates after the divergence of modern jawless vertebrates but before the origin of cartilagi-

nous fish. After the gene duplication, SPARC lost basic amino acids from domain I, while SPARCL1 obtained a large exon and increased the size of intrinsically disordered domain I. Significantly, this initial SPARCSPARCL1 divergence overlaps with the origin of vertebrate skeletal mineralization as suggested by the fossil record (fig. 1) [Donoghue and Sansom, 2002; Janvier, 2003; Donoghue et al., 2006].

Gene Duplication and Skeletal Mineralization

Cells Tissues Organs 2007;186:7–24

SPARCL1 Arose from SPARC by Genome Duplication The period when SPARCL1 arose from SPARC, which we estimated above, also overlaps with the extensive gene duplication events that occurred early in vertebrate evolution. It has been proposed that two rounds of whole genome duplication (WGD) played a critical role to shape the genomes of jawed vertebrates (2R-hypothesis) [Ohno, 1970]. Although this hypothesis has long been a matter of controversy, recent studies support the idea that at least one, probably two rounds of WGD took place in a short period of time, first in the stem vertebrates and second in stem jawed vertebrates (fig. 1) [Holland et al., 1994; Gu et al., 2002; McLysaght et al., 2002; Dehal and Boore, 2005; Meyer and Van de Peer, 2005]. Below we will mention only the 2R-hypothesis but our discussion is compatible with one round of WGD and subsequent large-scale segmental duplications instead of two rounds of WGD. The human SPARC gene resides on 5q31.3-q32, but SPARCL1 on 4q22.1. These two genes are also located on two different chromosomes in mammals, birds, and tele15

osts [Kawasaki and Weiss, 2006], which is consistent with the hypothesis that SPARCL1 arose from SPARC by WGD. In order to test this hypothesis, we investigated 410 genes on 5q31-qter around SPARC (52-Mb region between HINT1 and TRIM52), and found that 126 genes (31%) have a paralog on chromosome 4, 86 genes (21%) on chromosome 10, and much smaller numbers on the other chromosomes (data not shown). These results are consistent with a recent genome-wide analysis, which clearly detected a paralogous region of 5q on chromosomes 4 and 10, and less clearly on chromosomes 2 and 8 [Dehal and Boore, 2005]. Among the 126 genes that have a paralog on chromosome 4, 119 genes also possess one or more paralogs on different chromosomes. Significantly, 47 of the 119 genes have a paralog on chromosome 10. However, among these 119 genes, 114 genes show the highest sequence homology to the paralog on chromosome 4 (or equally high homology to other paralogs on different chromosomes). In contrast, only five genes (DND1, PLAC8L1, RUFY1, HNRPH1, and MGAT4B) showed higher sequence homology to a paralog on different chromosomes. For instance, among the three SPOCK genes, SPOCK1 (5q31) shows higher sequence homology to SPOCK3 (4q32.3) rather than SPOCK2 (10q22.1). These results show that many genes around SPARC have paralogs on chromosomes 4 and 10, supporting the idea that the many paralogs on chromosomes 4, 5, and 10 were created by WGD, not by tandem duplication. Furthermore, among these paralogs, two genes on chromosomes 4 and 5 appear to be especially close to each other, suggesting that the paralogs on these two chromosomes were generated relatively late. Next we investigate this assumption by phylogenetic analysis. Timing of the Genome Duplication Previous studies have shown four closely related paralogs (quartets) thought to have arisen by the two rounds of WGD. Such quartets exist on each of the four chromosomes 4, 5 (near SPARC), 10, and 8, and their phylogenetic relationship has been investigated for the PPP2R2B, DPYSL3, FGFR4, and F12 quartets (below, symbols of the gene on chromosome 5 represent the quartet name) [Suga et al., 1999; Escriva et al., 2002; Vienne et al., 2003]. We also analyzed the PDLIM4 and UNC5A quartets on the same four chromosomes and the TCF7, ANXA6, KCNIP1, and MXD3 quartets on chromosomes 4, 5, 10, and 2. It has been thought that, among the paralogs that arose by the two rounds of WGD, many genes have been lost from the genome and only a small fraction of the four original 16

Cells Tissues Organs 2007;186:7–24

duplicates persist today [Nadeau and Sankoff, 1997; Wolfe, 2001]. TCF7L1 (2p11.2), KCNIP3 (2q21.1), DPYSL2 (8p12), and PPP2R2A (8p21.2), members of four different quartets, reside on two different human chromosomes, whereas their frog orthologs are all located within a small chromosomal region (scaffold 30). The proximal location of these frog genes supports a previous study inferring that parts of human chromosomes 2 and 8 had been a contiguous region paralogous to chromosomes 4, 5, and 10 [Lundin et al., 2003]. Our analysis also suggests that this paralogous region was shuffled and split into two different chromosomes after the human-frog divergence. We analyzed the phylogenetic relationship of all these quartets together with tunicate orthologs. Tunicates are the closest relatives of vertebrates but separated before the WGD, and hence their genes serve as a suitable outgroup for this analysis [Philippe et al., 2005; Delsuc et al., 2006]. Among these ten quartets, the PDLIM4 and F12 quartets were excluded from this study, because the tunicate ortholog of the PDLIM4 quartet was not found and more than four homologs of F12 were identified in the database (data not shown). For each of the other eight quartets, we found a single tunicate ortholog that is separated from the four human genes with the longest branch (fig. 5), confirming that the divergence of tunicates and the stem vertebrates predates the duplication of these genes. These phylogenetic trees also show close relationships between paralogs on chromosomes 4 and 5 for seven quartets (PPP2R2C-PPP2R2B and FGFR3-FGFR4 with relatively high statistical support, and CRMP1-DPYSL3, UNC5C-UNC5A, LEF1-TCF7, ANXA5-ANXA6, and KCNIP1-KCNIP4 with lower support; see fig. 5). The only exceptions are MXD4 and MXD3, which do not show the closest relationship within the quartet (fig. 5). The close relationship between paralogs on chromosomes 4 and 5 corroborates previous studies for the PPP2R2C, FGFR4, and DPYSL3 quartets, as well as the F12 quartet that was not used in this study [Suga et al., 1999; Escriva et al., 2002; Vienne et al., 2003]. In contrast to the close relationship between paralogs on chromosomes 4 and 5, our trees show inconsistent topologies for those on chromosomes 10 and 2 (or 8). If four paralogs were generated by two rounds of WGD, their phylogenetic trees should show a symmetrical topology such as ((4,5),(8,10)). However, it is now known that asymmetrical trees such as ((4,5),8),10) are common to the quartets generated by the two rounds of WGD. After WGD, meiotic recombination may continue equally Kawasaki/Buchanan/Weiss

PPP2R2B (5q31-q32)

TCF7 (5q31.1) PPP2R2C (4)

91 78

65 95

PPP2R2B (5) PPP2R2A (8) PPP2R2D (10) PPP2R2 Ciona

0.02

0.01

DPYSL3 (5q32)

CRMP1 (4) DPYSL3 (5) DPYSL2 (8) DPYSL4 (10) DPYSL Ciona

61 50

ANXA6 (5q32-q34) ANXA5 (4) ANXA6 (5) ANXA8 (10) ANXA4 (2) ANXA Ciona

64 37

0.1

0.05

FGFR4 (5q35-qter)

KCNIP1 (5q35.1) 90 59

FGFR3 (4) FGFR4 (5) FGFR2 (10) FGFR1 (8) FGFR Ciona

0.05

UNC5A (5q35.2)

LEF1 (4) TCF7 (5) TCF7L2 (10) TCF7L1 (2) TCF7 Ciona

43 49

KCNIP4 (4) KCNIP1 (5) KCNIP2 (10) KCNIP3 (2) KCNIP Ciona

0.05

37 51

MXD3 (5q35.3)

UNC5C (4)

68 56

UNC5A (5) UNC5B (10) UNC5D (8) UNC5 Ciona

MXD1 (2) MXI1 (10) MXD4 (4) MXD3 (5) Not7 Ciona

0.05

0.1

Fig. 5. Phylogenetic trees of quartets present on chromosomes 4, 5, 10, and 2 or 8. The number in parentheses

represents the location of the representative gene on chromosome 5 (after quartet name) or the chromosome on which each gene resides (after gene symbol). These trees were constructed by the NJ method using uniform substitution rates among different sites and the Johns-Taylor-Thornton matrix model. Bootstrap values are indicated at the nodes. Ciona Not7 was found in GenBank (NP_001027611).

among the resulting four equivalent chromosomes for a considerably long period. The subsequent differentiation of duplicate chromosomes will terminate the recombination but this timing could be different among chromosomal regions [Aburomia et al., 2003]. If two successive WGD took place, the relationship among the eight equivalent chromosomes could become extremely complicated [Furlong and Holland, 2002]. There are some other hypotheses that explain asymmetrical topologies [Spring, 1997; Wolfe, 2001], although the inconsistency might be simply due to statistical errors. But however gene duplication happened, our results support the scenario that the two paralogs on chromosomes 4 and 5 became differentiated relatively late, after the second WGD. Based on this analysis, we assume that the ancient SPARC differentiated into two distinct genes in our linGene Duplication and Skeletal Mineralization

eage through the two rounds of WGD. SPARCL1 is one of the two genes that probably arose by the second WGD. Possibly, the two lamprey SPARC genes described above might have been generated by the first WGD. The second WGD is thought to have occurred in the stem jawed vertebrates, which is consistent with the result obtained by the phylogenetic analysis of SPARC and SPARCL1 (fig. 1, 4). Vertebrate Skeletal Mineralization and Gene Duplication Gene families evolve through the birth-and-death process: some new genes created by gene duplication stay in the genome for a long time, while others become inactivated or deleted from the genome [Nei, 2005; Nei and Rooney, 2005]. The SCPP gene family appears to elucidate Cells Tissues Organs 2007;186:7–24

17

this. Reiterative tandem gene duplications have generated many SCPP genes but not a few genes have been secondarily lost in some lineages, as we showed above (fig. 2). Another gene duplication mechanism is WGD, which will make numerous redundant genes available for evolution [Ohno, 1970]. Hence, it was previously proposed that there is a strong relationship between many redundant genes, created by the two rounds of WGD, and the phenotypic complexity of vertebrates [Holland et al., 1994; Sidow, 1996]. However, we also know that WGD is not rare in teleosts and amphibians [Ohno, 1970], suggesting that many early extinct vertebrates also had duplicated genomes. In addition, the fossil record suggests that many complex traits gradually appeared in early vertebrates, and the WGD postdated the acquisition of at least some complex characters that now use multiple genes generated by this event, such as fin-like structures that appeared in extinct jawless vertebrates [Donoghue and Purnell, 2005]. It is thus likely that complex traits evolved over the period of the two rounds of WGD. Among vertebrates with duplicate genomes, one lineage opportunistically used many redundant genes available at the time, coincident with many other genetic changes. The functionally differentiated genes present today are the surviving originally redundant genes that took on distinct functions before they could be destroyed by mutational decay. They have survived by developing new functions for use by new or old traits [Ohno, 1970; Zhang, 2003], or, alternatively, by the complementary loss of ancestral subfunctions [Force et al., 1999, 2005]. Skeletal mineralization is among the traits that coopted redundant genes generated by WGD; after the WGD, some of the duplicate genes for fibrillar collagens and integrins were employed for use in mineralized tissues [Kawasaki et al., 2004]. In addition, SPARCL1 arose from SPARC simultaneously with these genes, and then SCPP genes originated from SPARCL1 after it had obtained a larger intrinsically disordered domain I (fig. 1). Our results cannot resolve which evolved first, the SCPP genes or the mineralized skeleton. If SCPP genes arose earlier than the origin of the mineralized skeleton, these genes had probably been used for different functions at the time, and were subsequently co-opted for tissue mineralization. However, many tetrapod and teleost SCPP genes are highly specialized for mineralization, suggesting that these genes arose during the modification of mineralized tissues. We thus speculate that SPARC alone, or together with proteins other than SCPPs, may have been the key molecule in early skeletal mineralization, a possibility we will discuss next. 18

Cells Tissues Organs 2007;186:7–24

SPARC and Early Skeletal Mineralization In addition to dentin and bone, SPARC is also distributed in the non-mineralizing basal lamina of protostomes and deuterostomes as well as vertebrate dermis, where it associates with various collagens [Holland et al., 1987; Metsäranta et al., 1989; Porter et al., 1995; Timpl, 1996; Fitzgerald and Schwarzbauer, 1998; Martinek et al., 2002; Bradshaw et al., 2003]. In the basal lamina, this protein presumably binds polygonal meshwork forming type-IV collagens, and these proteins together maintain mechanical stability [Yan and Sage, 1999]. In dermis, SPARC facilitates the maturation of collagen fibrils, mainly consisting of type-I and type-III collagens, and reinforces their mechanical strength [Bradshaw et al., 2003]. Paleozoic jawless vertebrates developed mineralized dermal skeleton adjacent to the basal lamina [Reif and Richter, 2001]. Similar to the tooth, ancient mineralized skeleton consisted of bone, dentin, and enameloid (or their related tissues), and these tissues are thought to have developed through the interaction between epithelium and neuralcrest-derived ectomesenchyme [Schaeffer, 1977; Smith and Hall, 1990; Donoghue and Sansom, 2002; Donoghue et al., 2006]. We thus suggest that the tooth initially co-opted the gene-regulatory circuits that had been already used for the development of dermal skeleton, and hence SPARC, a gene expressed at a late stage of tooth development, was also expressed in the cells involved in ancient skeletal mineralization. SPARC as well as many other ECM proteins require calcium ions for protein-protein or proteincell interactions and maintain the integrity of ECM [Maurer et al., 1996]. We assume that these calciumbinding proteins all had been potentially available for early skeletal mineralization. Disordered Protein and Skeletal Mineralization The mature SCPPs originated from intrinsically disordered domain I of SPARC and SPARCL1. This domain does not fold into a rigid globular structure in a native state [Engel et al., 1987; Hambrock et al., 2003; Kawasaki et al., 2004]. Similar to domain I of these proteins, SCPPs have no or few cysteine residues, and hence most SCPPs cannot stabilize fixed globular structures through intramolecular disulfide bonds. Some SCPPs, including caseins and dentin/bone SCPPs, have been experimentally shown to be intrinsically disordered proteins [Creamer et al., 1981; Bhattacharyya and Das, 1999; Fisher et al., 2001]. Although this has not been shown experimentally for all SCPPs, several protein disorder prediction programs predict relatively long (130 amino acids or most part of the Kawasaki/Buchanan/Weiss

small SCPPs) disordered regions for all teleost and tetrapod SCPPs tested so far (data not shown) [Romero et al., 2001; Linding et al., 2003; Ward et al., 2004]. The intrinsically disordered proteins (or regions) typically have flexible unfolded structures, which facilitate interactions with various proteins, ions, and crystals at low affinities [Dunker et al., 2002]. These interactions include self-assembly and enzyme-substrate reactions. In particular, phosphorylation, glycosylation, and proteolysis predominantly occur within or adjacent to these regions, and such modifications can regulate the activity of these proteins [Dunker et al., 2002; Iakoucheva et al., 2004]. In many cases, initially intrinsically disordered regions adopt folded structures upon binding to target molecules (e.g. proteins and ions) [Uversky et al., 2000; Dyson and Wright, 2002]. Many of these properties common to intrinsically disordered proteins seem to be essential to the functions of SCPPs, particularly those of tetrapod dentin/bone SCPPs, which are especially similar to domain I of SPARCL1, and hence to the ancestral protein. Indeed, dentin/bone SCPPs associate with other ECM proteins such as collagens and integrins; these SCPPs bind to calcium ions and hydroxyapatite crystals, and these proteins are phosphorylated, glycosylated, and processed by proteases [Qin et al., 2004]. Long disordered regions are also consistently predicted within all the Pro/Gln-rich enamel/enameloid SCPPs. However, experimental analyses do not always support the idea that the large middle portion of AMEL consists of a disordered structure, but instead suggest that this portion contains extended polyproline II and/or -spiral structures [Matsushima et al., 1998; Le et al., 2006; Margolis et al., 2006; Robinson, 2006]. Among milk/saliva/ enamel SCPPs, AMEL is especially rich in Pro and Gln, and has Pro-Xaa-Yaa repeats (Xaa and Yaa represent any amino acids but Pro and Gln appear frequently) within the C-terminal half. This repeat is expected to adopt the polyproline-II helix [Williamson, 1994]. Although the polyproline-II helix is ordered and stiffer than disordered proteins, this structure is also extended and unfolded, and hence the majority of side chains are free to participate in protein-protein interactions, in a way resembling disordered structures [Rath et al., 2005]. Notably, an intrinsically disordered region is also present or predicted in two other important bone ECM proteins, BGLAP and matrix
-carboxyglutamic acid protein [Hauschka and Carr, 1982; Isbell et al., 1993; Uversky et al., 2000]. Experimental studies have suggested that the disordered structure of SPARC and BGLAP both change

into ordered structure (-helix) by binding to calcium ions [Hauschka and Carr, 1982; Engel et al., 1987], and the helical BGLAP show greatly increased affinity for hydroxyapatite. We thus speculate that disordered structures have an intrinsic potential to regulate mineral precipitation or crystallization, and that this property was employed for tissue mineralization in ancient jawless vertebrates. Intrinsically disordered or unfolded structures would allow SCPPs to evolve rapidly without changing their original property. Because SCPPs do not have functionally crucial side chains to form a tightly folded protein core, their amino acid sequences are less constrained than those of ordered proteins [Dunker et al., 2002]. Furthermore, with only a few exceptions, all introns in SCPP genes are phase 0, descended from the ancestral domain I of SPARCL1. In genes with all phase-0 introns (or, in fact, all phase-1 or -2 introns), neither exon duplication nor deletion changes the downstream reading frame, so that the original unfolded nature would be retained after such mutations. The number of exons varies even among orthologous SCPP genes as described above, showing that, on the evolutionary time scale, such mutations occur repeatedly. As a result of its long history, skeletal mineralization today uses many distinct SCPP genes with considerable sequence variation but shared protein characteristics. We presume that the structure of these genes and the intrinsically unfolded nature of their protein products are both important factors that have generated the considerable variation in SCPP sequences.

Gene Duplication and Skeletal Mineralization

Cells Tissues Organs 2007;186:7–24

Genetic Basis of Various Mineralized Tissues As a consequence of the birth-and-death evolution of the SCPP gene family, in the presence of selection for a favored trait, phenogenetic drift can occur [Kawasaki et al., 2005], i.e. the same conserved structure can be produced by different genes in different lineages [Weiss and Fullerton, 2000; Weiss and Buchanan, 2004]. The structure of teeth in both teleosts and tetrapods is similar in that body dentin is covered with a highly mineralized tissue, enamel or enameloid [Poole, 1967; Meinke and Thomson, 1983]. However, these two vertebrates use distinct sets of SCPPs for tooth mineralization; none of the seven fugu SCPP genes appear to be orthologous to any of the tetrapod SCPP genes [Kawasaki et al., 2005]. Thus, whereas the tooth has been inherited by teleosts and tetrapods from their common ancestor, different genes regulate tooth mineralization in these two lineages. We assume that this phenogenetic drift of skeletal mineralization involved two steps of genetic changes: the generation 19

of lineage-specific genes by gene duplication and the loss of a common ancestral gene. The two types, Pro/Gln-rich and acidic SCPP genes, arose before the divergence of rayfinned fish and lobe-finned fish by tandem gene duplication (fig. 1, 2). After their divergence, tandem gene duplication next created lineage-specific SCPP genes and their common ancestral genes have been deleted from the genome of at least one lineage. SPP1 remains in both of these lineages but has been lost in the frog and fugu. Furthermore, compared to mammals, frogs use smaller numbers of SCPP genes for both enamel and dentin/bone mineralization. These facts all suggest frequent gene duplication and loss of SCPP genes. In addition to the composition of ECM proteins, teleost enameloid is also developmentally different from tetrapod enamel [Meinke and Thomson, 1983; Smith, 1995]. The inner dental epithelial cells and mesenchymal cells (odontoblasts) both deposit enameloid ECM proteins, whereas only epithelial cells (ameloblasts) secrete enamel [Shellis and Miles, 1974; Sasagawa, 1995]. Moreover, fugu SCPP5, SPARC, and type-I collagen genes are expressed both in the inner dental epithelial cells and odontoblasts at the early secretory stage for enameloid and continuously in odontoblasts at the late secretory stage for dentin [Kawasaki et al., 2005]. The shared ECM proteins, particularly type-I collagen, the mineralization scaffold of both enameloid and dentin, show a close relationship between these two tissues. These findings imply that teleost enameloid arose through modification of dentin. Compared to enameloid, enamel is more specialized and its evolution cannot easily be explained by simple modifications of other mineralized tissues. We have thus hypothesized that enamel might have arisen through an intermediate tissue, which is similar to present teleost enameloid [Kawasaki et al., 2005]. Enamel and enameloid are both highly mineralized tissue but enameloid is closer to dentin; hence enameloid may be regarded as an intermediate tissue between dentin and enamel, as proposed previously [Hall, 2005]. There seem to be many other intermediate or unique mineralized tissues in modern vertebrates. Cementum is a bone-like collagenous tissue primarily surrounding the base of teeth of mammals and certain reptiles [Peyer, 1968; Halstead, 1974; Carlson, 1990]. However, recent investigations have revealed the expression of SCPP genes for both dentin/bone (DMP1, IBSP, and SPP1, but not DSPP) and enamel (AMBN but not AMEL or ENAM) in Hertwig’s epithelial root sheath that produces acellular cementum [Fong et al., 1996; Zeichner-David et al., 2003]. The composition of these proteins shows a unique feature 20

Cells Tissues Organs 2007;186:7–24

of this tissue. In addition, the opportunistic nature of selection can favor differing developmental pathways to some conserved, stable trait [True and Haag, 2001]. We have reported a unique fugu jaw wall that consists of dentin as implied from the expression of SCPP genes. However, this structure grows within mesenchymal condensation similar to bone, and hence is distinct from teeth that form between the epithelial and mesenchymal layers [Kawasaki et al., 2005]. These examples illustrate considerable variation among present vertebrate mineralized tissues, produced by the shifting of genetic resources made available by gene duplication and/or co-option, as well as by the use of different developmental pathways to similar ends. Ancient Mineralized Skeleton The fact that different mechanisms underlay the evolution of various unique mineralized tissues in modern vertebrates implies the possibility that early jawless vertebrates, which lived before or shortly after the origin of SPARCL1, and hence when no or only a few SCPP genes were available, may have also used unique ECM proteins and perhaps employed different developmental processes for skeletal mineralization. The differences among mineralization proteins may affect mineralization processes. Indeed, the mineralization process of enameloid is different between elasmobranchs and teleosts, suggesting that some mineralization proteins are different in these two clades [Sasagawa, 1998, 2002]. It is thus plausible that the high diversity of skeletal tissues found in early vertebrates may reflect the differences in ECM proteins and accompanying mineral crystallization processes used for their skeleton [also see Donoghue et al., 2006]. To test this hypothesis, it is critical to identify genes involved in tissue mineralization from cartilaginous fish.

Conclusions

The duplication histories of SCPP genes in various vertebrate lineages make it clear that there may be a weaker relationship between genotype and phenotype than is usually thought. At the onset or at a very early stage of skeletal mineralization, SPARC was co-opted for use in the production of this new trait. SPARCL1 originated from SPARC at around this period through WGD, and then many SCPP genes arose initially from SPARCL1 by tandem duplication. Hence gene duplication has played a significant role in skeletal mineralization, both as it first arose and in sustaining it in current lineages. Kawasaki/Buchanan/Weiss

Gene duplication provides new genetic materials, which might be recruited by an adaptive trait. Nevertheless, this is only one way among many other possible ways to produce genetic changes. SCPP genes, and other genes involved in regulating these genes or the tissue in which they are expressed, are thus a consequence of the evolution of skeletal mineralization, and this is a story that undoubtedly continues without end.

Acknowledgments We thank Dr. Yuko Ohta and Prof. Martin F. Flajnik at the University of Maryland, and the National Aquarium in Baltimore for help providing us the shark sample, Mr. Bill Swink at the Hammond Bay Biological Station for providing us lampreys, and Mr. Samuel Sholtis for critical discussion. This work was made possible by the financial support from awards SBR9804907, SBE0343442, and BCS-0343442 from the US National Science Foundation, and by research funds from the Pennsylvania State University to K.M.W.

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Cells Tissues Organs 2007;186:25–48 DOI: 10.1159/000102679

The Origin and Evolution of Enamel Mineralization Genes Jean-Yves Sire a Tiphaine Davit-Béal a Sidney Delgado a Xun Gu b a

UMR 7138, Université Pierre et Marie Curie-Paris 6, Paris, France; b Department of Genetics, Development, and Cell Biology, Iowa State University, Ames, Iowa, USA

Key Words Enamel
Evolution
Genomics
Mineralization
Tooth

Abstract Background/Aims: Enamel and enameloid were identified in early jawless vertebrates, about 500 million years ago (MYA). This suggests that enamel matrix proteins (EMPs) have at least the same age. We review the current data on the origin, evolution and relationships of enamel mineralization genes. Methods and Results: Three EMPs are secreted by ameloblasts during enamel formation: amelogenin (AMEL), ameloblastin (AMBN) and enamelin (ENAM). Recently, two new genes, amelotin (AMTN) and odontogenic ameloblast associated (ODAM), were found to be expressed by ameloblasts during maturation, increasing the group of ameloblast-secreted proteins to five members. The evolutionary analysis of these five genes indicates that they are related: AMEL is derived from AMBN, AMTN and ODAM are sister genes, and all are derived from ENAM. Using molecular dating, we showed that AMBN/AMEL duplication occurred 1600 MYA. The large sequence dataset available for mammals and reptiles was used to study AMEL evolution. In the N- and Cterminal regions, numerous residues were unchanged during 1200 million years, suggesting that they are important for the proper function of the protein. Conclusion: The evolutionary analysis of AMEL led to propose a dataset that will be useful to validate AMEL mutations leading to Xlinked AI. Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel 1422–6405/07/1861–0025$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/cto

Introduction

Living vertebrates possess a great diversity of mineralized elements, comprising not only endochondral and dermal bone (including osteoderms and scutes), mineralized cartilage, and teeth (dentin and enamel), but also scales, fin rays and otoliths, and egg shells [Huysseune and Sire, 1998; Sire and Huysseune, 2003]. The first mineralized elements, which have given rise to the current

Abbreviations used in this paper AIH1

amelogenesis imperfecta 1, hypoplastic/hypomaturation, X-linked AIH2 amelogenesis imperfecta 2, hypoplastic local, autosomal dominant AMBN ameloblastin AMEL amelogenin AMTN amelotin EMP enamel matrix protein ENAM enamelin gene KLK4 kallikrein-4 (prostase, enamel matrix, prostate) MMP20 matrix metallopeptidase 20 (enamelysin) MYA million years ago ODAM odontogenic, ameloblast associated (APIN, FLJ20513) SCPP secretory calcium-binding phosphoprotein SIBLING small integrin-binding ligand, N-linked glycoprotein SPARC secreted protein, acidic, cysteine-rich (osteonectin) SPARCL1 SPARC-like 1 (mast9, HEVIN) C4orf7 chromosome 4 open reading frame 7 (FDC-SP, MGC71894)

Dr. Jean-Yves Sire Equipe ‘Evolution et Developpement du Squelette’, UMR 7138 Université Pierre et Marie Curie-Paris 6, 7 quai St-Bernard, Case 5 FR–75252 Paris (France) Tel./Fax +33 1 44 27 35 72, E-Mail [email protected]

skeletal diversity in vertebrates, were identified as early as 500 million years ago (MYA) [Sansom et al., 1992, 1994; Janvier, 1996; Donoghue, 1998, 2001]. In fact, the occurrence of mineralized tissue in vertebrates was a major innovation, which was fundamental to the radiation of modern vertebrates in relation to the important roles of the skeletal elements in protection, predation and locomotion [Reif, 1982; Smith and Hall, 1990; Janvier, 1996; Donoghue, 2002; Donoghue and Sansom, 2002; Donoghue et al., 2006]. Our understanding of the mechanisms by which organisms form mineralized elements is still at a rudimentary stage, but we know that biomineralization is mediated by the organic matrix, either through its biological activity or in controlling nucleation, growth and microarchitecture of the mineral deposited [Carter, 1990]. It is assumed that the basic processes of biomineralization are common to all systems and that mineral formation by any individual biological system may diverge from this common pathway. This general definition applies to vertebrates in which the main skeletal elements derive from common ancestral elements [Huysseune and Sire, 1998; Sire and Huysseune, 2003] and there is growing evidence that most proteins currently involved in mineralization of skeletal tissues (bone, dentin, and enamel) also have diverged from a common ancestor [Kawasaki and Weiss, 2003; Kawasaki et al., 2004; Kawasaki and Weiss 2006]. The evolutionary analysis of genes coding for these ‘mineralizing’ proteins not only has the potential to provide insight into the debated question of the origin of mineralization in vertebrates and of its subsequent diversification, but could also bring important information for humans, as mutations of these proteins lead to genetic disorders (bone [Rowe, 2004]; dentin [Zhang et al., 2001], and enamel [Stephanopoulos et al., 2005]). This review is devoted only to our current knowledge on the origin and evolution of the genes coding for enamel matrix proteins (EMPs). The reader is referred to the paper by Kawasaki et al., published in this issue [pp 7–24], regarding the history of the other mineralizing proteins in vertebrates. In living and extinct vertebrates, teeth are protected by a hypermineralized tissue, either ‘true’ enamel (e.g. in tetrapods) or ‘enamel-like’ tissue, enameloid (e.g. in cartilaginous and ray-finned fish). These hard dental tissues are identified early in the history of the mineralized integument. They were present in the dermal skeleton of various lineages of jawless vertebrates [Ørvig, 1967, 1977; Reif, 1982; Smith and Hall, 1990; Janvier, 1996; Donoghue and Sansom, 2002; Donoghue et al., 2006]. Enamel 26

Cells Tissues Organs 2007;186:25–48

and enameloid are homologous tissues that correspond to different aspects of the same hypermineralization process [Donoghue et al., 2006]. Enamel has replaced enameloid in the lineage leading to tetrapods, probably by a process of heterochrony1 [Smith, 1995], but enameloid was conserved in two important lineages, chondrichthyans2 and actinopterygians3. The close evolutionary relationships, the similar features of the ameloblasts during their formation, and the same maturation process strongly indicate that both enamel and enameloid matrices could contain similar mineralizing proteins, and that some of them (if not all) were already present in toothrelated elements of early vertebrates, 500 MYA. Unfortunately, our knowledge on EMP genes is restricted to the tetrapods, and the road is still long before we will be able to test the hypothesis of an early origin of EMPs. In mammals, the enamel matrix is composed of three specific proteins secreted by ameloblasts: amelogenin (AMEL), which represents 90% of the matrix deposited, and enamelin (ENAM) and ameloblastin (AMBN), which are components of the remaining 10% organic matrix. Evolutionary analyses have indicated that these three EMPs constitute a family, which, itself, is included into a larger family, the secretory calcium-binding phosphoprotein (SCPP) family. This SCPP family comprises other Ca-binding proteins: some saliva proteins, milk caseins and small integrin-binding ligand, N-linked glycoproteins (SIBLINGs) [Fisher and Fedarko, 2003], which contain five dentin and bone proteins [Kawasaki and Weiss, 2003]. Interestingly, with the exception of AMEL that is located elsewhere (on sex chromosomes X and Y in placental mammals), all SCPP genes are located in two clusters on the same autosomal chromosome. This supports the idea that SCPP genes originated by tandem duplication followed by neofunctionalization. In humans, several types of amelogenesis imperfecta (AI), leading to enamel hyploplasia or hypomineralization, are related to mutations in AMELX (14 X-linked AI, AIH1, identified to date [Hart et al., 2002; Kim et al., 2004; Stephanopoulos et al., 2005]) or in ENAM (5 autosomal-dominant AI, AIH2 [Hart et al., 2003; Hu and Yamakoshi, 2003; Kim et al., 2005]) genes [review in Stephanopoulos et al., 2005]. In contrast, although being con-

1

Heterochrony: developmental changes in the timing of events, leading to changes in size and shape from an ancestral state. 2 Chondrichthyans: the cartilaginous fishes, including sharks, rays and chimaeras. 3 Actinopterygians: the ray-finned fish, which are the dominant group of vertebrates.

