Role of SIBLINGs on matrix mineralization: Focus on dentin matrix protein 1 (DMP1)

Journal of Oral Biosciences 54 (2012) 30–36

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Review

Role of SIBLINGs on matrix mineralization: Focus on dentin matrix protein 1 (DMP1) Satoru Toyosawa a,n, Sunao Sato a, Ryosuke Kagawa b, Toshihisa Komori c, Kazunori Ikebe b a

Department of Oral Pathology, Osaka University Graduate School of Dentistry, 1-8 Yamadaoka, Suita, Osaka 565-0871, Japan Department of Prosthodontics and Oral Rehabilitation, Osaka University Graduate School of Dentistry, 1-8 Yamadaoka, Suita, Osaka 565-0871, Japan c Department of Developmental and Reconstructive Medicine, Nagasaki University Graduate School of Biomedical Sciences, 1-7-1 Sakamoto, Nagasaki 852-8523, Japan b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 25 April 2011 Received in revised form 23 May 2011 Accepted 26 May 2011 Available online 15 February 2012

Dentin matrix protein 1 (DMP1) is a member of the SIBLING (Small Integrin-Binding Ligand, N-linked Glycoprotein) family of genetically related noncollagenous proteins in mineralized tissues. DMP1 was originally postulated to be dentin-specific, but its expression was later found to be present predominantly in osteocyte of bone. A unique feature of DMP1 is its unusually large number of acidic domains. Because of its highly acidic nature, DMP1 can bind to calcium, thereby regulating matrix mineralization. Protein chemistry analysis showed that full-length DMP1 is a precursor that is cleaved into NH2-terminal 37-kDa and COOH-terminal 57-kDa fragments. Functional analyses demonstrated that the COOH-terminal 57-kDa fragment of DMP1 is essential for mineralization and maturation of osteoblast to osteocyte. The phenotype of DMP1-null mice with hypophosphatemia led to the discovery of DMP1 mutations in autosomal-recessive hypophosphatemic rickets (ARHR). Both DMP1-null mice and individuals with ARHR exhibit elevated serum FGF23 that causes increased renal phosphate wasting, leading to hypophosphatemia. These findings indicate that regulation of matrix mineralization by DMP1 is coupled to renal phosphate homeostasis through FGF23 production by osteocyte. This review summarizes the current understanding of DMP1 focusing on bone formation and homeostasis. & 2012 Japanese Association for Oral Biology. Published by Elsevier B.V. All rights reserved.

Keywords: Dentin matrix protein 1 (DMP1) Mineralization Osteocyte Autosomal-recessive hypophosphatemic rickets (ARHR) Fibroblast growth factor 23 (FGF23)

Contents 1. 2. 3. 4. 5. 6.

7.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 The SIBLING family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 Gene structure and regulation of DMP1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 Protein structure of DMP1. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 Temporospatial localization of DMP1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Functions of DMP1. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 6.1. DMP1 promotes mineralization in vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 6.2. DMP1 function in bone in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 Conflict of interest . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35

1. Introduction Bone, dentin, and cementum are mineralized tissues that are composed of organic matrix and mineral contents (hydroxyapatite). The matrix consists of collagen (predominantly) and several

n

Corresponding author. Tel.: þ81 6 6879 2891; fax: þ 81 6 6879 2895. E-mail address: [email protected] (S. Toyosawa).

noncollagenous proteins. The collagen fibrils function as a scaffold for the deposition of hydroxyapatite crystals, whereas the noncollagenous proteins are believed to be involved in matrix mineralization. One category of noncollagenous proteins is termed the SIBLING (Small Integrin-Binding Ligand, N-linked Glycoprotein) family of genetically related proteins that are clustered on human chromosome 4 [1]. This family includes osteopontin (OPN), bone sialoprotein (BSP), dentin matrix protein 1 (DMP1), dentin sialophosphoprotein (DSPP), and matrix extracellular phosphoglycoprotein (MEPE). SIBLINGs are

1349-0079/$ - see front matter & 2012 Japanese Association for Oral Biology. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.job.2011.05.001

S. Toyosawa et al. / Journal of Oral Biosciences 54 (2012) 30–36

Exon No.

