Bone 107: 196-207

PMC: PMC5803363

PMID: 29313816

Osteopontin regulates dentin and alveolar bone development and mineralization

M Yadav+4 authors

1. Introduction

The periodontium is the attachment complex for the tooth, and includes cementum, PDL, and alveolar bone. Cementum is a mineralized tissue covering the tooth root that serves as an interfacial connective tissue between the tooth root dentin and the PDL [1,2]. While alveolar bone is continually remodeled by osteoblast, ostecyte, and osteoclast activities to adapt to mechanical loading associated with tooth function, cementum does not undergo physiological remodeling, but forms by continual slow apposition throughout life, mediated by activities of cementoblasts. The acellular cementum of the cervical tooth root is marked by concentrated depositions of ECM proteins including bone sialoprotein (BSP) and osteopontin (OPN), multifunctional phosphoproteins thought to regulate biomineralization and contribute to cell signaling [3–9]. The apposition and mineralization of acellular cementum is strongly regulated by local mineral metabolism, especially levels of pyrophosphate (PP_i), a potent inhibitor of hydroxyapatite crystal growth. PP_i levels are controlled by tissue-nonspecific alkaline phosphatase (Alpl/TNAP), progressive ankylosis protein (Ank/ANK), and ectonucleotide pyrophosphatase phosphodiesterase 1 (Enpp1/ENPP1), and genetic ablation of any of these factors has dramatic effects on acellular cementum formation [10–14]. Cellular cementum surrounds the apical root and its formation has been reported to be less influenced by mineral metabolism [11,13].

The “sandwich” arrangement of cementum/PDL/bone maintains flexibility of the tooth in the socket, provides a means for distribution of the forces from occlusion, allows for mechanoresponsive cells to direct remodeling and repair, and maintains vascular, lymphatic, and nerve supply to the periodontium. For proper maintenance of periodontal function, the PDL must be maintained as an unmineralized fibrous tissue, though it lies between two mineralized tissues (bone and cementum), harbors osteo- and cemento-progenitor cells, and is composed of a fibrillar collagenous matrix rich in pro-mineralization factors, such as the enzyme TNAP [1,13,15,16]. While regulators of cementum and bone mineralization have been partially defined, it remains unclear what factors are essential for maintaining the cementum-PDL and bone-PDL borders, and for preserving the PDL in its unmineralized state. It has been hypothesized that a balance of locally expressed regulators allow for continued cementum growth and adaptation of alveolar bone, while the PDL space is maintained. This question, while both clinically relevant and critical to understanding the basic biology of the periodontium, may also lie at the heart of the evolution of the periodontium (tooth in socket, or gomphosis joint), as the ability to promote and restrict mineralization in a site-specific fashion would be key for creating and maintaining these hard-soft interfaces. This evolutionary concept is supported by studies pointing to an ancient origin of cementum and documenting variable regulation of periodontal mineralization in extinct and extant species [17–21].

Our goal in this study was to identify factors contributing to the regulation of mineralization in the periodontium. As a first approach, we performed proteomic analysis on PDL from Ank^{^/}^{mice to identify factors altered (compared to normal controls) under conditions of rapid cementogenesis and subsequent PDL-bone remodeling. From this screening, we identified the protein OPN as a factor of interest. We then analyzed developmental expression of}Spp1/OPN in the periodontium, effects of Spp1 ablation on dental and periodontal formation, and potential in vivo functional role(s) of OPN in hypermineralization of cementum.

2. Materials and methods

2.1. Mice

Animal experiments complied with ARRIVE guidelines [22]. All animal experiments were approved by the Animal Care and Use Committee (ACUC) of the National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS, Bethesda, MD) the Institutional Animal Care and Use Committee (IACUC) of the Sanford Burnham Prebys Medical Discovery Institute (La Jolla, CA), or the IACUC of The Ohio State University (Columbus, OH). Previous publications described the generation, colony maintenance, and genotyping of mice null for the progressive ankylosis protein gene (Ank^{) on a mixed C57BL/6 and 129S1/SvlmJ background [}11,23], mice null for the osteopontin gene (Spp1^{) on a C57BL/6 background [}24,25], mice null for tissue-nonspecific alkaline phosphatase (Alpl^{) on a mixed C57BL/6 and 129S1/SvlmJ background [}26,27], and mice null for both Spp1 and Alpl (Alpl^;Spp1^{) on the same mixed background [}24,27]. Mice deficient for both Ank and Spp1 (Ank^;Spp1^{) were created by breeding double heterozygote males and females on a mixed C57BL/6, 129S1/SvlmJ, and 129/CD1 background. Three to six mice were analyzed per genotype (unless otherwise noted) at ages including 10, 14, 26, 30, 60, 90, and 240 days postnatal (dpn). Male and female mice did not show significant differences from one another, therefore, both were included in analyses. Mice were housed in standard mouse cages in a 12-hr light-dark cycle and with access to standard rodent chow and water ad libitum.}

2.2. Laser capture microdissection and protein extraction

Tissues from 60 dpn wild-type (WT) control and Ank^{mice (}n = 5 and 6, respectively) were used for laser capture microdissection (LCM). Mandibles fixed in Bouin’s solution, decalcified in AFS (acetic acid, formalin, and sodium chloride) solution, and embedded in paraffin were serial sectioned around the first molar region [4]. Tissues were deparaffinized by submerging twice in xylene for 2 min, followed by an additional xylene wash for 5 min. Tissues were then air dried and immediately microdissected. LCM of PDL was performed using combined infrared (IR) and ultraviolet (UV) laser cutting on an Arcturus XT Micro-dissection Instrument (Applied Biosystems, Waltham, MA, USA) with adjusted settings: UV cutting speed of 300 mm/s; IR laser power of 70–80 mW, duration of 20 ms, and spot size of 30 μm. The total micro-dissected areas of WT and Ank^{^/}^{PDL samples were calculated (average of 302 ± 123 μm}^{) and compared (}p = 0.66, t-test). The captured area was used to normalize the amount of tissue/protein for liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis. Caps containing the captured tissues were incubated with 30 μL of 8 M urea for 30 min at room temperature for protein extraction. Samples were sonicated and centrifuged briefly. Whole protein extracts were reduced, alkylated, and trypsin digested as described previously [28–30]. The resulting tryptic peptide samples were dried in a vacuum concentrator and reconstituted in 0.1% formic acid for analysis.

2.3. LC-MS/MS and bioinformatics analysis

Peptide mixtures were analyzed in an electron transfer dissociation (ETD)-enabled LTQ Orbitrap Velos mass spectrometer (Thermo Fisher Scientific, Bremen, Germany) connected to the EASY-nLC system (Proxeon Biosystem, West Palm Beach, FL, USA) through a Proxeon nanoelectrospray ion source in a data-dependent mode. Peptides were separated by a 2–90% acetonitrile gradient in 0.1% formic acid using a PicoFrit analytical column (20 cm × 75 μm internal diameter, 5 μm particle size; New Objective, Woburn, MA), at a flowrate of 300 nL/min for 45 min. The nanoelectrospray voltage was set to 1.7 kV with the source temperature at 275 °C. All instrumental parameters were set up in data-dependent acquisition mode. The Orbitrap analyzer acquired the full scans of MS spectra (m/z 300–1600) after accumulation to a target value of 1e6, with a resolution setting of r = 60,000. The 5 most intense peptide ions with charge states ≥2 were sequentially isolated to a target value of 5000 and fragmented in the linear ion trap by low energy collision induced dissociation (CID; normalized collision energy of 35%). The signal threshold to trigger an MS/MS event was set to 500 counts. Dynamic exclusion was enabled with an exclusion size list of 500, exclusion duration of 60 s, and repeat count of 1. An activation q = 0.25 and time of 10 ms were used.