Sire /Davit-Béal /Delgado /Gu

sidered as a candidate gene, AMBN was excluded from a causative role within the families studied [Mardh et al., 2001]. Since a few years, we focus our attention on EMP gene relationships (AMEL, AMBN, and ENAM), and more precisely on the origin and evolution of AMEL, the best known member of the family [Delgado et al., 2001; 2005; Sire et al., 2005; 2006; Delgado et al., in press]. Here, (i) we summarize these previous data, (ii) we provide new information on two newly identified genes, amelotin and APIN protein, that are expressed by the ameloblasts, (iii) we provide a date for EMP gene origin and discuss this result in the light of our knowledge of enamel and/or enameloid appearance in vertebrate evolution, and (iv) we show how evolutionary analysis of AMEL can help to identify structural features that might be important for the protein function, and to validate mutations responsible for genetic diseases.

Ameloblast Products: EMPs, and Amelotin and APIN Proteins

In mammals, the synthetic activity of ameloblasts is divided in two successive phases corresponding to two stages of enamel formation: secretion and maturation, separated by a transition stage. To our knowledge, during the former step ameloblasts deposit four proteins in the extracellular matrix: three EMPs (AMEL, ENAM, and AMBN) and a tooth-specific, calcium-dependent peptidase, MMP20 (= enamelysin) [Bartlett et al., 1998; Bartlett, 2004]. During the transition and maturation stages, ameloblasts have been shown to produce a fifth protein, kallikrein 4 (KLK4), a pleiotropic, calcium-independent protease, which is involved in the final proteolysis of the remaining organic matrix [Simmer et al., 1998; Hu et al., 2002; Simmer and Hu, 2002]. Recently, two novel genes were found to be also expressed by ameloblasts during tooth formation: amelotin (AMTN, but annotated UNQ689 in human genome build 36.2) [Iwasaki et al., 2005; Moffat et al., 2006b] and ODAM (‘odontogenic, ameloblast associated’, previously named APIN or FLJ20513) [Moffat et al., 2006a]. Can the proteins encoded by these two genes be considered EMPs? In other words, although being produced by ameloblasts, are AMTN and ODAM structural proteins playing a role in enamel matrix formation and/or mineralization? In rats, AMTN was localized to the basal lamina of maturation stage ameloblasts [Moffatt et al., 2006b]. This location seems to indicate a possible role of AMTN in cell The Origin and Evolution of Enamel Mineralization Genes

adhesion, and it also demonstrates the absence of AMTN participation in enamel matrix formation. In humans, ODAM was first identified from extracts of amyloid deposits obtained from calcifying epithelial odontogenic tumors [Solomon et al., 2003]. Transcripts of this gene were also found at a high level in gastric cancer [Aung et al., 2006]. In rats, ODAM is specifically expressed in ameloblasts during maturation stage [Moffatt et al., 2006a], but the location of the protein in the extracellular matrix remains to be shown. However, late expression during tooth formation does not mean that the secreted ODAM protein is not incorporated in the enamel matrix at the end of the mineralization process. Such a location would not be surprising if one considers that the protein was first isolated from calcifying tissues of odontogenic tumors [Solomon et al., 2003]. Therefore, if AMTN cannot be considered an EMP, the few data available to date do not permit to exclude ODAM from this family. Interestingly, these two genes are located in the same cluster as EMPs, and they share structural similarities with the members of this family (see below). This indicates that AMTN and ODAM were probably created after duplication of an ancestral EMP gene and, therefore, that they belong to the SCPP family [Kawasaki and Weiss, 2006]. In the following, we provide some data on these two newly identified ameloblast-secreted proteins, although concentrating on the evolutionary relationships of EMPs (AMEL, AMBN, and ENAM).

Evolutionary Relationships of AMTN, ODAM, and EMP Genes

EMPs are evolutionarily related, forming a gene family that belongs to a super-gene family called SCPP [Kawasaki and Weiss, 2003]. All SCPP genes probably derive from a common ancestor by gene duplications. The key gene could be SPARC-like1 (SPARCL1, also called HEVIN or SC1), which was created after a duplication of SPARC (osteonectin) [Kawasaki et al., 2004]. Four lines of evidence have permitted to establish SCPP relationships, and SPARCL1 may resemble the ancestral form of SCPP: (i) common gene structure and similar protein characteristics in the N-terminal region, (ii) in most SCPPs, presence of an SXY phosphorylation site encoded in the 3 region of the second coding exon, suggesting Ca-binding properties, (iii) location on the same chromosome, and (iv) presence of SPARCL1 on this chromosome, adjacent to the dentin-bone protein gene cluster [reviewed by Kawasaki and Weiss, 2006]. Cells Tissues Organs 2007;186:25–48

27

Ancestral Human Chimpanzee Rhesus monkey Mouse Rat Cow Pig Dog Elephant Opossum

exon2 | MKTTILLFCL LGTTQSLPKQ .RS....... ..S.R...-. .RS....... ..S.R...-. ...M...... ..S.....-. ...M...L.. ..SA...... ...VV..L.. ..SA....R. ..AA...... ..S.L...M. .......... ..S.L...M. ...M...LY. ..S.....A. .......... ..S....... ...AV..... ...I....Q.

exon3 LNPALGLPPT KLGPDQPTLL .K........ ..A...G..P .K........ .PA...E..P .K........ ..A...G..P ....S.V.A. .PT.G.V.P. .S....A.A. .PT.G.V.P. .....V.... ..V...A... F..V...... ..V...A..R .......... ..T.H.A... .......SAA ..V...A... .Y.GV....P ...LE..A.F

| NQQQPNQVFP ....S..... ....S..... ....S..... P......... T......... .P.......S .......... .......... .......... TP..S..L..

exon4 SLSQIPLTQM .......... .......... .......... .I.......L .I.......L ........H. .......... .......... .........L P.GL......

Ancestral Human Chimpanzee Rhesus monkey Mouse Rat Cow Pig Dog Elephant Opossum

exon5 | QTLPMTLGDL NIQHQLKPQM ..H.L...G. .V.Q..H.HV ..H.L...G. .V.Q..H.HV .AH.L...V. .LPQ..Q.H. H...F...P. .G.Q..Q... ....F...P. .G.Q..Q... ....LA..G. KV.Q..Q... ....L...A. .V.Q..Q... ....LS..V. .T.Q..Q... ..F.LN..G. T.KQ..QS.L .I.......T S.AP.VN...

exon6 LPIIVAQIGA ...F.T.L.. ...F.T.L.. ...F.T.L.. .......L.. .......L.. ..V...HF.. I.V...HL.. ..V...HL.. .......L.. ..VL......

| exon7 | QGAILSSEEL ..T....... ..T....... ..T....... ...L...... ...L...... ..T....... .......... H......... .......... ...VR.....

PMAPQIFTGL .---....S. .---....S. .---....S. .L.S...... .L.S...... QGTS..L... .ATR..L... .GS....... .......A.. .I........

exon8 LIQPLFPGAI LPTSQAGTNP I.HS....G. .......A.. I.HS....G. .......A.. I.HS....G. .......A.. ..H....... P.SG....K. ..H....... Q.SG.T.AK. IFH....... .....--A.. IFHT...... ..P.P--AK. IF........ ....P--A.. .......... ....L..AT. ....FGT..T ...G.S.IDA

Ancestral Human Chimpanzee Rhesus monkey Mouse Rat Cow Pig Dog Elephant Opossum

QSGGNPAIQG GA.V...T.. GA.V...T.. .A.V...... .A.AKAVN.. .A.AS..N.A .A.A...A.. .A.A...V.. .A........ .A.L.....R ........W.

exon8 GTDD-VFEAT .....D.AV. .....D.AV. .....D.AV. V...DDY.MS V...DDY.MS .....D.AS. .....D.DV. D....D.GV. D..S...GV. SP........

TPAGIQRATH .......S.. ...D...S.R .......S.. ....LR.... ....L..... .......GRP ....L..G.. A......G.. ....L..GMR I.V...K..-

TTEETTTEAP AI..A...SA AI..A...SA A.......*. ...G..IDP. ...G..MDP. .....P.GS. A......GS. ..Q...SGP. ..G.....S. --.GS.....

| ex9 NGTQ ..I. ..I. ..I. .R.. .R.K K.I. ..M. .... .EI. ...D

TPEGQLPTPS ..A.R..... ..A.R..... ..A.R..... .TP.HVT..G .TP.H-T..A ...DPFS... ..R.PF..S. ...SFST... ...KHPS.S. .S........

| LTLGTDLQLI ....P..H.L ....P..H.L ....P..H.L ....S..P.F ....S..P.F ....SN...L ....S....L F..AS....L ....S...QL FSV...M..M

exon 5 NPATGMPPGT (80) ...A..T... ...A..T... ...A..T... ...A.-.H.A ....-..H.. ...A...S.. ...I..V.SS ...A..AS.. ......A... T....LL..I

NAQDGALPAG (160) DV...S.... DV...S.... DV...S.... DV.N.V..TR DV.N....TR D..N.I.... D..N.IH... D..N.I.... EV.E.I.... .T..A.....

(214) (210) (210) (210) (213) (214) (212) (212) (212) (214) (211)

Fig. 1. Amelotin (AMTN): alignment of 10 complete mammalian amino acid sequences and of the putative an-

cestral sequence (shown at the top). Six sequences were inferred from DNA sequences retrieved in databases (blast search against sequenced genomes). These sequences were checked against three published complete coding sequences: human, accession number AY358528; mouse, AK017352, and rat, DQ198381. The pig sequence was obtained from the literature [Moffatt et al., 2006]. The putative ancestral sequence was calculated using PAUP 4.0 and MacCLADE 3.06. Vertical bars indicate the limits between exons. The signal peptide is in a box. The total number of residues in each protein is indicated at the end of each sequence. Unchanged residues are shown on a gray background. · · · · · = Identical residue; – – – = indel.

Recent studies on the origin and evolution of AMEL in tetrapods have extended our knowledge on EMP relationships [Sire et al., 2005, 2006]. A phylogenetic analysis using a large set of sequences demonstrated that AMEL and AMBN are sister genes, and that AMEL was created from a duplication of AMBN. In addition, it was shown that both genes are related to ENAM, which was recognized as a more ancient member of the EMP family. The calculation of putative ancestral sequences of EMP genes and the use of SPARCL1 as an outgroup were helpful for this phylogenetic analysis. Putative ancestral sequences permit to go back to the gene origin, while the whole dataset of sequences is less informative to reveal possible relationships. Indeed, although they are phylogenetically 28

Cells Tissues Organs 2007;186:25–48

related, EMP genes show large sequence variations when comparing evolutionary distant lineages. Moreover, since their creation, hundreds of million years ago, AMEL, AMBN, and ENAM have acquired specific functions and their sequences diverged rapidly. However, currently available sequences permit to calculate putative ancestral sequences of EMPs at the origins of the mammals only, i.e. when monotremes4 and therians5 diverged, 225 MYA [van Rheede et al., 2006]. Using amniote or tetrapod sequences was not possible because of the few sequences 4

Monotremes: egg-laying mammals: extant members are the echidnas and the duck-billed platypus. 5 Therians: marsupials and placental (eutherian) mammals.

Sire /Davit-Béal /Delgado /Gu

Ancestral Human Chimpanzee Orangutan Rhesus monkey Mouse Rat Cow Dog Tenrec Opossum

exon2 | MKTTILLGLL GATMSAPLIP ..II....F. ...L...... ..II....F. ...L...... ..II....F. ...L...... ..II....F. ...L...... ..II.....I ..SS.....S ..II.....I ...S.....T .R.L....I. .......... ...I...... .......... ..II...... ...S....T. .RAA....F. .VALA...L.

exon3 QHLLSASNSN .R.M...... .R.T...... .R.M...... .R.M...... .R.......H .R.......H ...M...... .R.M...... .......... .P.......R

| exon4 | ELLLNLNNAR LRPLQ--LQG ........GQ .L........ ........GQ .L........ ........GQ .L........ ........GQ .L..R..... ........GQ .L.....F.. ........GQ .L.....F.S .........Q .......... .........Q .Q..P..F.. .......... .L.....F.. ...MG.G... ..G.PPG..A

PANSLIPPFP .L..W....S .L..W....S .L..W....S .L..W....S AF..W..... AF..W..... .F..WF.... .F..W....S .F...T...S S..P..F.L.

exon5 GILQQQQQ-T .........A .........A .........A .V.......A .F.....-.A .L......QA .........N .........A ...H.....A .A.HHG--.P

QTPGLSQFSL (80) .I........ .I........ .I........ .I...A.... .VS.RP..T. .VS.HP..P. .V....P... .I........ .V..AP.... RP--.GL-.W

Ancestral Human Chimpanzee Orangutan Rhesus monkey Mouse Rat Cow Dog Tenrec Opossum

PTLDQFAGLV SA.......L SA.......L SA.......F SA.......F S..ES....F S..ES....F S.REW..... SA..R....F .SQ.L....F .S.GH.G...

exon5 KLAQRTQAAQ SF..GA..G. SF..GA..GH SF..GA..G. SF..GA..G. G...GG..G. GF..GG..G. SF..G...G. SF..G..VG. NF..ESRPG. R....S..V.

QEPSQPQTPQ VD.L.L...P VD.L.L...P VD.L.L...P VD...A...P PDL..Q...P PDF..Q...S LD........ .D.......P LDF......L ..APRL.ML.

| QNQQDPNQMI .T.PG.SHVM .T.PG.SHVM .T.PG..HV. .T.PG..HVM .T..SASP-M .T..-ASP-M .T.RG.KNVM .T..S..HVM .I..GT.PVL ......Y.I.

exon6 PYVFSFKVPQ .......M.. .......M.. .......M.. .......M.. S..VPV.... S..VPV.... .S..-..M.. .......M.. ..S....L.. .CF...G...

| DQAQMLQYYP E.G..F.... E.G..F.... E.G..F.... E.G..FE... ..T..F.... ..T..F.... E......... E......... E.T.R.H... VWG..VP...

exon7 VYMYRPWEQP (160) ...VL..... ...LL..... ...LL..... ..VLL..... ...LL..... ...LL..... ...FL..... ...LL....S .FL.F..... .CV.GA----

Ancestral Human Chimpanzee Orangutan Rhesus monkey Mouse Rat Cow Dog Tenrec Opossum

exon7 QQTPT--QLP QQAGQQQPEE ...VP..RS. ..TR...Y.. ...VP..RS. ..TR...Y.. ...VP..RS. ..TRE..Y.. ...VP..RS. P.TR...Y.. .-.V...SS. .HT...LF.. ...V...SS. ..T...LY.. ...VA...S. P.TRE.LF.K ...AP...S. P.T....F.. L..GPST... .......F.V .DP.L....S AL..PP..P.

Ancestral Human Chimpanzee Orangutan Rhesus monkey Mouse Rat Cow Dog Tenrec Opossum

NAGIFMPSNS S..V....T. S..V....T. ...V....T. S..VL...T. ...V...TT. .V.VST..T. .....I..T. .G...V..T. G..MSR..A. Y.......YP

PNQIPLPGQA ......T.E. .....F.... .....F.... .....F.... ......SR.V .....FSR.V ....FV...V ...T.F..RV .....F...T ..EV......

| exon8 | QVPFYTQFGY IPQQAQPVIP .I...A.... ...L.E.A.S .I...A.... ...L.E.ATS .....A.... ...L.E.A.. .I...D.... ...L.E.A.. .I...N...F A.P..E.GV. .I...N...F V....E.GV. .M....E... ....VE..M. .M........ V.V.VE..M. E.......E. .....D..L. ...S.PEL.C LT......L.

exon10 PKHSTTNIFA SPTDKTITPE ..P....V.T .AV.Q..... ..P....A.T .AV.Q..... ..P....V.T .AI.R.L... ..P....V.T .AI.R...AK ..P..D.F.T .GI.P..A...PD.G.F.T .EINP..A.L Q.P...IF.T .AV.PI..R. QTP....Y.. PAI.P..... ..P.IAT..T .NI.P.MD.. L..Q.A..L. ....NVVPL.

exon9 GGQQQIAFDP LRGTAPETPA .....L.... QL.....IAV .....L.... QL.....IAV .....L...T QL..D..IAV .....L.... QL.....IAV ....HL...S FV.......G ....HLVL.S FV.......G VE...PV... FL.....IA. .....L.L.. VL......VV ..H..L.... .I......TI ....EM...L ...DV..S..

| exon11 LMEEKTNTDS LKEP .P...DK... .R.. .P...DK... .R.. .P...***** **** FP...AK..G .R.. --.Q.VK... .R.. .P.Q.V.A.. .R.. .T.K.AK... .... ...K.AK..Y .... F..A.AT... .R.. .L..EI.P.L ....

| exon10 MPTEKVIPYT QKEMINLRHP (240) .S.GEE...L ...A..F..D .S.GEE...L G..A..F..D .S.GEE...L ...V..F..D .S.GEE...L ...V..F.RD ..V.GSLL.P ...P.SFK.D ..AVEGPL.P ...P.GFKQD ..A-E.S..L ......FQ.T .VRSR....L R..V..FK.A ..AGG..THS ...RT.S... ..I.N.L... .R..V..GY.

(279) (279) (279) (279) (279) (273) (278) (277) (279) (281) (276)

Fig. 2. ODAM (APIN protein): alignment of 10 complete mammalian amino acid sequences and of the putative ancestral sequence (shown at the top). Seven sequences were inferred from DNA sequences retrieved in databases (blast search against sequenced genomes and trace archive-Whole Genome Shotgun). The sequences were checked against three published complete coding sequences: human, NM17855; rat, DQ198380, and mouse, NM27128. For further information, see legend to figure 1. * = Unknown residue.

available in reptiles, and amphibians are not representative enough of EMP evolution in these lineages (see below). Here, we use the same approach to try to identify the origins of the two newly identified genes, AMTN and ODAM, with regard to the EMPs. Ten complete coding therian [a metatherian (opossum) + nine eutherian species] sequences of both genes were retrieved from data-

bases and the literature. The inferred protein sequences were aligned using CLUSTALX and hand-checked using the sequence alignment editor Se-Al 2.0 (fig. 1, 2). The putative ancestral sequences of therian AMTN and ODAM (i.e. 190 million years old [van Rheede et al., 2006]) were calculated with PAUP 4.0 (Sinauer, Sunderland, Mass., USA), taking into account the current mammalian phylogeny [Madsen et al., 2001; Murphy et al.,

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29

2001; Delsuc et al., 2002; van Rheede, 2006] using MacCLADE 3.06 (Sinauer; fig. 1, 2). Given the small number of sequences available and the lack of sequences of representative species in some important mammalian lineages (e.g. Perissodactyla, Insectivora, Xenarthra, and prototherians: platypus or echidna), it was not possible to perform an evolutionary analysis. However, some findings from these alignments reveal some interesting points. AMTN Analysis in Mammals The amino acid sequences (ranging from 210 to 214 residues) were easily aligned without including numerous gaps (fig. 1). The presence of large conserved regions when comparing eutherian and metatherian AMTN, and the large differences with the other members of the family suggest that this gene was created long before mammalian lineage divergence, which occurred 310 MYA [Murphy et al., 2001; Hedges, 2002]. As a consequence, a functional AMTN gene might be found in reptile genomes. The putative mammalian ancestral sequence comprised 214 residues. Four residues were lost during primate evolution. Only a few residues (22%) remained unchanged during mammalian evolution (47 out of 214, fig. 1). Such relaxed selective constraints on AMTN suggest that some polymorphism could be encountered in humans. This idea is supported by the comparison of chimpanzee and human sequences: four amino acids (1.9%) were substituted within a time period of 5–7 million years, which separates the two lineages [Kumar et al., 2005]. In addition, most of the unchanged residues are dispersed through the sequence. This means that the number of conserved positions will almost certainly drop when sequences from other mammalian and reptilian species become available [Sire, unpubl. res.]. However, three important features emerge from this alignment. (i) In the N-terminal region encoded by exon 2, 55% of the residues (10 out of 18) are unchanged. This region is similarly organized as in the other SCPPs, and it is mainly composed of the signal peptide, which plays an important role in the extracellular secretion of the proteins. (ii) In positions 55–58 (exon 4), an IPLT motif is conserved, which means that this predicted O-glycosylated site could be important for the function of AMTN. Two other predicted O-glycosylated sites (threonines) are also conserved, but isolated, in exon 8. (iii) In positions 116–120 (exon 6), a SSEEL motif is well conserved. This is a putative CK2 serine phosphorylation site [Moffatt et al., 2006b]. Surprisingly, in contrast to the condition observed in EMPs, there are no con30

Cells Tissues Organs 2007;186:25–48

served residues in the C-terminal region of AMTN. It is clear that a further study, including new mammalian and reptilian sequences, is necessary to reveal further details on gene ancestry and to perform an accurate evolutionary analysis. ODAM Analysis in Mammals The amino acid sequences, which contain 273–281 residues depending on the species, were easily aligned without including numerous gaps (fig. 2). The absence of large sequence variations and the large differences compared with the other members of the family indicate that ODAM, like AMTN and the EMPs, arose before the mammalian/reptilian split. The putative ancestral mammalian sequence comprised 279 amino acids. Regarding AMTN, only a few residues (16.8%) are unchanged (47 out of 279, fig. 2), and this low selective pressure suggests that some polymorphism could occur in human ODAM (seven amino acid variations, 2.5%), are found between human and chimpanzee). Most of the conserved residues are dispersed along the sequence, but four features emerged from this alignment: (i) the N-terminal region (signal peptide) in which 47% of residues (8 out of 17) are unchanged (exon 2); (ii) in positions 25–33 (exon 3 and beginning of exon 4), a SASNSxELL motif is well conserved; this is a probable phosphorylation site; (iii) in positions 147–150 (exon 7), a YYPV motif is kept unchanged, but its function remains to be discovered, and (iv) in contrast to AMTN, four residues are conserved in the C-terminal region (exon 11). Here too, further sequences from species representative of other tetrapod lineages are needed to perform an accurate evolutionary analysis. Relationships of Ameloblast-Expressed SCPP Genes The structure and organization of the two newly identified ameloblast-expressed genes, AMTN and ODAM, were compared to the putative ancestral sequences previously calculated for the three EMP genes and SPARCL1 (fig. 3). A previous analysis of the putative ancestral sequences of EMPs had shown that: (i) AMEL exon 4 was created during eutherian evolution (it is present in some eutherian lineages only), and two additional exons 8 and 9, that are unique to the mouse and rat, were created by duplication of exons 4 and 5 [Bartlett et al., 2006a]; (ii) AMBN exons 8 and 9 have appeared in primates only, as duplications of exon 7; Sire /Davit-Béal /Delgado /Gu

Exons 2

AMEL 54 2

AMBN 54

cestral coding sequences calculated for the EMP genes (AMEL, AMBN, and ENAM), the two other ameloblast-expressed SCPP genes (ODAM and AMTN) and SPARCL1, the supposed SCPP ancestor. The reference to exon number on top of the boxes is that of the human sequences. Empty boxes indicate exons lacking in the basal mammalian taxa. The nucleotide number of each exon is indicated within the boxes (not to scale). Dark gray = Signal peptide.

5

4

48

45 4

3

4

5

42

2

3

4

ODAM 51

42

48

ENAM 54

2

3

AMTN 54

87

2

SPARCL1 54

6

7

438

3 7 8 9 10 11 12

6

5

48 117 232 39

54

45

3

Fig. 3. Gene structure of the putative an-

3

6

579

8

7

9

264 63 54 6

5

13

60 45 45

7

2865 9

8

10 11

234 48 105 48 72 162 27 4

5

6

8

9

271

8

7

66 90 36 27 3

4

5

177

1029

72

6

(iii) ENAM exon 3 can be considered as homologous to exon 2 of the other genes, and (iv) although considered the probable ancestor of SCPPs, the N-terminal organization of SPARCL1 is different from that of the EMPs, except for the first coding exon, exon 2 [Sire et al., 2005; 2006]. The structural comparison of the six putative ancestral genes, i.e. EMPs, AMTN and ODAM, and SPARCL1, confirms the previous findings that only the first three coding exons share similarities (fig. 3). As already shown for human genes, the strongest similarity of the ancestral sequences concerns exon 2 (exon 3 in ENAM), which encodes a well-conserved signal peptide and the first two residues of the protein [Kawasaki and Weiss, 2003; Kawasaki et al., 2004]. The two following exons in EMPs and ODAM are small and of roughly the same size (42–54 and 42–48 bp, respectively), with the third exon (exon 4 for ENAM) ending with an SXE phosphorylation motif. In mammals, such an organization is not observed in AMTN and in SPARCL1, which exhibit a larger third exon (87 and 177 bp, respectively), and which lack an SXE motif. The sizes of exon 3 in chicken and teleost fish SPARCL1 are small (54–57 bp), similar to the size of SPARC exon 3. This suggests that SPARCL1 originally had a small exon 3. However, in the absence of data for SPARCL1 in amphibians, crocodiles and squamates (lizards and snakes) we cannot claim that a small exon 3 was the condition when actinopterygian and sarcopterygian lineages separated. Our alignment (not shown) indicates

that the third exon in AMTN could correspond to the two short exons 3 and 4 in the other genes. The phylogenetic position of AMTN suggests that this exon could have been created by a fusion of these two short exons (see below). The mere comparison of gene organization already suggests that these genes belong to a single family [Kawasaki and Weiss, 2003]. With the exception of AMTN, the structure of which is somewhat different from the four other genes, the N-terminal region of EMPs and ODAM is similar. In addition, the organization of ODAM is more similar to that of ENAM, which suggests closer relationships of ODAM with ENAM than with the other genes (fig. 3). Since 2002, the study of EMP (and SCPP) relationships has highly benefited from gene mapping in humans, and new data have progressively accumulated in other tetrapod species (but unfortunately mainly in mammals) [http://www.ncbi.nlm.nih.gov/]. In humans, SCPP genes are located on chromosome 4, on which they form two clusters, separated by 15 Mb: the dentin and bone protein cluster (4q22, approximately 375 kb), to which SPARCL1 is adjacent, and the saliva, milk and ameloblast-secreted protein cluster (4q13, approximately 770 kb; fig. 4). The only exception is AMEL, two copies of which are found on the sex chromosomes. The most important copy, which encodes 90% of the transcripts, resides on chromosome X (fig. 4). In humans AMELX is located in antisense in the intron 1 of the ARHGAP6 gene. As AMEL belongs to the EMP family, it is clear that it was translo-

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Cells Tissues Organs 2007;186:25–48

31

SULT1E1 CSN1S1 STATH Chr. X

Chr. 4

HTN1 LOC401137

Milk, salivary, and enamel gene cluster

M I D 1

Xp22

ODAM C4orf7 CSN3 SMR3B PROL1 MUC7 AMTN AMBN EMAM IGJ

H C C S

4q13 15Mb

4q22

A 2 R

H G A P 6

NUDT9 Fig. 4. Location of the ameloblast-ex-

pressed SCPP genes and of SPARCL1 on human chromosomes. ENAM, AMBN, AMTN, ODAM, and SPARCL1 are located on chromosome 4, in two clusters separated by 15 Mb. AMEL is the only SCPP found elsewhere, on the sex chromosomes. The most important AMEL copy is on chromosome X, located in antisense within ARHGAP6 intron 2. SCPP genes are identically oriented on chromosome 4.

SPARCL1 DSPP Dentin, and bone DMP1 gene IBSP cluster MEPE (SIBLINGS) SPP1 PKD2

cated from the ‘EMP family’ chromosome to another chromosome (ARHGAP6 gene intron), either immediately after its duplication, or during a particular event, which occurred some time after a tandem duplication. ENAM, AMBN, and AMTN are adjacent genes on chromosome 4, while ODAM is located between C4orf7 (follicular dendritic cell secreted peptide) and LOC401137 (a hypothetical protein), at some distance from the three ameloblast-expressed genes, and separated from them by some salivary protein and milk casein genes (fig. 4). This syntheny is conserved in the few mammalian species for which genes are mapped [http://www.ncbi.nlm.nih.gov/ map view/]. In birds, which lost teeth approximately 80– 100 MYA [Huysseune and Sire, 1998], the SIBLING genes are found in syntheny, while the enamel, saliva, and milk 32

Cells Tissues Organs 2007;186:25–48

A M E L

M S L 3

protein gene cluster is lacking [Kawasaki and Weiss, 2006]. In amphibians (Xenopus) the syntheny is roughly conserved, but some mineralizing protein genes, known to be important in mammals, are apparently lacking. However, this absence could be related to the currently incomplete assembly of this frog genome [Kawasaki and Weiss, 2006]. The five ameloblast-expressed genes (ENAM, AMBN, AMTN, ODAM, and AMEL) were created by tandem duplication from a common ancestor [Kawasaki and Weiss, 2003, Kawasaki et al., 2004; Kawasaki and Weiss, 2006]. These duplications were probably asymmetric, i.e. after each duplication one copy kept the former function of the protein and did not diverge much from the ancestral sequence, while the other copy differentiated rapidly and Sire /Davit-Béal /Delgado /Gu

acquired new functions (neofunctionalization) [Chung et al., 2006; Steinke et al., 2006]. These functions were positively selected, but they are still to be uncovered for most of these genes. This finding is deduced from comparison of the gene structure (fig. 3) and from sequence analysis (fig. 1, 2 and Sire et al. [2006]). Indeed, the roughly conserved features of the N-terminal region suggest not only a common origin but also some functional similarities (they are all ameloblast-expressed proteins). In contrast, the rest of the sequence (the largest part) houses the specificities of each protein (i.e. its proper functions) and, therefore, is strongly divergent. The specific function of each protein could reside either in this whole sequence, as for instance for most part of the region coded by AMEL exon 6 [Sire et al., 2006] (see below), or in some particular important loci, as for instance the conserved motifs that emerge from the alignment of AMTN and ODAM mammalian sequences (fig. 1, 2). The next questions now are: how are these ameloblastexpressed genes related and which evolutionary scenario can be proposed for their origins in vertebrates?