1

2

3

4

5

OPN

Non-coding

Signal+AA

SSEE

PPPP

SSEE

BSP

Non-coding

Signal+AA

SSEE

PPPP

SSEE

DMP1

Non-coding

Signal+AA

SSEE

PPPP

SSEE

DSPP

Non-coding

Signal+AA

MEPE

Non-coding

Signal+AA

PPPP

31

SSEE

6

RGD

RGD

RGD

RGD

(SSD) n

RGD

SSEE

Fig. 1. Gene organization of the SIBLING family. The SIBLING family includes osteopontin (OPN), bone sialoprotein (BSP), dentin matrix proetin1 (DMP1), dentin sialophosphoprotein (DSPP), and matrix extracellular phosphoglycoprotein (MEPE). They share the following common features. The exons (boxes) are separated by type 0 introns. The first exon is non-coding. The second exon contains the start codon, the hydrophobic signal peptide and the first two amino acids of the mature protein (AA). Exons 3 and 5 frequently contain consensus sequences for serine phosphorylation (SSEE). Exon 4 can be relatively proline-rich. The integrin-binding tripeptide, Arg-Gly-Asp (RGD), is found within the last one or two large exons. Proteolytic cleavage of SIBLINGs (arrowheads) by specific proteases (thrombin [64], SKI-1 [65], bone morphogenetic protein 1 (BMP1) [30,66], cathepsin B [67]) is thought to be important for their activation. DSPP contains tandem repeats of the phosphorylated nominal Ser-Ser-Asp (SSD) tripeptide.

defined as small, soluble RGD motif containing, integrin-binding ligands to distinguish them from other extracellular matrix proteins such as laminin and collagen. Although SIBLINGs’ amino acid sequences have little similarity except in limited regions of the molecules, they do share the following common features (Fig. 1) [1]: (1) Their genes are clustered along chromosome 4q21–23 in the human and chromosome 5q21 in the mouse. (2) They have conserved motifs in individual exons. (3) They share similarities in post-translational modifications such as phosphorylation, glycosylation, and proteolytic processing. One SIBLING identified with the help of recombinant DNA techniques was originally termed AG1 [2] but was later renamed dentin matrix protein 1 (DMP1) [3]. The DMP1 cDNA clone was obtained by screening a rat cDNA library prepared from the odontoblast–pulp fibroblast complex of incisors. The DMP1 amino acid sequence contains an unusually large number of acidic domains, a property that implicates it as a possible participant in regulating mineralization [2]. This hypothesis is supported by in vitro observations of DMP1-overexpressing cells [4] and in vivo findings from DMP1-null mice [5,6]. Recent studies have demonstrated DMP1 mutations as the cause of a novel human disease, autosomal-recessive hypophosphatemic rickets (ARHR) [7,8]. In this review, we summarize the current understanding of DMP1 in terms of gene and protein characteristics, temporospatial expression, and biological functions in bone.

2. The SIBLING family The term ‘‘SIBLING’’ refers to the gene family’s simple biochemical and genetic characteristics, but not to functional activity [1]. Individual exons of genes in this family share some similar properties (Fig. 1). For example, exon 1 is always noncoding, exon 2 contains the leader sequence and codons for the first two amino acids of the mature protein, exon 3 usually contains the consensus sequence (Ser-Ser-Glu-Glu: SSEE) for casein kinase II (CKII) phosphorylation, exon 4 is usually somewhat proline-rich

(among the acid proteins, this part is the only significantly positively charged domain), and exon 5 usually contains a CKII phosphorylation site and is the site of the only two known splice variants. The majority of each protein is encoded by the last one or two exons, which contain the integrin-binding Arg-Gly-Asp (RGD) tripeptide. Another point of similarity within the SIBLING family is that all members appear to contain at least one important controlled proteolysis site (arrowheads in Fig. 1). OPN, BSP, DMP1, and DSPP are similar in that they are phosphorylated, sulfated sialoproteins containing an abundance of acidic amino acids. OPN and DSPP are rich in Asp and BSP is rich in Glu. DMP1 is rich in Asp and Glu. These four members were discovered to be trapped in high numbers within mineralized matrices and have a role in nucleating hydroxyapatite crystals. In contrast, MEPE appears to be more distantly related in that it is not acidic in character although it is likely to have a phosphorylated COOH-terminus and is believed to inhibit mineralization [9–11].