Protein identification was performed with the Sequest search engine (Thermo Finnigan, San Jose, CA, USA; version 1.4.0.288) employing Pro-teome Discover version 1.3 (Thermo Fisher Scientific). The MS/MS spectra (msf) generated from raw files were searched against the UniProt Mouse Protein Database (released 22 January 2014; 51,116 entries) with parameters set to a maximum of one missing cleavage, with a parent ion tolerance of 10.0 ppm for MS search and a fragment ion mass tolerance of 1.0 Da. Oxidation of methionine (+16 Da) was set as a variable modification, and carbamidomethylation in cysteine residues (+57 Da) was set as a fixed modification. For label-free protein quantification, the data files were analyzed in Scaffold Q+ (version Scaffold_4.6.1, Proteome Software, Inc., Portland, OR, USA). The quantitative value (normalized spectral counts) was obtained with the protein thresholds set at a minimum 99% probability to achieve a false discovery rate (FDR) b1%, and containing at least one peptide with thresholds established at a minimum 60% probability, XCorr cutoffs +1 > 1.8, +2 > 2.2, +3 > 2.5, and +4 > 3.5. Resulting spectrum count values were used to analyze the distribution of identified proteins throughout the samples. Proteins with at least one valid value were considered for further analysis. Independent samples t-test was applied for testing for differences in protein intensities between WT and Ank^{^/}^{sample groups. Protein ratios were calculated from the average of normalized spectral protein intensities.}

2.4. Microcomputed tomography

For quantitative microcomputed tomography (micro-CT) analysis, dissected and formalin fixed hemi-mandibles (n = 3–7 per genotype at ages 30, 60, 90, and 240 dpn, unless otherwise noted) were scanned in 70% ethanol on a Scanco μCT 50 (Scanco Medical, Brüttisellen, Switzerland) with parameters: 70 kVp, 76 μA, 0.5 Al Filter, 900 ms integration time, and 6 μm voxel size. DICOM images were uploaded to AnalyzePro 1.0 (AnalyzeDirect, Overland Park, KS, USA) and calibrated to five known densities of hydroxyapatite (mg/cm^{HA). Mandibles were reoriented to a standard position in 3D space using first molar anatomical landmarks: In mid-molar sagittal section, a line through mesial and distal cemento-enamel junctions (CEJ) was made parallel to the transverse plane (}Supplementary Fig. S1A); in mid-root (halfway from CEJ to apex) transverse section, mesial and distal roots were made parallel to the sagittal plane (Supplementary Fig. S1B); in mid-root coronal section, the mesial root canal was made parallel to the z-axis (Supplementary Fig. S1C). The region of interest (ROI) for analysis (first molar and surrounding bone) was then defined by cropping the mandible 480 μm mesial and 240 μm distal from the most mesial and distal aspects of the first molar cusps, respectively. This ROI was defined to include the mesial rise of the alveolar bone in the diastema and the interproximal bone between the first and second molars (Supplementary Fig. S1D–F). Enamel was segmented at a threshold of 1600 mg/cm^{, and dentin/cementum and bone at 650 mg/cm}^{. Pulp and PDL spaces were segmented semi-automatically using AnalyzePro. Tissue segments were analyzed for total volume and mineral density. Further subdivisions of mandibular bone were made for refined analysis, including 7 components: basal bone, buccal/lingual alveolar (tooth-associated) bone, and cervical/mid/apical alveolar bone (}Supplementary Fig. S1G–I). Basal bone was defined as mandibular bone located inferior to the most apical aspect of the distal root. The buccal-lingual divide was determined by mid-sagittal section, and the cervical/mid/apical regions were defined by dividing the length of the distal root (CEJ to apex) into three equal sections. Root dentin/cementum, pulp, and PDL tissues were subdivided into equal halves or thirds for additional analyses. Tooth width was measured as mesial-distal and buccal-lingual linear distances at the CEJ. Root lengths for mesial and distal roots were measured as linear distances from CEJ to apices.

2.5. Histology, immunohistochemistry, and in situ hybridization

Samples used for histology were prepared as described previously [4]. In brief, harvested hemi-mandibles were fixed in Bouin’s solution, decalcified in AFS solution, and serial 6 μm paraffin sections were prepared by microtome. Histological sections were stained by hematoxylin and eosin (H&E), toluidine blue (TB), alcian blue/nuclear fast red (AB-NFR), or picrosirius red (PR). Tartrate-resistant acid phosphatase (TRAP) staining for osteoclast-like cells was performed according to manufacturer’s instructions (Wako Chemical, Japan) as previously described [3,8,31]. TRAP-positive osteoclast-like cells were enumerated on alveolar bone surfaces over the inner circumference of the first molar mesial root socket in mandibles sectioned in the coronal orientation. Positive controls for TRAP staining included sections from mouse ligature-induced periodontitis featuring numerous osteoclasts on alveolar bone and tooth root surfaces [8].

Immunohistochemistry (IHC) was performed on deparaffinized histological sections using an avidin-biotinylated peroxidase (ABC) based kit (Vectastain Elite, Vector Labs, Burlingame, CA, USA) with a 3-amino-9-ethylcarbazole (AEC) substrate (Vector Labs) to produce a red-brown product. Primary antibodies included: rabbit polyclonal LF-150 anti-mouse fibromodulin (FMOD; Dr. Larry Fisher, NIDCR, Bethesda, MD), rabbit polyclonal LF-175 anti-mouse osteopontin (OPN; Dr. Larry Fisher, NIDCR, Bethesda, MD) [4,11,27] and goat polyclonal anti-human ENPP1 (Abcam, Cambridge, MA, USA). Hematoxylin was used as a counterstain.

In situ hybridization (ISH) with antisense Spp1, Col1a1, Ibsp, Bglap, and Enpp1 probes was performed on deparaffinized histological sections and visualized with fast red dye (Advanced Cell Diagnostics, Newark, CA), using hematoxylin as a counterstain, as previously described [32].

2.6. Histomorphometry

Acellular cementum thickness was measured on WT control and Spp1^{^/}^{mouse first molar mesial roots at 14, 30, 90, and 240 dpn (}n = 3–6 for both genotypes unless otherwise stated). At 14 dpn, measurements were made at a distance of 100 μm from the cemento-enamel junction (CEJ), while at later ages, measurements were made at 300 μm from the CEJ on these longer roots. At ages older than 14 dpn, cellular cementum area was measured in central mesial root sections where buccal and lingual cellular cementum could be observed. Acellular cementum thickness was measured at 300 μm from the CEJ in WT control, Spp1^{^/}^,Ank^{^/}^{, and}Spp1^{^/}^;Ank^{^/}^{double knock-out mouse first molar mesial roots at 60 dpn (}n = 5–6 for all genotypes). Linear measurements were performed using a digital slide scanner and Digital Image Hub software (version 4.0.7; Leica Biosystems, Nussloch, Germany), while cellular cementum area was measured using ImageJ software (version 1.49d; National Institutes of Health, Bethesda, MD, USA). Statistical analyses were performed using GraphPad Prism (version 6.01; La Jolla, CA. USA).

2.7. Quantitative real-time polymerase chain reaction

For analysis of WT and Spp1^{^/}^{mouse periodontal tissues, first mandibular molars were extracted at 26 dpn from}n = 3 mice (for each genotype) and total RNA was collected and analyzed as described above, according to manufacturer instructions. The qPCR array was a 96-well plate that featured a customized set of 84 target gene primers (see Supplementary Table S1 for full list), and included 5 housekeeping genes, positive PCR controls, and reverse transcriptase controls in every plate, and amplification results were analyzed and statistics performed using proprietary software (RT2 Profiler PCR Array Data Analysis version 3.5; Qiagen).

2.8. Statistical analysis

Results are expressed as mean ± standard deviation (SD) in the graphs presented. Data were analyzed using Student’s (independent samples) t-test or one-way analysis of variance (ANOVA) with post-hoc Tukey test for multiple comparisons, where p-values < 0.05 were considered statistically significant. Statistical analyses were computed using GraphPad Prism (version 6.01 for Windows GraphPad Software, La Jolla, CA, USA).

2.1. Mice

2.2. Laser capture microdissection and protein extraction

2.3. LC-MS/MS and bioinformatics analysis

2.4. Microcomputed tomography

2.5. Histology, immunohistochemistry, and in situ hybridization

2.6. Histomorphometry

2.7. Quantitative real-time polymerase chain reaction

2.8. Statistical analysis

3. Results

3.1. Proteomic analysis of PDL in Ank^{versus control mice}

The progressive ankylosis protein knock-out (Ank^{) mouse features rapidly growing acellular cementum, yet maintains the width of PDL, apart from ectopic “cementicles” sometimes present in the cervical PDL space [}11,14]. We used this model for proteomic identification of factors involved in regulation of mineralization in the periodontium and maintenance of cementum-PDL-bone borders. Fixed and paraffin-embedded 60 dpn WT and Ank^{mandible tissues were used, with sections of first molar teeth deparaffinized and prepared for laser capture microdissection (LCM) of PDL tissue (}Fig. 1A, B). Total proteins identified included 763, with 496 proteins found in both WT and Ank^{mouse PDL samples, 130 exclusive to WT, and 137 exclusive to}Ank^{mice (}Fig. 1C). Proteins N1.5-fold different with statistical significance (p < 0.05) in Ank^{vs. WT PDL samples are plotted in a Volcano plot in}Fig. 1D and are listed in Supplementary Table S2. Four factors were significantly increased in Ank^{mice (ranging from 0.8 to 6.6-fold), whereas 10 factors were significantly decreased (ranging −0.7 to −5.0-fold).}

An external file that holds a picture, illustration, etc.
Object name is nihms927401f1.jpg