AMEL and the Evolutionary Origin of EMP Genes

The current knowledge on the relationships and evolutionary origin of EMPs was acquired in several steps, and this study represents the last (but not least) one. This story can be briefly reconstructed as follows. In 2001, Delgado et al. showed a high sequence similarity of the 5 region (exon 2, which mainly encodes the signal peptide) of AMEL, SPARC, and SPARCL1, suggestive of a common origin of this region after duplication. Using a molecular-clock method to estimate SPARC/ SPARCL1 divergence, these authors proposed that AMEL exon 2 was created 1600 MYA (i.e. at the end of the Precambrian). This meant that AMEL could have been present before the origin of vertebrates, 530 MYA [Shu et al., 1999, 2003], and of the first evidence of mineralized elements in euconodonts, 500 MYA [Sansom et al., 1992; 1994; Janvier, 1996]. Two years later, taking advantage of the availability of the sequenced human genome and gene mapping, Kawasaki and Weiss [2003] convincingly demonstrated that (i) EMPs comprise a subfamily, (ii) EMP, milk casein, and salivary protein families together are regrouped into a cluster on chromosome 4, forming a larger family, and (iii) this family also contains the SIBLING gene cluster, which is located in another locus on the same chromosome. The SCPP family was now a fact. The Origin and Evolution of Enamel Mineralization Genes

Another chapter was added to the story when SPARCL1 was proposed to be the common ancestor of SCPP genes on the basis of its location, adjacent to the SIBLING cluster on chromosome 4, and of the structure of its Nterminal region [Kawasaki et al., 2004]. Therefore, although SPARC still remains at the origin of the mineralizing protein gene story, it was SPARCL1 that gave rise to the SCPP gene ancestor. SPARC is present in both protostomes and deuterostomes6, where it influences cell behavior and interactions with the extracellular matrix, rather than being involved in the generation of mineralized tissues. Several runs of duplications, and subsequent sub- and/or neofunctionalization have occurred and led to the current diversity of this family. Using a molecularclock method, the divergence date between SPARC and SPARCL1 was found to be inferior or equal to the current divergence date of cartilaginous fishes (estimated at 528 8 56 MYA using molecular dating [Kumar and Hedges, 1998]). This led to the conclusion that the SCPP genes probably emerged after this date [Kawasaki et al., 2004]. This dating is more recent than the 1600 MYA previously calculated by Delgado et al. [2001]. Taken together, these findings suggest that AMEL is more distantly related to SPARC and/or SPARCL1 than hitherto believed before, and that at least five duplication events took place from SPARC to AMEL [Sire et al., 2006]: SPARC ] SPARCL1 ] SCPP ancestor ] ENAM ] AMBN ] AMEL Below, we briefly review the current scenario for EMP gene relationships, which was established in the course of studies dealing with AMEL origins [Sire et al., 2005, 2006]. The previously published dataset is completed by additional information on AMTN and ODAM (fig. 1, 2), with the aim to clarify the relationships of all ameloblastsecreted SCPP proteins. The Evolutionary Origin of AMEL This study was performed in three steps: Step 1: Evolutionary Analysis of AMEL Sequences in Tetrapods A total of 80 AMEL sequences (including mammals, reptiles, and amphibians) were compiled (published se6

Protostomes and deuterostomes: the two main divisions of bilateria mostly comprising animals with bilateral symmetry and three germ layers (endoderm, mesoderm, and ectoderm).

Cells Tissues Organs 2007;186:25–48

33

AMEL 71

AMBN

56

ODAM 70

AMTN ENAM SPARCL1

Fig. 5. Phylogenetic analysis (distance analysis with maximum likelihood using neighbor-joining method) of the five ameloblastexpressed SCPP genes (AMEL, AMBN, AMTN, ODAM, and ENAM) based on the 5 region (288 bp) of their putative ancestral sequences. The ancestral sequence of SPARCL1, the probable ancestor of SCPP genes, was used to root the tree. Bootstrap values are indicated (1,000 replicates).

quences, sequences retrieved in the databases, and new sequences; see Sire et al. [2006] for the species list). The sequences were aligned as described above for AMTN and ODAM, and a putative AMEL ancestral sequence was calculated using PAUP 4.0. The conserved versus variable regions were determined and used for the next step. Step 2: Search for Sequence Similarity in Databases A PSI-blast search (National Center for Biotechnological Information) of statistically significant similar peptides was performed in GenBank [Sire et al., 2006]. The well-conserved regions of the putative ancestral AMEL were used, i.e. the N-terminal region: exon 2 (signal peptide), exon 3, exon 5, and beginning of exon 6. Sequence similarities were detected with AMBN, then with ENAM and, finally, with SPARCL1. It is noteworthy that the first non-AMEL sequence to be found using PSI-blast was crocodile AMBN, indicating that the latter is closer to ancestral AMEL than mammalian AMBN. This would mean that crocodile AMBN is more conservative of an ancestral state, and could have been subjected to a slower rate of evolution than mammalian AMBN after reptile/ mammal divergence. At this time (July 2004), neither AMTN nor ODAM sequences were available in databases [Sire et al., 2005]. Step 3: Sequence Analysis The putative ancestral sequences of AMEL, AMBN, ENAM, and SPARCL1 were calculated as described above for AMTN and ODAM. The dataset comprised AMEL 34

Cells Tissues Organs 2007;186:25–48

sequences, 30 AMBN, 28 ENAM, and 20 SPARCL1 (entire and partial sequences), and those obtained here from 10 AMTN and 10 ODAM (fig 1, 2). The N-terminal region of SPARCL1 was only used because EMPs and the other SCPPs are supposed to be derived from this region [Kawasaki et al., 2004]. The N-terminal regions of these putative ancestral sequences were aligned to the same region of AMEL (i.e. the first 62 residues, from exon 2 to the TRAP proteolytic site at the beginning of exon 6) with CLUSTALX and hand-checked using Se-Al 2.0. The phylogenetic analysis was performed using maximum likelihood (neighbor-joining method) in PAUP 4.0 and the tree was rooted on SPARCL1, since this is the probable ancestor of the SCPPs. This analysis confirms with a good statistical support the previous finding that AMEL and AMBN are sister genes [Sire et al., 2006] (fig. 5). The two newly identified ameloblast-expressed genes, ODAM and AMTN, appear as two sister genes (this is well supported statistically), and their group is the sister group of the AMEL/AMBN group. ENAM is the sister gene of the two groups AMEL/AMBN + ODAM/AMTN, and SPARCL1 is the sister gene of the three. However, the relationships of ENAM and SPARCL1 are not strongly supported by our bootstrap analysis. This phylogenetic analysis means that AMEL/AMBN and ODAM/AMTN have a common ancestor, which was probably issued from a duplication of the ENAM ancestor, itself deriving from a copy of the SPARCL1 ancestor. This phylogeny corresponds to our relatively weak knowledge of ameloblast-expressed genes and must be interpreted with caution. Indeed, even though a large number of sequences were used, most of them are from mammals, and even from eutherians only. Only a few AMEL and AMBN sequences are available in reptiles and amphibians, and no ENAM, AMTN, and ODAM sequences are known in these lineages. This lack of data in non-mammalian lineages does not allow to obtain representative putative ancestral sequences at the amniote and tetrapod levels. This means that the phylogenetic signal (i.e. gene relationships) is probably reduced by (i) the long evolutionary period (hundreds of million years) that separates each gene from its closest relative, (ii) the different evolution rate for each gene in each lineage, and (iii) the rapid divergence of some gene regions in relation to their proper functions. This phylogeny will become more accurate in the near future, when more ameloblast-expressed SCPP gene sequences will be known in reptiles and amphibians. Nevertheless, the present analysis supports AMBN/AMEL relationships and the hypothesis that both genes derive from ENAM. It furthermore indiSire /Davit-Béal /Delgado /Gu

Fig. 6. Current probable scenario for the origin and evolution of SCPP genes and, in particular, of ameloblast-expressed genes (AMEL/AMBN, AMTN/ODAM, and ENAM). Early in deuterian evolution, SPARC duplicated into SPARCL1. During successive rounds of genome and gene duplication, SPARCL1 and its descendants were copied several times on the same chromosome, giving rise to two clusters: the ameloblast-expressed/milk/saliva protein gene cluster and the bone/dentin protein gene cluster (SIBLINGs). The ENAM ancestor duplicated from an SCPP ancestor and one ENAM copy was duplicated again, giving rise to the ancestors of AMBN/AMEL and of AMTN/ODAM. After its duplication from AMBN, AMEL was translocated to another chromosome.

SPARC Vertebrate SPARC ancestor

Chr.5

SPARCL1 DSPP, DMP1, IBSP MEPE, SPP1

SIBLING ancestor SPARCL1 ancestor

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STATH, HTNs

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ENAM ancestor

AMTN AMBN AMBN ancestor AMEL

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cates that ODAM and AMTN could also be derived from ENAM. This implies that an additional duplication event has occurred between ENAM and the other ameloblastexpressed SCPP genes (fig. 6). A preliminary, schematic scenario for SCPP evolution and for the place of the ameloblast-secreted actors (to which AMTN and ODAM are now added) can be drawn, but the story is far from complete (fig. 6). In particular, the relationships between SPARCL1 and the two gene clusters (SIBLINGs and enamel-milk-saliva protein genes), and among the SIBLINGs are not established. In contrast, within the salivary SCPPs, histatins 1 and 3 derive from statherin duplication, and the latter was created from a copy of a milk casein ancestor (CSN1S2) [Kawasaki and Weiss, 2003]. The evolutionary story of salivary SCPPs is relatively recent (they are known in some eutherians only), while the origin of milk caseins is more ancient in mammalian evolution. Indeed,
-, - and -caseins are identified in the milk of metatherians (marsupials) [Ginger et al., 1999; Stasiuk et al., 2000]. Milk casein family members are also evolutionarily related and, given their structural similarity with EMP genes, the ancestral Ca-sensitive casein gene was probably derived from the duplication of an EMP [Kawasaki and Weiss, 2003], which remains to be found (fig. 6). In summary, depending on the branches of the tree, SCPP relationships are either strongly or weakly supported. Strong relationships are: SPARC/SPARCL1; STATH/ HTHs; CSN/STATH/HTHs; AMEL/AMBN, and AMEL/ AMBN/ENAM. In contrast, there are (i) no clear rela-

tionships established within the SIBLING cluster, and between this cluster and SPARCL1; (ii) no clearly identified connection between CSNs and EMPs; (iii) weak (lack of non-mammalian sequences) relationships between ODAM/AMTN, and ENAM/ODAM/AMTN, and (iv) no clear relationship between the ameloblast-expressed genes (AMEL/AMBN, ODAM/AMTN, and ENAM) and SPARCL1. Sequencing these SCPP genes in non-mammalian species [reptiles (crocodiles, lizards, and snakes) and amphibians (salamanders, caecilians, and frogs)] will help to improve our knowledge on the relationships in the family.

The Origin and Evolution of Enamel Mineralization Genes

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Dating of AMBN/AMEL Duplication Now that AMEL and AMBN are clearly established sister genes, the last questions are: was the ancestral gene AMBN or AMEL and is it possible to date this duplication event? The stronger support to AMBN ancestry is indirectly suggested by the location of AMEL on sex chromosomes. Indeed, it is difficult to imagine that an AMEL copy (that would have become AMBN) was translocated by mere chance, on the chromosome housing the other SCPP genes, and close to ENAM, their close relative. In contrast, the close location of AMBN and ENAM on the same autosomal chromosome (fig. 4) strongly supports that AMBN was created from a copy of ENAM, and, as a consequence, that AMEL originated after a duplication of the ancestral AMBN, and then translocated to another chromosome. One could argue that AMEL translocation 35

AMBN Human AMBN Mouse AMBN Crocodile AMBN Xenopus AMEL Xenopus AMEL Crocodile AMEL Mouse AMEL Human

Evolutionary distance

a

0.6

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y = 874.03x R2 = 0.7

400 350 300 250

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Fig. 7. a Linearized tree obtained from the phylogenetic analysis of AMBN and AMEL sequences in human, mouse, crocodile, and Xenopus. The calibration time used is: human/mouse: 90 MYA; human/crocodile: 310 MYA; human/Xenopus: 360 MYA [Hedges, 2002]. b Linear regression of time versus distance (y-x). Each point has two evolutionary distances of AMBN and AMEL. The duplication time of AMBN/AMEL can be estimated when we add the evolutionary distance of duplication to this linear equation, i.e. it occurred 1 600 MYA.

occurred after its duplication from the ENAM ancestor and that the copy remained close to ENAM and differentiated into AMBN. This scenario cannot be maintained since the similarities found in gene organization (fig. 3) and in amino acid pattern indicate that AMBN is closer to ENAM than AMEL is. Therefore, AMBN is the ‘mother’ of AMEL and not the opposite. In summary, all ameloblast-expressed genes are phylogenetically related, and ENAM could be the ancestor of all of them. AMEL, which codes for the major protein of the forming enamel matrix in mammals (90% of the protein content) is the youngest EMP gene. This strongly suggests that AMEL divergence after AMBN duplication was an important innovation for enamel, at least in mammals. To date, the relationships of EMP genes with SPAR36

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CL1 are difficult to establish and more data are needed to test the hypothesis of SPARCL1 ancestry. The availability of AMEL and AMBN sequences in various mammalian species, in a crocodile and in an amphibian (Xenopus) allowed to envisage a molecular dating of AMBN/AMEL duplication. A phylogenetic tree was inferred from the amino acid sequences using the neighbor-joining method (fig. 7a). From the phylogeny, it is apparent that the duplication event was much earlier than the speciation events such as the mammal/amphibian split, or the mammal/reptile split, and roughly two times of these events. To give an approximate estimate of when this duplication event occurred, we utilized the molecular dating technique developed by Gu et al. [2002], calibrated by the fossil record: primate/rodent split (around 90 MYA), mammal/reptile split (310 MYA), and amniote/amphibian split (360 MYA) [Hedges, 2002]. Our results are as follows. 1 If the amniote/amphibian split is used alone, the date of duplication (T) = 627 MYA. 2 If the mammal/reptile split is used alone: T = 896 MYA. 3 If the primate/rodent split is used alone: T = 480 MYA. 4 If all three calibrations are used: T = 682 MYA. This is a molecular dating of gene duplication, so it should be compared to other molecular date profiling [Gu et al., 2002]. Here, (2) and (3) are unreliable because the distance between human-mouse or human-crocodile differs considerably in AMBN/AMEL genes. In contrast (1) is mostly reliable and (4) takes the average, but both give similar results, i.e. AMBN/AMEL duplication occurred 1600 MYA (fig. 7b). This result confirms the previous dating of AMEL origins during the Precambrian period [Delgado et al., 2001]. A major peak of genome and gene duplication occurred around 700–500 MYA [Gu et al., 2002]. Therefore, like many developmental genes, EMPs were duplicated during this period, which preceded vertebrate diversification and skeletal mineralization. In summary, two unrelated molecular dating methods of EMP origins (SPARC/SPARCL1 divergence date: Delgado et al. [2001] and AMBN/AMEL duplication date: this study) indicate that the genes encoding them were created from several duplication rounds that have occurred before the currently accepted dates of the appearance of the first vertebrates in the fossil record (1 600 MYA). In contrast, the molecular dating of SPARC/SPARCL1 divergence proposed by Kawasaki et al. [2004] supports an emergence of EMPs after the diSire /Davit-Béal /Delgado /Gu

vergence of cartilaginous fish (approximately 500 MYA Kumar and Hedges [1998]). The knowledge of the divergence date of SPARC/SPARCL1 is of importance as SPARCL1 is considered the probable ancestor of SCPPs. However, the apparent different evolutionary rates of SPARC and SPARCL1 in various taxa, together with the fact that various gene regions were compared within each species or each clade, does not allow an accurate prediction of the divergence date. Indeed, these two paralogs share a well-conserved C-terminal region which is not easy to differentiate from one gene to the next in the vertebrate species examined. In contrast, their N-terminal region is not only extremely different but also, when comparing this region in various species, difficult to align due to a large number of sequence variations. Nevertheless, the N-terminal region of SPARCL1 is considered the probable ancestor of SCPPs. The divergence date of AMBN/AMEL seems to be more reliable because the relationships of these two genes are now well established. Also, the presence of enamel-like tissues in early vertebrates indicates that the divergence of SCPP genes might have preceded the origin of vertebrate tissue mineralization. It is important to realize the following. (i) The molecular dating of AMBN/AMEL duplication does not indicate the presence of these molecules in forming enamel, 600 MYA. After the duplication, several dozens of millions of years were probably necessary before one copy acquired its new function (new gene structure and new expression). This divergence could have occurred before, during or after the vertebrate diversification, reported to be in the Cambrian as demonstrated in the fossil record. Moreover, genetic evidence suggests that most animal phyla evolved dozens of millions of years before they started to leave behind fossil evidence, although this is debated by paleontologists. Given the lack of a temporal association between the birth of a gene (e.g. AMEL 600 MYA) and the advent of mineralized ‘teeth’ 150–100 millions of years later, the confidence in the assigned dating should be softened. (ii) Tissue mineralization could not have occurred if the necessary tools were not already present. This implies that EMPs could have had other functions before the first enamel/enameloid tissues mineralized and before EMPs were recruited for mineralization later in vertebrate evolution. This novel trait (mineralization) therefore probably evolved by employing already existing materials.

The Origin and Evolution of Enamel Mineralization Genes

Enamel/Enameloid and the Origin of EMPs

Morphological studies of enamel and enameloid in living taxa have shown that they are different in their mode of formation. The enamel organic matrix is secreted by the ameloblasts, and contains enamel-specific proteins. In contrast, enameloid organic matrix is mostly deposited by odontoblasts and contains a large amount of collagen, but the ameloblasts contribute to its formation, too [Prostak and Skobe, 1984; Sasagawa, 1984; Prostak and Skobe, 1988; Prostak et al., 1993; Sasagawa, 1995, 2002]. However, in functional teeth, the structure of both tissues is similar, i.e. highly mineralized with only a little organic matrix left (!5%). Given the same location, the same final structure and the same evolutionary origin, most authors have considered enamel and enameloid as homologous tissues. Enamel and enameloid matrices are only partially mineralized when laid down, and their final hardness is acquired during a second stage, maturation, during which the matrix is lost through the activity of proteolytic enzymes. This process creates space, allowing mineral crystal growth to eventually achieve a highly mineralized structure. Because they are highly mineralized, enamel and enameloid are easily recognizable in the fossil record and their relationships can be traced back deep in vertebrate evolution. The question of which tissue appeared first, enamel or enameloid, has been long debated and it is not clearly answered yet. It is, however, accepted that enamel progressively replaced enameloid during evolution in various lineages (e.g. in tetrapods) [Smith, 1995; Donoghue, 2002; Donoghue and Sansom, 2002; Donoghue et al., 2006]. Odontoblasts progressively reduced their production of loose collagenous matrix, which characterizes forming enameloid, while ameloblast activity increased with the secretion of large amounts of enamel-specific products at the dentin surface. This evolutionary ‘transition’ between enameloid and enamel was, in fact, probably an enameloid-dentin transition, as recently demonstrated in the ontogeny of caudate amphibians [Davit-Béal et al., 2007]. However, enamel did not replace enameloid in all vertebrate lineages. A particular type of enameloid is present in chondrichthyans (cartilaginous fish [Prostak et al., 1993; Sasagawa, 2002]), and this supports an ancient origin for this tissue, at least for the gnathostome lineage. Enamel and enameloid were certainly present in basal actinopterygians (ray-finned fish), as in polypterids and lepisosteids [Sire et al., 1987; Sire, 1990, 1994, 1995]. This supports the idea that enamel was already present in early osteichthyans, which also indicates an ancient origin. Cells Tissues Organs 2007;186:25–48

37

Fig. 8. Chordate relationships and the origin of the mineralized skeletal elements in vertebrates (adapted from

Shimeld and Holland [2000]). Chordates are deeply anchored in the Precambrian era (1700 MYA). The acquisition of a mineralized skeleton, a major event for vertebrate radiation, occurred 600–500 MYA, a period which post-dates the two genome duplications [Gu et al., 2002]. Bone and dental tissues are clearly recognized in early, jawless vertebrates, 450 MYA. Skeletal diversification in jawed vertebrates was next favored by the appearance of new genes after tandem duplication.

Enamel is absent in more derived actinopterygian taxa (teleost fish), which possess enameloid only [Sasagawa, 1984; Prostak and Skobe, 1984; Sasagawa, 1995]. The large evolutionary distance between all living representatives of these chondrichthyan and actinopterygian lineages (430–420 MYA, respectively, in the fossil record: Janvier [1996]) explains why the current structure of these enameloids is so different. Enamel and enameloid appear, therefore, to be merely grades of a hypermineralized tissue that has evolved independently in a number of vertebrate lineages [Donoghue, 2001]. The origin of these tissues can be traced back in early vertebrates, along with the appearance of a bony mineralized skeleton, one of the four main vertebrate 38

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character acquisitions, together with neural crest cells and their derivatives, neurogenic placodes, and an elaborate segmented brain (fig. 8). These vertebrate innovations appeared after the divergence between tunicates7 (Ciona) and craniates8 (recent genetic evidence indicates that tunicates could be closer to vertebrates than cephalochordates [Graham, 2004]), and probably after the divergence between craniates and vertebrates as witnessed by the fossil record. The absence of mineralized tissues in living hagfish and lampreys is probably primitive [Jan7

Tunicates: subphylum of chordates that feed by siphoning plankton through a filter. 8 Craniates: animals with skull.

Sire /Davit-Béal /Delgado /Gu

Jawed vertebrates

Jawless vertebrates

Reptiles Actinopterygian Mammals Lizards fish Chondrichthyans Amphibians Crocodiles Birds

Lampreys 0

Fig. 9. Enamel/enameloid tissues during

vertebrate evolution (as reported in the fossil record), and current knowledge of the presence of EMP and SCPP genes in vertebrate lineages. Enamel-like tissues are identified in early vertebrates, the euconodonts, and they display a different evolutionary history in the various lineages. Enameloid was conserved in chondrichthyan and actinopterygian lineages, but disappeared in amniotes. The early presence of enamel/enameloid tissues in vertebrate evolution strongly suggests that EMP divergence predates this time (1500 MYA). However, there is a large gap between this theoretical EMP presence in early vertebrate lineages and the current knowledge of the genes coding for these proteins, which is restricted to the tetrapod level (350 MYA). SCPPs are known, however, from actinopterygian fish.

Triassic 250

Euconodonts

Permian 300

Amniotes Ostracoderms

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Devonian 400 Silurian 450 Ordovician 500 Cambrian 550

Teeth: Enameloid, enamel Dermal skeleton: Enameloid, enamel Conodont apparatus: Enamel-like tissue

EMP SCPP

EMPs?

Million years

vier, 1996]. Indeed, the most ancient vertebrates discovered in the Lower Cambrian of China (530 MYA), Haikouichthys (which looks like a hagfish) and Myllokunmingia (which looks like a lamprey), possessed a skeleton composed of unmineralized cartilage only [Shu et al., 1999, 2003]. The first mineralized elements encountered in vertebrates are the tooth-like organs (conodont apparatus) composed of enamel-like and dentine tissue found in euconodonts, fossil marine vertebrates known from the Middle Cambrian (500 MYA) to the Late Triassic (230 MYA) [Sansom et al., 1992, 1994; Janvier, 1996; Donoghue, 1998, 2001] (fig. 9). These minute comb-shaped denticles are located at the entrance of the pharynx (viscerocranium). Bone appears to be absent from these elements [Donoghue, 1998]. Enamel, or enameloid, is clearly identified in the skeleton of early jawless vertebrates (e.g. pteraspidomorphs, heterostracans, thelodonts, and ‘ostracoderms’) from the Early Ordovician (480 MYA) to Late Devonian (380 MYA) periods and of jawed vertebrates (early chondrichthyans and osteichthyans) [Janvier, 1996; Donoghue et al., 2006] (fig. 8). The earliest skeleton was a dermal skeleton comprising odontodes (tooth-like elements consisting of enameloid and dentine), ornamenting dermal

plates composed of acellular bone [Sansom et al., 2005]. It is noteworthy that our current knowledge of early vertebrates reveals a gap of 30 million years between the appearance of the first vertebrates (530 MYA) and the first evidence of vertebrate mineralized elements (500 MYA). It is clear that numerous gene families expanded by gene duplication in the vertebrate stem lineage (in particular gene families encoding transcription factors and signaling molecules) [Shimeld and Holland, 2000]. The acquisition of the mineralized skeleton followed the increased genetic complexity (two genome duplications and several gene duplications) which occurred early in vertebrate evolution (during the Precambrian and Cambrian periods) [Dehal and Boore, 2005; Panopoulou and Pouska, 2005]. These large scale genomic events facilitated the evolutionary success of the vertebrate lineage and, probably, led to the diversification of several members of the SCPP family. Additional tandem duplications certainly occurred during the long period of vertebrate evolution and resulted in new gene differentiation and in a further diversification of SCPPs into new biological functions (fig. 8). The presence of enamel and enameloid tissues in early vertebrates strongly suggests that EMPs (and some other SCPPs) were present in these tissues at least 500 MYA

The Origin and Evolution of Enamel Mineralization Genes

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39

(fig. 9). This would mean that SCPPs diversified earlier. The hypothetical date of this diversification could be not so distant from the molecular dating of EMP origins (1600 MYA) if we consider that the duplication could have occurred long before the divergence of function/expression of the copies, and that vertebrates possessing a mineralized skeleton could have lived dozens of millions of years before any evidence of them in the fossil record. However, although structurally well-identified enameloid and enamel tissues are present in the teeth of chondrichthyans, actinopterygians, and basal sarcopterygians, EMP genes are known in tetrapods only (fig. 9). However, this statement relates to genes only; there is evidence from immunohistochemical studies or Southern hybridization that AMEL and/or ENAM proteins could be present in sharks [Slavkin et al., 1983; Herold et al., 1989], teleost fish [Lyngstadaas et al., 1990], polypterids [Zylberberg et al., 1997] and lungfish [Satchell et al., 2000]. Whilst the data on EMP genes (mainly in model mammals) slowly accumulated over a period of approximately 15 years, the last years witnessed a rapid increase in our knowledge, mainly because of genome sequencing in numerous species, and in particular in mammals. To date eight well-covered mammalian genomes are available and seven additional genomes are provided at a low coverage level (see http:/www.ensembl.org/). The current mammalian genome project aims to add 11 mammalian species to this list in a phylogenetic perspective (http:/ www.broad.mit.edu/mammals). Therefore, within the next few months, we will have access to at least 26 mammalian genomes and, potentially, will be able to perform evolutionary analyses of any gene in the mammalian lineage. Opposite to this large covering of mammalian phylogeny, our knowledge of non-mammalian EMPs is, unfortunately, much less advanced (fig. 10). We can see two reasons: (1) the lack of sequenced genomes and (2) the divergence of EMP sequences. The Lack of Sequenced Genomes In toothed reptiles (crocodiles, snakes, and lizards), there is still no sequenced genome available, although the reptilian (sauropsid) lineage is the lineage closest to mammals (fig. 10). However, AMEL sequences are available in a crocodile [Toyosawa et al., 1998], in a snake [Ishiyama et al., 1998], and in two lizards [Delgado et al., 2006; Wang et al., 2006], and AMBN has been sequenced in a crocodile [Shintani et al., 2002]. At present, there are no data on reptilian ENAM but, fortunately, we will soon have access to a lizard genome (Anolis carolinensis genome is being sequenced). However, sequencing a croco40

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dile genome (a representative of the lineage closest to birds) would be also extremely interesting for evolutionary analyses. In amphibians, AMEL [Toyosawa et al., 1998] and AMBN [Shintani et al., 2003] have been sequenced in the pipid frog Xenopus laevis, and an AMEL sequence is available in another frog (Rana pipiens [Wang et al., 2005a]). Moreover, sequencing of a pipid genome (Silurana tropicalis) is well advanced (fig. 10). Surprisingly, although as expected AMEL and AMBN are present in this genome, to our knowledge ENAM has not been found yet [Kawasaki and Weiss, 2006]. It is questionable whether this EMP is really absent from this frog genome. Indeed, on the one hand our evolutionary analysis indicates that ENAM is the oldest representative of the EMP family and, on the other hand, ENAM plays important roles in enamel structure and organization as illustrated by AIH2 resulting from ENAM mutations. It is also clear that pipids have a well-formed enamel [Sato et al., 1986]. Therefore, this ‘lack’ is probably related to the fact that the pipid genome is still not entirely (or correctly) assembled. One should also take into consideration that pipids are highly derived anurans and, as a consequence, EMPs could be divergent compared to more basal amphibian species. Sequencing another frog, salamander/newt or caecilian genome would be, therefore, highly informative for evolutionary analysis. No EMP is known in basal sarcopterygians, i.e. lungfish and coelacanth, nor in basal actinopterygians (polypterids and lepisosteids), and there is no sequenced genome available nor sequencing project running. However, these taxa possess enamel and they belong to lineages that are crucial to improve our understanding of EMP relationships and evolution. In contrast to this lack of data, the genome has been sequenced in four teleost species, and several SCPPs were identified. However, teleosts are derived actinopterygian lineages, and the long evolutionary distance (1 420 million years) between actinopterygians and tetrapods explains the difficulty encountered when trying to identify homology between teleost and tetrapod SCPP genes [Kawasaki et al., 2005]. For instance, no EMP gene can be related to these SCPPs. No SCPP is known in chondrichthyans (sharks and rays). Here too, the long evolutionary distance (1 430 million years) between cartilaginous fish and tetrapods could lead to problems when trying to identify homologous genes, but the syntheny conservation of SCPP genes could help [Kawasaki et al., 2005; Kawasaki and Weiss, 2006].

Sire /Davit-Béal /Delgado /Gu

Fig. 10. Current knowledge of EMP genes in vertebrates. To date only two EMPs are characterized at the tetrapod level (AMBN and AMEL). ENAM is only known in mammals. The lack of data in non-mammalian lineages is clearly related to the absence of sequenced genomes. SCPP genes are identified in teleost fish, but the large evolutionary distance makes their relationships to EMPs uncertain. EMP genes on gray background are potentially accessible to sequencing. Question marks indicate lineages in which sequencing of EMP genes might be a priority to improve our understanding on their origin and evolution. * = Large DNA regions (Whole Genome Shotgun) have been sequenced in a lizard (A. carolinensis).

The Divergence of EMP Sequences The difficulty to find EMP (and other SCPP) genes using PCR or RT-PCR resides in their variability. Indeed, except for the short N-terminal region that is relatively well conserved in each member of the family, the largest part of the sequence is variable. For instance, although they probably conserve their main function, most of the mammalian AMEL exon 6 sequences (the largest part of AMEL) cannot be accurately aligned with the homologous region in reptiles and amphibians due to numerous substitutions and indels [Sire et al., 2006]. These highly variable sequences indicate that SCPPs are intrinsically disordered proteins [Dunker et al., 2001; Kawasaki et al., 2005] and there are only a few conserved residues. Therefore, the only means to find EMPs in evolutionary distant species, such as basal sarcopterygians or actinopterygians, is to study sequenced genomes or sequences of large DNA regions suspected to house these genes. For example, in a teleost fish (fugu), several SCPP genes were identified in a DNA region corresponding to the SIBLING cluster in mammals, meaning that the syntheny of the SIBLING cluster is conserved between fish and tetrapods [Kawasaki et al., 2005; Kawasaki and Weiss, 2006]. These SCPP genes were found not based on their similarity with known SCPP sequences but because they are located adjacent to SPARCL1, and because they share some structural features with tetrapod SCPPs. Fish SCPP genes are

so different from tetrapod SIBLINGs that no homology could be recognized. Fish SCPP genes are expressed during tooth formation [Kawasaki et al., 2005] but one can wonder whether they play the same function as EMPs. Moreover, SIBLINGs (DSPP, DMP1, IBSP, and SPP1) are known to be expressed during tooth matrix formation in tetrapods [Fisher and Fedarko, 2003; Qin et al., 2004]. EMP genes could also be conserved in other regions of the teleost fish genome, but they remain to be discovered. Indeed, morphological studies strongly support that EMPs are present in the enamel-like tissue (ganoine) of basal actinopterygian lineages, polypterids and lepisosteids [Sire et al., 1987; Sire, 1994; 1995]. To date the information available for the three EMP genes largely relates to mammals and the few sequences available (or planned to be so) in other tetrapods are not sufficient to perform an evolutionary analysis at this level (fig. 10).