3. Gene structure and regulation of DMP1 DMP1 sequences from several species have been reported: human [12], pig [13], bovine [14], rat [2,15], mouse [16], caiman [17], and chicken [18]. Southern blot analyses of genomic DNA from the rat [15], caiman [17], and chicken [18] produced banding patterns consistent with the existence of a single copy of a gene encoding for DMP1. The locus of the DMP1 gene has been mapped to 5q21 in the mouse using interspecific backcross mapping [3], and to 4q21.2–21.3 in the human by fluorescent in situ hybridization [19]. The human, pig, bovine, rat, mouse, and caiman genes were found to be composed of six exons. In the chicken DMP1 gene, mammalian/reptilian exons 4 and 5 appear to be missing (Fig. 2). Exon 1 is composed solely of the 50 untranslated sequence in all species. Exon 2 contains the first 54 bp of the coding sequence including the signal peptide sequences in all species. Exon 6 is

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S. Toyosawa et al. / Journal of Oral Biosciences 54 (2012) 30–36

Exon No.

1

2

3

4

54 48 33

Human (mammal) 5’UT

6

48

1356 (bp)

ATG 54 48 33

Caiman (reptile)

5

TAG 357

3’UT

1479 (bp) 3’UT

5’UT

54 36

Chicken (bird)

3’UT

ATG

TAA 1197 (bp) 3’UT

5’UT ATG

TGA

3’UT

Fig. 2. Gene organization of DMP1 in the mammal, reptile, and bird. Green-, blue-, and red-colored rectangles indicate translated exon regions while gray-colored rectangles indicate regions that are untranslated (UT). Coding lengths of exons in base pairs are indicated above the boxes. ATG is the initiation codon, and TAG, TAA, or TGA are the stop codons.

the largest exon (about 1200–1500 bp) and comprises 80% of the coding region. Mounting evidence suggests that variant forms of DMP1 may exist as a result of differential RNA splicing. Some clones from the human molar cDNA library did not have exon 5 [12]. The same alternatively spliced form was identified in the mouse and rat DMP1 mRNA [15,16]. Further, exon 5 can vary eight-fold in size, from 45 bp in the mouse to 357 bp in the caiman [17]. The observed variation of exon 5 and the absence of exons 4 and 5 in the chicken [18] imply that these exons play a role that is not critical to the function of DMP1. In the regulation of DMP1 transcription, two cis-regulatory fragments were identified, one in the proximal region between 2.4 and þ 4.5 kb and the other more distally between  2.4 and 9.6 kb [20,21]. Analysis of transgenic mice bearing reporter genes driven by these promoters showed that osteocyte-specific gene expression was found in the distal promoter ( 7892 to þ4439 bp), but not in the proximal promoter ( 2433 to þ4439 bp), in postnatal animals [20]. However, the proximal promoter (  2433 to þ4439 bp) could support gene expression in rapidly forming osteoblasts and pre-osteocytes in the embryo. In the tooth, the distal promoter (  9624 to þ4439 bp) supports gene expression in odontoblasts identical to the endogenous DMP1 gene, while the proximal promoter (  2433 to þ4439 bp) could stimulate gene expression in odontoblasts only during the early stage of odontogenesis [21]. Thus, the proximal promoter seems to control the early stages of DMP1 expression, and the distal promoter seems to control its later expression. DMP1 promoter sequences have potential responsive elements for several transcription factors involved in controlling hard tissue formation. These include the sites for Runx2, AP-1, SP-1, MSX, etc. [15,22]. Among these transcription factors, AP-1 was shown to potentially play a significant role in the transcriptional regulation of the DMP1 gene in MC3T3-E1 (preosteoblastic) cells, while it was not required in the later stages of mineralization [23]. Transient transfection studies showed forced expression of DMP1 under control of Runx2, and gel shift assays indicated potential response elements for Runx2 in the DMP1 promoter region in vitro, although it may be through indirect mechanisms [24].