Fig. 1

Proteomic analysis of PDL in Ank^{^/}^{versus control mice and confirmation. Histology sections of 60 dpn}Ank^{^/}^{first mandibular molar (M1) tooth (A) before and (B) after laser capture microdissection (LCM) of periodontal ligament (PDL) tissues on lingual and buccal aspects. (C) Proteomic analysis of PDL tissues collected from 60 dpn}Ank^{^/}^{and WT mice (}n = 6 and 5, respectively) identifies 137 peptides exclusively localized to Ank^{^/}^{mouse PDL, 130 peptides exclusively localized to WT mouse PDL, and 496 peptides detectable in both. (D) Volcano plot of proteomic analysis indicating proteins identified as significantly increased (upper right quadrant) or significantly decreased (upper left quadrant). Identities of these proteins and their fold-change differences are shown in}Supplementary Table S1. (E) Immunohistochemistry for OPN in 60 dpn WT molar tissues shows intense OPN localization in acellular cementum (AC) and alveolar bone (AB). (F) In situ hybridization for Spp1 in 60 dpn WT molars shows minimal constitutive expression in cementoblasts (Cb) or osteoblasts (Ob) at this age. In Ank^{^/}^{mice at 60 dpn, (G) strong immunolocalization of OPN to thick AC, AB surface, and ectopic cementicles (yellow star) is mirrored by (H) intense expression of}Spp1 mRNA in Cb and Ob. DE = dentin. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Among the decreased factors in Ank^{mice was fibromodulin (FMOD), a small leucine-rich proteoglycan (SLRP) known to modulate collagen fibril formation in the PDL and implicated in periodontal homeostasis [}33–35]. Immunohistochemistry for FMOD revealed strong staining patterns in both Ank^{and WT periodontia at 60 dpn (}Supplementary Fig. S2A, B).

A pool of potential candidates for periodontal regulation was identified a priori, including ECM proteins, proteoglycans, and factors known to regulate biomineralization and/or be involved in PDL biology (Supplementary Table S3). This list allowed for confirmation of the approach and analysis, as several proteins previously identified in PDL were confirmed, including collagen type 1 (COL1A1), collagen type XII (COLXIIA) [36], decorin (DCN) [33], tissue nonspecific alkaline phosphatase (ALPL; also called TNAP, TNSALP, and TNALP) [11,13], and periostin (POSTN) [37,38]. Among these a priori identified factors, the ECM protein osteopontin (OPN) was detected in Ank^{mouse PDL samples, but was below detection limits in WT controls. We analyzed the presence of OPN in WT and}Ank^{mouse periodontia using IHC. In WT periodontia at 60 dpn, OPN was present in the acellular cementum layer and alveolar bone (}Fig. 1E). Ank^{mice featured intense staining for OPN in the thick cementum, and near cementum and alveolar bone surfaces at 60 dpn (}Fig. 1G). The ectopic cementicles in Ank^{mouse PDL showed intense OPN localization (yellow star in}Fig. 1G). In situ hybridization (ISH) for the OPN-encoding gene, Spp1, revealed little expression in WT cementoblasts or alveolar bone osteoblasts at 60 dpn (Fig. 1F), however intense expression in both cementoblasts and osteoblasts of Ank^mice.

OPN was further analyzed for its functional importance in dental development because of its identified functions as a negative regulator of hydroxyapatite mineralization in vitro [39–42], a modulator of bone mineral density in vivo [24,25,43–46], and a response protein to ectopic calcification in several soft tissues [47–50].

3.2. Osteopontin is expressed in the periodontium during tooth root development

To provide insights on the potential role(s) of OPN in periodontal tissue formation, we analyzed WT periodontal tissues for localization of Spp1 mRNA during development using ISH. During early root formation at 14 dpn, Spp1 was present in numerous osteoblasts of the alveolar bone, new cementoblasts near the apical root (though only a minority of cementoblasts along the entire root), and some odontoblasts (Fig. 2A, B). At 26 dpn, Spp1 was expressed in bone cells, few cementoblasts along the acellular cementum, many cementoblasts surrounding the cellular cementum, and cementocytes in the cellular cementum (Fig. 2C, D). This Spp1 expression pattern is distinct from that of two other osteoblast/cementoblast markers, Ibsp and Alpl, in the dental/periodontal region. Unlike the transience of Spp1 mRNA, the closely related gene Ibsp was continuously expressed by cementoblasts along the entire root surface (Fig. 2E, F) [8]. Another marker of cementoblasts and regulator of cementum, Alpl, was widely expressed in the periodontium after root initiation (Fig. 2G, H) [13].

An external file that holds a picture, illustration, etc.
Object name is nihms927401f2.jpg

Fig. 2

Osteopontin is expressed in the periodontium during development. In situ hybridization (ISH) for Spp1 mRNA (red colour) was used to determine cellular expression during mouse molar root formation and function (n = 3–5 for each age). (A, B) In the developing first molar (M1) root at 14 dpn, Spp1 is present in numerous osteoblasts (Ob) of the alveolar bone (AB) and is expressed by new cementoblasts (Cb) near the apex. Spp1 mRNA is not detected in the cells of the periodontal ligament (PDL). (C, D) At 26 dpn, where cellular cementum (CC) is forming, Spp1 is associated with bone cells and Cb surrounding the cellular cementum (CC). (E, F) Ibsp is continuously expressed by Cb all along the root surface, as well as by many Ob of the AB. (G, H) Alpl is widely expressed in the periodontium after root initiation, including in Cb, Ob, odontoblasts (Od), and cells in the PDL. DE = dentin. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

3.3. Loss of OPN affects dental and periodontal development and mineralization

Though OPN knockout (Spp1^{^/}^{) mice have been employed in some studies on dentoalveolar biology [}27,51,52], to our knowledge, no detailed analysis of dental development in these mice has been previously reported. To identify potential physiological function(s) of OPN in dentoalveolar tissues, we analyzed mice at ages from 14 to 240 dpn.

By micro-CT and histology, similar cellular and tissue organization was observed in Spp1^{^/}^{and WT control mouse dentoalveolar tissues over this range of ages (representative images at 30 dpn shown in}Fig. 3A–J). Histomorphometric measurements at 14–240 dpn revealed no significant differences in acellular cementum thickness (lingual or buccal aspects) in Spp1^{^/}^{vs. WT mice (}p > 0.05 in independent samples in all t-tests)(Fig. 3K), while Spp1^{^/}^{cellular cementum showed significantly increased area at both 30 (}p < 0.05) and 90 (p < 0.01) dpn (Fig. 3L). Quantitative micro-CT analysis was performed to further explore potential changes in three dimensional (3D) tissue volumes (as shown in Supplementary Fig. S1 and described in the Materials and Methods). These analyses revealed significantly increased volumes of dentin/cementum (necessarily analyzed together because of the inability to reliably segment them by density using these scan parameters), mandibular bone, and enamel in Spp1^{^/}^{vs. WT mice (}Fig. 3M, N and Supplementary Fig. S3A–F). As dentin/cementum volume significantly increased (20.7% and 32.9% at 30 and 90 dpn, respectively), dental pulp volume significantly decreased, supporting more rapid dentin accumulation in Spp1^{^/}^{mice. PDL volume around the first molar was significantly diminished in}Spp1^{^/}^{vs. WT mice by 90 dpn (}Fig. 3N and Supplementary Fig. S3F). Additional volumes and linear measurements of Spp1^{^/}^{and WT mouse molars, and ROI subdivision of dentin/cementum, bone, and PDL are described in the Appendix and}Supplementary Fig. S3 and S4.