The Origin and Evolution of Enamel Mineralization Genes

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What Can the Evolutionary Analysis of EMP Genes Tell Us? The Case of AMEL

AMEL Evolution AMEL is the main component of forming enamel and it plays crucial roles in enamel structure and mineralization [Diekwisch et al., 1993; reviews in Bartlett et al., 41

2006b; Margolis et al., 2006]. Mutations of the encoding gene lead to AIH1 [Hart et al., 2002; Kim et al., 2004]. Given this importance it is not surprising that AMEL is the best-known EMP. Over the past years, AMEL studies on model animals have provided information on the gene structure and supposed functions of the various regions of the protein [Fincham et al., 1991; Fincham and Moradian-Oldak, 1995; Greene et al., 2002]. AMEL is subject to posttranslational modifications [Fincham and Moradian-Oldak, 1993] and it self-assembles to form nanospheres that are involved in enamel mineralization [Wen et al., 2001; Snead, 2003; Du et al., 2005; Veis, 2005]. The N- and C-terminal regions interact with mineral [Aoba et al., 1989; Aoba, 1996; Hoang et al., 2002; Paine et al., 2003; Snead, 2003] and are involved in adhesion with the ameloblast surface through membrane proteins (e.g. Cd63, annexin A2, and Lamp1 [Wang et al., 2005b; Tompkins et al., 2006]). AMEL interacts also with some keratins in ameloblasts through ligand-binding properties located in the N-terminal region [Ravindranath et al., 1999, 2000, 2001, 2003]. Some splice products have been proposed to be signaling molecules [Veis et al., 2000; Veis, 2003]. From these studies, increasing evidence accumulates to support the idea that the N-terminal, and to a lesser degree the C-terminal, regions are the most important regions for proper AMEL function. This importance is also revealed by several AIH1, caused by mutations modifying the functioning of these regions. The question of a possible role for the central variable region (encoded by most of exon 6) is completely ignored. Is it useless? Certainly not. Evolutionary analyses indicate that this core region of the protein, although intrinsically disordered, could be responsible for the well-ordered microstructure of enamel [Delgado et al., 2005; Sire et al., 2005; 2006]. More data are still needed to understand the relationships between structure and function of this region and, more generally, to reveal the amino acid positions and regions that could play an essential role. As an alternative to biochemical and in vitro approaches, an evolutionary analysis of mammalian AMEL was performed using 56 sequences constituting a dataset representative of mammalian diversity [Delgado et al., 2005]. Here, we summarize and complete these results in proposing two alignments (fig. 11): one, illustrated with 20 sequences of the N- and C-terminal regions only, reveals the numerous well-conserved residues that are important for the proper function of the protein (interactions with the cell membrane and/or with mineral crystals). The other alignment, comprising 51 sequences, is 42

Cells Tissues Organs 2007;186:25–48

centered in the variable central region of exon 6, which houses, in mammals, a hot spot of mutation. The putative ancestral sequence has been calculated for both alignments. Briefly, this evolutionary analysis reveals the following points. (i) A total of 56 residues (out of 74 in the full-length sequence) have remained unchanged in the N- and C-terminal regions of AMEL during mammalian evolution, i.e. during 225 million years [van Rheede et al., 2006] (fig. 11a). This indicates that strong functional constraints act on these amino acids, meaning that they certainly play, either alone or with other conserved residues, an important role. Most variants are found in the C-terminal region of exon 5. (ii) The hot spot of mutation (large insertions/deletions of residues) has appeared recently in mammals, and independently in several lineages (fig. 11b). Insertions are found in basal primates (lemurs), in tree shrews, in basal rodents (squirrel and guinea pig), in bovids (cow and goat) and cervids (deer), in only one family of carnivores (ursids), in bats (Macrochiroptera), in insectivores (hedgehog), in afrotherians (elephant shrew), and in marsupials (opossums). The perissodactyls (e.g. horse) and prototherians (platypus and echidna) are the only important lineages in which such large insertions are absent. These insertions contain a variable number of three amino acid (triplet) repeats (e.g. PIQ-PMQ-PLQ). These triplet repeats range from two (in the tree shrew) to 12 (in a fruit bat), in which a total of 36 residues (108 bp) are inserted. Within some lineages, e.g. bovids, the number of repeats can vary in closely related species (8 repeats in the African buffalo, 7 in cattle, and 5 in the other members of the family). It is noteworthy that AMELY, that is expressed at a low level in forming enamel (less than 10% [Salido et al., 1992]), does not show insertions in this region. This illustrates the separate evolution of the two AMEL copies on sex chromosomes [Girondot and Sire, 1998], AMELY being subjected to the particular mode of evolution of the Y chromosome [Iwase et al., 2001; Lahn et al., 2001; Iwase et al., 2003]. The lack of triplet insertions in AMELY versus AMELX exon 6 allows to easily discriminate males from females in lineages possessing the hot spot of mutation, e.g. bovids [Weikard et al., 2006] and ursids [Yamamoto et al., 2002]. Large deletions (69 residues) are found in dolphin, Weddell seal, panda and roundleaf bat (Microchiroptera). However, we do not know whether these indels have a consequence on enamel microstructure in these species [Delgado et al., 2005]. It is clear, however, that the conservation of such large indels during evolution has no negative results on enamel function as protective tissue. Sire /Davit-Béal /Delgado /Gu

AMEL_Ancestral Human Squirrel_monkey Lemur Galago Mouse Guinea_pig Squirrel Goat Cow Pig Horse Dog Flying_fox Hedgehog Elephant Tenrec Hyrax Opossum Wallaby Platypus

exon2 | MGTWILLACL LGAAFAMPLP .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... ..T....... .......... .......... .......... .....S.... .......... .....S.... .......... .....S.... ........S. ......I... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .R....L... ......I... .R....L... ......I... .......T.. I.....I...

exon3 PHPGHPGYIN .......... .......... ......A... .......... ....S..... .......... .......... .......... .......... .......... S......... .......... .......... .......... .......... .......... .......... .......... .......F.. ...A......

| exon5 | FSYEVLTPLK WYQNMIRQQY .......... ...S-..PP. .......... ...S...PA. .......... ...S...PP. ........V. ...S.L.PP. L......... ...S....P. .......... ...S....P. .....I..F. ........P. .....P.... ...S...HP. .......... ...S...HP. .......... .......HP. .......... ...SL...P. .......... .......HP. ....****** *********. .......... .......PP. .......... ........P. .......... .....L..P. ........I. .......-P. .......... ...S.M.HE. .......... ...S.M.-.. .........Q .....K....

exon6 -----exon6 | ex7 PSYGYEPMGG WLHHQIIPVL DLPLEAWPAT DKTKREEVD .......... .......... ..T.....S. ......... .......... .......... .......... --------.......... .......... .......... --------.......... .......... .......... --------.......... .......... E......... ......... .......... .V...V.... .......... ......... .......... .......... .......... ......... .......... .........V .VL..D.... ......... .......... .........V .......... ......... T......... .........V .......... ......... T......... .......... .......... ......... .......... .......... .......... ......... .......... .......... .......... ......... .......... .......... .......... ......... .......... .......... .......... ......... .......... .......... ...M...... ......... ........S. .......... .......... ......... .......... .......... .M--...... ......... .......... .......... ********** ********* .......... .......... .....Q.... ......... a

AMEL_Ancestral Human Orangutan Squirrel_monkey Lemur Galago Marmoset Tree_shrew Flying_lemur Mouse Hamster Guinea_pig Squirrel Goat Sheep Cow African_buffalo Japanese_serow Deer Pig Hippopotamus Dolphin Porpoise Horse Tapir Rhinoceros Wolverine River_otter Dog Arctic_fox Gray_seal Weddell_seal Canada_lynx Tiger Brown_bear Panda Flying_fox Fruit_bat Roundleaf_bat Hedgehog Shrew Armadillo Elephant Manatee Tenrec Golden_mole Elephant_shrew Hyrax Opossum Aquatic_opossum Platypus Echidna

PNLPQPAQQP ....P..... ...LP..... ....P..... ....P..... ....P..... ....P..... ....P..... S.I.M...P. ..I.PS.... ..I.PS.... ....PTS... ....P..... ....L..... *******... ....L..... ....L..... ....L..... ********.. ....L..... ..F.L..... .H..V......F.V..... ....P.V... .HF.P..... ....P.V... ******.... ....L..... ....L..... *****..... ....L..... ....L..... *****..... AT..L..... ....L..... ....L..... ...LP..... ...LP..... T..LP..... S...A..... ..V.P..... ..V.P.V... ....P.I... ....P.I... .H..P.V... AH..P-V... .H..P.V... ....P.I... ......G... ......GH.. S......... S.....G...

Q-PQ--PHQP .......... .......... .......... .......... .......... .......... .......... ..L....... P.....S... S.....S... ......H... ......S... ......H... ..S....... .......... .......... .......... .......... .......... .........H ........-. .......... .......... .......... .......... ......A... ......A... .......... .......... .......... .......... .......... .......... .......... ........H. .......... -......... .......... .......... .......... .......... .......Q.. .......... .......... ..Q....... .......... .......... .Q........ .Q..PQ.... ..QP...... ..KP..T.R.

IQ-------M......... M......... M......... M.PMQPMQPM M.PMQPMQPM M......... ..PIQPIQ.. M......... M......... M......... ..PIQPIQPI M.PMQPMQPM L.PLQPMQPL L.PLQPLQPL L.PHQPLQPM L.PHQPLQPM L.PLQPMQPL L.PLQPLQPL L......... L......... --........ M......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... ..PIQPIQPI .......... ..PIQPIQPI ..PQQPVHPI .......... M.PMAPMQPM .......... .......... M......... .......... MP........ MH........ M.PMQPMHPM L......... ..PIQPIQPI ..PIQPIQPI .......... ..........

---------.......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... PLQ....... PLQPLQ.... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... PIQPIQPMQP .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... PMQPMQPMQ. PLQPMQPMQP .......... ..........

YQ----PQPP .........V .........V F.......TV F........V F........V F.......TV F........V .........V F.QPFQ..AI F.QPFQ...I F.QPFPT..V F.QPFQ..SI F.......SI F.......SI F.......SI F.......SI F.......SI F.......SI F........V F........I --....--.V .........V FH.......V F........V F........V F........V F........V F........V F........V F........V F........V F........V F........I F........V F........V F.......HV F.....---F........V .........V F........A F........A .........V .........V .........V F.PIQ.H..V ........SV .........V .........A .........A F.......V. F.......F.

Fig. 11. Alignment of AMEL amino acid sequences in representative mammals. a Well-conserved N- and C-terminal regions in 20

species. Exon 4 is not represented because it is lacking in several species. Partial sequences were removed from this alignment. Vertical bars indicate the limits between exons. Unchanged residues are shown on a gray background. b Central region of exon 6

The Origin and Evolution of Enamel Mineralization Genes

---------.......... .......... .......... QPIQPIQPIQ QPIQ...... .......... .......... .......... .......... .......... QPIQPIQ... QPMQPVQ... QPLQPLQ... QPLQPLQ... QPMQPLQPLQ QPMQPLQPLQ QPLQPLQ... QPLQPLQ... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... Q......... .......... QPIQPIQPIQ QPQSPVHSMQ .......... Q......... .......... .......... .......... .......... .......... .......... H......... .......... QPIQPMQPMQ QPMQPMQPMQ .......... ..........

--------PQ .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .........K ........-.......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... .......... ........-MQPMQPIQ.. .........L ........-.......... .......... .......... .......... ........Q. .......... .......... .......... .......... .......... MQ........ .......... ..........

APVHPMQPLP P......... P......... P......... P.L....... P......... S......... T......... P.....H... S.L......A S.L......A S.L..I.... P.L.SLH... P....I.... S....I.... P....I.... P....I.... P....I.... P....I.... S.M..I...L P.M..I...L -------..L -....I.... P.L..I.... P....I.... P....IH... P.M..I.... P....I.... P....I.... P.M..I...L P....I.... P--------S.M..I...L S.M..I...L P.M..I.... -------... P....I.... S....I.... -------... S.M.---... P.M..I.... P.M..I.--P....I.... P....I.... P......... P.M....... QSM....... P.M..I.... T...AVL... T...AVL... ..A...P.M......P.M-

PQ-PPLPPMF .......... .......... .......... ....H...L. ........L. .......... .......... ........L. ........L. ........L. ...QA..... ....H..... ......L.I. ........I. ........I. ........I. ........I. ........I. .......... ......S... .......... .......... ........I. ........I. ........I. .......... .......... .......... .E........ .......... .......... .......... .......... .......... .......... ........L. ........L. .......... ......H.I. Q......... --......V. .......... .......... .......... ......S... .......... ........I. .......... .......... .......... ....Q.....

b

in 51 species emphasizing the region considered a hot spot of mutation. This region is characterized by amino acid triplet insertions or deletions. Identical sequences were not included in these alignments: e.g. human = chimpanzee and rhesus monkey; mouse = rat; cow = European bison. · · · · · = Identical residue; – – – = indel; * = unknown residue.

Cells Tissues Organs 2007;186:25–48

43

Fig. 12. Amino acid sequence of human amelogenin highlighting the residues which remained unchanged during the 225 million years of mammalian diversification. The importance of amino acids is inferred from the alignment of 60 mammalian sequences representative of the main lineages, as partially shown in figure 11. Exon 4 (14 residues) was not included because it is missing in several species studied. Signal peptide is on gray background. The protein sequence (191 amino acids) is numbered from methionine (1). Bold characters (n = 75) indicate residues unchanged in mammals, italics (n = 35) residues that can be substituted by an amino a{tb}cid from the same group only, small roman characters residues that can be substituted, characters on gray background (n = 5) residues that are known so far to lead to amelogenesis imperfecta when substituted, and underlined characters indicate (n = 31) residues that are unchanged in amniotes (mammals and reptiles) [Delgado et al., in press].

(iii) Although this central region of AMEL exon 6 is variable, it maintains its richness in proline (30%) and glutamine (20%) in all sequences studied. This means that this region is also subject to a functional constraint but that this selective pressure probably acts on the general conservation of the P and Q richness rather than on specific amino acid positions. This strongly suggests that this region could be subject to polymorphism in humans. (iv) The origin of the largest of AMEL exon 6 has to be found in the repeats of nine nucleotides coding for three residues (triplets) PXQ or PXX [Delgado et al., 2005]. These repeats have not been blurred by substitutions during at least 310 million years of amniote evolution, because such triplet repeats have been identified in crocodile AMEL [Sire et al., 2006]. The triplet insertions found in the hot spot mutation in mammals are probably reminiscent of this mechanism. These repeats are to be found, probably, in the origin of AMEL after AMBN duplication, and also constitute the originality of AMEL compared to the other EMPs and to ameloblast-secreted SCPPs in general. This leads to the hypothesis that AMEL divergence consisted of the loss of most of the C-terminal region of the AMBN ancestor and of the development of exon 6 (probably from AMBN exon 5) through several runs of PXQ triplet repeats. This new protein was posi44

Cells Tissues Organs 2007;186:25–48

tively selected during enamel evolution in vertebrates because this hydrophobic region, rich in P and Q, improved the resistance of enamel to wear and microbreaks. This could explain why today AMEL represents 90% of the forming enamel matrix in mammals. Validation of Mutations and Important Residues The evolutionary analysis of AMEL in mammals reveals 170 residues (out of 191) that are certainly important for a correct function of AMEL because they have remained unchanged during 225 million years of evolution (fig. 12). The number of conserved residues is reduced to 34 when reptilian AMELs are added to this analysis [Delgado et al., in press]. These 34 positions conserved during 310 million years of amniote evolution are considered crucial residues for enamel formation. All of them are located in the N- and C-terminal regions of AMEL, known to play an important role in relation with the environment (interactions with the ameloblast surface and/or with the mineral crystals). The residues conserved only in mammals could indicate that they play new, important roles for enamel formation in this lineage. As a consequence of their long-lasting conservation, substitution of the important amino acids revealed in this study could result in enamel defects (AIH1) when substiSire /Davit-Béal /Delgado /Gu

tuted in humans (fig. 12). The five substitutions leading to AIH1 are validated when using the mammalian, and four of them when using the amniote dataset. Therefore, this list of conserved residues in the human AMEL sequence (fig. 12) can be useful for the clinical diagnosis of AIH1 since it helps to validate any human AMEL mutation, which could be suspected for AIH1.

Conclusion

Although the origin of enamel can be traced back to early vertebrates, at least 500 MYA in the fossil record, our knowledge of enamel mineralization genes is still restricted to the tetrapod level (350 MYA) for AMEL and AMBN, and to the mammalian level (225 MYA) for ENAM. The difficulty encountered when looking for EMP genes in the vertebrate lineages that diverged earlier in evolution (i.e. chondrichthyans, 430 MYA, and actinopterygians, 420 MYA) resides in their high sequence variations (intrinsically disordered proteins) and in the lack of sequenced genomes in basal lineages such as lungfish, polypterids and sharks, which do not allow looking for EMP genes using syntheny. Our approach using putative ancestral sequences could help to obtain data in closely related but not in evolutionary distant lineages. Molecular dating of AMBN/AMEL duplication indicates that EMP genes probably appeared at the end of the Precambrian era (1600 MYA) after several rounds of genome/gene duplications that took place in this period. ENAM was created first, then AMBN and AMEL. After AMBN duplication, one copy lost a large part of the ancestral 3 region and accumulated PXQ repeats. These events gave rise to a new protein: AMEL. AMEL was then positively selected (and constrained), probably because it

improved enamel microstructure and thickness: it is now the major protein forming enamel in amniotes. The AMEL story is relatively well established now, but some details will be undoubtedly added when the evolutionary analyses in amphibians and reptiles will be achieved. Such a study will probably open the door to access the AMEL sequence in lungfish, the sister group to tetrapods. In contrast to our knowledge on AMEL, the other ameloblast-secreted SCPP proteins (AMBN, ENAM and the newly identified AMTN and ODAM) are poorly known. Efforts have to be made towards better knowledge of the relationships and evolution of these proteins, and the current genome sequencing programs will certainly be of great value in this quest. It is clear that evolutionary analyses are necessary not only for thorough knowledge of each protein (i.e. its origin, relationships, and mode of evolution) but also because they provide insights into residues that play important roles for the correct function of the protein. In addition, as illustrated with AMEL, sequence datasets obtained in a phylogenetic perspective will be helpful to validate mutations responsible for genetic diseases in humans.

Acknowledgments We are grateful to Ann Huysseune (Ghent University, Belgium), J. Hu and J.P. Simmer (University of Michigan School of Dentistry, Ann Arbor, Mich., USA), N. Takahata (Graduate University for Advanced Studies, Kanagawa, Japan), and K. Kawasaki (Pennsylvania State University, University Park, Pa., USA) for helpful remarks and suggestions. We thank J.P. Simmer for his kind invitation for J.Y.S. to present this review to the 2006 Symposium of the International Association for Dental Research in Brisbane, Australia.

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The Origin and Evolution of Enamel Mineralization Genes

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Cells Tissues Organs 2007;186:49–59 DOI: 10.1159/000102680

Evolutionary History of Sex-Linked Mammalian Amelogenin Genes Mineyo Iwase Satoko Kaneko Hielim Kim Yoko Satta Naoyuki Takahata Department of Biosystems Science, Graduate University for Advanced Studies, Hayama, Japan

Key Words Amelogenin
Evolution
Mammalian sex chromosomes
Polymorphism
Pseudoautosomal boundary

Abstract Amelogenin (AMEL) arose prior to the emergence of tetrapods and transposed into an intron of the Rho GTPase-activating protein 6 gene. In the mammalian lineage leading to eutherians, a pair of homologous autosomes with this nested gene structure fused with the then already differentiating sex chromosomes by suppressing homologous recombination. As sex-chromosomal differentiation extended to the fused region, a pair of homologous AMEL genes too differentiated from each other in two steps; first in the 5 region (the promoter region to transposon MER5 in intron 2) and second in the remaining 3 region. This resulted in gametologous AMELX and AMELY in the eutherian sex chromosomes. Although the early differentiation of the 5 region between AMELX and AMELY is consistent with the lowered expression level of AMELY, there is no indication for deterioration of AMELY at the amino acid level. Rather, both AMELX and AMELY in particular lineages might undergo positive selection, followed by negative selection to preserve established function. Based on patterns and levels of AMELX and AMELY polymorphisms in the human population, it is also argued that a recombination cold spot near AMELX might be related to the cause of the ancient pseudoautosomal boundary. Copyright © 2007 S. Karger AG, Basel

M.I. and S.K. contributed equally to this work.

© 2007 S. Karger AG, Basel 1422–6405/07/1861–0049$23.50/0 Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/cto

Introduction

Amelogenin (AMEL) is a structural enamel protein and plays a major role in amelogenesis (dental enamel formation) by controlling crystallization of calcium hydroxyapatite. In eutherians, AMEL comprises more than 90% of the total enamel extracellular matrix proteins that contain enamelin (ENAM) and ameloblastin (AMBN) as well [Hu et al., 2005]. It was found that the AMEL gene belongs to the secretory calcium-binding phosphoprotein (SCPP) family [Kawasaki and Weiss, 2003]. Except

Abbreviations used in this paper

AMBN AMEL AMELX AMELY ARHGAP6 CDY ENAM LD MER5 MYA PAR RBMY SAPARCL1 SCPP SNP SPARC SRY VCY

ameloblastin amelogenin X-linked amelogenin Y-linked amelogenin Rho GTPase-activating protein 6 chromodomain Y enamelin linkage disequilibrium medium reiterated frequency repeat 5 million years ago pseudoautosomal region RNA-binding motif Y SPARC-like 1 secretory calcium-binding phosphoprotein single nucleotide polymorphism secreted protein, acidic, cysteine rich sex-determining region Y variable charge Y

Dr. Naoyuki Takahata Department of Biosystems Science Graduate University for Advanced Studies (Sokendai) Hayama, Kanagawa 240-0193 (Japan) Tel. +81 46 858 1502, Fax +81 46 858 1542, E-Mail [email protected]

for AMEL, all SCPP genes are clustered in tandem on chromosome 4 in humans and chicken. The SCPP genes are thought to have originated from domain I of secreted protein, acidic, cysteine-rich (SPARC)-like 1 (SPARCL1) and this gene was in turn descended from the entire SPARC prior to the emergence of teleosts [Kawasaki and Weiss, 2003, 2006]. The three enamel protein genes likely arose after the divergence between actinopterygians (ray-finned fish such as zebrafish and fugu) and sarcopterygians (lobe-finned fish such as coelacanth and lungfish), but before the emergence of tetrapods [Kawasaki et al., 2004]. At present, there is no information about the presence or absence of AMEL in lobe-finned fish. Like AMBN, AMEL is found only in amphibians, reptiles and mammals [Ishiyama et al., 1998; Toyosawa et al., 1998; Delgado et al., 2005; Sire et al., 2005]. However, unlike ENAM and AMBN both of which are located within the SCPP gene cluster, any tetrapod AMEL gene thus far identified resides in the intron
40-kb upstream from exon 2 of the Rho GTPase-activating protein 6 gene (ARHGAP6). The evolutionary history of tetrapod AMEL has been eventful since the origin. For instance, the absence of chicken AMEL within or adjacent to ARHGAP6 suggested that AMEL has disappeared from the chicken genome [Kawasaki et al., 2004; Kawasaki and Weiss, 2006]. The same may apply to AMEL in turtles that have been toothless for 1100 million years [Toyosawa et al., 1998]. AMEL was initially autosomally coded. However, in the ancestral lineage of eutherian mammals, a pair of homologous autosomes encoding AMEL translocated and became the short arms of the sex chromosomes, 1100 but !180 million years ago (MYA) [Graves, 2002]. Subsequently, in this newly added region of the sex chromosomes, homologous recombination was inhibited and X-Y chromosomal differentiation took place in a stepwise fashion, one stratum at a time [Lahn and Page, 1999; Skaletsky et al., 2003]. Such stepwise differentiation of the eutherian sex chromosomes has left an ancient pseudoautosomal boundary within the second intron of AMEL [Iwase et al., 2001, 2003]. It is speculated that the Y-linked amelogenin gene (AMELY), although containing no detrimental mutations, is not under strong functional constraint. It appears that the expression of AMELY is too low to recover the function of the more active X-linked homolog (AMELX) if this gene is inactivated by mutation, as in recessive Xlinked amelogenesis imperfecta [Lagerström et al., 1990; Toyosawa et al., 1998]. Indeed, without any obvious effect, AMELY has been lost in rodents [Lau et al., 1989; Nakahori et al., 1991], and the locus is polymorphic with re50

Cells Tissues Organs 2007;186:49–59

spect to partial deletion in some human populations [Chang et al., 2007]. However, it is also speculated that AMELY could benefit males through a delay in primary tooth development and weaning [Lahn et al., 2001; but see Yamamoto et al., 2002; Delgado et al., 2005]. Functional importance of AMELY thus remains to be elucidated. In this study, we first review the origin and subsequent evolution of tetrapod AMEL genes. For eutherian AMEL genes, special attention is paid to the intimate relationship between the sex-chromosomal differentiation and the divergence between gametologous AMELX and AMELY genes. Second, carrying out molecular phylogenetic analyses, we examine roles of natural selection for and against eutherian AMELX and AMELY genes. Third, we determine the genomic DNA sequences of AMELX and AMELY for samples of humans with different ethnic backgrounds and examine patterns and levels of polymorphism. We also examine linkage disequilibrium (LD) or lowered recombination frequency surrounding the human AMELX locus based on our own sequence data and the public HapMap data.

Material and Methods Genomic DNA Sources of Human AMELX/AMELY and PCR Amplification Human genomic DNAs used for AMELX were taken from 32 unrelated individuals (19 males and 13 females) in a worldwide sample in Coriell Cell Repositories (http://locus.umdnj.edu/ccr/): 12 Africans (6 Biaka Pigmies, 4 Mbuti Pygmies, and 2 Blacks), 8 Eastern Europeans (4 Adygeis and 4 Russians), 4 Middle Easterns (Druze), 4 Asians (Ami), and 4 Amerinds (2 Mayan and 2 Karitiana). The repository numbers are NA10469–10473, 10492– 10496, 10968, 10969, 10975, 10976, 11521–11524, 13607–13610, 13617–13620, 13820, 13838, 13849, 13877, 14537, and 14661. Those DNAs were used as templates for PCR amplification of AMELX (6513 bp). The PCR primers were designed within the promoter region and at the end of exon 6. The upper and lower primer sequences are 5-GCTGCTGAGAACATGGAA-3 and 5CACTTCCTCCCGCTTGGTCTTGTC-3, respectively. Similarly, 18 unrelated males were used for AMELY: 5 Africans (1 Biaka Pigmy, 3 Mbuti Pygmies, and 1 Black), 4 Eastern Europeans (2 Adygeis and 2 Russians), 1 Middle Eastern (Druze), 4 Asians (Ami), and 4 Amerinds (2 Mayan and 2 Karitiana). The repository numbers are NA 10470, 10492, 10494, 10495, 10968, 10969, 10975, 10976, 11522, 13607–13610, 13619, 13620, 13820, 13838, and 14537. The PCR primers of genomic AMELY sequences (6627 bp) were designed within the promoter region and at the end of exon 5. Unfortunately, exon 6 had to be excluded for technical reasons. The upper and lower primer sequences are 5-AAGTGCCATGTGGTGAATTAGG-3 and 5-GTACCAGAGCATGATAAGA-3, respectively. Each PCR reaction mixture was 50
l in volume and contained 15 ng of genomic DNA, Platinum PCR SuperMix High

Iwase/Kaneko/Kim/Satta/Takahata

ARHGAP6 exons 2–13

ARHGAP6 exon 1

Human (X chromosome) AMEL Opossum (chromosome 7) Deletion of exon 1 Chicken (chromosome 1)

Fig. 1. The presence or absence of the

AMEL gene within the Rho GTPase-activating protein 6 gene (ARHGAP6) in humans, opossums, chicken, Xenopus tropicalis and Tetraodon nigroviridis. In chicken, it appears that exon 1 and the following intron of ARHGAP6 are deleted together with AMEL.

Xenopus (chromosome: unknown)

Fugu (chromosome 3)

Fidelity (Invitrogen, San Diego, Calif., USA), and 0.2
M of each primer. The PCR condition was as follows: denaturation at 94 ° C for 2 min; 34 cycles at 94 ° C for 15 s, 59 ° C for 30 s, and 68 ° C for 5 min. The PCR products were purified using ExoSAP-IT (United States Biochemical, Cleveland, Ohio, USA) and sequenced directly. In the case of heterozygous AMELX, PCR products were cloned into pCR-XL-TOPO with TOPO XL PCR Cloning Kit (Invitrogen). Sequencing reactions were performed with BigDye
Terminator v1.1 and v3.1 Cycle Sequencing Kits (Applied Biosystems, Foster City, Calif., USA) and analyzed on ABI PRISM 377, 3100, and 3730 DNA sequencer (Applied Biosystems). To avoid sequencing errors, PCR products or plasmid DNAs were read twice in both directions. Sequences were also confirmed by independent PCRs. These fragmental sequences were assembled by DNASIS (Hitachi, Yokohama, Japan).

100 kb

the average pairwise nucleotide differences  [Nei and Li, 1979],  [Watterson, 1975], the normalized  –  that is defined as D [Tajima, 1989], and linkage disequilibrium that is defined as r 2 [Hill and Robertson, 1968]. The HapMap data based on 90 Yorubas (30 parent-offspring trios) in Ibadan and Nigeria (www.hapmap.org) were retrieved. A total of 2,851 single nucleotide polymorphism (SNP) genotypes in X chromosome region were downloaded from the February 2006 public release of the HapMap data. After excluding SNPs with minor variants and missing genotypes, the remaining 1,036 SNPs were used for linkage disequilibrium (LD) map construction by the method of Haploview 3.32 [Barrett et al., 2005].

Results and Discussion Data Analyses All DNA sequence alignments were made by Clustal X [Thompson et al., 1997] and then manually checked. For phylogenetic analyses, the neighbor-joining method [Saitou and Nei, 1987] was used. To evaluate functional importance of AMELX and AMELY, the genomic sequences of primate and non-primate mammalian amelogenin genes were determined [Iwase et al., 2003] and homologous sequences available in the National Center for Biotechnology Information database were retrieved. Unfortunately, most of mammalian AMELX sequences used in Delgado et al. [2005] were unavailable in the DNA sequence database, so that only 20 sequences were used. The ratio (f) of the non-synonymous to synonymous substitutions per site was computed along individual branches in the reconstructed neighbor-joining tree by the least square method [Zhang et al., 1998]. For the human population data, DnaSP v.4.10 [Rozas et al., 2003] was used to compute some summary statistics concerning polymorphism:

Evolutionary History of Sex-Linked Mammalian Amelogenin Genes

AMEL and ARHGAP6 The AMEL gene in eutherians, opossums and African clawed toads resides in a large intron of ARHGAP6 in the opposite orientation (fig. 1). By contrast, the chicken and fugu genome databases show no trace of AMEL in their genomes. As mentioned earlier, AMEL is a member of the SCPP family and all the members except AMEL are clustered in one chromosome [Kawasaki and Weiss, 2003, 2006]. It is therefore likely that AMEL too originally arose in that cluster and happened to transpose into an intron of ARHGAP6, thereby making a nested gene structure. There are two possible explanations for the presence or absence of AMEL in different tetrapod genomes. One inCells Tissues Organs 2007;186:49–59

51

AMELY

>50 MYA

Stratum 3, p < 0.1 Ancient PAR

AMELX

AMELY

<50 MYA

Stratum 3, p > 0.1

Stratum 4, p > 0 New PAR

AMELX

Fig. 2. Evolutionary stratum and a rela-

tively recent X-chromosomal inversion encompassing AMELX. Strata 3 and 4 began to be formed in the ancestral lineage of eutherians, about 100 MYA, and in the ancestral lineage of simian primates (New and Old World monkeys), 150 MYA, respectively. The gametologous ARHGAP6 on the human Y chromosome has since disappeared. The inversion of 3- to 4-Mb harboring AMELX is X-chromosome specific and must have occurred after the formation of stratum 4. p = The nucleotide differences per site between X-Y homologous regions.

AMELY

Present PAR

vokes a single transposition in the ancestral lineage of tetrapods. The absence of AMEL in chicken and possibly in toothless turtles can then be explained by secondary loss of the gene. The disappearance of chicken AMEL is consistent with the absence of exon 1 and intron 1 of chicken ARHGAP6 (fig. 1). Alternatively, AMEL might transpose twice independently: one in the amphibian lineage and the other in the ancestral mammalian lineage. Although there is no need to invoke loss of AMEL in birds, this alternative becomes much less parsimonious than the first when we explain (1) why the genomic position of mammalian and amphibian AMELs is the same and (2) how AMEL came to exist in reptiles, e.g. caimans [Toyosawa et al., 1998]. Hence, it is concluded that all tetrapod AMELs in ARHGAP6 have experienced a single transposition.