4. Protein structure of DMP1 DMP1 sequences have been reported from several mammal [2,12–16], reptile [17], and bird species [18]. In all species, DMP1 begins with a hydrophobic leader sequence composed of 16–21

amino acids. The acidity of the DMP1 molecule in all species is due to high quantities of Asp (12–13%) and Glu (12–15%) [18]. The average isoelectric point (pI) of DMP1s is calculated at 3.9 from their amino acid compositions. The conservation of high acidity supports the view that the Asp and Glu residues are required for mineralization of the extracellular matrix. The most highly conserved amino acid in DMP1 is Ser (17–23%) [18], which is an essential residue of the sequence motif recognized by the casein kinases I and II and the target of phosphorylation catalyzed by these enzymes [25]. Moreover, the Ser positions and indeed all of the phosphorylation motifs are highly conserved among all DMP1s sequenced thus far [18]. DMP1 amino acid sequences deduced from rat DMP1 cDNA indicated 65 potential phosphorylation sites [2]. Phosphate analysis of the DMP1 extracted from rat long bone revealed that a full-length DMP1 contained 53 phosphates [26]. Because phosphates are negatively charged, their large numbers in DMP1, as well as DMP1’s acidic nature, suggest a high calcium ion-binding capacity as is necessary for matrix proteins participating in mineralization. Recombinant rat DMP1, 90–100 kDa in size, has been synthesized [27,28], while the native forms of DMP1 isolated from dentin and bone were 37-kDa and 57-kDa fragments [26]. Peptide sequencing of these fragments demonstrated that 37-kDa fragments originated from the NH2-terminal region while the 57-kDa fragments were from the COOH-terminal part of DMP1. It also indicated that rat DMP1 is proteolytically cleaved at four bonds: Phe189-Asp190, Ser196-Asp197, Ser233-Asp234, and Gln237-Asp238 [26,29]. Recently, a native full-length form (105 kDa) of DMP1 has been detected in the extracellular matrix of bone and dentin at a considerably lower level than those of the processed 37-kDa and 57-kDa fragments [29]. The uniform position of cleavage sites at the NH2-terminal peptide bonds of Asp residues (X–D bonds) may indicate involvement of a single proteinase. Bone morphogenetic protein 1 (BMP1)/tolloid-like metalloproteinase cleaved one site (Ser212Asp213 of mouse DMP1 corresponding to Ser196-Asp197 in the rat) of the above four possible sites, leading to the production of 37-kDa and 57-kDa fragments [30]. It has been suggested that PHEX (a phosphate-regulating gene with homologies to endopeptidases on the X chromosome) might contribute to DMP1 processing, since PHEX and DMP1 are highly expressed in osteocytes [28,31] and PHEX has demonstrated preference for cleavage sites with Asp (D) in the S10 position [32]. However, the coexpression of PHEX and DMP1 in 293EBNA had no apparent effect on DMP1 cleavage, suggesting that PHEX may not be required for DMP1 processing [33]. Instead, the processing of

S. Toyosawa et al. / Journal of Oral Biosciences 54 (2012) 30–36

DMP1 in transfected CHO cells was blocked by a furin protease inhibitor [33]. DMP1 specifically binds to the N-telopeptide region (EMSYGYDEKSAGVAVP) of type I collagen, and the two collagen-binding sites (349DSESSEEDR357 and 424SEENRDSDSQDSSR437) of DMP1 have been identified [34]. The binding of DMP1 to type I collagen was shown to be involved in the accelerated assembly of collagen fibrils and apatite deposition at the collagen fibril in vitro [34]. A recent study showed that glycosaminoglycans (mainly chondroitin 4-sulfate) are linked to the NH2-terminal 37-kDa fragment of DMP1 via Ser74, located in the Ser74-Gly75 dipeptide [35], and the dipeptide’s flanking regions are highly conserved among a wide range of species from reptiles to mammals [18], suggesting that the glycosaminoglycans-linked form may have functional significance.