An external file that holds a picture, illustration, etc.
Object name is nihms927401f3.jpg

Fig. 3

Loss of OPN affects dental and periodontal tissues. Panels A-F show representative 2D micro-CT images from WT control and Spp1^{^/}^{mouse mandibles at 30 dpn. Compared to WT first molar (M1) in (A) coronal, (B) sagittal, and (C) axial/transverse planes of section, (D-F) age-matched}Spp1^{^/}^{tissues in the same section planes exhibit longer roots, thicker dentin, and thicker alveolar bone (AB) and basal bone (BB) of the mandible. WT and}Spp1^{^/}^{M1 at 14, 30, 60, 90, and 240 dpn were analyzed using hematoxylin and eosin (H&E) staining (}n = 2–6 per genotype per age), with representative images from 30 dpn shown here. (G-J) Tooth root formation and tissue organization appear similar in WT and Spp^{^/}^{mice in the completed root at 30 dpn, including formation of acellular cementum (AC), periodontal ligament (PDL), alveolar bone (AB), dentin (DE), and cellular cementum (CC). CC is outlined in red dotted lines in H and J to denote borders. (K) 2D measurements at 14–240 dpn reveal no significant differences (}p > 0.05) in buccal or lingual AC in Spp1^{^/}^{vs. WT first mandibular molars (p > 0.05 in independent samples in all}t-tests). (L) Spp1^{^/}^{cellular cementum exhibits significantly increased area vs. WT at both 30 (*}p < 0.05) and 90 (** p < 0.01) dpn, though no differences at 240 dpn (p > 0.05). Graphs show individual measurements (circles, squares) and means (black line) ± standard deviation, and statistical results reflect only comparisons made between same side CC (buccal or lingual) only at the same age. Quantitative micro-CT analysis of tissue volumes at (M) 30 dpn (n = 4–5) and (N) 90 dpn (n = 3–6), reveals statistically significant changes in several tissues in Spp1^{^/}^{mice compared to WT controls. Boxes indicate numerical percent difference in enamel (gray), dentin/cementum (green), pulp (yellow), periodontal ligament (red), buccal alveolar bone (blue), lingual alveolar bone (pink), and basal bone (orange). Statistical results of independent samples t-test indicated by: n.s. = not significant, p > 0.05; * p < 0.05; ** p < 0.01; ***}p < 0.001. Absolute values are shown in graphs in Supplementary Fig. S3. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Quantitative micro-CT analysis was used to analyze tissue mineral densities at 30 and 90 dpn. Mineral density of Spp1^{^/}^{dentin/cementum was significantly increased compared to WT at 90 dpn, and pulp tissue density also showed significant increases over WT at both time points (}Fig. 4A, B). Enamel mineral density was found not to be significantly different (Fig. 4C) between genotypes. Overall mandibular bone density and PDL density were significantly greater in Spp1^{^/}^{mice vs. WT at both ages (}Fig. 4D, E). Tissue densities were further analyzed by subdivision. As observed for volume analysis above, collected density data suggested generalized changes rather than specific regional changes in cervical and apical dentin/cementum, buccal and lingual alveolar bone and basal bone, and upper, middle, and lower alveolar bone and PDL tissue densities (Supplementary Fig. S5).

An external file that holds a picture, illustration, etc.
Object name is nihms927401f4.jpg

Fig. 4

OPN regulates mineral density in dentoalveolar tissues. Quantitative micro-CT was employed to measure tissue mineral density in mg/cm^{hydroxyapatite (HA) at ages 30 and 90 dpn. Compared to WT controls,}Spp1^{^/}^{mice exhibit trends of increased mineral density in (A) dentin/cementum, (B) dental pulp, (D) mandibular bone, and (E) periodontal ligament (PDL), though no difference is observed in (C) enamel. Graphs show individual measurements (circles, squares) and means (black line) ± standard deviation. Statistical results of independent samples t-test indicated by: * p < 0.05; ** p < 0.01; ****}p < 0.0001.

Evaluation of a small cohort (n = 2 WT, n = 3 Spp1^{^/}^{) of mice at the advanced age of 240 dpn showed that volumes and densities of dentin/cementum, pulp, enamel, mandibular bone, and PDL were similar between}Spp1^{^/}^{and WT mice (}Supplementary Fig. S6A–J). While differential changes in WT and Spp1^{^/}^{dentoalveolar tissues over time had apparently reduced differences documented at earlier ages, one out of three}Spp1^{^/}^{mice featured nearly complete pulp obliteration by ectopic mineralized tissue (}Supplementary Fig. S6K–P). Histology of this 240 dpn Spp1^{mouse revealed an abnormal accumulation of ECM in the pulp space featuring few cells and no apparent organization consistent with dentin or bone (}Supplementary Fig. S6Q–T).

These analyses support a non-redundant role for OPN in regulating development and mineralization of dentin and alveolar bone, especially at early ages, and a role in regulating tissue properties of dental pulp and PDL.

3.4. Altered gene expression in periodontal tissues of Spp1^mice

Because of significant changes identified in volumes and densities of dentoalveolar tissues in Spp1^{^/}^{vs. WT mice, we analyzed whether there were alterations in gene markers for dentoalveolar cells. Localization of}Col1a1, an early marker of osteoblasts and odontoblasts, was similar in Spp1^{^/}^{and WT mice, marking odontoblasts, osteoblasts of the alveolar bone and basal bone at 14 and 26 dpn, and PDL cells at the later age (}Supplementary Fig. S7A–L). Localization of Ibsp, a marker for osteoblasts and cementoblasts showed similar localization on alveolar and basal bone surfaces and tooth root surfaces of Spp1^{^/}^{and WT mice at both ages (}Supplementary Fig. S7M–X). Bglap, a later marker for osteoblast and odontoblasts also showed similar localization, intensity, and positive cell numbers in in Spp1^{^/}^{and WT mouse molars (}Supplementary Fig. S7Y–JJ).

Unlike dentin and bone, loss of OPN appeared to have no substantial effect on development of acellular cementum or cementum-PDL interface, raising the possibility of compensatory factors acting in overlapping fashion in the periodontium. Ablation of Spp1 in mice was previously shown to cause elevated circulating PP_i (an inhibitor of mineralization) by reducing Alpl and elevating Ank and Enpp1 expression in osteoblasts of the postcranial skeleton [24,25,45]. In order to identify potential local compensators for loss of Spp1/OPN, we performed qPCR array on periodontal tissues (cementum-associated PDL) harvested from Spp1^{^/}^{and WT mice at 26 dpn. The qPCR array included primer pairs for 84 target genes, chosen to target mineral metabolism regulators (including}Alpl, Ank, and Enpp1), genes associated with ECM and mineralization, transcription factors, and signaling pathway elements (see Supplementary Table S1 for full list). This qPCR approach identified only 7 genes significantly up- or down-regulated by 2-fold or more (including Spp1) in Spp1^{^/}^{vs. WT mice, while 93% of genes (78/84 total) in this array showed no change (}Supplementary Table S4). Factors significantly increased included Mmp3 (5.35-fold) and AP-1 (3.39-fold), and those decreased included Phex (−2.06-fold), Nfic (−2.25-fold), Bglap (−3.10-fold), and Abcc6 (−3.45-fold). Based on known functions of these factors, the observed patterns of expression did not indicate a likely factor that might compensate for OPN in terms of regulating cementum mineralization. Moreover, in the Spp1^{^/}^{mouse PDL, no alterations were identified in expression of}Alpl, Ank, or Enpp1, or in closely related SIBLINGs including Ibsp, Dmp1, Dspp, or Mepe (Supplementary Table S4).

3.5. OPN does not regulate cementogenesis in Ank^mice

Investigations into osteoclast numbers revealed no difference in tartrate resistant (TRAP) positive cells in alveolar bone around the first molar teeth in Spp1^{^/}^{vs. WT mice (Appendix Results and}Supplementary Fig. S8). Additionally, genetic ablation of Spp1 in the Alpl^{^/}^{mouse model of severe hypophosphatasia (HPP) indicated that OPN did not play a major role in inhibition of acellular cementum in those mice (Appendix Results and}Supplementary Figs. S9 and S10).

While OPN was indicated to have some developmental role in regulating both dentin and alveolar bone, histological analysis of Spp1^{^/}^{mice did not indicate a critical, non-redundant physiological role for OPN in regulating acellular cementum. Because the functions of OPN in skeletal biology became more apparent under conditions where mice were physiologically challenged (e.g. bone unloading), we asked whether OPN played a role under the challenge of hypermineralizing conditions that promote increased acellular cementum, by examining}Ank^/^;Spp1^{^/}^{double-deficient mice.}

Comparison of Ank^{^/}^{to WT mice confirmed the statistically significant 11 to 13-fold increase in acellular cementum thickness at 60 dpn (}p < 0.05; Fig. 5A–E). However, Ank^/^;Spp1^{^/}^{double-knockout mice did not exhibit additionally increased cementum thickness over}Ank^{^/}^{mice (}p > 0.05; Fig. 5E), indicating that increased in Spp1/OPN in Ank^{^/}^{mice did not act in functional repression of cementum growth. Micro-CT analysis indicated that while both}Spp1^{^/}^andAnk^{^/}^{mice exhibited increased dentin/cementum and alveolar bone volumes compared to WT controls, that double-knockout mice did not feature additional gains in either tissue (}Fig. 5F, G).