52

Cells Tissues Organs 2007;186:49–59

Stratum 3, p = 0.2

Stratum 4, p = 0.1

At present

XLEMA Inversion

Even after sex chromosomes evolved independently in some reptiles, birds and mammals, AMEL and ARHGAP6 had remained to be autosomal. It is only in eutherian mammals that a pair of homologous autosomes that carried these and all other linked genes were added or translocated to the telomeric end of the pseudoautosomal region (PAR) of the sex chromosomes. As a consequence, subsequent evolution of eutherian AMEL became intimately related to evolution of the sex chromosomes. AMEL Differentiation in the Eutherian Sex Chromosomes The original mammalian sex chromosomes arose from a pair of homologous autosomes, 1200 MYA before the divergence of monotherians [Ohno, 1967; Graves, 2002]. One possible cause for this sex-chromosomal difIwase/Kaneko/Kim/Satta/Takahata

100 100

Human X Chimpanzee X

99

Squirrel monkey X Ring-tailed lemur X

73

Cattle X

98 98

Pig X Horse X

88

100 Human Y Chimpanzee Y

100 86

Squirrel monkey Y Ring-tailed lemur Y

41

Cattle Y

66 93

Pig Y Horse Y House shrew AMEL Opossum AMEL

a

0.05

Human X

100 99

Chimpanzee X Squirrel monkey X

100

Fig. 3. Phylogenetic relationships of mammalian AMELX and AMELY sequences rooted by house shrew and opossum AMELs [accession numbers: AB287298 and AB287299]. Not only the coding but also intron sequences were used wherever they can be aligned. Open diamonds stand for differentiation points between gametologous AMELX and AMELY by recombination inhibition. a The 5 region located in evolutionary stratum 3 [Lahn and Page, 1999], including partial intron sequences (483 bp). b The 3 region located in evolutionary stratum 4, including partial intron sequences (770 bp). a, b The number near a node stands for the bootstrap value in 1,000 replications.

100 Human Y Chimpanzee Y

100 98

Squirrel monkey Y Ring-tailed lemur X

100

95

Ring-tailed lemur Y Cattle X

41 87

Cattle Y Pig X Pig Y

100

83

Horse X 98

Horse Y House shrew AMEL Opossum AMEL

b

0.05

ferentiation is that two or more genes that determine complex sex characters evolved in one chromosome and that homologous recombination among these genes was inhibited [Nei, 1969 and see a later discussion]. The earliest inhibition of homologous recombination appears to be responsible for forming the oldest so-called ‘evolutionary stratum 1’ manifested in the long arm of the X chromosome [Lahn and Page, 1999]. Later, after the divergence between eutherians and metatherians, a pair of homologous autosomes that harbored AMEL fused with the

original mammalian sex chromosomes and became the short arms [Graves, 2002]. The proximal part of the short arms adjacent to stratum 2, which was deposited near the centromeric region by the time of the chromosomal fusion, was then subjected to recombination inhibition in the stem lineage of eutherians 1100 MYA. However, the distal part of the short arms was permitted to recombine until the emergence of simian primates (New and Old World monkeys), about 50 MYA [Martin, 1993; Takahata, 2001]. Thus, during the period of about 50 MYA, the

Evolutionary History of Sex-Linked Mammalian Amelogenin Genes

Cells Tissues Organs 2007;186:49–59

53

1.67 0.29

Fig. 4. The ratio (f) of per site nonsynony-

mous to synonymous substitutions along branches in the neighbor-joining tree of eutherian AMELXs and AMELYs with opossum AMEL [Hu et al., 1996, accession number: AB287299] as an outgroup. The exon 3, 5 and 6 sequences are used for computing per site synonymous substitutions, whereas only the exon 6 sequences are used for computing per site nonsynonymous substitutions. The ratio f is then calculated as the ratio of the per site nonsynonymous substitutions in exon 6 to the per site synonymous substitutions in exons 3, 5 and 6. The total number of synonymous sites is 132.8 in the three exons, and the number of nonsynonymous sites is 268.3 in exon 6. The symbol G means no synonymous substitutions. Significance levels of f ! 1 are indicated by asterisks (* 0.01 ! p ! 0.05, ** p ! 0.01).

Cells Tissues Organs 2007;186:49–59

Human X Chimpanzee X

0.30 Rhesus monkey X

Squirrel monkey X 14.4 1.84 Human Y 0 Chimpanzee Y 0.53 Squirrel monkey Y 0 Mouse X 0.73 0 0.87 Rat X 0.16 Golden hamster X 0.81 Guinea pig X

0.07*

0.50

0.73

5.33 0.28



Goat X Goat Y 0.25 Cattle X 0.18** 0.70

2.01 1.55 0.17**

0.35* 0.30**

Cattle Y

Pig X Pig Y Horse X Horse Y Opossum AMEL

0.02

proximal part (stratum 3) accumulated substantial sequence differences, yet the distal part was still allelic or constituted the ancient PAR in which the X and Y chromosomes could pair and recombine in meiosis (fig. 2). The junction between these proximal and distal parts is marked by transposon medium reiterated frequency repeat 5 (MER5) within intron 2 of AMELX and regarded as an ancient pseudoautosomal boundary [Iwase et al., 2001, 2003]. The phylogenetic analysis of eutherian AMELX and AMELY genes shows that the 5 region (upstream from MER5) differentiated before the eutherian radiation, while the 3 region (downstream from MER5) differentiated independently within individual eutherian orders (fig. 3). In primates, differentiation of the 3 region occurred after the divergence between prosimians and simian primates, but before the splitting between New and Old World monkeys. Since exons 1 and 2 in the 5 region are largely untranslated, it is naturally found that the phylogenetic relationship in the 3 region is identical to the one previously studied based on the amino acid or intron 3 sequences [Huang et al., 1997; Toyosawa et al., 1998]. Differentiation of AMELX and AMELY is likely a result, rather than a cause, of recombination inhibition in the short arms. It was argued that SRY (sex-determining region Y) and RBMY (RNA-binding motif Y) are candi54

0.40



date genes for recombination inhibition [Iwase et al., 2003]. In this respect, it is interesting to note the presence of nine gene families in the human Y ampliconic region or massive repeat units [Skaletsky et al., 2003]. These gene families including RBMY are expressed exclusively or predominantly in testes and many of them are implemented in spermatogenesis or sperm production. The families originated either from proto-XY gene pairs in the original mammalian sex chromosomes or from retroposition or transposition of autosomal genes. The emergence of such genes as CDY (chromodomain Y) and VCY (variable charge Y) coincides with formation of stratum 3, about 100 MYA, and stratum 4, about 50 MYA, respectively [Bhowmick et al., 2006]. For these reasons, it is tempting to speculate that these ampliconic genes or male-specific genes in the human Y amplicons have somehow been involved in stepwise differentiation of the eutherian sex chromosomes as well as in that of AMELX and AMELY. Selection for and against AMELX and AMELY As aforementioned, AMELY genes may not be under strong functional constraint or may be even on the way to dead genes or pseudogenes. To examine this possibility, we estimated synonymous (bS) and nonsynonymous (bN) substitutions that have accumulated in individual Iwase/Kaneko/Kim/Satta/Takahata

Table 1. Polymorphism at the human AMELX and AMELY loci and the average nucleotide differences per site

(p distances) from the chimpanzee ortholog

Nucleotide sites, bp Segregating sites Haplotypes , % [Nei and Li, 1979] , % [Watterson, 1975] D [Tajima, 1989] p distances, %

5 region in evolutionary stratum 3

3 region in evolutionary stratum 4

Both regions

AMELX

AMELY

AMELX

AMELX

3,229 0 1 0 0 0 1.24

3,942 6 9 0.055 0.035 0.491 0.69

2,571 10 12 0.064 0.089 –0.831 0.86

AMELY 3,398 6 8 0.022 0.042 –1.381 1.27

6,513 16 18 0.058 0.056 0.121 0.76

AMELY 6,627 6 8 0.012 0.022 –1.381 1.25

The sample size of AMELX and AMELY is 45 and 18, respectively. 1 Not significant (p > 0.1).

branches of the AMELX and AMELY gene tree (fig. 4). The ratio (f) of bN/bS is an indicator of selective pressure for nonsynonymous substitutions relative to synonymous substitutions both of which have accumulated for the same period of evolutionary time. The value of f ranges from 0 to 1 under the neutral theory of molecular evolution [Kimura, 1983]. Since the neutral theory assumes negligible roles of positive selection at the molecular level, the smaller the f value, the stronger the negative pressure against nonsynonymous substitutions. A caveat is that although the neutral mutation rate per se may differ between the X- and Y-linked genes [Ebersberger et al., 2002], the f value is independent of the mutation rate. It is therefore sensible to compare f values of various genes irrespective of their chromosomal locations. On the other hand, if positive selection operates for nonsynonymous substitutions, the f value may become 11. However, since all nonsynonymous sites in a gene are unlikely to be subjected to positive selection, the observation of f 1 1 averaged over the nonsynonymous sites in a given gene tends to be a very conservative criterion for detecting positive selection. Unexpectedly, along the common ancestral lineage leading to humans and chimpanzees, f becomes 11 in both AMELX and AMELY (fig. 4). This enhanced rate of nonsynonymous substitutions can also be visible in human AMELY. Similarly, in the ancestral lineage of Ruminantia (cattle and goats) or in that of Perissodactyla (horses), f 1 1 is found before the divergence between their gametologous AMELX and AMELY. Although these f values are subject to large sampling errors Evolutionary History of Sex-Linked Mammalian Amelogenin Genes

and are not significantly greater than 1 in most of the cases, it is suggested that positive selection operated in particular lineages of eutherian amelogenin genes. The remaining lineages show conservation of amelogenin genes at the amino acid level. In particular, AMELY is highly conserved in cattle and horses. On the other hand, AMELX in rodents exhibits relatively high f values. This observation raises two possibilities: relaxation of functional constraint and positive selection for some nonsynonymous substitutions. In the absence of AMELY in rodents, the latter possibility appears more likely than the former. In any case, there is no indication for preferential deterioration of AMELY at the amino acid level. Rather, like AMELX, existing AMELY genes have experienced positive selection, followed by negative selection. Polymorphism of Human AMELX and AMELY We examined the DNA sequences of 45 AMELX genes (each about 6.5 kb) and 18 AMELY genes (each about 6.6 kb) for a worldwide sample taken from the human population (table 1). As expected, almost all observed polymorphic or segregating sites are due to single nucleotide substitutions and occur in introns 1 and 2. Exceptionally, two substitutions are found in the coding region. One is a synonymous substitution in exon 6 of AMELX that is shared by different ethnic groups. The other is a nonsense mutation in exon 5 of AMELY that is represented by a single Asian male (Ami) in the Y chromosome sample. In addition, there is only one insertion/deletion polymorphism (4 bp) in intron 1 that is represented by a single chromoCells Tissues Organs 2007;186:49–59

55

some in Druze. Thus, although human AMELX and AMELY experienced positive selection in the past, they are well conserved in the present-day human population. The nucleotide differences per site (p) between human and chimpanzee orthologs are uniformly distributed over the 5 and 3 regions (table 1), suggesting no regionspecific, differential mutation rates. However, the overall p-distances are significantly greater in the comparison of AMELY (1.25%) than of AMELX (0.76%). These values are in agreement with previous estimates if high occurrences of C to T mutations at CpG sites are excluded [Ebersberger et al., 2002; Jobling et al., 2004]. The relatively large p value for AMELY supports the notion of male-driven hypothesis of molecular evolution [Miyata et al., 1987]. Provided that the sex ratio is 1, AMELY evolves with male mutation rate rm whereas one third of AMELX in a population evolves with rm and two thirds with female mutation rate rf. Thus, we have p(Y ) = 2trm

(1a)

p(X ) = 2t(rm + 2rf )/3

(1b)

in which p(Y ) = 1.25%, p(X) = 0.76% and t is the divergence time between humans and chimpanzees. Eliminating t in (1), we obtain the ratio of male to female mutation rates ( = rm/rf ) of about 2.6. The extent of polymorphism measured by nucleotide diversity  and  is also more or less homogeneous over the 5 and 3 regions. The theoretical formula of  [Nei, 1987] suggests that the ratio of X chromosomal to autosomal  is given by (1 + /2)/(1 + ).

(2)

Formula (2) takes into account the differences in both population sizes and mutation rates between autosomal and X-linked genes. If  = 2.6, the ratio becomes 0.64. Even if the  value is as large as suggested by other studies [see Jobling et al., 2004 for review], the expected ratio must be 10.5. A typical value of  for human autosomes is as low as 0.088% [Yu et al., 2002] and implies a relatively small effective size in the human demographic history [Takahata, 1993]. With 0.088% for autosomal , the expected  value for human X-linked genes ranges from 0.044 to 0.056% and is in agreement with the observed value in the 5 and 3 regions (table 1). In our sample of human AMELY, there is no segregating site in the 5 region, but six in the 3 region. However, since the difference in the extent of polymorphism between the 5 and 3 region is not statistically significant, we compare the expected  value with the observed 56

Cells Tissues Organs 2007;186:49–59

0.012% over the two regions. The ratio of Y chromosomal to autosomal  is given by /{2(1 + )}.

(3)

This ratio becomes 0.36 for  = 2.6 and 0.5 for larger . Thus, the  value in formula (3) ranges from 0.032 to 0.044%. Although the observed 0.012% is below this range and may be lowered by either positive or negative selection at completely linked sites, there is no significant difference between these expected and observed values. LD and Ancient Pseudoautosomal Boundary When carrying out a human population survey of AMELX and AMELY, we hypothesized that molecular mechanisms responsible for making the ancient pseudoautosomal boundary within the amelogenin gene may still somehow affect patterns and levels of the present-day polymorphism. We first examined LD at pairs of polymorphic sites in a specified region. We measured nonrandom association at such a pair of sites by r 2 [Hill and Robertson, 1968] or the absolute square root r. These values cannot be large when segregating sites under study are not intermediate in frequency in a sample. We ignored this fact and took the average over all pairs of polymorphic sites. Indeed, the average r value for AMELY becomes as small as 0.19 despite the absence of recombination. On the other hand, the average value for AMELX is 0.15 in the 5 region, 0.37 in the 3 region and 0.21 in the entire region. The r value is slightly larger in the 3 region than in the 5 region, reflecting some excess of rarefrequency segregating sites in the former region (or D ! 0 in table 1). Clearly, it is necessary to examine LD in a large sample as well as in a large chromosomal scale, because formation of evolutionary stratum is a chromosome-wide phenomenon. To this end, we used the HapMap data encompassing X-linked ARHGAP6 of 570 kb length. The LD analysis for the African population (fig. 5) shows the presence of a strong LD block in the large intron of ARHGAP6. The block is largest within the surrounding region of 3 Mb length and can be taken as evidence for the presence of a recombination cold spot in ARHGAP6. Since almost the same pattern is obtained in the European and Asian populations (data not shown), the phenomenon does not seem to stem from the human demography, but from genomic causes. We presume that when stratum 3 was formed 100 MYA, the cold spot in ARHGAP6 already existed in the eutherian sex chromosomes. We then hypothesize that this cold spot was used Iwase/Kaneko/Kim/Satta/Takahata

Fig. 5. LD map in the region of 10.6–13.6 Mb from the short arm end of the human X chromosome that includes

AMELX and ARHGAP6. The top bar indicates positions of genotyped SNPs. Solid triangles indicate strong LD blocks defined as in Gabriel et al. [2002]. The lower panel is a magnification of a region surrounding ARHGAP6.

to determine the proximal end of the ancient PAR that consists of the current PAR and stratum 4 (fig. 2). However, it is to be noted that the boundary of this ancient PAR occurs within AMELX and does not exactly correspond to the distal end of the cold spot (fig. 5): Actually the cold spot is included in the centromeric end of the ancient PAR. Nonetheless, we may argue that the cold spot was fortuitously involved in the determination of the ancient pseudoautosomal boundary. Curiously, the gene orientation or the centromeretelomere polarity of AMELX and ARHGAP6 in humans, chimpanzees and rhesus monkeys is reversed compared with that of AMELY and deteriorated A RHGAP6 in their Y chromosomes as if the 5 region of AMELX disrupts the continuity of stratum 4 (fig. 1, 2). This may result from a small X chromosomal inversion of !3 to 4 Mb length [Ross et al., 2005]. Indeed, the gene orientation of AMELX

in cattle in the National Center for Biotechnology Information database is just opposite to that of humans, chimpanzees and rhesus monkeys. It thus appears that the inversion occurred after the eutherian radiation, !100 MYA, but before the divergence between hominoids and Old World monkeys, 130 MYA. More precisely, since the extent of sequence differentiation in the 3 region between human AMELX and AMELY is the same as that of stratum 4, we may conclude that the inversion should not predate the formation of stratum 4 (fig. 2). In other words, the inversion that occurred during the time period from 30 to 50 MYA is unlikely to be the cause of recombination inhibition for stratum 4.

Evolutionary History of Sex-Linked Mammalian Amelogenin Genes

Cells Tissues Organs 2007;186:49–59

57

Acknowledgments

Conclusions

In this article, we tried to elucidate the entire history of the amelogenin gene from its origin in the tetrapod ancestor to the subsequent evolution in the lineage leading to humans. Molecular phylogenetic and populationgenetic analyses revealed that the gene has experienced various evolutionary events such as transposition, translocation, positive selection and sex differentiation. We also showed that the eutherian amelogenin locus inscribes footprints of stepwise differentiation of the sex chromosomes and chromosomal rearrangements such as translocation and inversion.

We thank Prof. J. Simmer for his kind invitation to the 2006 IADR Symposium entitled Mineralization Genes of Bones and Teeth: The New Evolutionary Synthesis. We also thank Drs. K. Kawasaki and J.-Y. Sire for their comments on an early version of this paper. This work is supported in part by the Japan Society for Promotion of Science Grant 12304046 to N.T.

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Tajima, F. (1989) Statistical method for testing the neutral mutation hypothesis by DNA polymorphism. Genetics 123: 585–595. Takahata, N. (1993) Allelic genealogy and human evolution. Mol Biol Evol 10: 2–22. Takahata, N. (2001) Molecular phylogeny and demographic history of humans; in Tobias, P.V., M.A. Raath, J. Moggi-Cecchi, G. A. Doyle (eds): Humanity from African Naissance to Coming Millennia. Johannesburg, Firenze/Witwatersrand University Press, pp 299–305. Thompson, J., T. Gibson, F. Plewniak, F. Jeanmougin, D. Higgins (1997) The CLUSTAL_ X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876–4882. Toyosawa, S., C. O’hUigin, F. Figueroa, H. Tichy, J. Klein (1998) Identification and characterization of amelogenin genes in monotremes, reptiles, and amphibians. Proc Natl Acad Sci USA 95: 13056–13061.

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Cells Tissues Organs 2007;186:60–69 DOI: 10.1159/000102681

Unraveling the Molecular Mechanisms That Lead to Supernumerary Teeth in Mice and Men: Current Concepts and Novel Approaches Rena N. D’Souza a Ophir D. Klein b a

Department of Biomedical Sciences, Baylor College of Dentistry, Texas A&M University Health Science Center, Dallas, Tex., and b Department of Pediatrics, University of California, San Francisco, Calif., USA

Key Words Fibroblast growth factors
Runx2
Sprouty
Supernumerary teeth
Transgenic mice

Abstract Supernumerary teeth are defined as those that are present in excess of the normal complement of human dentition and represent a unique developmental anomaly of patterning and morphogenesis. Despite the wealth of information generated from studies on normal tooth development, the genetic etiology and molecular mechanisms that lead to congenital deviations in tooth number are poorly understood. For developmental biologists, the phenomenon of supernumerary teeth raises interesting questions about the development and fate of the dental lamina. For cell and molecular biologists, the anomaly of supernumerary teeth inspires several questions about the actions and interactions of transcription factors and growth factors that coordinate morphogenesis, cell survival and programmed cell death. For human geneticists, the condition as it presents itself in either syndromic or non-syndromic forms offers an opportunity to discover mutations in known or novel genes. For clinicians faced with treating the dental complications that arise from the presence of supernumerary teeth, knowledge about the

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

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basic mechanisms involved is essential. The purpose of this manuscript is to review current knowledge about how supernumerary teeth form, the molecular insights gained through studies on mice that are deficient in certain tooth signaling molecules and the questions that require further research in the field. Copyright © 2007 S. Karger AG, Basel

Abbreviations used in this paper

BMP CCD Eda FGF3–10 FGFR1, 2 M1 MSX1 PAX6, PAX9 RUNX2 SHH Spry2, 4 Wnt

bone morphogenic protein cleidocranial dysplasia ectodysplasin gene fibroblast growth factors 3–10 fibroblast growth factor receptors 1, 2 first molar homeo box, msh-like 1 paired box genes 6, 9 runt-related gene 2 sonic hedgehog sprouty genes 2, 4 an oncogene

Dr. Rena N. D’Souza Department of Biomedical Sciences, Baylor College of Dentistry Texas A&M University System, Health Science Center, 3302 Gaston Avenue Dallas, TX 75246 (USA) Tel. +1 214 828 8260, Fax +1 214 828 8375, E-Mail [email protected]

Introduction

The Dental Lamina

In contrast to the reduced monophyodont rodent dentition in which only incisors and molars are present, the diphyodont human dentition exhibits all four tooth classes at maturity. The deciduous complement of 20 teeth consists in each quadrant two incisors, one canine and two molars that are shed during childhood while the permanent or secondary dentition consists of 32 teeth, namely, two incisors, one canine, two premolars and three molars in each quadrant. Molecular and genetic studies performed in the past two decades have shown that the patterning of dentition is a highly complex process that provides a valuable developmental model for studying genes that control three-dimensional patterning and morphogenesis. Taken together, these advances have improved our understanding that the precise control of the number, position, size and shape of teeth within the maxilla and mandible requires signaling between odontogenic (tooth-specific) epithelium and mesenchyme. Such interactions involve diffusible morphogens and nuclear transcription factors that operate within parallel signaling pathways to give rise to an exquisitely patterned dentition. That such a complex process frequently goes awry is not surprising, and patterning defects in human dentition occur often and are characterized by alterations in the number, size and shape of teeth. The importance of the genetic control of the patterning of human dentition is underscored by the findings that mutations in genes that encode the transcription factors PAX9 and MSX1 lead to non-syndromic tooth agenesis or the congenital absence of teeth, among the most commonly inherited disorders in humans [hypodontia (OMIM 106600)]. Another condition affecting the patterning of dentition is the presence of supernumerary teeth, a relatively rare condition that is characterized by the presence of more than the normal complement of 20 deciduous and 32 permanent teeth. In contrast to advances made in our understanding of the genetic etiology of tooth agenesis and its underlying molecular mechanisms, not as much has been reported about the genetic and molecular defects that lead to supernumerary tooth development. The objectives of this review are to survey classical literature concerning the etiology, frequency and classification of supernumerary teeth, to highlight molecular insights gained from the use of genetically engineered mouse models, and to propose new research opportunities in human molecular genetics.

Knowledge about the origin, development and fate of the dental lamina is critical to our understanding of how supernumerary teeth arise. In humans, the primitive oral cavity or stomatodeum becomes apparent around day 25 of development and is demarcated laterally by the first pair of branchial (pharyngeal) arches [Krause and Jordan, 1965; Nanci, 2003]. The latter consist of an external layer of ectoderm, an intermediate layer of lateral plate mesoderm and an internal layer of endoderm. Microscopically, the stomatodeum is lined by a two- to three-cell layer of epithelial cells that overlies a layer of embryonic connective tissue derived from cranial neural crest ectomesenchyme. Around day 37 of embryonic development, the presumptive maxilla and mandible become visible as each is covered by a continuous layer of thickened epithelium, called the primary epithelial band. Shortly after, this band of tissue divides into two distinct structures, the vestibular lamina and the dental lamina. While both structures are composed of highly proliferative cells that rapidly grow into surrounding ectomesenchyme, cells of the vestibular lamina enlarge and undergo programmed cell death to form a cleft (the space between the cheek and the arch). In contrast, the dental lamina remains proliferative, extending outgrowths into the ectomesenchyme but only at future sites of odontogenesis. The epithelial outgrowth or tooth bud then progresses from the cap to the bell stages of morphogenesis but remains attached to the dental lamina through a stalk-like extension called the lateral lamina. During the bell stage and prior to the differentiation of odontoblasts and ameloblasts, the dental lamina and the lateral lamina that connects the tooth organ to the overlying oral epithelium fragment into small clusters of cells. These cells normally undergo programmed cell death but sometimes persist to form structures called epithelial pearls. The formation of eruption cysts from these epithelial remnants is known to interfere with the normal eruption pathway of the underlying tooth organ [Moskow and Bloom, 1983; Pindborg, 1970]. Thus far, studies on the fate of the dental lamina have been limited to histologic and ultrastructural analyses that suggest that the sequence of events leading to its degeneration is initiated by underlying mesenchyme [Khaejornbut et al., 1991]. The primary dental lamina that gives rise to the deciduous dentition is also responsible for the formation of the succedaneous (permanent) incisors, canines and premolars. These develop as lingual extensions of the dental lamina and are clearly visible in a coronal section through a developing human jaw. The three permanent molars that

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Fig. 1. Panel describing the dental complications and the sequence of treatment for a 17-year-old male affected by CCD. a, b Panoramic radiograph and tracing. CI = Central incisor; L = lateral incisor; C = canine; P = premolar; 6 = permanent first molar; 7 = permanent second molar; 8 = permanent third molar; D = deciduous first molar; E = deciduous second molar. The presence of multiple supernumerary teeth creates several problems. Note that the only permanent teeth which have erupted are the maxillary left central incisor and all first molars. Multiple retained deciduous teeth are also present. c Preoperative intraoral frontal photo-

do not have deciduous predecessors develop from an outgrowth of the primary dental lamina backward into the jaw ectomesenchyme. There is relatively little known about the genetic factors that control the fate of the dental lamina. Future studies on the patterns of differential gene expression within this specialized epithelium and its surrounding mesenchyme are needed to increase our understanding of the molecular mechanisms that lead to aberrations like missing teeth or the formation of supernumerary teeth. 62

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graph showing that the majority of permanent teeth are unerupted. d Surgical exposure and extraction of all supernumerary teeth and deciduous teeth with the exception of the right deciduous second molar, which was not extracted for orthodontic anchorage. e Intraoral view of mandibular arch, showing lip bumper appliance. Orthodontic traction, through bonded attachments with chains, will be applied to promote eruption of the remaining teeth. A series of orthodontic interventions are planned to restore normal occlusion.

Supernumerary Teeth in Humans

Supernumerary teeth exist in excess of the normal complement of deciduous and permanent teeth and are easily diagnosed by clinical examination and/or radiographs. Most common among the supernumeraries are mesiodens, or teeth that appear on the palatal side between the maxillary incisors. Accessory canines and premolars are typically located within the arch form while D’Souza /Klein

supernumerary molars develop buccal to the permanent molars or distal to the third molars. Supernumerary teeth can present unilaterally, bilaterally, singly or multiply, and in the maxilla and/or mandible. Non-syndromic cases of supernumerary teeth are reported to occur with varying frequencies in different ethnic populations while a few cases of inherited or familial forms have been reported [Burzynski and Escobar, 1983; Yusof, 1990]. Supernumerary teeth can also be associated with other organ anomalies [D’Souza et al., 2006]. Syndromes that involve supernumeraries include X-linked conditions like focal dermal hypoplasia syndrome and orofaciodigital syndromes types I and III, autosomal recessive disorders like steroid dehydrogenase deficiency and RothmundThomson syndrome, and autosomal dominant conditions that include syndromic cleft lip and palate, cleidocranial dysplasia (CCD), Gardner syndrome and NanceHoran syndrome [for review, see Zhu et al., 1996]. Although the presence of supernumerary teeth can greatly compromise esthetics, the condition is not lifethreatening. Several predictable dental complications can arise that range from mild to severe, depending on the number and location of supernumeraries. These include but are not limited to: dentigerous cysts; malocclusion due to impaction and pressure on adjacent teeth; resorption of bone; pericoronitis and impingement on nerves leading to paresthesia and/or pain [Bodin et al., 1978; Primosch, 1981; Burzynski and Escobar, 1983]. Figure 1 describes the sequence of treatment involved in correcting malocclusion associated with CCD. The overall patterns of presentation and incidence of supernumerary teeth support the various theories that are proposed to explain how the condition arises. One theory hypothesizes that these teeth are derived from remnants of the dental lamina that fail to degenerate and become reactivated to form accessory tooth organs. Another theory proposes that the dental lamina, while intact, continues to proliferate due to a failure of programmed cell death, which may be brought on by defects in signaling between epithelium and mesenchyme. Finally, the possibility that supernumerary teeth arise from the division of a single tooth bud is supported by a few case reports. One describes differences in the mesiodistal width of central incisors depending on unilateral or bilateral occurrence of mesiodens. Another report describes gemination of a deciduous incisor on the same side of a mesiodens [Stellzig et al., 1997]. From a developmental and molecular viewpoint, each theory is plausible and can explain the origin of different types of supernumerary teeth.

When using animal models to study changes in tooth number, it is important to keep in mind that tooth number varies dramatically among species, presumably as a functional adaptation in response to environmental pressures [Line, 2003]. Both humans and rodents have reduced mammalian dentition in comparison to the ancestral eutherian formula, in which up to three incisors, one canine, four premolars, and three molars can occur in each dental quadrant. This primitive formula can be seen in some extant mammals, such as certain species of mole. Compared with humans, the adult mouse dentition is severely reduced. Each dental quadrant contains three molars and one incisor, separated by an edentulous region called a diastema (fig. 2), and no premolars or canines. The absence of teeth in the adult mouse diastema does not reflect a lack of tooth development during embryogenesis. Transient epithelial primordia originate in the mouse diastema and reach the bud stage before regressing [Peterkova et al., 2002]. These primordia are presumably evolutionary remnants of the developmental program for tooth formation in species that have teeth between the incisors and molars. Although the diastema buds initially appear quite robust, they do not progress to the cap stage. In the mandibular antemolar region, two primordia are detected anterior to the presumptive first molar (M1). The more anterior of these structures is thought to regress completely, whilst the more posterior of these, adjacent to M1, may be partially absorbed into the developing M1. The mouse maxilla has not been examined in as much detail as the mandible, but the maxillary diastema is believed to contain as many as seven buds [Peterkova et al., 2000]. The reason for the different number of buds in the maxilla and mandible is not well understood. The elimination of the diastema tooth buds involves apoptosis [Tureckova et al., 1996; Peterkova et al., 2000; Peterkova et al., 2003], but it is not known if cell death is a primary event in failure of the diastema tooth bud to develop, or if it is secondary to failure of the bud to progress to the cap stage. Supernumerary diastema teeth have been found in a few mutant mouse strains, including mice null for the fibroblast growth factor (FGF) antagonists Spry2 or Spry4 [Klein et al., 2006] (fig. 2), Polaris hypomorphs [Zhang et al., 2003], mice that overexpress ectodysplasin (Eda) or its receptor [Mustonen et al., 2003; Pispa et al., 2004], Pax6 mutants [Kaufman et al., 1995], and mice that are

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Mouse Models of Supernumerary Teeth

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Fig. 2. Views of the mouse diastema and the presence of diastema teeth in Spry2 null. Side views of the molar and diastema region of wild-type (left) and Spry2-null (right) mice. M1, M2, M3 = 1st, 2nd, and 3rd molars. White arrow points to a diastema tooth.