33

phosphorylated, with negative-charged protein involved in matrix mineralization. Accurate determination of DMP1 expression involving the replacement of DMP1 exon 6 with a LacZ reporter gene showed a signal in hypertrophic chondrocytes, osteoblasts, osteocytes, pulp cells, odontoblasts, cementoblasts, and ameloblasts, but not in any soft tissues such as the brain, during both the embryonic and postnatal stages [41]. However, several reports have demonstrated the presence of DMP1 in non-mineralized tissues such as the brain, salivary glands, kidney, liver, muscle, pancreas, and some tumors [42–47].

6. Functions of DMP1 6.1. DMP1 promotes mineralization in vitro

5. Temporospatial localization of DMP1 Transcripts of the DMP1 gene were originally detected by Northern blot in rat odontoblasts by George et al. [2]. The same group also demonstrated highly restricted DMP1 mRNA expression in mature secretory odontoblasts using in situ hybridization [3,36]. Later, other groups showed that DMP1 mRNA was also present in other mineralized tissue cells including osteoblasts, cementoblasts, and ameloblasts by in situ hybridization [16,37]. In the chicken, which secondarily lost the tooth-forming ability, DMP1 mRNA is detected only in bone tissues, indicating its important role in bone rather than dentin [18]. In the bones of chickens and rats, DMP1 mRNA is predominantly expressed in osteocytes, and not in osteoblasts, and this protein is localized in the bone matrix surrounding osteocytes and their processes [28] (Fig. 3). Fracture healing is a suitable model in which what types of cells express DMP1 is determined because a large number of osteoblasts or chondrocytes are observed during fracture healing. Using this model, Toyosawa et al. [38] confirmed that DMP1 was expressed in osteocytes but not in osteoblasts or chondrocytes. Kalajzic et al. [39] also showed DMP1 expression in osteocytes, but not in osteoblasts, in embryonic 18-day calvaria, postnatal 13-day long bone, and 21-day jawbone. In contrast, other study during various stages of development showed that from embryonic to postnatal stage, DMP1 firstly appeared in hypertrophic cartilage and then in osteoblasts, and later DMP1 was expressed strongly in osteocytes [24]. A critical increase in the expression of casein kinase II in osteocytes developing from osteoblasts [40] correlated with the presence of DMP1 in the bone matrix surrounding the developing osteocytes. Considering that DMP1 contains numerous consensus sites for phosphorylation by casein kinases I and II [2], when situated in a surrounding matrix of osteocytes it would become highly

DMP1 localization at sites of mineralization tissues, along with a longstanding hypothesis that phosphorylated proteins of bone and dentin are involved in the mineralization process [48], have promoted studies on the effects of DMP1 on this activity. Boskey et al. [49] demonstrated that recombinant DMP1 promoted hydroxyapatite formation in a gelatin gel system. DMP1 was shown to nucleate the formation of hydroxyapatite in vitro in a multistep process that began by DMP1-binding calcium ions and initiating mineral deposition, and the intermolecular assembly of DMP1’s acidic clusters into a beta-sheet template was essential for the mineral nucleation [50]. Further, in vitro mineralization studies demonstrated that apatite was deposited only at collagen-bound DMP1 sites [34]. Full-length recombinant DMP1, post-translationally modified native full-length DMP1, and its COOH-terminal fragment isolated from bone were all able to nucleate hydroxyapatite in the presence of type I collagen, although the NH2-terminal fragment of DMP1 (amino acid residues 1–334) inhibited hydroxyapatite formation and stabilized the amorphous phase [51]. Gericke et al. [52] showed that NH2-terminal and COOH-terminal fragments of DMP1 were both promoters of hydroxyapatite formation and growth, while a chondroitin sulfate-linked NH2-terminal fragment of DMP1 was an inhibitor. Tartaix et al. [53] reported complicated data demonstrating that the non-phosphorylated form of prokaryote recombinant DMP1 worked as a hydroxyapatite nucleator while its phosphorylated form had no effect, and that phosphorylated eukaryote recombinant full-length DMP1 inhibited mineralization whereas highly phosphorylated COOH-terminal fragments of DMP1 extracted from rat bone initiated mineralization. Narayanan et al. [4] showed that overexpression of DMP1 in both MC3T3-E1 cells and C3H10T1/2 (pluripotent embryonic undifferentiated mesenchymal) cells enhanced the onset of mineralization and the size of the mineralized nodule when compared with the