An external file that holds a picture, illustration, etc.
Object name is nihms927401f5.jpg

Fig. 5

Increased OPN does not regulate acellular cementum formation in Ank^{^/}^{mice. First mandibular molars at 60 dpn were analyzed using hematoxylin and eosin (H&E) staining, in situ hybridization (ISH) for}Enpp1, and immunohistochemistry (IHC) for ENPP1 (n = 3–4 per genotype per assay). Compared to (A) WT and (B) Spp1^{^/}^{mouse molars, (C)}Ank^{^/}^{mouse molars exhibit dramatically increased acellular cementum (AC) covering the cervical root. (D) Genetic ablation of}Spp1/OPN in Ank^{^/}^;Spp1^{^/}^{mice does not lead to increased AC. (E) Histological measurements reveal a statistically significant 11 to 13-fold increase in}Ank^{^/}^{vs. WT AC, however no difference in}Ank^{^/}^vs.Ank^{^/}^;Spp1^{^/}^{mice. (F) Micro-CT analysis shows increased dentin/cementum volume in}Spp1^{^/}^andAnk^{^/}^{molars compared to WT, however, no further increase in}Ank^{^/}^;Spp1^{^/}^{mice compared to single knockout models. (G) Micro-CT analysis shows increased alveolar bone (AB) volume in}Spp1^{^/}^andAnk^{^/}^{molars compared to WT, however, no further increase in}Ank^{^/}^;Spp1^{^/}^{mice compared to single knock-out models. Graphs in panels}E-G show individual measurements (circles, squares, and triangles) and means (black line) ± standard deviation. Statistical results of one-way ANOVA and post-hoc Tukey test indicated by: * p < 0.05; ** p < 0.01; *** p < 0.001, **** p < 0.0001. Compared to (H) WT control and (I) Spp1^{^/}^{mice, both (J)}Ank^{^/}^{and (K)}Ank^{^/}^;Spp1^{^/}^{mouse periodontal tissues exhibit increased}Enpp1 mRNA in cementoblasts (Cb) lining the AC, as well as some osteoblasts (Ob) of the AB. While ENPP1 protein is detectable at low levels in Cb in (L) WT control and (M) Spp1^{^/}^{teeth, the protein is over-expressed near the root surface in both (N)}Ank^{^/}^{and (O)}Ank^{^/}^;Spp1^{^/}^{mice. DE = dentin.}

Lacking ANK and OPN expression locally and systemically, other PP_i regulators like ENPP1 could be compensating in controlling cementogenesis. Enpp1 mRNA expression was increased in cementoblasts and alveolar bone osteoblasts in both Ank^{^/}^andAnk^/^;Spp1^{^/}^{double-knockout mice (}Fig. 5H–K). Furthermore, ENPP1 protein was dramatically increased directly adjacent to the expanded cementum layer, around cementicles, and in some regions of bundle bone on the alveolus (Fig. 5L–O).

3.4. Altered gene expression in periodontal tissues of Spp1^mice

3.5. OPN does not regulate cementogenesis in Ank^mice

Fig. 5

4. Discussion

OPN is a member of the Small Integrin-Binding Ligand N-linked Glycoprotein (SIBLING) protein family of mineralized tissue-associated proteins, along with closely related proteins including BSP [7]. OPN regulates mineralization in vitro and in vivo [7,44,53,54], and is a constituent of the ECM of both cementum and alveolar bone in the periodontal complex [4,6,55–59]. In this study, we identified OPN as a candidate of interest in maintaining periodontal cementum-PDL-bone borders based on proteomic analysis of the Ank^{^/}^{mouse model that features increased cementum formation. The functional importance of OPN in the periodontium in vivo was analyzed using several animal models. We determined that loss of OPN in}Spp1^{^/}^{mice increased cellular cementum formation, volume and mineral density of dentin/cementum and alveolar bone, and increased the tissue densities while decreasing the volumes of both PDL and dental pulp. However, acellular cementum apposition was not altered in}Spp1^{^/}^{mice vs. controls. In}Ank^{^/}^{mice, a mouse model of hypercementosis that features increased local OPN and reduced PP}_i concentrations, genetic ablation of Spp1 did not further exacerbate excessive cementogenesis. Based on these multiple approaches, we conclude that OPN has an important and non-redundant role in regulating mineralization in the periodontium, though its importance lies with cellular cementum and at the PDL-bone border and influencing tissue density properties of the PDL and bone, and not by directly controlling acellular cementum apposition.

4.1. Role of OPN in regulation of dentoalveolar development and mineralization

This study began as a proteomic analysis to identify “molecular tools” employed to regulate mineralization in the cementum-PDL-bone periodontal complex. As our model, we chose the Ank^{^/}^{mouse, wherein the acellular cementum grows rapidly due to deficient extracellular PP}_i, and where the challenge to the existing hard-soft borders may spur induction of mineral regulatory factors. Increased presence of OPN was identified by proteomic analysis in Ank^{^/}^{mouse periodontal tissues. OPN is abundant in mineralized tissues (bone, dentin, and cementum), and has been implicated in modulating mineralized tissue formation [}5,7,9,60]. The amino acid sequence of OPN is highly conserved, especially in functional domains including a polyaspartic acid (polyD) motif, RGD integrin-binding sequence, serine/threonine phosphorylation sites, and a thrombin cleavage site [60]. In vitro studies revealed that OPN regulates both initiation and growth of hydroxyapatite crystals, and that this inhibitory ability relies on calcium-binding properties of the polyD sequence and serine phosphorylation [40,54,61]. Importantly, in vivo studies confirmed that OPN regulates mineral growth in both physiological and pathological situations [24,44,47,50], and ex vivo experiments with Spp1^{^/}^{osteoblasts further supported the role of OPN in direct regulation of mineralization rather than osteoblast differentiation [}53]. The mineral-regulating function of OPN has been implicated to be ancient and highly conserved based on genetic and evolutionary analyses [62,63], and observations that OPN plays a similar role in wider contexts, e.g. avian eggshell calcification [64].

Developmental analysis revealed normal dentoalveolar appearance and organization in Spp1^{^/}^{mice compared to controls. This was not surprising in light of the “apparently normal phenotype of mice lacking OPN” described by those who created the model and were anticipating effects in the skeleton [}65]. However, subsequent in vivo studies by that group and others teased out skeletal functions for OPN, including subtle but detectable effects on bone mineral crystallinity [44] and bone mechanical properties [66], and the capacity for OPN to contribute to skeletal pathology [24,25].

We found that lack of OPN promoted more rapidly forming dentin and cellular cementum in mice. Mineral density of dentin/cementum was significantly increased in Spp1^{^/}^{mice compared to WT. These volume and density changes became more apparent between 30 and 90 dpn. Histology and ISH indicated normal odontoblasts expressing}Col1a1 and Bglap. In total, these dentin changes suggest a role for OPN in regulating the pace of dentinogenesis and influencing the extent of tissue mineralization. This is consistent with previous studies that supported a role for ECM phosphoproteins in directing circumpulpal dentin mineralization [67–69], or demonstrated OPN interaction with matrix vesicles in mantle dentin [6,27]. Moreover, these observations of OPN effects on dentin are supportive of findings pointing to increased OPN deposition in pathologies of dentin hypomineralization, e.g. X-linked hypophosphatemia [70–72]. A previous report employing energy dispersive spectroscopy analysis on Spp1^{^/}^{mice found no differences in calcium or phosphorus content in teeth or alveolar bone [}73]. We speculate that our current approach of high resolution micro-CT on multiple ages of extensively backcrossed Spp1 mice was well equipped to find these sorts of quantitative volume and mineralization differences. Interestingly, the only report to date associated with a dentin phenotype in Spp1^{^/}^{mice relates to deficiency in reparative dentin formation, possibly due to defective migration and/or differentiation of new odontoblasts [}52].

Loss of OPN affected mandibular bone in a similar fashion to dentin, with increased bone volume and mineral density at both 30 and 90 dpn. Osteoblasts appeared normal in number and location, consistent with a previous in vitro study [53]. These changes indicate a role for OPN in controlling bone matrix mineralization, consistent with reports of increased mineralization in long bones of Spp1^{^/}^{mice [}24,44]. These mandibular bone changes closely paralleled observations from Spp1^{^/}^{long bones, including changes in cortical bone (increased cortical bone thickness and mineral density), trabecular bone (increased bone volume fraction, increased trabecular thickness, increased trabecular number, decreased trabecular spacing), and decreased osteoid volume, compared to WT controls [}25]. These long bone changes were more apparent at 90 dpn, as documented for mandibular bone here, and fold-changes in mandibular bone were greater than those documented in long bones.