Fig. 3. Perturbation of signaling pathways can lead to supernumerary teeth by affecting epithelial-mesenchymal interactions. This model summarizes some of the critical signaling interactions between the enamel knot (EK) in the dental epithelium (DE) and the condensing dental mesenchyme (CDM) in a cap-stage molar tooth germ. It is based on data from both gene expression studies and manipulations of tooth germs in vitro [Kettunen et al., 2000; Kratochwil et al., 2002] and from genetic studies on knockout mice [Kassai et al., 2005; Klein et al., 2006]. Arrows indicate a stimulatory effect. The symbol indicates an inhibitory effect of one molecule on the expression of another when solid, and inhibition of signaling by a ligand when dashed. Wnt signal-

ing induces epithelial FGFs, which in turn induce mesenchymal FGFs via MSX1 and RUNX2. Mesenchymal FGFs induce SHH in the epithelium. FGF signaling to the epithelium and mesenchyme is blocked by SPRY2 and SPRY4, respectively. Loss of function of Spry2 or Spry4 leads to supernumerary tooth development by upregulating FGF signaling. Wnt signaling may also induce ectodysplasin (EDA), a molecule that can lead to supernumerary teeth when overexpressed. Ectodin/WISE is a putative inhibitor of both BMP and Wnt signaling, and loss of ectodin/WISE function leads to supernumerary teeth. The precise signaling pathways modulated by ectodin/WISE function have yet to be elucidated.

null for ectodin/WISE, a bone morphogenic protein (BMP) and/or Wnt inhibitor [Kassai et al., 2005]. Interestingly, mutations in the gene encoding Polaris affect sonic hedgehog (SHH) signaling [Murcia et al., 2000; Huangfu et al., 2003; Liu et al., 2005], and mutations in

the Eda pathway can be partially rescued by increasing FGF signaling in organ culture [Pispa et al., 1999]. Thus, modulation of pathways that are necessary for molar and incisor development, such as those initiated by SHH, FGFs, and BMPs, can lead to the development of diastema

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D’Souza /Klein

teeth in mice. Pax6 mutants have also been reported to have supernumerary incisors [Quinn et al., 1997]. Another major difference between mouse and human dentition is that mice have only a single set of teeth, whereas in humans the first set of teeth (primary or deciduous teeth) is replaced by a permanent set during childhood. The generation of progressive cycles of teeth appears to be the primitive (ancestral) condition for vertebrates. For example, lower vertebrates like fish replace their teeth throughout life. Most mammals, including humans, have only one cycle of tooth replacement, in which a deciduous (primary) dentition is supplanted by a permanent (secondary) dentition. The secondary dentition develops from lingual buds off of the deciduous lamina. Mice, in contrast, have only one dentition. The mouse therefore provides a simplified model for tooth formation in humans.

The development of diastema teeth could potentially be due to events either at the placode or at the bud stage. An expansion of the dental lamina at an early stage could lead to the development of more teeth. Later events, such as survival of the diastema bud, can also lead to diastema teeth [Klein et al., 2006]. Because recent work has focused on signaling events at the bud-cap transition leading to survival of the diastema bud, we have focused on this below. Whether the diastema bud normally has a functional enamel knot is still a matter of controversy. It appears, though, that in mutants with diastema teeth, the presence of a functional enamel knot is essential for maintenance of the diastema bud. The pathways that are important in this process are diagrammed in figure 3, and they include members of the FGF, EDA, BMP and Wnt signaling families [reviewed in Tucker and Sharpe, 2004]. As an example of how changes in the function of one of these pathways can lead to development of diastema teeth, we focus below on the role of the FGF signaling pathway in this process. The FGF signaling pathway appears to mediate epithelial-mesenchymal interactions at several stages of tooth morphogenesis in mammals and other vertebrates [Thesleff and Sharpe, 1997; Jernvall and Thesleff, 2000; Mandler and Neubüser, 2001; Jackman et al., 2004]. In the developing mouse molar, at least five different FGF ligands (FGF3, FGF4, FGF8, FGF9, and FGF10) and two receptors (FGFR1 and FGFR2) are expressed in complex,

overlapping patterns in the epithelium and/or mesenchyme [Niswander and Martin, 1992; Neubüser et al., 1997; Kettunen et al., 1998; Kettunen and Thesleff, 1998; Kettunen et al., 2000]. Molar development is thought to be initiated by signaling via FGF8 [Trumpp et al., 1999], but Fgf8 does not appear to have a function later in tooth development, because Fgf8 expression is not detected at later stages [Kettunen and Thesleff, 1998]. Subsequent development of the tooth is thought to depend on signaling via other FGFs (fig. 3). This hypothesis has been based primarily on gain of function studies in organ culture and gene expression analyses. Epithelial cell proliferation and morphogenesis throughout the cap and bell stages is stimulated by FGF3 and FGF10 produced in the dental mesenchyme from the bud stage onwards [Jernvall et al., 1994; Kettunen and Thesleff, 1998; Kettunen et al., 2000]. These FGFs signal to the epithelium by activating the ‘b’ isoforms of FGFR1 and FGFR2, which are expressed exclusively in the epithelium [Kettunen et al., 1998]. Conversely, FGF4 and FGF9 produced in the epithelium are presumed to be key mediators of enamel knot activity in coordinating tooth morphogenesis. These signals bind to and activate the ‘c’ isoforms of FGFR1 and FGFR2 expressed in the mesenchyme [Kettunen et al., 1998]. Their major function is to maintain Fgf3 and Fgf10 expression in the dental mesenchyme, which – as discussed above – is thought to be critical for sustaining tooth morphogenesis. Some of the downstream targets of this FGF signaling cascade have been identified (fig. 3). Of particular interest, two genes known to be involved in dental anomalies in humans – Msx1 and Runx2 – are thought to be transcriptional targets of FGF signaling [Bei and Maas, 1998; Åberg et al., 2004]. In addition to being a mesenchymal target of FGF signaling, Runx2, the gene responsible for CCD, has been proposed to directly induce mesenchymal Fgf3 expression [Åberg et al., 2004]. Thus, Runx2 appears to be a critical link in the epithelial-mesenchymal FGF signaling loop. In patients with CCD, which is caused by haploinsufficiency for RUNX2 [Mundlos et al., 1997], supernumerary teeth arise from the permanent teeth, representing a third dentition [Jensen and Kreiborg, 1990]. The underlying molecular mechanism is proposed to result from an incomplete resorption of the dental lamina of the secondary dentition [Lukinmaa et al., 1995]. In Runx2 homozygous null mutant mice, molar development ceases at the bud stage, a time during which there is normally strong expression of Runx2 in the dental mesenchyme [D’Souza et al., 1999]. However, in Runx2 heterozygote mice there appears to be the beginning of successional tooth development: lingual epithelial buds

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Molecular Insights

65

500 µm

a

500 µm

b

500 µm c

500 µm d

Fig. 4. Runx2 mutant tooth phenotype and rescue of arrest in Runx2/Twist-1 double heterozygote mutant mice. a E 16.5 WT. b E 16.5 Runx2 (–/–). c E 16.5 Twist-1 (+/–). d E 16.5 Runx2 (+/–)/Twist-1 (+/–). E = Embryonic

day; WT = wild-type.

with active Shh signaling are present [Wang et al., 2005]. Thus, in both mice and humans, an important role of the Runx2 protein appears to be prevention of excess budding of successional laminae. Interestingly, both FGF receptor levels and Runx2 activity are modulated by Twist-1 [Bialek et al., 2004; Guenou et al., 2005], a transcription factor involved in Saethre-Chotzen syndrome, and patients with this condition have been reported to have dental anomalies [Goho, 1998]. The relief of a functional antagonism between Runx2 and Twist-1 proteins leads to the onset of osteoblast differentiation [Bialek et al., 2004], suggesting a potential mechanism for the formation of supernumerary 66

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teeth in human CCD. Presently, experiments are underway to evaluate whether the formation of accessory tooth buds in Runx2 homozygous null mice are due to a relative overabundance of Twist-1. The latter may result in a prolonged survival of the dental lamina. Tooth development progresses normally in Runx2/Twist-1 double heterozygote mutant mice, suggesting that the in vivo genetic interaction between the two molecules is critical for tooth morphogenesis (fig. 4). Interactions between Runx2 and Twist-1 proteins may thus modulate a variety of events in both development and homeostasis. For each of the major intercellular signaling pathways in development, antagonists have been identified. SignalD’Souza /Klein

ing via FGFRs is inhibited by a molecule called Sprouty (spry), which was first identified in a screen for mutations that affect tracheal branching in Drosophila melanogaster [Hacohen et al., 1998]. Because Sprouty is a negative feedback regulator of FGF signaling, the FGF pathway affects the expression of its own antagonist and thereby limits the range over which FGF signaling is active. When subsequent experiments in Drosophila showed that spry also regulates epidermal growth factor receptor signaling and other receptor tyrosine kinase pathways, the notion arose that spry is a general inhibitor of receptor tyrosine kinase signaling pathways [Casci et al., 1999; Kramer et al., 1999; Reich et al., 1999]. Four mouse genes have sequence similarity to Drosophila spry, and all have human orthologs [de Maximy et al., 1999; Minowada et al., 1999]. Three of the four mouse Sprouty genes, Spry1, Spry2, and Spry4, are expressed at various stages of embryonic development, whereas Spry3 expression has been detected only in the adult [Minowada et al., 1999]. As Sprouty gene expression is induced by FGF signaling, it is observed in association with FGF signaling centers throughout the embryo in numerous developing organs, including the brain, lungs, digestive tract, kidneys and limb buds [Minowada et al., 1999; Zhang et al., 2001]. Sprouty family members act intracellularly to negatively regulate FGF and other receptor tyrosine kinase signaling through diverse biochemical mechanisms, primarily via effects on the mitogen-activated protein kinase pathway [Dikic and Giordano, 2003; Guy et al., 2003; Kim and Bar-Sagi, 2004]. In the tooth buds that form in the wild-type embryonic diastema, the genetic program that normally controls progression from the bud to the cap stage is not active. One mechanism by which diastema bud development is normally suppressed is via inhibition of FGF gene expression, including Fgf4 in the enamel knot and Fgf3 in the dental mesenchyme. Sprouty genes are required to prevent diastema tooth development even though there is little or no FGF gene expression in wildtype diastema buds. It is likely that the normal function of Spry2 is to prevent the relatively low level of signaling via FGF10 produced in diastema bud mesenchyme from inducing/maintaining Shh expression. Likewise, the normal function of Spry4 in the mesenchyme is to prevent any epithelial FGF signals, including FGF4 and FGF9 produced in the adjacent M1 tooth germ, from inducing/maintaining Fgf3 expression. As a result of the combined activities of Spry2 and Spry4, the diastema bud regresses and there are no teeth in the adult diastema. Supernumerary Teeth

Future Challenges and Directions

As is evident from the discussion above, the molecular events that lead to the prolonged survival or proliferation of the dental lamina, and possibly to abnormal division of the tooth bud, are complex and warrant further analysis. Elucidating the precise etiology of supernumerary tooth development will require the use of multifaceted approaches that involve both mouse and human genetic studies. The availability of individuals and families with non-syndromic forms of supernumerary teeth offer a valuable resource to identify genes and underlying mutations that give rise to this condition. Furthermore, information gained from understanding the precise pathogenesis of supernumerary tooth development can be translated to regenerative strategies aimed at bioengineering replacement tooth forms.

Acknowledgments This work is dedicated to the memory of Prof. Robert Gorlin, who provided much inspiration for this work. These studies were supported by grants from the National Institutes of Health to R.N.D’S. (R01 DE 013368) and a Pediatric Scientist Development Program award (K12 HD00850) to O.D.K. The work by O.D.K. was performed in the laboratory of Dr. Gail Martin at the University of California at San Francisco and supported by grants from the National Institutes of Health to Dr. Martin and O.D.K. The authors are grateful to Dr. Martin for mentorship and intellectual guidance, to Dr. Hitesh Kapadia for sharing his clinical insights and data from a patient with CCD, and to Ms. Adriana Cavender for her help in the preparation of the manuscript.

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Cells Tissues Organs 2007;186:70–77 DOI: 10.1159/000102682

Disorders of Human Dentin P. Suzanne Hart a Thomas C. Hart b a

National Human Genome Research Institute, and b Section of Dental and Craniofacial Genetics, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, Md., USA

Key Words Dentin
Dentin dysplasia
Dentinogenesis imperfecta
DSPP
Small integrin-binding ligand N-linked glycoprotein

Abstract Dentin, the most abundant tissue in teeth, is produced by odontoblasts, which differentiate from mesenchymal cells of the dental papilla. Dentinogenesis is a highly controlled process that results in the conversion of unmineralized predentin to mineralized dentin. By weight, 70% of the dentin matrix is mineralized, while the organic phase accounts for 20% and water constitutes the remaining 10%. Type I collagen is the primary component (185%) of the organic portion of dentin. The non-collagenous part of the organic matrix is composed of various proteins, with dentin phosphoprotein predominating, accounting for about 50% of the non-collagenous part. Dentin defects are broadly classified into two major types: dentinogenesis imperfectas (DIs, types I–III) and dentin dysplasias (DDs, types I and II). To date, mutations in DSPP

Abbreviations used in this paper

DD DI DGP DPP DSP DSPP EDS OI SIBLING

dentin dysplasia dentinogenesis imperfecta dentin glycoprotein dentin phosphoprotein dentin sialoprotein dentin sialophosphoprotein Ehlers-Danlos syndrome osteogenesis imperfecta small integrin-binding ligand N-linked glycoprotein

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/cto

have been found to underlie the dentin disorders DI types II and III and DD type II. With the elucidation of the underlying genetic mechanisms has come the realization that the clinical characteristics associated with DSPP mutations appear to represent a continuum of phenotypes. Thus, these disorders should likely be called DSPP-associated dentin defects, with DD type II representing the mild end of the phenotypic spectrum and DI type III representing the severe end. Copyright © 2007 S. Karger AG, Basel

Introduction

Dentin defects are broadly classified into two major types: dentinogenesis imperfectas (DIs, types I–III) and dentin dysplasias (DDs, types I and II). This classification originally proposed in 1973 was based solely upon clinical characteristics and radiographic features without knowledge of the underlying molecular pathophysiology [Shields et al., 1973]. Inheritance of dentin defects is typically autosomal dominant, although autosomal recessive and X-linked cases of dentin defects associated with syndromic conditions are reported. Dentin defects occur as a feature in a number of syndromes, including osteogenesis imperfecta [Steiner et al., 2005], Ehlers-Danlos syndrome (EDS) [De Coster et al., 2007], tumoral calcinosis [Benet-Pagès et al., 2005; Ichikawa et al., 2005; Specktor et al., 2006] and hypophosphatemic rickets [Qin et al., 2004] as well as potentially newly described syndromes [Cauwels et al., 2005; De Coster et al., 2006]. Clinical features associated with DI and DD include brown-blue or opalescent brown appearance of teeth. Enamel fractures off of the dentin,

Dr. P. Suzanne Hart National Human Genome Research Institute, National Institutes of Health Office of the Clinical Director, Building 10 CRC Room 3-2551, 10 Center Drive Bethesda, MD 20892 (USA) Tel. +1 301 594 6051, Fax +1 301 496 7157, E-Mail [email protected]

facilitating attrition of dentin. Radiographically, crowns appear bulbous, pulp chambers are small or obliterated and roots may be narrow with small or obliterated canals. Both primary and secondary teeth may be affected. As the underlying genetic defects are being identified, it is now evident that variable expressivity accounts for some of the differences in clinical expression.

Table 1. Genes underlying dentin defects

Disorder

Location

Gene

DI type I

17q21.33 7q21.3 4q22.1 4q22.1 unknown 4q22.1

COL1A1 COL1A2 DSPP DSPP unknown DSPP

DI type II DI type III DD type I DD type II

Dentinogenesis

Dentin, the most abundant tissue in teeth, is produced by odontoblasts, which differentiate from mesenchymal cells of the dental papilla, and forms the foundation for enamel formation [Arana-Chavez and Massa, 2004]. Dentinogenesis is a highly controlled process that results in the conversion of unmineralized predentin to mineralized dentin. Dentin is heavily mineralized (70% of the matrix by weight) with the organic phase accounting for 20% of the matrix, and the remaining 10% being water [Rajpar et al., 2002]. Type I collagen is the primary component of the organic portion of dentin, accounting for over 85% of the organic phase. The non-collagenous part of the organic matrix is composed of various proteins, with dentin phosphoprotein (DPP) predominating, accounting for about 50% of the noncollagenous part. Odontoblasts differentiate during the crown formation stage of tooth development. The mature odontoblasts line the pulp chamber as a layer of single cells and secrete an organic matrix into the surrounding space. Predentin is an unmineralized zone of 10–40
m that separates the mineralized dentin and the odontoblasts. The odontoblasts secrete type I collagen into the predentin layer where the collagen molecules aggregate into fibrils. At the same time, Ca2+ ions are brought to the mineralization front along with various proteins, such as DPP and dentin sialoprotein (DSP). DPP then serves as a nucleator of mineralization and induces apatite formation. As they secrete the organic matrix, the odontoblasts migrate towards the center of the dental papilla, elongating as they migrate, leaving cytoplasmic extensions, known as the odontoblast process. This section of mineralized tissue is called mantle dentin and is typically 150
m thick. Primary dentin forms as the odontoblasts increase in size and secrete collagen in smaller quantities. The smaller extracellular space results in more tightly packed collagen fibrils that are then mineralized. After completion of root formation, secondary dentin is formed, but the rate of formation is slower than that of primary dentin Dentin Disorders

and not uniform across the tooth. Tertiary dentin is synthesized in response to external stimuli such as caries or trauma. Histologically dentin is permeated by dentinal tubules that run from the pulpal wall to the dentinoenamel junction. The diameter of the tubules is 3.0
m at the pulpal wall, but only 0.06
m at the dentinoenamel junction. These tubules contain odontoblastic processes and dentinal fluid, which contains a variety of proteins including transferrin, tenascin, albumin and proteoglycans [Linde and Goldberg, 1993].

Dentinogenesis Imperfecta

In DI, both the deciduous and permanent teeth are clinically affected, appearing blue-gray or amber brown and opalescent, although the deciduous teeth are often more severely affected. Radiographically, bulbous crowns, narrow roots, and small or obliterated pulp chambers and root canals are seen. Histologically, the dentin has a dysplastic appearance with irregular dentinal tubules and areas lacking dentin tubules. Because of the defect in dentin, enamel is easily broken off, exposing the underlying dentin, leading to accelerated attrition. Three subtypes of DI have been recognized (table 1; fig. 1). DI Type I Individuals with DI type I have a syndromic form of DI. In addition to having DI, they also have osteogenesis imperfecta (OI), an autosomal dominant disorder of bone fragility. OI is further classified into collagenous and non-collagenous forms. All four collagenous OI subtypes can have DI as a feature. DI is more common in types III and IV. DI only occurs in cases of OI due to dominant negative effects (such as missense mutations). In some cases, DI may be the most penetrant clinical finding [Pallos et al., 2001]. Thus, it is important for health care providers to ask about histories of bone fractures (especially Cells Tissues Organs 2007;186:70–77

71

DI type I c

b

DI type II

a

d

DI type III

e

g

DD type I

DD type II

f

j

i

h

Fig. 1. Clinical and radiographic features of DI types I–III (a–g) and DD types I and II (h–j). a–c Individuals with DI type I. In this family, affected members also had OI type I due to a missense alteration in COL1A1 (p.Gly559Cys). a, b The constriction of the crown in the cervical region and almost complete obliteration of the pulp chambers. c The enamel has fractured from the central

and lateral incisors, exposing the underlying yellow-brown dentin. d, e An individual with DI type II. f, g An affected male from the Brandywine cohort. h The extremely short roots characteristic of DD type I. i The phenotype in deciduous teeth that resembles DI type II, but the permanent teeth are normal in color. j The thistle-tube deformity commonly seen in DD type II.

with minimal trauma), joint hyperextensibility, short stature, hearing loss and scleral hue in all patients exhibiting DI. Approximately 90% of individuals with OI types I–IV have an identifiable mutation in COL1A1 or COL1A2, the two genes that encode the chains of type I procollagen [Steiner et al., 2005].

DI Type II The presence of DI without other etiologically related clinical findings (i.e. non-syndromic DI) is classified as DI type II. The clinical and radiographic tooth phenotype is indistinguishable from that seen in syndromic DI type I. Penetrance is almost complete and de novo mutations are

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Hart /Hart

rare. The estimated incidence of DI type II is between 1/6,000 and 1/8,000 [Witkop, 1957]. A few families with DI type II have also had hearing loss, but it is not clear how the mutation in DSPP causes the hearing loss. DI Type III DI type III (OMIM 125500) was initially described in the Brandywine, triracial isolate in southern Maryland. The phenotype in this extended kindred can be traced to a sea captain from Liverpool who came to Maryland in 1732 [Witkop, 1965]. This isolate has the highest reported frequency of dental defects in the world, with approximately 6% having DI [Kim et al., 2005]. Overall, 1 in 15 individuals in this population is estimated to have DI [Witkop, 1957]. In addition to the abnormalities in tooth color and size seen in DI type II, very large pulp chambers and multiple pulp exposures are seen in deciduous teeth in approximately 2% of the Brandywine cohort. Enamel pitting has also been reported in some members of the Brandywine cohort [Levin et al., 1983]. These large pulp chambers led to the designation DI type III, however not all affected family members demonstrate large pulp chambers. Other unrelated families have been reported with these clinical features, demonstrating that the features associated with DI type III are not unique to the Brandywine cohort, and may represent variable clinical expression. Asymptomatic radiolucencies are sometimes seen in teeth with significant attrition. The Brandywine cohort also segregated juvenile periodontitis as an autosomal dominant trait [Boughman et al., 1986]. Linkage analysis supported the existence of a separate gene for juvenile periodontitis, located approximately 17 cM from the locus for DI type III.

Dentin Dysplasia

In DD, the permanent teeth are typically normal in color, but the pulp chambers are obliterated by abnormal dentin. DD Type I DD type I (OMIM 125400) is the rarest of the human dentin disorders, with an estimated incidence of 1 in 100,000. In DD type I, the crowns of deciduous and permanent teeth are normal in shape, form and color, but have short roots with high mobility leading to early exfoliation; frequent periapical radiolucencies are also noted.

Dentin Disorders

DD Type II In DD type II (OMIM 125420), the deciduous teeth have features of DI type II, but the permanent teeth appear normal. Radiographically, pulp cavities show thistle-tube deformity and commonly have pulp stones. Unlike DD type I, root length is normal and frequent periapical radiolucencies are not observed.

Genes Involved in Dentin Formation

COL1A1 and COL1A2 Type I collagen, which comprises the bulk of the organic phase of dentin, is a triple helix containing two pro1(I) and one pro2(I) chain. The two chains are encoded by the COL1A1 and COL1A2 loci, at 17q21.3-q22 and 7q22.1, respectively. Assembly of the collagen triple helix begins at the C-terminus and proceeds towards the N-terminus. Thus mutations at the C-terminus are typically associated with a more severe phenotype than those nearer the N-terminus [Steiner et al., 2005]. OI type I usually results from mutations in COL1A1 that result in haplosufficiency of the pro1(I) chain. Haploinsufficiency leads to half of the amount of type I collagen. No abnormal collagen is produced. These individuals do not have DI. Individuals with OI type I with DI (OI type IB in the classification of Sillence et al. [1979]) have missense mutations in COL1A1. A missense mutation is a nucleotide substitution in the coding region of a gene that changes a nucleotide codon so that it codes for a different amino acid in the protein, resulting in an abnormal collagen fibril. Incorporation of an abnormal collagen fibril interferes with normal triple helical formation and produces OI through a dominant negative effect. In this case, three fourths of the collagen produced are abnormal. Missense mutations also underlie OI types II, III, and IV. The ultimate phenotype depends upon the particular substitution, with substitutions of glycine in the pro1(I) chain by arginine, valine, glutamic acid, aspartic acid, and tryptophan usually resulting in OI type II if they occur in the carboxylterminal 70% of the triple helix. DI may or may not be present in these cases of OI due to dominant negative effects. Recently, a patient with type I EDS, a disorder characterized by joint hypermobility, skin hyperextensibility and tissue fragility, in whom a C to T transition (c.934C1T; p.R134C) in COL1A1 was identified, was shown to have structural dentin defects despite a clinically unaffected dentition [De Coster et al., 2007]. This same mutation has been reported in 2 additional type I EDS patients, but no histological and ultraCells Tissues Organs 2007;186:70–77

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structural studies have been performed on teeth from those patients [Nuytinck et al., 2000]. SIBLINGs The SIBLING (small integrin-binding ligand N-linked glycoprotein) family of proteins includes dentin sialophosphoprotein (DSPP), osteopontin [also known as SPP1 (secreted phosphoprotein)], IBSP (integrin binding sialoprotein), DMP1 (dentin matrix protein 1), and MEPE (matrix extracellular phosphoglycoprotein) [Fisher et al., 2001]. Although all members were originally isolated from tooth or bone, it is now apparent that at least some SIBLINGs are expressed in non-mineralized tissues. The SIBLING genes are syntenic (on the same chromosome), clustered in a 0.37-Mb region on chromosome 4q. The encoded proteins contain an integrin binding tripeptide (Arg-Gly-Asp) and several conserved N-glycosylation and phosphorylation sites. The expression levels of the different SIBLINGs vary greatly between tissues. For example, OPN is expressed in bone at a level 70 times higher than that in dentin [Butler et al., 2003]. Although all SIBLINGS are thought to function in the mineralization process, their expression outside of mineralized tissues suggests additional roles. The recent finding of DSPP expression in embryonic kidney and lung has led to the hypothesis that DSPP may regulate branching morphogenesis [Alvares et al., 2006]. The only two SIBLINGS known to be associated with human genetic disorders are DMP1 and DSPP. Mutations in DMP1 were recently shown to underlie autosomal recessive hypophosphatemic rickets [Feng et al., 2006; Lorenz-Depiereux et al., 2006]. DSPP DSPP is located at 4q22.1 and consists of five exons spanning 8,343 bp. DSPP is expressed in dentin at levels that are hundreds of times higher than that of other tissues. Three distinct protein products result from proteolytic cleavage of the initial 1253-amino acid protein product, DSPP: DSP (amino acids 16–374), dentin glycoprotein (DGP; amino acids 375–462), and DPP (amino acids 463–1253; fig. 2). The initial cleavage by a yet unidentified protease releases DPP, a process that occurs almost immediately upon formation of DSPP [Yamakoshi et al., 2006]. Matrix metalloproteinase (MMP) 20 then cleaves DSP-DGP to generate DSP and DGP, while MMP2 catalyzes other cleavages on DSP. DPP is a highly repetitive protein with a great degree of phosphorylation and is thought to be involved in the nucleation and control of mineralization in dentin [George et al., 1996]. DPP contains repeats of aspartic acid (Asp) and phosphoserine 74

Cells Tissues Organs 2007;186:70–77

(Pse) mainly as (Asp-Pse-Pse) and (Asp-Pse). There is extensive variability in the number of repeats in normal individuals [Fisher, pers. commun.]. After synthesis and cleavage, DPP moves quickly to the mineralization front and binds to collagen. DSP is a heavily glycosylated protein that forms covalent dimers by intermolecular disulfide bridges [Yamakoshi et al., 2005], but whose function is not entirely known. DGP was recently isolated from porcine odontoblasts and found to contain four phosphorylated serines and one N-glycosylated asparagine. Given the posttranslational modifications and amino acid conservation across species, it has been proposed that DGP plays a role in dentin biomineralization [Yamakoshi et al., 2005]. DSPP knockout mice develop abnormalities similar to those seen in DI type III [Sreenath et al., 2003]. Histologically, large areas of unmineralized dentin were seen as well as an irregular border between the unmineralized and mineralized dentin. In addition, there were areas of partially mineralized dentin between areas that were fully mineralized. To date only mutations in DSPP have been found to underlie DI types II and III and DD type II. Although 11 mutations have been reported in the literature, two of the published disease-causing mutations are actually normal sequence variants: p.R68W and Del1160–1171; ins1198– 1199 (table 2). Holappa et al. [2006] found the p.R68W change in 15% of Finnish controls. Our laboratory found p.R68W with allele frequencies of 6 and 16% in Caucasian and African-American controls, respectively. We have also studied a branch of the Brandywine family reported by Dong et al. [2005]. Although we identify the same alteration in the hypervariable region, there is considerable variation in this region in normal controls, with similar deletions and insertions found. This normal variation is apparent when trying to align the genomic sequence with the DSPP cDNA reference sequence [Beattie et al., 2006]. We identified a second nucleotide change in the genomic sequence (g.49C1T) of the DSPP gene in this family, predicted to result in a p.P17S missense alteration, that segregates with the phenotype and causes endoplasmic reticulum retention of the protein in a functional assay. Thus, we propose that the p.P17S substitution is the true causative mutation in the Brandywine family. Of the nine confirmed DSPP mutations, all occur within the DSP region of DSPP. It should be pointed out that the effect of the mutation on the protein is simply a prediction, as more than one consequence may be possible. For example, the c.52G1T alteration might result in the missense alteration, p.V18F. On the other hand, this alteration occurs as Hart /Hart

c.16T>G

c.44C>T c.49C>A or T

c.52G>T c.133C>T c.135+1C>G or A

c.52–3C>G or A

aa 1–17

aa 18–43

aa 46–374

aa 463–1253

Fig. 2. Diagram of the DSPP gene and corresponding protein structure. Mutations are shown above the genomic structure. Gray areas correspond to untranslated regions. The white box corresponds to the signal peptide [amino acids (aa) 1–15]. DSP is shown as black regions (aa 16–382). DGP, represented by a stippled box, corresponds to aa 383–462. The highly repetitive protein, DPP, is shown as a striped box (aa 463–1253).