bone

Fig. 3. mRNA expression and protein localization of DMP1 in bone: (a) osteoblast is present on the surface of bone (arrows) while osteocytes are located within osteoid and bone matrix (hematoxylin and eosin); (b) DMP1 mRNA is expressed in osteocytes, but not in osteoblasts; and (c) DMP1 is localized in bone matrix surrounding osteocytes and their processes.

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mock cells. Further, Feng et al. [24] reported that DMP1 expression is associated with bone nodules using rat primary osteoblastic cells. 6.2. DMP1 function in bone in vivo To study the function of DMP1 in vivo, DMP1-null mice were created by replacing exon 6 with a lacZ reporter gene [41]. DMP1null embryos and newborns displayed no apparent abnormal phenotype although they had a modest increase in the size of hypertrophic chondrocyte zone and in long bone diameter, suggesting that DMP1 is not essential for early skeletal development [41]. Ye et al. [5] reported that DMP1-null mice during the postnatal development phase (3 days–12 months) showed a chondrodysplasia-like phenotype with a highly expanded growth plate and delayed secondary ossification centers, indicating that DMP1 is essential for normal postnatal chondrogenesis. Ling et al. [6] reported that DMP1-null mice (4 and 16 weeks) showed osteomalacia with decreased mineral content compared to backgroundmatched heterozygous and wild-type controls, and indicated a key role for DMP1 in bone mineralization. Interestingly, DMP1-null mice had significantly reduced serum concentrations of phosphate as contrasted with heterozygous and wild-type mice [6]. This finding led to the discovery of the DMP1 mutation in autosomalrecessive hypophosphatemic rickets (ARHR). ARHR is a condition characterized by rickets and osteomalacia with elevated FGF23 (fibroblast growth factor 23) serum levels and hypophosphatemia; the abnormalities are also seen in DMP1-null mice [7,8]. FGF23, a phosphate-regulating factor that is secreted hormonally from osteoblast/osteocyte, causes increased renal phosphate wasting, leading to hypophosphatemia [54]. DMP1-null mice had increased FGF23 mRNA expression in osteocytes and defects in the maturation of osteoblast into osteocyte [7], indicating that DMP1 is critical for regulating FGF23 and osteocyte maturation. A high-phosphate diet cured rickets but not osteomalacia in DMP1-null mice [7]. ARHR patients exhibit skeletal deformities [7,8,55–57] as well as dental defects [56–58]. Patients also experience hypophosphatemia as a consequence of FGF23-induced renal phosphate wasting as well as low levels of serum 1,25-dihydroxyvitamin D [1,25(OH)2D], whereas serum calcium, parathyroid hormone (PTH), and urinary calcium excretion are normal [7,8,55–57]. There have been several reports of ARHR families with mutations of the DMP1 gene (1A4G [7,8], 55-1G4 C [8], 98G4A [55], IVS51G 4A [57], 362delC [8], 485delT [56], 1484–1490del [7])