Elucidating the mechanism for OPN knockout effects on bone is not so straightforward, as numerous previous studies strongly implicate OPN in osteoclast-mediated bone remodeling. After initial studies on Spp1^{^/}^{mice revealed relatively normal skeletal development, “challenge” experiments on these mice further clarified OPN functions, including recruitment of osteoclasts for long bone remodeling under unloading conditions [}65,74,75]. Intrinsic functional defects were identified in Spp1^{^/}^{osteoclasts [}76], such as hypomotility and decreased resorptive capability related to loss of osteoclast-derived secretion of OPN into resorption pits [66,77]. For osteoclast function in alveolar bone, OPN was also shown to be important in force-induced bone and tooth resorption [73] and unloading-induced horizontal tooth drift in mouse molars [51], where tooth movement is facilitated by osteoclast recruitment to remodel alveolar bone. Analysis of osteoclast location and appearance in and Spp1^{^/}^{mouse periodontal tissues under orthodontic loading revealed reduced osteoclast recruitment and bone resorption, and fewer multinucleated and more mono- and bi-nucleated TRAP-positive cells compared to WT, indicating defective precursor cell fusion in the absence of OPN [}78]. In alveolar bone of Spp1^{^/}^{mice from 14 to 240 dpn, we observed no difference in osteoclast numbers. While we did not find differences in cell numbers in normally developing}Spp1^{^/}^{vs. WT mice, one could speculate that defective osteoclast recruitment may only be apparent under conditions of heightened necessity. Presence of similar numbers of osteoclasts does not confirm similar activity and resorptive capability, and we did not test this due to a number of previous reports on this aspect of osteoclast biology [}66,76,77].

We also detected density increases in the dental pulp and PDL of Spp1^{^/}^{vs. WT mice, starting at 30 dpn and increasing by 90 dpn. While it cannot be conclusively determined by micro-CT whether this increased density is attributable to increased calcium phosphorus content, this interpretation is consistent with the observed trends in dentin and bone, as well as with known functions for OPN in regulating mineral maturation in hard tissues [}44] and preventing ectopic calcification in soft tissues [47,48,50,79,80]. One Spp1^{^/}^{mouse at the advanced age of 240 dpn exhibited near total pulp obliteration by disorganized calcified tissue. While one might speculate that increased tissue density of pulp and PDL in}Spp1^{^/}^{mice may predispose these tissues to ectopic calcification, additional studies are necessary to clarify tissue changes and the role of OPN.}

4.2. Osteopontin and cementum formation

Studies over the past two decades have revealed that acellular cementum apposition is strongly controlled by factors that dictate mineralization. For example, genetic mutation or ablation of any of the primary regulators of local PP_i (i.e. Alpl/TNAP, Ank/ANK, and Enpp1/ENPP1) has dramatic effects on acellular cementum growth [10–14, 81]. Acellular cementum formation is also heavily influenced by other perturbations in mineral metabolism such as altered systemic inorganic phosphate (P_i), or introduction of mineralization inhibitors including bisphosphonates and matrix gla protein (MGP) [82–87]. Loss of BSP, an ECM protein closely related to OPN that functions as a nucleator of hydroxyapatite crystal growth, severely hampers acellular cementum formation [3,8,88]. In all of these examples, cellular cementum formation was less disturbed by alterations, though its mineralization status can be altered by changes negatively affecting bone mineralization.

Identification of OPN as a potential negative regulator of acellular cementum mineralization seemed to fit into this developing paradigm of positive and negative regulators of cementum acting as a system of molecular checks and balances. While we found increased cellular cementum in Spp1^{^/}^{mouse molars, we found no differences in acellular cementum growth. While the qPCR approach to identify local compensators for OPN did not reveal likely stand-ins, some intriguing downstream effects were revealed. OPN has been identified as a substrate for the peptidase PHEX [}71,89], and mRNA for this enzyme was significantly downregulated in Spp1^{^/}^{periodontia, though no feedback between the two factors has been previously reported.}Bglap/OCN, another marker for both cementum and bone was also down-regulated, but its functions in periodontal development and mineralization remain poorly understood [6,56,90–92]. It is possible that the up to 5-fold increased circulating PP_i reported in Spp1^{^/}^{mice [}24] works to repress cementum growth, compensating for the absence of OPN.

Beyond considering the role of OPN in normal development of periodontal tissues, we further challenged the cementum-PDL border to test whether OPN functions became apparent under these circumstances. If the increased OPN in Ank^{^/}^{mice was a compensatory attempt to counteract rapid cementogenesis, then genetic ablation of}Spp1 would be expected to exacerbate the hypercementosis. However, Ank^{^/}^;Spp1^{^/}^{double-knockout mice did not exhibit increased acellular cementum thickness over}Ank^{^/}^{mice, refuting this hypothesis. In this context, up-regulation of other factors such as}Enpp1/ENPP1 (increasing local PP_i) likely contributes to cementum regulation.

There are some important limitations and caveats to the current study that should be kept in mind when considering OPN in the dentoalveolar tissues. Selection of 60 dpn for proteomic analysis of Ank^{^/}^{PDL limits our results to one snapshot, where additional experiments at different ages could identify additional or alternative factors. While we did probe mineralization of alveolar bone and dentin using micro-CT, mouse cementum was not able to be similarly tested. Though we cannot rule out subtle changes in cementum mineralization, all of the micro-CT, histology, and histomorphometry data indicated that}Spp1^{^/}^{mice display no significant changes indicative of periodontal alteration or dysfunction during the time period of the study.}

Another limitation extending to all studies of OPN in dentoalveolar tissues to date is that we largely do not know the forms of OPN that are produced and/or deposited in these tissues. OPN is produced as low- and high-phosphorylated forms, and the phosphorylation status of OPN from dentoalveolar tissues and cells has never been reported. This question is relevant to the current study because the degree of serine/threonine phosphorylation has been shown to be critical (and proportional) to the potency of OPN to regulate hydroxyapatite crystal growth [39,49,60,89,93,94]. While OPN produced by osteoblasts and isolated from bone has sometimes been shown to be highly phosphorylated, there is great variability between reports [9,60,95–97]. It is possible that OPN derived from dental-periodontal cells is relatively less phosphorylated. TNAP, an enzyme that hydrolyzes PP_i and is expressed by skeletal and dental cells [98,99], dephosphorylates OPN, rendering it less effective at regulating mineralization [43,100]. Periodontal tissues are extremely rich in TNAP/ALP activity [11,13,27,101–104], pointing to the possibility that OPN exists in a relatively dephosphorylated state. Intriguingly, experiments by Christensen and colleagues indicated that phosphorylation status of the C-terminal portion of OPN had direct effects on the ability of the RGD domain to bind integrins, raising the possibility that dephosphorylated OPN may be more effective as an attachment and signaling protein [105]. Clearly, observed changes in dentin, bone, pulp, and PDL indicate that lack of OPN is altering dentoalveolar mineralization, though the question could be asked whether locally produced and circulating OPN [106] have different roles in this context.

Lastly, we observed variability in penetrance or expressivity of the phenotype resulting from targeting the Spp1 allele in mice. Significant differences in dentoalveolar tissues of Spp1^{^/}^{vs. control mice were documented in the main cohorts that were analyzed in detail at 30 and 90 dpn. These mice were maintained on a C57BL/6 genetic background [}25]. However, when additional studies were initiated to explore interactions of Spp1/OPN with other alleles, and these experiments required crossing C57BL/6 Spp1 mice with a different background, the resulting mixed background had the effect of confounding the phenotype resulting from the loss of Spp1 (data not shown). Previous reports on Spp1^{^/}^{mice have detected varying severities in skeletal and other phenotypes using different origins of C57BL/6 mice [}24,25,51–53,65,78], or mixed/hybrid mouse backgrounds [47,66,73–75,107–109]. It is well established that genetic background can have profound effects on phenotypes (including bones and teeth) of genetically engineered mice [110–117]. In accordance with previous findings, we propose that this background-dependent variability in penetrance and expressivity may arise from modifiers that are differentially expressed in different mouse strains [118–120]. Potential modifiers may include genes that overlap with, compensate for, or antagonize OPN, e.g. SIBLINGs like Ibsp, Dmp1, Dspp, and Mepe, mineral metabolism regulators like Alpl, Ank, or Enpp1, or additional factors directing systemic mineral metabolism. Further studies to identify gene modifiers may elucidate the functions of OPN and its interactions with other regulators of dentoalveolar development. In addition, accumulating data from genome-wide association studies (GWAS) on humans, including the precision medicine “All of Us” initiative, may help to translate findings from mouse models to humans by clarifying the role of mutations in SPP1/OPN and other SIBLING family proteins in regulating mineralized tissues in health and disease states.