Table 2. Mutations in DSPP resulting in dentin defects

Protein

cDNA

Genomic

Diagnosis

References

Exon 2

p.Y6D p.A15V p.P17T p.P17S

c.16T>G c.44C>T c.49C>A c.49C>T

g.16T>G g.44C>T g.49C>A g.49C>T

DD-II DI-II DI-II DI-II

Rajpar et al. [2002] Malmgren et al. [2004] Xiao et al. [2001] this report [2007]

Intron 2

p.V18_Q45del

c.52-3C>G c.52-3C>A

g.1194C>G g.1194C>A

DI-II

Kim et al. [2004] Holappa et al. [2006]

Exon 3

p.V18F or p.V18_Q45del

c.52G>T

g.1197G>T

DI-II DI-III DI-III

Xiao et al. [2001] Kim et al. [2005] Song et al. [2006] Holappa et al. [2006]

p.Q45X

c.133C>T

g.1278C>T

DI-II

Zhang et al. [2001] Song et al. [2006]

c.135+1G>A

g.1281G>A

DI-II

Xiao et al. [2001]

c.202A>T

g.1480A>T

DI-II

Malmgren et al. [2004]

DI-III

Dong et al. [2005]

Intron 3

p.V18_Q45del a

Exon 4

p.R68W

Exon 5

Del1160-1171 Ins1198-1199a

All numbering assumes the A of the ATG start codon as nucleotide 1. The reference is NM_014208. This alteration has been shown to be a normal variant and is not disease causing.

a

the first nucleotide of exon 3 and may thus disrupt splicing, causing skipping of exon 3 (p.V18_Q45del). Site-directed mutagenesis was used to introduce each of the nine mutations and the p.R68W polymorphism into the human DSP mRNA which were then transfected into odontoblast cells. The ability of the cells to secrete DSP was evaluated by Western blot analysis. This functional assay demonstrated that with the exception of the nonsense mutation, p.Q45X, all other mutations are associated with

endoplasmic reticulum retention, consistent with a dominant negative mutation [Choi et al., submitted]. The finding that the same mutation produces different phenotypes suggests that factors other than the specific DSPP mutation contribute to the ultimate phenotype. There are some families with dentin defects linked to 4q22.1 in which no DSPP mutation can be identified [Beattie et al., 2006]. Family A in the paper by Malmgren et al. [2004] is also an example of linkage to 4q22.1 with-

Dentin Disorders

Cells Tissues Organs 2007;186:70–77

75

out the underlying defect identified since the predicted causative mutation (p.R68W) has since been shown to be simply a non-synonymous single nucleotide polymorphism that segregated with the phenotype. Whether these families have undetected mutations in DSPP, e.g. intronic changes, promoter alterations, deletions of exons, or mutations in the hypervariable region which is typically not analyzed, or have involvement of another gene in the 4q22.1 region remains to be determined. Proteases As noted above, MMP2 and MMP20 were recently shown to be involved in the processing of DSPP [Yamakoshi et al., 2006], specifically to generate DSP and DGP. Mutations in MMP20 result in amelogenesis imperfecta without an obvious dentin abnormality [Kim et al., 2005; Ozdemir et al., 2005]. It has been suggested that molecular redundancy may explain this lack of phenotype [Yamakoshi et al., 2006]. MMP8 was recently identified as the major collagenase in human dentin [Sulkala et al., 2007]. Other metalloproteases synthesized by odontoblasts include MMP9, and membrane-bound MMP14 (MT1-MMP). KLK4 has also been detected in dentin extracts but its role in processing dentinal proteins is not known [Yamakoshi et al., 2006], but its expression by odontoblasts might be associated with the hypermineralization of enamel above the dentinoenamel junction [Fukae et al., 2002]. Other Genes Expressed in Dentin A variety of other genes are expressed in dentin. Many of these genes, if mutated, would not be expected to give rise to isolated enamel defects. Patients with EDS type VIIC, who have a defect in the ADAMTS2 gene which encodes the enzyme responsible for removing the N-terminal propeptide in procollagen types I–III, exhibit dentin defects [De Coster et al., 2006]. Recently, HMGB1 (high mobility group box 1) was shown to be highly expressed in

dentin at regions undergoing mineralization [Sugars et al., 2007]. HMGB1–/– mice die within 24 h of birth due to hypoglycemia [Calogero et al., 1999]. PHEX, mutated in Xlinked hypophosphatemic rickets (XLH), encodes an endopeptidase postulated to play a role in the processing of DMP1 and DSPP [Qin et al., 2004]. Loss of function mutations in the gene encoding fibroblast growth factor 23 (FGF23), a phosphaturic protein, result in autosomal recessive hyperphosphatemic familial tumoral calcinosis (OMIM 211900) [Benet-Pagès et al., 2005] while gain of function mutations produce autosomal dominant hypophosphatemic rickets (OMIM 193100) [ADHR Consortium, 2000]. Hyperphosphatemic familial tumoral calcinosis can also result from mutations in GALNT3 which encodes a glycosyltransferase involved in initiating O-glycosylation [Ichikawa et al., 2005; Specktor et al., 2006]. The demonstration that both hyper- and hypophosphatemic disorders have dentin defects underlies the importance of phosphate homeostasis in normal dentin development.

Proposal for New Classification of Dentin Defects

The finding of differing dentin phenotypes within a family highlights the variable expressivity often associated with autosomal dominant disorders. The clinical phenotypes associated with DSPP mutations appear to represent a continuum of phenotypes. Thus, these disorders should likely be called DSPP-associated dentin defects, with DD type II representing the mild end of the phenotypic spectrum and DI type III representing the severe end.

Acknowledgement This research was supported by the Intramural Research Program of the NIH, NHGRI and NIDCR.

References ADHR Consortium (2000) Autosomal dominant hypophosphatemic rickets is associated with mutations in FGF23. Nat Genet 26: 345– 348. Alvares, K., Y.S. Kanwar, A. Veis (2006) Expression and potential role of dentin phosphophoryn (DPP) in mouse embryonic tissues involved in epithelial-mesenchymal interactions and branching morphogenesis. Dev Dyn 235: 2980–2990. Arana-Chavez, V.E., L.F. Massa (2004) Odontoblasts: the cells forming and maintaining

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Boughman, J.A., S.L. Halloran, D. Roulston, S. Schwartz, J.B. Suzuki, L.R. Weitkamp, R.E. Wenk, R. Wooten, M.M. Cohen (1986) An autosomal-dominant form of juvenile periodontitis: its localization to chromosome 4 and linkage to dentinogenesis imperfecta and Gc. J Craniofac Genet Dev Biol 6: 341–350. Butler, W.T., J.C. Brunn, C. Qin (2003) Dentin extracellular matrix (ECM) proteins: comparison to bone ECM and contribution to dynamics of dentinogenesis. Connect Tissue Res 44: 171–178.

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Calogero, S., F. Grassi, A. Aguzzi, T. Voigtlander, P. Ferrier, S. Ferrari, M.E. Bianchi (1999) The lack of chromosomal protein Hmg1 does not disrupt cell growth but causes lethal hypoglycaemia in newborn mice. Nat Genet 22: 276–280. Cauwels, R.G., P.J. De Coster, G.R. Mortier, L.A. Marks, L.C. Martens (2005) Dentinogenesis imperfecta associated with short stature, hearing loss and mental retardation: a new syndrome with autosomal recessive inheritance? J Oral Pathol Med 34: 444–446. De Coster, P.J., R.M.H. Verbeeck, V. Holthaus, L.C. Martens, A. Vral (2006) Seckel syndrome associated with oligodontia, microdontia, enamel hypoplasia, delayed eruption, and dentin dysmineralization: a new variant. J Oral Pathol Med 35: 639–641. De Coster, P.J., M. Cornelissen, A. De Paepe, L.C. Martens, A. Vral (2007) Abnormal dentin structure in two novel gene mutations [COL1A1, Arg134Cys] and [ADAMTS2, Trp795-to-ter] causing rare type I collagen disorders. Arch Oral Biol 52: 101–109. Dong, J., T.T. Gu, L. Jeffords, M. MacDougall (2005) Dentin phosphoprotein compound mutation in dentin sialophosphoprotein causes dentinogenesis imperfecta type III. Am J Med Genet 132A: 305–309. Feng, J.Q., L.M. Ward, S. Liu, Y. Lu, Y. Xie, B. Yuan, X. Yu, F. Rauch, S.I. Davis, S. Zhang, H. Rios, M.K. Drezner, L.D. Quarles, L.F. Bonewald, K.E. White (2006) Loss of DMP1 causes rickets and osteomalacia and identifies a role for osteocytes in mineral metabolism. Nat Genet 38: 1310–1315.. Fisher, L.W., D.A. Torchia, B. Fohr, M.F. Young (2001) Flexible structures of SIBLING proteins, bone sialoprotein, and osteopontin. Biochem Biophys Res Commun 280: 460– 465. Fukae, M., T. Tanabe, T. Nagano, H. Ando, Y. Yamakoshi, M. Yamada, J.P. Simmer, S. Oida (2002) Odontoblasts enhance the maturation of enamel crystals by secreting EMSP1 at the enamel-dentin junction. J Dent Res 81: 668–672. George, A., L. Bannon, B. Sabsay, J.W. Dillon, J. Malone, A. Veis, N.A. Jenkins, D.J. Gilbert, N.G. Copeland (1996) The carboxyl-terminal domain of phosphophoryn contains unique extended triplet amino acid repeat sequences forming ordered carboxyl-phosphate interaction ridges that may be essential in the biomineralization process. J Biol Chem 271: 32869–32873. Holappa, H., P. Nieminen, L. Tolva, P.-L. Lukinmaa, S. Alaluusua (2006) Splicing site mutations in dentin sialophosphoprotein causing dentinogenesis imperfecta type II. Eur J Oral Sci 114: 381–384. Ichikawa, S., K.W. Lyles, M.J. Econs (2005) A novel GALNT3 mutation in a pseudoautosomal dominant form of tumoral calcinosis: evidence that the disorder is autosomal recessive. J Clin Endocrinol Metab 90: 2420– 2423.

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Kim, J.-W., S.-H. Nam, K.-T. Jang, S.-H. Lee, C.-C. Kim, S.-H. Hahn, J.C.-C. Hu, J.P. Simmer (2004) A novel splice acceptor mutation in the DSPP gene causing dentinogenesis imperfecta type II. Hum Genet 115: 248–254. Kim, J.-W., J.C.-C. Hu, J.-I. Lee, S.-K. Moon, Y.-J. Kim, K.-T. Jang, S.-H. Lee, C.-C. Kim, S.-H. Hahn, J.P. Simmer (2005) Mutational hot spot in the DSPP gene causing dentinogenesis imperfecta type II. Hum Genet 116: 186–191. Kim, J.W., J.P. Simmer, T.C. Hart, P.S. Hart, M.D. Ramaswami, J.D. Bartlett, J.C. Hu (2005) MMP-20 mutation in autosomal recessive pigmented hypomaturation amelogenesis imperfecta. J Med Genet 42: 271–275. Levin, L.S., S.H. Leaf, R.J. Jelmini, J.J. Rose, K.N. Rosenbaum (1983) Dentinogenesis imperfecta in the Brandywine isolate (DI type III): clinical, radiologic, and scanning electron microscopic studies of the dentition. Oral Surg Oral Med Oral Pathol 56: 267–274. Linde, A., M. Goldberg (1993) Dentinogenesis. Crit Rev Oral Biol Med 4: 679–728. Lorenz-Depiereux, B., M. Bastepe, A. BenetPagès, M. Amyere, J. Wagenstaller, U. Müller-Barth, K. Badenhoop, S.M. Kaiser, R.S. Rittmaster, A.H. Shlossberg, J.L. Olivares, C. Loris, F.J. Ramos, F. Glorieux, M. Vikkula, H. Jüppner, T.M. Strom (2006) DMP1 mutations in autosomal recessive hypophosphatemia implicate a bone matrix protein in the regulation of phosphate homeostasis. Nat Genet 38: 1248–1250. Malmgren, B., S. Lindskog, A. Elgadi, S. Norgren (2004) Clinical, histopathologic, and genetic investigation in two large families with dentinogenesis imperfecta type II. Hum Genet 114: 491–498. Nuytinck, L., M. Freund, L. Lagae, G.E. Pierard, T. Hermanns-Le, A. De Paepe (2000). Classical Ehlers-Danlos syndrome caused by a mutation in type I collagen. Am J Hum Genet 66: 1398–1402. Ozdemir, D., P.S. Hart, O.H. Ryu, S.J. Choi, M. Ozdemir-Karatas, E. Firatli, N. Piesco, T.C. Hart (2005) MMP20 active-site mutation in hypomaturation amelogenesis imperfecta. J Dent Res 84: 1031–1035. Pallos, D., P.S. Hart, J.R. Cortelli, S. Vian, J.T. Wright, J. Korkko, D. Brunoni, T.C. Hart (2001) Novel COL1A1 (G559C) associated with mild osteogenesis imperfecta and dentinogenesis imperfecta. Arch Oral Biol 46: 459–470. Qin, C., O. Baba, W.T. Butler (2004) Post-translational modifications of SIBLING proteins and their roles in osteogenesis and dentinogenesis. Crit Rev Oral Biol Med 15: 126–136. Rajpar, M.H., M.J. Koch, R.M. Davies, K.T. Mellody, C.M. Kielty, M.J. Dixon (2002) Mutation of the signal peptide region of the bicistronic gene DSPP affects translocation to the endoplasmic reticulum and results in defective dentine biomineralization. Hum Mol Genet 11: 2559–2565.

Shields, E.D., D. Bixler, A.M. El-Kafrawy (1973) A proposed classification for heritable human dentine defects with a description of a new entity. Arch Oral Biol 18: 543–553. Sillence, D.O., A. Senn, D.M. Danks (1979) Genetic heterogeneity in osteogenesis imperfecta. J Med Genet 16: 101–116. Song, Y., C. Wang, B. Peng, X. Ye, G. Zhao, M. Fan, Q. Fu, Z. Bian (2006) Phenotypes and genotypes in 2 DGI families with different DSPP mutations. Oral Surg Oral Med Pathol Oral Radiol Endod 102: 360–374. Specktor, P., J.G. Cooper, M. Indelman, E. Sprecher (2006) Hyperphosphatemic familial tumoral calcinosis caused by a mutation in GALNT3 in a European kindred. J Hum Genet 51: 487–490. Sreenath, T., T. Thyagaragan, B. Hall, C. Longenecker, R. D’Souza, S. Hong, J.T. Wright, M. MacDougall, J. Sauk, A.B. Kulkarni (2003). Dentin sialophosphoprotein knockout mouse teeth display widened predentin zone and develop defective dentin mineralization similar to human dentinogenesis imperfecta type III. J Biol Chem 278: 24874–24880. Steiner, R.D., M.G. Pepin, P.H. Byers (updated 1-28-05) Osteogenesis imperfecta; in GeneReviews at GeneTests: Medical Genetics Information Resource (database online). Copyright, University of Washington, Seattle. 1997–2006. http://www.genetests.org (accessed 10-1-06). Sugars, R., E. Karlström, C. Christersson, M.-L. Olsson, M. Wendel, K. Fried (2007) Expression of HMGB1 during tooth development. Cell Tissue Res 327: 511–519. Sulkala, M., T. Tervahartiala, T. Sorsa, M. Larmas, T. Salo, L. Tjäderhane (2007) Matrix metalloproteinase-8 (MMP-8) is the major collagenase in human dentin. Arch Oral Biol 52:121–127. Witkop, C.J., Jr. (1957) Hereditary defects in enamel and dentin. Acta Genet 7: 236–239. Witkop, C.J., Jr. (1965) Genetic diseases of the oral cavity; in Tiecke, R.W. (ed) Oral Pathology. New York, McGraw Hill, pp 786–843. Xiao, S., C. Yu, X. Chou, W. Yuan, Y. Wang, L. Bu, G. Fu, M. Qian, J. Yang, Y. Shih, L. Hu, B. Han, Z. Wang, W. Huang, J. Liu, Z. Chen, G. Zhao, X. Kong (2001) Dentinogenesis imperfecta 1 with or without progressive hearing loss is associated with distinct mutations in DSPP. Nat Genet 27: 201–204. Yamakoshi, Y., J.C.-C. Hu, M. Fukae, H. Zhang, J.P. Simmer (2005) Dentin glycoprotein: the protein in the middle of the dentin sialophosphoprotein chimera. J Biol Chem 280: 17471–17479. Yamakoshi, Y., J.C.-C. Hu, T. Iwata, K. Kobayashi, M. Fukae, J.P. Simmer (2006) Dentin sialophosphoprotein is processed by MMP-2 and MMP-20 in vitro and in vivo. J Biol Chem 281: 38235–38243. Zhang, X., J. Zhao, C. Li, S. Gao, C. Qui, P. Liu, G. Wu, B. Qiang, W.H.Y. Lo, Y. Shen (2001) DSPP mutation in dentinogenesis imperfecta Shields type II. Nat Genet 27: 151–152.

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Cells Tissues Organs 2007;186:78–85 DOI: 10.1159/000102683

Enamel Formation and Amelogenesis Imperfecta Jan C.-C. Hu Yong-Hee P. Chun Turki Al Hazzazzi James P. Simmer University of Michigan School of Dentistry, Ann Arbor, Mich., USA

Key Words Amelogenesis imperfecta
Amelogenin
Enamel
Enamelin
Enamelysin
Kallikrein 4

Abstract Dental enamel is the epithelial-derived hard tissue covering the crowns of teeth. It is the most highly mineralized and hardest tissue in the body. Dental enamel is acellular and has no physiological means of repair outside of the protective and remineralization potential provided by saliva. Enamel is comprised of highly organized hydroxyapatite crystals that form in a defined extracellular space, the contents of which are supplied and regulated by ameloblasts. The entire process is under genetic instruction. The genetic control of amelogenesis is poorly understood, but requires the activities of multiple components that are uniquely important for dental enamel formation. Amelogenesis imperfecta (AI) is a collective designation for the variety of inherited conditions displaying isolated enamel malformations, but the designation is also used to indicate the presence of an enamel phenotype in syndromes. Recently, genetic studies have demonstrated the importance of genes encoding enamel matrix proteins in the etiology of isolated AI. Here we review the essential elements of dental enamel formation and the results of genetic analyses that have identified disease-causing mutations in genes encoding enamel matrix proteins. In addition, we provide a fresh perspective on the roles matrix proteins play in catalyzing the biomineralization of dental enamel. Copyright © 2007 S. Karger AG, Basel

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/cto

Introduction

In this presentation we focus on the developmental biology and genetics of dental malformations. For related articles, the reader is referred to a number of excellent reviews on dental enamel formation [Simmer and Fincham, 1995; Smith, 1998; Fincham et al., 1999; Nanci, 2003] and on inherited enamel defects [Witkop and Sauk, 1976; Witkop, 1988; Hu et al., 2005; Stephanopoulos et al., 2005].

Abbreviations used in this paper

AI AIHHT AMBN AMELX AMELY AMTN DLX3 KLK4 LAMA3 LAMB3 LAMC2 MMP20

amelogenesis imperfecta amelogenesis imperfecta hypoplastic-hypomaturation with taurodontism ameloblastin gene amelogenin gene on X chromosome amelogenin gene on Y chromosome amelotin gene distal-less homeobox 3 kallikrein 4 gene laminin
3 gene laminin 3 gene laminin 2 gene enamelysin gene

Dr. James P. Simmer Department of Biologic and Materials Sciences University of Michigan Dental Research Laboratory, 1210 Eisenhower Place Ann Arbor, MI 48108 (USA) Tel. +1 734 975 9318, Fax +1 734 975 9329, E-Mail [email protected]

Amelogenesis Imperfecta

Inherited enamel defects that occur in the absence of a generalized syndrome are collectively designated as amelogenesis imperfecta (AI). The range of enamel malformations observed in patients with AI is classified according to the thickness, hardness and smoothness of the affected enamel. Differences in these parameters are believed to reflect differences in the timing, during amelogenesis, when the disruption occurred. During tooth formation, enamel first appears on the surface of recently deposited dentin, at the forming dentinoenamel junction. Flaws in the dentinoenamel junction can result in an enamel layer that tends to shear from the underlying dentin. During the secretory stage of amelogenesis, the thickness of the enamel layer increases by appositional growth: the continuous deposition of enamel proteins on the existing enamel surface, which is accompanied by the radial movement of the formative cells (ameloblasts) away from the point of secretion and is associated with the elongation of enamel crystallites. Insufficient appositional growth and associated crystal elongation leaves the enamel layer pathologically thin, or hypoplastic. The most severe form of hypoplastic AI is enamel agenesis, where there is almost no clinical or radiographic evidence of enamel. The teeth are yellowish brown in color, rough in texture, and widely spaced. In normal tooth development, when the enamel crystals achieve their final length (and the enamel layer itself achieves its final thickness), the organic matrix separating individual enamel crystallites is degraded and reabsorbed. The enamel layer then hardens or ‘matures’ as mineral deposits on the sides of the crystals until adjacent crystallites contact. A failure to properly remove the organic matrix and promote the hardening of the enamel layer leads to pathologically soft or hypomaturation forms of AI. The dental crowns are of normal size and contact adjacent teeth, but the mottled, brownish-yellow enamel is soft and has a radiodensity approaching that of dentin. In the third classification type, hypocalcified AI, the failure in mineralization is most extreme. The enamel layer may be of normal thickness, but is rough and soft and wears away quickly following tooth eruption. Patients with hypocalcified enamel form calculus rapidly and develop acute and chronic periodontitis. Often patients with inherited enamel defects experience difficulty maintaining oral hygiene, have lower self-esteem due to the poor appearance of their teeth, and perceive themselves as having an inferior quality of life [Coffield et al., 2005]. An understanding of tooth formation provides some insight into the reasons why inherDental Malformations

ited enamel malformations are manifested as they are into hypomaturation, hypocalcified, and hypomaturation types.

Early Tooth Formation

Teeth form as a result of a series of interactions between oral epithelium and neural crest-derived ectomesenchyme [Thesleff and Sharpe, 1997; Cobourne and Sharpe, 2003; Tucker and Sharpe, 2004]. Early tooth development advances through initiation and bud stages. Genetic disturbances in early odontogenesis can arrest tooth development causing familial tooth agenesis [for a review please see D’Souza and Klein, pp 60–69, this issue]. During the cap stage, the primary enamel knot serves as a signaling center [Thesleff and Jernvall, 1997]. In multi-cusped teeth, secondary enamel knots guide the differentiation of the enamel epithelium at each cusp tip during the bell and crown formation stages [Thesleff et al., 2001; Matalova et al., 2005]. During the crown formation stage, cellular differentiation is completed and fully differentiated odontoblasts and ameloblasts deposit dentin and enamel matrices. Odontoblasts differentiate first and secrete a collagen-rich predentin matrix directly beneath the epithelial-derived basal lamina, but dentin mineralization does not occur until the basement membrane material is removed. Differentiating ameloblasts (pre- and presecretory ameloblasts) start expressing small amounts of enamel proteins even before the basal lamina disintegrates. As the basal lamina fragments, ameloblasts send cytoplasmic projections through the gaps. Dentin starts to mineralize with the disappearance of the basal lamina, as the apical surfaces of ameloblasts associate with the superficial collagen fibrils of the mantle dentin. Patches of enamel matrix proteins appear in quantity on the irregular dentin surface and enamel mineralization begins. As the (now) secretory ameloblasts recede, the patches grow larger and merge until a continuous and uniform layer of initial enamel is deposited. This initial enamel layer is aprismatic, i.e. not separated into rod and interrod enamel. Secretory ameloblasts form a novel cell extension called a Tomes’ process at their secretory ends. This extension has secretory and nonsecretory regions and provides the architectural basis for organizing enamel crystals into rod and interrod enamel, which differ from each other in the orientation of their crystals [Meckel et al., 1965; Cevc et al., 1980; Fejerskov and Thylstrup, 1986].

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79

Fig. 1. Ameloblast changes during enamel formation. The epithelial cells of the inner enamel epithelium (1) rest on a basement membrane containing laminin. These cells increase in length as differentiating ameloblasts above the predentin matrix (2). Presecretory ameloblasts send processes through the degenerating basement membrane as they initiate the secretion of enamel proteins on the villous surface of mineralizing dentin (3). After establishing the dentinoenamel junction and mineralizing a thin layer of aprismatic enamel, secretory ameloblasts develop a secretory specialization, or Tomes’ process. Along the secretory face of the Tomes’ process, in place of the absent basement membrane, secretory ameloblasts secrete proteins at a mineralization front where the enamel crystals grow in length (4). Each enamel rod follows a retreating Tomes’ process from a single ameloblast. At

The Basal Lamina

Prior to the secretory stage of dental enamel formation, the inner enamel epithelium rests on a basal lamina containing type IV collagen and the glycoprotein laminin (types 1 and 5), which are common to the basal lamina of many epithelial tissues [Salmivirta et al., 1997]. Laminin 5 is comprised of three polypeptides (
3, 3, and 2) expressed from LAMA3, LAMB3, and LAMC2. Junctional epidermolysis bullosa is a syndrome featuring blistering skin tissues with AI, and is caused by defects in LAMB3 [Buchroithner et al., 2004]. While syndromic enamel defects are not the focus of this presentation, this disease illustrates the importance of the basal lamina to enamel formation, and yet, degradation of the basal lamina is necessary for amelogenesis to occur. A novel basal 80

Cells Tissues Organs 2007;186:78–85

the end of the secretory stage, ameloblasts lose their Tomes’ process and produce a thin layer of aprismatic enamel (5). At this point the enamel has achieved its final thickness. During the transition stage, the ameloblasts undergo a major restructuring that diminishes their secretory activity and changes the types of proteins secreted (6). KLK4 is secreted, which degrades the accumulated protein matrix and amelotin (AMTN) is secreted as part of the new basement membrane. During the maturation stage ameloblasts modulate between ruffled and smooth-ended phases (7). Their activities harden the enamel layer by promoting the deposition of mineral on the sides of enamel crystals laid down during the secretory stage. The histology of the developing tooth is adapted from Uchida et al. [1991].

lamina containing the tooth-specific component amelotin (AMTN) is established at the onset of the maturation stage and persists in the junctional epithelium following tooth eruption [Moffatt et al., 2006].

The Secretory Stage of Amelogenesis

Ameloblasts secrete enamel proteins on top of and around existing crystallites (dentin crystals initially and enamel crystals thereafter) and into the space that was previously occupied by the basal lamina. Secreted enamel proteins concentrate along the ameloblast secretory membrane and create a mineralization front, which differs from those of dentin and bone by lacking an unmineralized layer (i.e. there is no pre-enamel analogous to Hu/Chun/Al Hazzazzi/Simmer

predentin or osteoid). The mineralization front retreats with the Tomes’ process as the enamel crystals grow in length (
4 m/day) [Risnes, 1986], and the ameloblasts continue their secretion of enamel proteins (fig. 1). Enamel crystals elongate in diurnal cycles leaving incremental lines (in some mammals, including humans) corresponding to daily increases in the enamel layer. Throughout the secretory stage enamel crystals grow primarily in length and the enamel layer thickens. Disturbances during the secretory stage cause the final enamel layer to be thinner, or hypoplastic. Systemic or environmental disturbances cause horizontal bands of enamel hypoplasia related to the timing of the disturbance [Hillson and Bond, 1997; Reid and Dean, 2000]. Ultimately, it is the final length of the enamel crystals that determines the thickness of the enamel layer as a whole, and the final length of the crystals is determined by the length of time ameloblasts continue in the secretory stage, or how long they continue to secrete enamel proteins (sustain appositional growth). Disturbances during the secretory stage of amelogenesis impede crystal elongation and result in pathologically thin, or hypoplastic enamel. Defects in the genes encoding enamel proteins tend to disturb the secretory stage elongation of enamel crystals and cause hypoplastic forms of AI [Hu and Yamakoshi, 2003; Kim et al., 2006].

Amelogenin (AMELX, Xp22.3)

The most abundant enamel matrix protein is amelogenin. It comprises approximately 80–90% of total enamel protein [Fincham et al., 1999]. Amelogenin is secreted as a variety of isoforms, the major one having a molecular weight around 25 kDa. Following its secretion, amelogenin may quickly pass through the mineralization front [Nanci et al., 1998] and assemble into nanospheres, about 20 nm in diameter, that are thought to regulate crystal spacing [Fincham et al., 1994; Fincham and Simmer, 1997]. In humans, amelogenin is expressed from genes on the sex chromosomes: AMELX and AMELY [Lau et al., 1989; Nakahori et al., 1991], but only 10% of the amelogenin mRNA transcripts are expressed from AMELY [Salido et al., 1992]. Mutational analyses have identified 15 AMELX mutations in kindreds afflicted with X-linked AI (fig. 2). X-linked forms of hypoplastic and hypomaturation AI often show a distinctive phenotype in affected females, where the enamel displays alternating vertical bands of normal and defective enamel. This phenotype is called ‘lyonization’, Dental Malformations

2 1

3/4

5

2 1-18

7

6 3

4

8

5

19-34 34-48 49-64

14 AMELX 15

10 12 9 11 13 6

7

65-204

205

Number Mutation References 1 p.0 [Lagerström et al., 1991] 2 p.M1T [Kim et al., 2004] 3 p.W4S [Kim et al., 2004] 4 p.W4X [Sekiguchi et al., 2001b] 5 p.I5-A8delinsT [Lagerstrom-Fermer et al., 1995] 6 p.T51I [Lench and Winter, 1995] [Aldred et al., 1992] 7 p.P52fsX53 [Lench et al., 1994] 8 p.P52R [Kida et al., 2007] [Collier et al., 1997] 9 p.P70T [Hart et al., 2000] [Ravassipour et al., 2000] 10 p.H77L [Hart et al., 2002] 11 p.H129fs187 [Sekiguchi et al., 2001a] 12 p.Y141fsX187 [Greene et al., 2002] 13 p.P158fsX187 [Lench and Winter, 1995] [Kindelan et al., 2000] p.L181fsX187 14 [Hart et al., 2002] 15 p.E191X [Lench and Winter, 1995] Fig. 2. AMELX mutations causing X-linked AI. The intron/exon structure of AMELX is shown at the top. The seven AMELX exons are indicated by numbered boxes; the six introns are the lines connecting the exons. The numbers below each exon show the range of amino acids encoded by that exon. Characterized AMELX mutations are numbered in bold starting at the 5 end of the gene. The 15 known mutations and their references are listed below the gene diagram. Mutation 1 was a large deletion that is not shown on the gene diagram. The abbreviations for the mutations indicate the predicted effect of the mutation on the amelogenin protein. For example, the mutation 3 abbreviation (p.W4S) indicates that in the mutant protein tryptophan at amino acid 4 is changed to serine, while for mutation 4 (p.W4X) the tryptophan codon was changed to a translation termination codon, truncating the protein.

and results from the alternative inactivation of either the normal or the defective X chromosome in different cohorts of enamel-forming cells [Berkman and Singer, 1971]. In males the enamel is much more severely affected, with brown teeth having little or no enamel covering the dentin.

Cells Tissues Organs 2007;186:78–85

81

1 2 19-41 57-70

ENAM 1

2

Number

3 4 5 67 158-178

3

4 5 6

7

1-18

42-56

71-157

8

9

10

179-196

197-1142

Mutation

References

1

p.K53X

[Mårdh et al., 2002] [Kim et al., 2006]

2

p.M71-Q157del

[Kim et al., 2005a]

3

p.A158-Q178del

[Rajpar et al., 2001] [Urzua et al., 2005]

[Kida et al., 2002] [Hart et al., 2003a] [Kim et al., 2005a] [Ozdemir et al., 2005a] 5 p.S246X 6 p.V340_M341insSQYQYCV [Ozdemir et al., 2005a] [Hart et al., 2003c] 7 p.P422fsX448 [Ozdemir et al., 2005a] 4

p.N197fsX277 or p.R179-N196del

Fig. 3. ENAM mutations causing autosomal dominant AI. The

intron/exon structure of ENAM is shown at the top. The range of enamelin amino acids encoded by a given exon is shown above or below that exon. The 7 ENAM mutations characterized to date are numbered in bold starting at the 5 end of the gene and refer to the mutations listed below.

Ameloblastin (AMBN, 4q13.3)

Ameloblastin comprises roughly 5% of enamel protein. It has an apparent molecular weight of 65–70 kDa, which includes one or two sulfated O-linked glycosylations [Yamakoshi et al., 2001]. N-terminal ameloblastin cleavage products accumulate in the sheath space separating rod and interrod enamel [Uchida et al., 1995], which may help maintain rod boundaries. Ameloblasts in the Ambn–/– mice detach from the surface of the developing teeth, suggesting a potential function in the adhesion of the ameloblasts to the forming enamel [Fukumoto et al., 2004]. Other interpretations are possible, however. In the Ambn–/– mice ameloblasts actually fail to attach to mineralized dentin, as there is no enamel layer and any attachment apparatus requiring an enamel surface would be expected to fail even if Ambn were not involved in the mechanism. No human AMBN mutations outside of apparent polymorphisms have been identified in AI kindreds, presumably because its pattern of inheritance is autosomal recessive and occurs only rarely [Kim et al., 2006]. 82

Cells Tissues Organs 2007;186:78–85

Enamelin (ENAM, 4q13.3)

Enamelin is the largest enamel protein (
200 kDa), and also the least abundant (3–5%) of the three major structural proteins in developing enamel. Enamelin is a glycosylated, phosphorylated protein that is rapidly cleaved following its secretion. The intact protein is only observed at the mineralization front, suggesting it plays a role in crystal elongation [Hu et al., 1997, 2000]. Enamelin gene mutations are perhaps the most significant single contributing factor in the etiology of AI, causing autosomal dominant forms of AI, although other currently unidentified genes are certain to contribute to autosomal dominant forms of AI [Hart et al., 2003b]. ENAM mutations so far identified in AI kindreds are listed in figure 3.