Signal peptide

Glycosaminoglycan Cleavage site attachment

NH2-

Protein

(Fig. 4). Transient transfection analysis of DMP1 mutants causing ARHR showed that the Met1Val (1A4G) DMP1 mutant was not sorted to the secretory pathway, and that the 1484–1490del mutant demonstrated replacement of the 18 COOH-terminal amino acid residues with 33 non-native residues, likely compromising DMP1 processing [59]. Protein chemistry showed that rat DMP1 was proteolytically processed into NH2-terminal 37-kDa and COOH-terminal 57-kDa fragments at the critical cleavage bond Ser196-Asp197 [26]. In vitro studies showed that the substitution of residue Asp213 (corresponding to Asp197 in rat DMP1) by Ala213 at a cleavage site blocked the processing of mouse DMP1 in HEK293 cells [60]. Sun et al. [61] generated transgenic mice having a mutant DMP1 (Asp213 replaced by Ala213) but no wild-type DMP1 in osteoblasts under a 3.6-kb rat alpha 1(I) collagen (Col1) promoter [62]. The substitution of Asp213 by Ala213 in mouse DMP1 blocked the cleavage of DMP1 in bone and overexpression of mutant DMP1 had no effect on the DMP1-null background [61]. The COOHterminal 57-kDa fragment has a highly phosphorylated domain as a hydroxyapatite nucleator [51,53]. In addition, the discovery of ARHR, in which DMP1 has a normal NH2-terminal 37-kDa fragment and a mutant COOH-terminal 57-kDa fragment (1484–1490del) [7], suggests that the COOH-terminal 57-kDa fragment may be the primary functional domain of DMP1. To study the in vivo role of the COOH-terminal 57-kDa fragment, transgenic mice overexpressing full-length DMP1 or the 57-kDa fragment in osteoblasts controlled under the 3.6-kb rat Col1 promoter were generated [33]. Overexpression of both transgenes had no effect on the skeleton of wild-type background; however, overexpression of both transgenes on the DMP1-null background resulted in the rescue of skeletal abnormalities including osteomalacia, abnormal osteocyte maturation, elevated serum FGF23 levels, and hypophosphatemia, indicating that the COOH-terminal 57-kDa fragment is the functional domain of DMP1 [33,63]. Although there is a little information of dental abnormalities in ARHR, the ARHR patients with 485delT (leading to a premature stop codon in exon 6) showed wider pulp chamber, thinner dentin and enamel [56], the ARHR patients with IVS5-1G4A showed dental abnormalities [57], and the ARHR patients with 1484–1490del showed dentin and enamel defect similar to a dentinogenesis imperfecta (DI) III-like phenotype [58]. These dental abnormalities may be due to systemic hypophosphatemia and local deficient mineralization by mutant DMP1. However, there is no denying that local FGF23 signaling may affect dentin abnormalities since DMP1-null mice showed increased FGF23 mRNA expression also in odontoblasts [58].

37 kDa

Conserved C-terminus

57kDa

-COOH

Start codon Gene

Mutation

E1

TGG ATG AG AG E2 E3 E4 E5 AC AA TAG GTG

Stop codon CCC CCT

TGCCTATCACAA E6

CC# CC#

1A>G 55-1G>C 98G>A IVS5-1G>A 362delC 485delT

TGC#######AA

1484-1490del

Fig. 4. Schematic presentation of wild-type DMP1 protein and the DMP1 gene mutation identified in ARHR. DMP1 is cleaved into secreted NH2-terminal 37-kDa and COOH-terminal 57-kDa fragments. DMP1 mutation sites in ARHR are shown relative to the known proteolytic cleavage site.

S. Toyosawa et al. / Journal of Oral Biosciences 54 (2012) 30–36

7. Concluding remarks It has been almost two decade since the initial discovery of DMP1 in a tooth cDNA library by George et al. [2]. At the beginning, DMP1 was considered merely to be a component of noncollagenous matrix and generated little interest. However, studies over the last decade have increased the understanding of DMP1 as a local and systemic regulator of mineralization and phosphate regulation. During this period, there have been several landmark studies of DMP1, including those related to the gene’s molecular evolution, promoter, osteocyte-specific expression, and proteolytic cleavage processes, as well as the phenotype of DMP1-null mice with hypophosphatemia and elevated serum FGF23 and the discovery of DMP1 mutations in ARHR. These outcomes will provide new insight into the biology of mineralization and potential therapeutics.

Conflict of interest No potential conflicts of interest are disclosed.

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