4.1. Role of OPN in regulation of dentoalveolar development and mineralization

4.2. Osteopontin and cementum formation

Supplementary Material

1

Click here to view.^{(40K, docx)}

2

Click here to view.^{(17K, docx)}

3

Click here to view.^{(15K, docx)}

4

Click here to view.^{(18K, docx)}

5

Click here to view.^{(17K, docx)}

S1

Click here to view.^{(19M, tif)}

S10

Click here to view.^{(19M, tif)}

S2

Click here to view.^{(19M, tif)}

S3

Click here to view.^{(19M, tif)}

S4

Click here to view.^{(19M, tif)}

S5

Click here to view.^{(19M, tif)}

S6

Click here to view.^{(23M, tif)}

S7

Click here to view.^{(20M, tif)}

S8

Click here to view.^{(19M, tif)}

S9

Click here to view.^{(19M, tif)}

1

Click here to view.^{(40K, docx)}

2

Click here to view.^{(17K, docx)}

3

Click here to view.^{(15K, docx)}

4

Click here to view.^{(18K, docx)}

5

Click here to view.^{(17K, docx)}

S1

Click here to view.^{(19M, tif)}

S10

Click here to view.^{(19M, tif)}

S2

Click here to view.^{(19M, tif)}

S3

Click here to view.^{(19M, tif)}

S4

Click here to view.^{(19M, tif)}

S5

Click here to view.^{(19M, tif)}

S6

Click here to view.^{(23M, tif)}

S7

Click here to view.^{(20M, tif)}

S8

Click here to view.^{(19M, tif)}

S9

Click here to view.^{(19M, tif)}

Acknowledgments

This research was supported by grant AR 066110 to BLF from the National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) of the National Institutes of Health (NIH, Bethesda, MD), grant DE 12889 to JLM from the National Institute of Dental and Craniofacial Research (NIDCR)/NIH, and the Intramural Research Program of NIAMS (MJS). We thank Dr. Susan Rittling (Forsyth Institute, Boston, MA, USA) for allowing access to and study of Spp1 mice. We thank Dr. Kristina Zaal (Light Imaging Section, NIAMS/NIH) for assistance in slide scanning and Nasrin Kalantaripour and Kathryn Hemstreet (NIAMS/NIH) for additional histological sectioning. We thank the editor and anonymouse reviewers for their suggested revisions that improved this manucript.

^{Division of Biosciences, College of Dentistry, The Ohio State University, Columbus, OH, USA}

^{National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS), National Institutes of Health (NIH), Bethesda, MD, USA}

^{Department of Prosthodontics and Periodontics, Division of Periodontics, Piracicaba Dental School, University of Campinas, São Paulo, Brazil}

^{Department of Dental Materials, São Leopoldo Mandic Research Center, Campinas, São Paulo, Brazil}

^{Sanford Children’s Health Research Center, Sanford Burnham Prebys Medical Research Institute, La Jolla, CA, USA}

^{Corresponding author at: Division of Biosciences, College of Dentistry, The Ohio State University, 4163 Postle Hall, 305 W. 12th Avenue, Columbus, OH, USA.}ude.uso@4001.retsof (B.L. Foster)

Abstract

The periodontal complex is essential for tooth attachment and function and includes the mineralized tissues, cementum and alveolar bone, separated by the unmineralized periodontal ligament (PDL). To gain insights into factors regulating cementum-PDL and bone-PDL borders and protecting against ectopic calcification within the PDL, we employed a proteomic approach to analyze PDL tissue from progressive ankylosis knock-out (Ank^{^/}^{) mice, featuring reduced PP}_i, rapid cementogenesis, and excessive acellular cementum. Using this approach, we identified the matrix protein osteopontin (Spp1/OPN) as an elevated factor of interest in Ank^{^/}^{mouse molar PDL. We studied the role of OPN in dental and periodontal development and function. During tooth development in wild-type (WT) mice,}Spp1 mRNA was transiently expressed by cementoblasts and strongly by alveolar bone osteoblasts. Developmental analysis from 14 to 240 days postnatal (dpn) indicated normal histological structures in Spp1^{^/}^{comparable to WT control mice. Microcomputed tomography (micro-CT) analysis at 30 and 90 dpn revealed significantly increased volumes and tissue mineral densities of}Spp1^{^/}^{mouse dentin and alveolar bone, while pulp and PDL volumes were decreased and tissue densities were increased. However, acellular cementum growth was unaltered in}Spp1^{^/}^{mice. Quantitative PCR of periodontal-derived mRNA failed to identify potential local compensators influencing cementum in}Spp1^{^/}^{vs. WT mice at 26 dpn. We genetically deleted}Spp1 on the Ank^{^/}^{mouse background to determine whether increased}Spp1/OPN was regulating periodontal tissues when the PDL space is challenged by hypercementosis in Ank^{^/}^mice.Ank^{^/}^;Spp1^{^/}^{double deficient mice did not exhibit greater hypercementosis than that in}Ank^{^/}^{mice. Based on these data, we conclude that OPN has a non-redundant role regulating formation and mineralization of dentin and bone, influences tissue properties of PDL and pulp, but does not control acellular cementum apposition. These findings may inform therapies targeted at controlling soft tissue calcification.}

Keywords: Extracellular matrix, Mineralization, Periodontal tissues, Tooth development, Dental cementum, Bone biology

Abstract

Footnotes

The authors declare that they have no conflicts of interest with the contents of this article.

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bone.2017.12.004.