The Maturation Stage of Amelogenesis

At a certain point in the developmental program, ameloblasts undergo a transition that not only reduces their secretion of enamel proteins, but also initiates the secretion of kallikrein 4 (KLK4), a serine protease that degrades the organic matrix, facilitating its removal from the extracellular compartment. These changes terminate the growth of enamel crystallites in length, and vastly accelerate their growth in width and thickness. The degradation and removal of growth-inhibiting enamel proteins exposes the sides of the thin crystals to ion deposition. The predominant site of mineral deposition shifts from the enamel surface to the entire thickness of enamel. The pace of mineralization quickens and the crystallites thicken until they press against one another [Smith, 1998]. The maturation stage for the human permanent dentition, during which the crystallites grow in width and thickness, takes about 3–6 years. This process is necessary to harden the enamel layer, and is directed by modulating ameloblasts that cycle through smooth and ruffle-ended phases. Fluoride is incorporated into crystal structure during the maturation stage. Disturbances during the maturation stage of amelogenesis result in pathologically soft (hypomaturation) enamel of normal thickness. Despite the rapid degradation of matrix proteins during the transition and early maturation stages, a basal lamina is successfully erected at the base of the maturation stage ameloblasts (fig. 1). One component of this basal lamina has recently been cloned and is named amelotin (AMTN) [Iwasaki et al., 2005; Moffatt et al., 2006]. Hu/Chun/Al Hazzazzi/Simmer

Amazingly, a role for the gene encoding amelotin (UNQ689) in enamel formation was predicted before it was cloned, based in part upon its location on chromosome 4q next to AMBN in the SCPP (secretory calciumbinding phosphoprotein) gene cluster [Kawasaki and Weiss, 2006]. Only recently discovered, no AMTN mutations have yet been associated with AI.

Autosomal Recessive Hypomaturation AI: MMP20 and KLK4 Defects

Enamelysin (MMP20) is expressed by ameloblasts throughout the secretory stage and into early maturation; KLK4 is expressed later, starting in transition/early maturation and continuing through tooth eruption [Hu et al., 2002; Simmer et al., 2004]. Despite there being important differences in the timing of their expression, MMP20 and KLK4 mutations cause a similar phenotype in humans: autosomal recessive pigmented hypomaturation AI (fig. 4). The thickness of the enamel layer is within normal limits, but it is stained and aberrantly soft. This MMP20–/– phenotype is surprising. Mmp20–/– mice displayed obvious secretory stage defects, such as hypoplastic (thin) enamel lacking normal prism structures [Caterina et al., 2002]. MMP20 is predominantly expressed during the secretory stage when crystal elongation is occurring and is thought to be necessary for that process, but the human enamel phenotype is not obviously hypoplastic. The finding of hypomaturation AI in the MMP20–/– kindreds underscores the potential importance of the slow degradation of enamel proteins and the thickening of enamel crystals away from the enamel surface during the secretory stage.

2

1 1 1-42

2

3

43-125

4

5

6

MMP20 7

8

Number Mutation 1 p.H226Q 2 g.IVS6-2A>T

2 1-20

Number 1

3 21-75

451-483

References [Ozdemir et al., 2005b] [Kim et al., 2005b] 1

1

10

9

126-174 175-216 217-270 271-318 319-363 364-416 417-450

KLK4

4 5 76-158 159-204

Mutation p.W153X

6 205-254

References [Hart et al., 2004]

Fig. 4. MMP20 and KLK4 mutations causing autosomal recessive AI. The intron/exon structures of the enamelysin (MMP20) and kallikrein 4 (KLK4) genes are shown. Mutations in the enamel proteases cause a pigmented hypomaturation AI. Mutations in both alleles of either of these protease genes cause the disease. The MMP20 mutation abbreviated g.IVS6–2A 1T indicates that a splice junction in intron 6 was defective. The defect is predicted to cause the RNA transcripts to be degraded.

dento-osseous syndrome caused by a DLX3 mutation was classified as AIHHT [Dong et al., 2005; Wright, pp 86–93, this issue].

Summary and Conclusion

Mutations in the homeobox gene DLX3 (17q21) cause tricho-dento-osseous syndrome, which is characterized by enamel hypoplasia with root taurodontism, kinky, curly hair at birth and increased thickness and density of the cranial bones [Price et al., 1998; Haldeman et al., 2004]. Enamel hypoplasia and taurodontism are consistently found, but the non-dental features may be absent [Price et al., 1999]. DLX3 mutations were ruled out in a family with isolated AIHHT, suggesting that AIHHT and tricho-dento-osseous syndrome are two genetically distinct conditions. Recently, an apparent case of tricho-

Epithelial-mesenchymal interactions guide tooth development and lead to the differentiation of odontoblasts and pre-ameloblasts. Following early predentin formation, the basal lamina beneath pre-ameloblasts disintegrates and dentin mineralization begins. Enamel proteins are secreted onto the dentin surface and an enamel mineralization front is established near the ameloblast distal membrane, at the previous site of the basal lamina. During the secretory stage enamel crystallites grow primarily in length. Secretory stage defects are associated with hypoplastic or thin enamel. AMELX and ENAM mutations cause X-linked and autosomal dominant AI, respectively. During the maturation stage, enamel proteins are degraded and removed and the enamel layer hardens as the crystallites grow in width and thickness. MMP20 and KLK4 mutations cause autosomal recessive

Dental Malformations

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AI Hypoplastic-Hypomaturation with Taurodontism (AIHHT): DLX3

83

pigmented hypomaturation AI, which is characterized by the retention of enamel proteins and a reduction in enamel hardness. Candidate gene approaches to determine the genetic causes of isolated AI currently show a 25% success rate, suggesting that more AI candidate genes need to be identified [Kim et al., 2006].

Acknowledgments We thank Chris Jung for his help in creating figure 1, and Dr. Charlie Smith for his critical review of the manuscript. This investigation was supported by United States Public Health Service Research Grants DE12769, DE15846, and DE11301 from the National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, Md., USA.

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Cells Tissues Organs 2007;186:86–93 DOI: 10.1159/000102684

The Molecular Control of and Clinical Variations in Root Formation Tim Wright Department of Pediatric Dentistry, University of North Carolina School of Dentistry, Chapel Hill, N.C., USA

Key Words Cementum
Dentin
Development
Dysplasia
Genetics
Taurodontism

Abstract Roots of teeth perform critical functions to anchor the teeth in the jaws and transmit the masticatory forces in such a way as to minimize fracture and wear of the dentition. Tooth root development involves a variety of cell types, epithelial-mesenchymal interactions, the enumeration of specialized extracellular matrices, processing of these matrices and strict control over the microenvironment to allow the cementum and dentin to mineralize. While many of the specific molecular mechanisms involved in root formation remain poorly understood, our knowledge of these events and pathways has advanced markedly over the past decade. The molecular bases of many hereditary conditions having associated dental root anomalies are now known. Therapeutic approaches based on the molecular biology of root formation have and will continue to emerge and be translated into improved clinical care. The purpose of this study was to review our knowledge regarding developmental defects of root formation, the molecular mechanisms involved, and the impact of root variants on clinical dentistry.

Introduction

The dental root structure serves a variety of critical functions that are integral to normal tooth function. The root and its associated tissues provide support for the clinical crown, the means of attaching the crown to the jaw, help transmit and absorb forces placed on the clinical crown and serve as a conduit for the vascular and nervous tissues for the tooth. Root formation involves a series of complex processes involving epithelial- and mesenchymal-derived tissues that interact through molecular signaling, develop terminally differentiated cells that secrete unique extracellular matrices, and control the microenvironment so the root-associated tissues can mineralize [Slavkin et al., 1989; Thomas 1995; ZeichnerDavid et al., 2003]. The bulk of the root structure is comprised of dentin, which is covered externally by cementum. The cementum is anchored to the alveolar bone via Sharpies fibers (collagen fibers of the periodontal ligament that are imbedded in the alveolar bone or cementum) and the periodontal ligament. The unique structure and composition of the tooth root allow it to optimally

Abbreviations used in this paper

Copyright © 2007 S. Karger AG, Basel

This study was supported by NIDCR Grant 12879.

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/cto

AI ED OMIM TDO

amelogenesis imperfecta ectodermal dysplasia Online Mendelian Inheritance in Man tricho-dento-osseous syndrome

Dr. Tim Wright Department of Pediatric Dentistry, School of Dentistry, Brauer Hall 7450 The University of North Carolina Chapel Hill, NC 27599 (USA) Tel. +1 919 966 2739, Fax +1 919 966 7992, E-Mail [email protected]

Normal root formation requires specialized cells that are of both epithelial and mesenchymal derivation. The epithelial root sheath is derived from the enamel epithelium that proliferates apically from the cervical loop of the developing dental crown. These proliferating epithelial cells form a sheath that serves as the template for and definition of each tooth’s specific root morphology. For example, the epithelial sheath provides the template for establishing the multi-rooted morphology typically seen in posterior teeth. Multiple roots are formed when the epithelial root sheath invaginates inwards towards the pulp at the location of what is to become the future sight of the root furcation. Deviations in this process lead to a variety of morphological root variances such as supernumerary roots, pyramidal-shaped roots, and taurodontism. The specific molecular triggers that signal the epithelial root sheath to invaginate remain unknown, however, a variety of abnormalities in root number and morphology arise from abnormal root sheath activity. Delay or failure of the epithelial root sheath to invaginate results in the apical displacement of the furcation area resulting in taurodontism (Latin for bull teeth) [Keith, 1913]. The level at which the root sheath invaginates determines the degree of severity of the taurodontism (fig. 1) [Shaw, 1928]. Alternatively, abnormal invagination can result in surpernumerary or auxiliary root structures (fig. 2). As the root sheath proliferates apically, it interacts with the underlying mesenchymal cells. Molecular signaling

from the epithelial cells is believed to instruct the mesenchymal-derived pulpal cells to differentiate into odontoblasts that will then begin to produce dentin [Ten Cate, 1996]. The epithelial root sheath becomes fenestrated and mesenchymal cells contact the dentin where they begin to produce cementum at the future site of the cemento-dentinal junction [Thomas, 1995; Diekwisch, 2001]. A variety of mouse studies indicate numerous transcription and growth factors are expressed by cells involved in root development that likely play a critical role in orchestrating formation of this specialized appendage. For example, Shh, Dlx2 and Patched2 are expressed by the epithelial root sheath cells [Lézot et al., 2000; Nakatomi et al., 2006]. The transcription factors Nfic, Gli1, Patched1, and Smoothened are all expressed primarily by the mesenchymal cells [Steele-Perkins et al., 2003; Nakatomi et al., 2006]. Mice lacking Nfic expression show a near complete loss of molar root development [Steele-Perkins et al., 2003]. A variety of factors involving bone morphogenic proteins, growth differentiation factors, transforming growth factor-
, and genes involved in the tumor necrosis factor pathway are also known to be expressed and important in root development [Gao et al., 1998; Morotome et al., 1998; Miard et al., 1999; Thomadakis et al., 1999; Tucker et al., 2004]. Overexpression of transforming growth factor-
1 during dentinogenesis [driven by DSPP (the dentin sialophosphoprotein gene) promoter] also results in a lack of normal root formation in a transgenic mouse model [Thyagarajan et al., 2001]. In addition to the numerous signaling and growth factors, there are many extracellular matrix proteins produced by the odontoblasts and cementoblasts during development of the dental root. The expression of enamelrelated proteins, and most specifically amelogenin, during root formation has long been and remains controversial [Hammarstrom, 1997; Slavkin et al., 1988; Diekwisch, 2001; Bosshardt and Nanci, 2004; Janones et al., 2005]. Proteinases are also critical for normal root development with matrix metalloproteinases and other proteinases functioning to process the organic matrix [Sakuraba et al., 2006]. Additionally, alkaline phosphatase is highly expressed and is thought to help create the appropriate phosphate environment necessary for cementum formation [Beertsen et al., 1999; Nociti et al., 2002; Foster et al., 2006]. Given the complexity of dental root development, it is not surprising that there are many examples of both genetic and environmental conditions that result in altered root morphology and/or composition. The quality and quantity of the dental root have significant treatment im-

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perform its diverse functions of support, energy dissipation and neurosensory activity via the pulp that courses through the central portion of the root. Formation of the dental root is under strict molecular regulation that plays a fundamental role in determining cell lineage and fate, tissue composition and structure, and morphology [Nakatomi et al., 2006]. While our understanding of the genes and specific processes related to tooth initiation and crown formation have advanced tremendously over the past several decades [Thesleff, 2003; Ohazama and Sharpe, 2004], root development remains less well understood. There are numerous known clinical variants and hereditary conditions that result in a variety of root-related deviations and developmental defects illustrating the critical molecular regulation of root development. The purpose of this study is to review our knowledge regarding root formation, its molecular controls, and hereditary conditions known to affect root development.

Root Formation

87

Fig. 1. Taurodontism ranges in severity,

becoming increasingly severe with increasing delay of invagination of the epithelial root sheath. The degree of taurodontism can be quantified based on the crown-body (CB) to root (R) ratio.

Fig. 2. These two primary mandibular mo-

lars each have a supernumerary root variant (three roots as opposed to the more typical two roots) due to aberrant invagination of the epithelial root sheath.

plications in periodontal disease, orthodontics, treatment of trauma and restorative dentistry. In the following section some of the more common human variants of defective root development are reviewed.

Developmental Defects of the Root

There are numerous hereditary conditions that can affect the shape, size, and composition of the dental roots. Many syndromes and chromosomal anomalies have associated tooth and dental root phenotypes. Therefore, the 88

Cells Tissues Organs 2007;186:86–93

following section is not meant to be an exhaustive review but rather will highlight some of the critical components for root formation discussed in the previous section. Taurodontism There are many diverse conditions that are associated with taurodontism [Online Mendelian Inheritance in Man (OMIM) No. 272700] that result from the delayed invagination of the epithelial root sheath creating an elongated pulp chamber. Normally invagination of the epithelial root sheath occurs at the height of the furcation where it fuses to form the template and control the morWright

Fig. 3. The maxillary primary molars in this child with X-linked hypohidrotic ED both exhibit taurodontism (arrows) as viewed in this panoramic radiograph.

Fig. 4. This young adult affected with TDO displays severe generalized taurodontism, generalized thin enamel, and a marked increase in the mandibular bone as viewed in this panoramic radiograph.

phology of multi-rooted teeth. This root/pulp anomaly occurs in the normal population with a prevalence that varies substantially from one race to anther ranging from 1.5 to 48% of the population [MacDonald-Jankowski and Li, 1993; Toure et al., 2000]. Taurodontism is known to occur with many syndromes and chromosomal anomalies such as in individuals with Down syndrome, sex chromosome aneuploidy and hypodontia [Jaspers and Witkop, 1980; Seow and Lai, 1989; Alpoz and Eronat, 1997]. Given the purported abnormal epithelial root sheath function in the etiology of taurodontism it is not surprising that many conditions affecting epithelial-derived tissues also are associated with taurodontism. The ectodermal dysplasias (EDs) are a clinically and genetically diverse group of conditions characterized by defects in at least two ectodermally derived appendages (e.g. teeth, hair, nails, and sweat glands). The most common form is X-linked hypohidrotic ED (OMIM 305100) that is caused by mutations in ED1 (ectodysplasin gene). Phenotypically similar autosomal dominant and recessive hypohidrotic EDs result from mutations in the ED1

receptor (EDAR; OMIM 129490 and 224900). These genes are active in the tumor necrosis factor pathway and are critical in both early and late events in odontogenesis. The prevalence and severity of taurodontism associated with ED is variable (fig. 3) [Crawford et al., 1991]. The amelogenesis imperfectas (AIs) are a group of genetically heterogeneous hereditary conditions affecting primarily the enamel. These conditions were classified into distinct AI subtypes by Witkop [1989] based on mode of inheritance and the phenotype of enamel and the presence of taurodontism. Those AI types having taurodontism as a phenotypic feature were designated hypoplastic/hypomaturation AI with taurodontism (AI type IV) [Witkop, 1989]. Many cases originally reported as AI with taurodontism (OMIM 104510) were later reported to be individuals affected with the tricho-dento-osseous syndrome (TDO, OMIM 19032), due to the difficult clinical delineation between these conditions [Seow, 1993; Collins et al., 1999; Price et al., 1999]. Several studies indicate that the prevalence of taurodontism in AI cases is similar to that seen in unaffected controls [Seow, 1993;

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89

Collins et al., 1999]. The different AI conditions are known to be caused by a variety of allelic and non-allelic mutations in a variety of genes including AMELX, ENAM, KLK4 and MMP20 [Wright, 2006]. There appear to be numerous AI types with molecular etiologies that remain to be discovered [Hart et al., 2003]. While the AI-associated gene products are critical for normal enamel formation, they do not appear essential for normal root formation given that none of the AI phenotypes resulting from these mutations is known to have abnormal root morphology or structure and the dentin and cementum appear grossly normal. Marked taurodontism of the molar teeth (fig. 4) is a highly penetrant feature of TDO which is caused by mutations in the Distal-less homeobox gene DLX3. TDO derives its name from the three predominantly affected tissues, tricho-hair, dento-teeth, and osseous bone, which show variable severity in expression of the trait [Wright et al., 2000]. One family having a DLX3 mutation with no apparent hair or bone manifestations has been reported as being AI with taurodontism [Dong et al., 2005], however our evaluation of a large family with this same DLX3 mutation (CTdel) shows a marked hair phenotype suggesting this represents a TDO variant. Whether there is a distinct condition characterized by AI and taurodontism phenotypes remains controversial. While the expression of taurodontism in people with DLX3 mutations is variably expressed, the high penetrance of this trait suggests that DLX3 is an important regulator providing some degree of temporal control over the invagination of the epithelial root sheath. Variations in Root Size and Structure There are numerous hereditary conditions associated with abnormal development of the root that affect the size and structure. For example, dentinogenesis imperfectas (OMIM 125490 and 166240) and dentin dysplasia type II (OMIM 125420) frequently exhibit abnormal root morphology and often have a diminished root size (fig. 5). Mutations in the type 1 collagen genes (COL1A1 and COL1A2) and DSPP are associated with the dentinogenesis imperfecta phenotype with roots that are commonly described as being short and constricted [Pallos et al., 2001; Malmgren et al., 2004]. Additionally, the EhlersDanlos conditions that are associated with collagen mutations (Ehlers-Danlos type I, collagen V mutation, OMIM 13000: Ehlers-Danlos type IV, collagen III mutation, OMIM 13050, and Ehlers-Danlos type VI, lysyl hydroxylase mutation, OMIM 225400) also can be associated with abnormalities in dentin, root formation, and in 90

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Fig. 5. This radiograph shows the shortened and minimally sepa-

rated root morphology commonly seen in dentinogenesis imperfecta type I and type II.

some cases periodontal disease [Pope et al., 1992; Karrer et al., 2000]. Collectively, these conditions illustrate the importance of collagens and other extracellular matrix proteins in determining root morphology. Dentin dysplasia type I (OMIM 125400) is a rare apparently autosomal dominant condition characterized by crowns that appear clinically normal. However, after the initial layer of normal dentin is formed, a cycle of odontoblast death followed by new odontoblast recruitment again followed by odontoblast death occurs during repeated attempts at dentin formation. This pattern leads to a unique histological appearance of the dentin (cascading waterfall; fig. 6a) and markedly diminished root development (fig. 6b). The molecular etiology for this condition remains unknown at this time. Hypophosphatasia (OMIM 146300 and 24150) can be inherited as an autosomal dominant or recessive trait and is caused by mutations in ALPL (tissue nonspecific alkaline phosphatase gene). The dentin can be hypomineralized, have an increased Tomes granular layer, and the root surfaces lack a normal cellular and acellular cementum covering [Hu et al., 2000; van den Bos et al., 2005]. Consequently the teeth are frequently exfoliated prematurely with little or no root resorption. Loss of function of the tissue-nonspecific alkaline phosphatase alters the normal hydrolysis of pyrophosphate, which is an inhibitor of apatite crystal growth that is thought to be critical for normal cementum development [Nociti et al., 2002; van den Bos et al., 2005]. Wright

a

b

Fig. 6. a This mineralized thin section of a tooth viewed with transmitted light microscopy shows the lack of normal root structure and the classical histological cascading waterfall dentin phenotype characteristic of dentin dysplasia type I. b Dentin dysplasia type I teeth frequently show pulp obliteration, only rudimentary root formation, and periapical abscess formation (note second primary molars) as illustrated in this panoramic radiograph.

Normal formation of the dental root is critical to the function and longevity of the dental apparatus. Variable root formation results in numerous developmental variants that can challenge virtually every aspect of dental therapy. The loss of or a decrease in root structure compromises the support of the clinical crown (decreased crown to root ratio) and will likely result in decreased tooth longevity [Grossmann and Sadan, 2005]. The diverse morphology of the dental roots, multiple supernumerary roots, accessory canals and difficulty in manipulating canals for cleansing and obturation during conventional root canal therapy continue to provide challenges for endodontic therapy. Marked root dilacerations and divergences can create surgical challenges, too. The management of traumatized teeth frequently hinges on the stage of root development, with immature roots markedly decreasing the prognosis for non-vital teeth. Recent advances in approaches to revascularize the dental pulp and allow for continued root development in non-vital teeth having incomplete root formation show promise to enhance root formation [Banchs and Trope, 2004]. While still experimental, these approaches indicate a sustained ability for root formation even in teeth with necrotic pulp tissue, suggesting cells with root-forming potential remain viable at the cervical loop area even after the pulpal tissue becomes necrotic. Although the ability to enhance root formation presents multiple new therapeutic approaches to managing teeth with incomplete root forma-

tion, the problems of manipulating non-vital teeth with complete root formation remains a marked challenge. Periodontal therapy is frequently directed at maintaining or regenerating a healthy attachment between the tooth root, alveolar bone, and oral epithelium [Popowics et al., 2005]. Although our knowledge of the molecular controls and developmental mechanisms of cementum formation is expanding, only a few current therapies are directed at stimulating cementogenesis [Foster and Somerman, 2005; Bartold et al., 2006; Zeichner-David, 2006]. Despite a lack of clear developmental mechanisms, enamel proteins have been investigated extensively and are commercially available to enhance cementum formation [Suzuki et al., 2005; Kanazashi et al., 2006]. As our understanding of the molecular controls operating during cementogenesis is elucidated, this knowledge will likely be rapidly translated to new and more effective clinical therapies [Alvarez-Perez et al., 2006]. As the quest to grow a tooth continues, our knowledge of the molecular controls of root formation becomes absolutely essential for successfully completing this sojourn. The investigation of human variants in root formation, the use of a variety of animal models and basic molecular biological approaches are providing the materials and tools that are driving the advancement of our knowledge of root development. Acquisition of this new knowledge will direct many novel potential therapeutic applications regarding diagnostics, tissue engineering, biomimetics (e.g. improved implant systems), and dental trauma management to name just a few.

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Clinical Implications

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Wright, J.T. (2006) The molecular etiologies and associated phenotypes of amelogenesis imperfecta. Am J Med Genet 140: 2547–2555. Wright, J.T., J.A. Price, D. Bowden, K. Kula, D. Dawson, T.C. Hart (2000) Phenotypic variability of the tricho-dento-osseous syndrome associated with a DLX3 homeobox gene mutation; in Goldberg, M., A. Boskey, C. Robinson (eds): Chemistry and Biology of Mineralized Tissues. Proceedings of the Sixth International Conference. Rosemont, American Academy of Orthopedic Surgeons, pp 27–31. Zeichner-David, M. (2006) Regeneration of periodontal tissues: cementogenesis revisited. Periodontol 2000 41: 196–217. Zeichner-David, M., K. Oishi, Z. Su, V. Zakartchenko, L.S. Chen, H. Arzate, P. Bringas, Jr. (2003) Role of Hertwig’s epithelial root sheath cells in tooth root development. Dev Dyn 228: 651–663.

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Author Index Vol. 186, No. 1, 2007

Al Hazzazzi, T. 78 Buchanan, A.V. 7 Chun, Y.-H.P. 78 Davit-Béal, T. 25 Delgado, S. 25 D’Souza, R.N. 60 Gu, X. 25 Hart, P.S. 70 Hart, T.C. 70 Hu, J.C.-C. 78 Iwase, M. 49

Kaneko, S. 49 Kawasaki, K. 7 Kim, H. 49 Klein, O.D. 60 Satta, Y. 49 Simmer, J.P. 4, 78 Sire, J.-Y. 25 Takahata, N. 49 Weiss, K.M. 7 Wright, T. 86

Subject Index Vol. 186, No. 1, 2007

Amelogenesis imperfecta 78 Amelogenin 49, 78 Cementum 86 Dentin 70, 86 – dysplasia 70 Dentinogenesis imperfecta 70 Development 86 DSPP 70 Dysplasia 86 Enamel 25, 78 Enamelin 78 Enamelysin 78 Evolution 25, 49 Fibroblast growth factors 60 Gene duplication 7 Genetics 86 Genome duplication 7

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

Accessible online at: www.karger.com/cto

Genomics 25 Kallikrein 4 78 Mammalian sex chromosomes 49 Mineralization 25 Mineralized tissue 7 Polymorphism 49 Pseudoautosomal boundary 49 Runx2 60 Skeletal mineralization 7 Small integrin-binding ligand N-linked glycoprotein 70 Sprouty 60 Supernumerary teeth 60 Taurodontism 86 Tooth 25 Transgenic mice 60 Vertebrate evolution 7

Patent Watch Cells Tissues Organs 2007;186:95

United States Patents Title: In-situ Injection and Materials Screening Device Inventors: Trevor G. Frank, Fremont, CA Keith A. Hall, San Jose, CA William H. Chandler, Jr., Milpitas, CA Thomas Boussie, Menlo Park, CA Thomas J. Crevier, San Jose, CA Leonid Matsiev, San Jose, CA Christopher Goh, San Francisco, CA Assignee: Symyx Technologies, Inc., Santa Clara, CA Patent Number: US 6,841,127 B2 Title: Bioreactor Inventors: Thomas Wechsler, Zurich (CH) Ulrich Baer, Heiden (CH) Christian Oehr, Herrenberg (DE) Thomas Graeve, Stuttgart (DE) Assignee: Sefar AG, Rueschlikon (CH) Patent Number: US 6,844,187 B1 Title: Microfabricated Tissue as a Substrate for Pigment Epithelium Transplantation Inventors: Harvey A. Fishman, Menlo Park, CA Mark Blumenkranz, Portola Valley, CA Stacey F. Bent, Palo Alto, CA Christina Lee, San Francisco, CA Philip Huie Jr., Cupertino, CA Daniel V. Palanker, Sunnyvale, CA Patent Number: US 2005/0027356 A1 Title: Conditioned Compositions for Tissue Restoration Inventor: Stephen F. Badylak, Pittsburgh, PA Patent Number: US 2005/0025838 A1 Title: Cryopreservation Medium for Primate Embryo Stem Cells and Cryopreservation Method Inventors: Norio Nakatsuji, Kyoto (JP) Hirofumi Suemori, Kyoto (JP) Isao Asaka, Chiba (JP) Assignee: Asahi Techno Glass Corporation, Funabashi-shi (JP) Patent Number: US 2005/0026133 A1 Title: Isolation and Mobilization of Stem Cells Expressing VEGFR-1 Inventors: Shahin Rafii, Great Neck, NY Larry Witte, Stormville, NY Patent Number: US 2005/0026220 A1

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

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Title: Optimizing Culture Medium for CD34 Hematopoietic Cell Expansion Inventors: Shiaw-Min Hwang, Hsin-City (TW) Chi-Hsien Liu, Hsin-Chu City (TW) Chao-Ling Yao, Hsin-Chu City (TW) I-Ming Chu, Hsin-Chu City (TW) Tzu-Bou Hsieh, Hsin-Chu City (TW) Patent Number: US 2005/0032122 A1 Title: Materials and Methods to Produce Stem Cells Inventors: Steve Kah Weng Oh, Singapore (SG) Andre Boon Hwa Choo, Singapore (SG) Patent Number: US 2005/0032208 A1

European Patents Title: Bioartifizielles, primär vaskularisiertes Testgewebe für pharmakologische Testverfahren, Verfahren zu dessen Herstellung und Testverfahren unter dessen Verwendung sowie Bioreaktor zur Durchführung der Verfahren Inventor: Walles, Thorsten, Dr. med., 30163 Hannover (DE) Applicant: Walles, Thorsten, Dr. med., 30163 Hannover (DE) Patent Number: EP 1 500 697 A2 Title: Bioartifizielle Haut, die genetisch modifizierte Fibroblasten aufweist Inventor: Machens, Hans-Günther, DE–23538 Lübeck (DE) Applicant: Machens, Hans-Günther, DE–23538 Lübeck (DE) Patent Number: EP 1 504 087 A2 Title: Biomaterials for Nerve Reconstruction and Process for Producing the Same Inventors: Yamaguchi, Isamu, Tsukuba-shi, Ibaraki 305-0047 (JP) Taguchi, Tetsushi, Tsukuba-shi, Ibaraki 305-0032 (JP) Tanaka, Junzo, Tsukuba-shi, Ibaraki 305-0032 (JP) Shinomiya, Kenichi, Tokyo 114-0014 (JP) Itoh, Soichiro, Bunkyo-ku, Tokyo 112-0005 (JP) Fukuzaki, Hironobu, Tatsuno-shi, Hyogo 305-0047 (JP) Oka, Yoichi, Tsukuba-shi, Ibaraki 305-0061 (JP) Applicants: Japan Science and Technology Agency Kawaguchishi, Saitama 332-0012 (JP) National Institute for Materials Science, Tsukuba-shi, Ibaraki 305-0047 (JP) Taki Chemical Co., Ltd., Kakogawa-shi, Hyogo 675-0125 (JP) Patent Number: EP 1 493 454 A9

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Conference Calendar

19.7.–21.7.2007 San Antonio, Tex. USA

XVIth Ovarian Workshop

Information: www.biosymposia.org/content16510.html

15.8.–18.8.2007 Houston, Tex. USA

Advances in Tissue Engineering

Information: www.tissue.rice.edu

19.8.–23.8.2007 Boston, Mass. USA

CELLutions SUMMIT

Information: www.CELLutionsSUMMIT.com

19.8.–24.8.2007 Budapest Hungary

XIXth International Symposium of Morphological Sciences

Information: www.ismsbudapest.com

4.9.–7.9.2007 London England

TERMIS-Europe

Information: www.termis.org

10.9.–12.9.2007 Cracow Poland

EMT 2007 Meeting – 3rd International Meeting on: Epithelial-Mesenchymal Transition Co-organized by TEMTIA and Marie-Curie Epiplastcarcinoma RTN network

Information: www.mtci.com.au/Temtia.html

26.9.–29.9.2007 Freiburg i.Br. Germany

49th Symposium of the Society for Histochemistry A Platform for Talents

Information: Martin Werner and Christoph Peters University of Freiburg/Br., D–79104 Freiburg i.Br. E-Mail [email protected]

11.10.–13.10.2007 Essen Germany

Biomaterials 2007 10th Essen Symposium on Biomaterials and Biomechanics 2007 Reserch development and clinical application – From bench to bedside

Information: Secretariat: Universitätsklinikum Essen Zentrum für Innere Medizin, Klinik für Kardiologie Westdeutsches Herzzentrum Essen Hufelandstrasse 55, DE–45122 Essen, Germany Tel.: +49 (0)201 723-4804, Fax: +49 (0)201 723-5404 E-Mail: [email protected]

18.10.–20.10.2007 Leipzig Germany

3rd World Congress on Regenerative Medicine

Information: www.regmed.org

8.11.–9.11.2007 Tokyo Japan

TERMIS-Asia-Pacific

Information: www.termis.org

14.2.–17.2.2008 Berlin Germany

8th International Conference on New Trends in Immunosuppression and Immunotherapy

Information: www.kenes.com/immuno

23.6.–27.6.2008 Porto Portugal

TERMIS-Europe

Information: www.termis.org

7.11.–8.11.2008 Taipei Taiwan

TERMIS-Asia-Pacific

Information: www.termis.org

6.12.–10.12.2008 San Diego, Calif. USA

TERMIS-North America

Information: www.termis.org

31.8.–3.9.2009 Seoul Korea

TERMIS-World Congress

Information: www.termis.org

© 2007 S. Karger AG, Basel Fax +41 61 306 12 34 E-Mail [email protected] www.karger.com

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