Footnotes

References

1. Foster BL, Nociti FH, Jr, Somerman MJ. Tooth root formation. In: Huang GTJ, Thesleff I, editors. Stem Cells, Craniofacial Development and Regeneration. Wiley-Blackwell; 2013. pp. 153–177. [PubMed]
2. Foster BL, Somerman MJ. Cementum. In: McCauley LK, Somerman MJ, editors. Mineralized Tissues in Oral and Craniofacial Science: Biological Principles and Clinical Correlates. Wiley-Blackwell; Ames, IA: 2012. pp. 169–192. [PubMed]
3. Foster BL, et al Mineralization defects in cementum and craniofacial bone from loss of bone sialoprotein. Bone. 78(2015):150–164.[Google Scholar]
4. Foster BLMethods for studying tooth root cementum by light microscopy. Int J Oral Sci. 2012;4(3):119–128.[Google Scholar]
5. Staines KA, MacRae VE, Farquharson CThe importance of the SIBLING family of proteins on skeletal mineralisation and bone remodelling. J Endocrinol. 2012;214(3):241–255.[PubMed][Google Scholar]
6. McKee M, Zalzal S, Nanci AExtracellular matrix in tooth cementum and mantle dentin: localization of osteopontin and other noncollagenous proteins, plasma proteins, and glycoconjugates by electron microscopy. Anat Rec. 1996;245(2):293–312.[PubMed][Google Scholar]
7. Fisher L, Fedarko NSix genes expressed in bones and teeth encode the current members of the SIBLING family of proteins, Connect. Tissue Res. 2003;44(Suppl. 1):33–40.[PubMed][Google Scholar]
8. Foster BL, et al Deficiency in acellular cementum and periodontal attachment in bsp null mice. J Dent Res. 2013;92(2):166–172.[Google Scholar]
9. Qin C, Baba O, Butler WPost-translational modifications of sibling proteins and their roles in osteogenesis and dentinogenesis. Crit Rev Oral Biol Med. 2004;15(3):126–136.[PubMed][Google Scholar]
10. Foster BL, Nociti FH, Jr, Somerman MJThe rachitic tooth. Endocr Rev. 2014;35(1):1–34.[Google Scholar]
11. Foster BL, et al Central role of pyrophosphate in acellular cementum formation. PLoS One. 2012;7(6):e38393.[Google Scholar]
12. Nociti FJ, et al Cementum: a phosphate-sensitive tissue. J Dent Res. 2002;81(12):817–821.[PubMed][Google Scholar]
13. Zweifler LE, et al Counter-regulatory phosphatases TNAP and NPP1 temporally regulate tooth root cementogenesis. Int J Oral Sci. 2015;7(1):27–41.[Google Scholar]
14. Foster BL, et al The progressive ankylosis protein regulates cementum apposition and extracellular matrix composition. Cells Tissues Organs. 2011;194(5):382–405.[Google Scholar]
15. Beertsen W, McCulloch C, Sodek JThe periodontal ligament: a unique, multifunctional connective tissue. Periodontol. 13(1997):20–40.[PubMed][Google Scholar]
16. Seo B, et al Investigation of multipotent postnatal stem cells from human periodontal ligament. Lancet. 2004;364(9429):149–155.[PubMed][Google Scholar]
17. Fong RK, et al Dental histology of Coelophysis bauri and the evolution of tooth attachment tissues in early dinosaurs. J Morphol. 2016;277(7):916–924.[PubMed][Google Scholar]
18. LeBlanc AR, et al Mineralized periodontia in extinct relatives of mammals shed light on the evolutionary history of mineral homeostasis in periodontal tissue maintenance. J Clin Periodontol. 2016;43(4):323–332.[PubMed][Google Scholar]
19. LeBlanc AR, Reisz RRPeriodontal ligament, cementum, and alveolar bone in the oldest herbivorous tetrapods, and their evolutionary significance. PLoS One. 2013;8(9):e74697.[Google Scholar]
20. Luan X, et al The mosasaur tooth attachment apparatus as paradigm for the evolution of the gnathostome periodontium. Evol Dev. 2009;11(3):247–259.[Google Scholar]
21. McIntosh J, et al Caiman periodontium as an intermediate between basal vertebrate ankylosis-type attachment and mammalian “true” periodontium. Microsc Res Tech. 2002;59(5):449–459.[PubMed][Google Scholar]
22. Kilkenny C, et al Improving bioscience research reporting: the ARRIVE guidelines for reporting animal research. PLoS Biol. 2010;8(6):e1000412.[Google Scholar]
23. Gurley K, et al Mineral formation in joints caused by complete or joint-specific loss of ANK function. J Bone Miner Res. 2006;21(8):1238–1247.[PubMed][Google Scholar]
24. Harmey D, et al Elevated skeletal osteopontin levels contribute to the hypophosphatasia phenotype in Akp2(−/−) mice. J Bone Miner Res. 2006;21(9):1377–1386.[PubMed][Google Scholar]
25. Yadav MC, et al Ablation of osteopontin improves the skeletal phenotype of phospho1(−/−) mice. J Bone Miner Res. 2014;29(11):2369–2381.[Google Scholar]
26. Narisawa S, Frohlander N, Millan JLInactivation of two mouse alkaline phosphatase genes and establishment of a model of infantile hypophosphatasia. Dev Dyn. 1997;208(3):432–446.[PubMed][Google Scholar]
27. Foster BL, et al Tooth root dentin mineralization defects in a mouse model of hypophosphatasia. J Bone Miner Res. 2013;28(2):271–282.[Google Scholar]
28. Salmon CR, et al Global proteome profiling of dental cementum under experimentally-induced apposition. J Proteome. 141(2016):12–23.[Google Scholar]
29. Flores IL, et al EEF1D modulates proliferation and epithelial-mesenchymal transition in oral squamous cell carcinoma. Clin Sci (Lond) 2016;130(10):785–799.[PubMed][Google Scholar]
30. Salmon CR, Giorgetti AP, Paes Leme AF, Domingues RR, Kolli TN, Foster BL, Nociti FHMicroproteome of dentoalveolar tissues. Bone. 101(2017):219–229.[PubMed][Google Scholar]
31. Xu H, Snider TN, Wimer HF, Yamada SS, Yang T, Holmbeck K, Foster BLMultiple essential MT1-MMP functions in tooth root formation, dentinogenesis, and tooth eruption. Matrix Biol. 2016;52–54:266–283.[Google Scholar]
32. Zweifler LE, et al Role of PHOSPHO1 in periodontal development and function. J Dent Res. 2016;95(7):742–751.[Google Scholar]
33. Wang L, et al Fibromodulin and biglycan modulate periodontium through TGFbeta/BMP signaling. J Dent Res. 2014;93(8):780–787.[Google Scholar]
34. Leong NL, et al Age-related adaptation of bone-PDL-tooth complex: Rattus-Norvegicus as a model system. PLoS One. 2012;7(4):e35980.[Google Scholar]
35. Matias M, et al Immunohistochemical localization of fibromodulin in the periodontium during cementogenesis and root formation in the rat molar. J Periodontal Res. 2003;38(5):502–507.[PubMed][Google Scholar]
36. MacNeil R, et al Expression of type I and XII collagen during development of the periodontal ligament in the mouse. Arch Oral Biol. 1998;43(10):779–787.[PubMed][Google Scholar]
37. Rios H, et al Periostin is essential for the integrity and function of the periodontal ligament during occlusal loading in mice. J Periodontol. 2008;79(8):1480–1490.[Google Scholar]
38. Horiuchi K, et al Identification and characterization of a novel protein, periostin, with restricted expression to periosteum and periodontal ligament and increased expression by transforming growth factor beta. J Bone Miner Res. 1999;14(7):1239–1249.[PubMed][Google Scholar]
39. Boskey AL, et al Post-translational modification of osteopontin: effects on in vitro hydroxyapatite formation and growth. Biochem Biophys Res Commun. 2012;419(2):333–338.[Google Scholar]
40. Boskey AL, et al Osteopontin-hydroxyapatite interactions in vitro: inhibition of hydroxyapatite formation and growth in a gelatin-gel. Bone Miner. 1993;22(2):147–159.[PubMed][Google Scholar]
41. Goldberg HA, et al Binding of bone sialoprotein, osteopontin and synthetic polypeptides to hydroxyapatite, Connect. Tissue Res. 2001;42(1):25–37.[PubMed][Google Scholar]
42. Goldberg HA, Hunter GKThe inhibitory activity of osteopontin on hydroxyapatite formation in vitro. Ann N Y Acad Sci. 760(1995):305–308.[PubMed][Google Scholar]
43. Narisawa S, Yadav MC, Millán JLIn vivo overexpression of tissue-nonspecific alkaline phosphatase increases skeletal mineralization and affects the phosphorylation status of osteopontin. J Bone Miner Res. 2013;28(7):1587–1598.[Google Scholar]
44. Boskey AL, et al Osteopontin deficiency increases mineral content and mineral crystallinity in mouse bone, Calcif. Tissue Int. 2002;71(2):145–154.[PubMed][Google Scholar]
45. Harmey D, et al Concerted regulation of inorganic pyrophosphate and osteopontin by akp2, enpp1, and ank: an integrated model of the pathogenesis of mineralization disorders. Am J Pathol. 2004;164(4):1199–1209.[Google Scholar]
46. Johnson K, et al Linked deficiencies in extracellular PP(i) and osteopontin mediate pathologic calcification associated with defective PC-1 and ANK expression. J Bone Miner Res. 2003;18(6):994–1004.[PubMed][Google Scholar]
47. Steitz S, et al Osteopontin inhibits mineral deposition and promotes regression of ectopic calcification. Am J Pathol. 2002;161(6):2035–2046.[Google Scholar]
48. Speer M, et al Inactivation of the osteopontin gene enhances vascular calcification of matrix Gla protein-deficient mice: evidence for osteopontin as an inducible inhibitor of vascular calcification in vivo. J Exp Med. 2002;196(8):1047–1055.[Google Scholar]
49. Jono S, Peinado C, Giachelli CPhosphorylation of osteopontin is required for inhibition of vascular smooth muscle cell calcification. J Biol Chem. 2000;275(26):20197–20203.[PubMed][Google Scholar]
50. Wesson J, et al Osteopontin is a critical inhibitor of calcium oxalate crystal formation and retention in renal tubules. J Am Soc Nephrol. 2003;14(1):139–147.[PubMed][Google Scholar]
51. Walker CG, et al Osteopontin is required for unloading-induced osteoclast recruitment and modulation of RANKL expression during tooth drift-associated bone remodeling, but not for super-eruption. Bone. 2010;47(6):1020–1029.[Google Scholar]
52. Saito K, et al Osteopontin is essential for type I collagen secretion in reparative dentin. J Dent Res. 2016;95(9):1034–1041.[PubMed][Google Scholar]
53. Holm E, et al Osteopontin mediates mineralization and not osteogenic cell development in vitro. Biochem J. 2014;464(3):355–364.[PubMed][Google Scholar]
54. Hunter GK, et al Nucleation and inhibition of hydroxyapatite formation by mineralized tissue proteins. Biochem J. 1996;317(Pt 1):59–64.[Google Scholar]
55. MacNeil R, et al Role of two mineral-associated adhesion molecules, osteopontin and bone sialoprotein, during cementogenesis. Connect Tissue Res. 1995;33(1–3):1–7.[PubMed][Google Scholar]
56. Bronckers A, et al Immunolocalization of osteopontin, osteocalcin, and dentin sialoprotein during dental root formation and early cementogenesis in the rat. J Bone Miner Res. 1994;9(6):833–841.[PubMed][Google Scholar]
57. Bosshardt D, et al Developmental appearance and distribution of bone sialoprotein and osteopontin in human and rat cementum. Anat Rec. 1998;250(1):13–33.[PubMed][Google Scholar]
58. McKee M, Nanci APostembedding colloidal-gold immunocytochemistry of noncollagenous extracellular matrix proteins in mineralized tissues. Microsc Res Tech. 1995;31(1):44–62.[PubMed][Google Scholar]
59. Somerman M, et al Expression of attachment proteins during cementogenesis. J Biol Buccale. 1990;18(3):207–214.[PubMed][Google Scholar]
60. Sodek J, Ganss B, McKee MDOsteopontin. Crit Rev Oral Biol Med. 2000;11(3):279–303.[PubMed][Google Scholar]
61. Hunter GK, Kyle CL, Goldberg HAModulation of crystal formation by bone phosphoproteins: structural specificity of the osteopontin-mediated inhibition of hydroxyapatite formation. Biochem J. 1994;300(Pt 3):723–728.[Google Scholar]
62. Kawasaki K, Weiss KMEvolutionary genetics of vertebrate tissue mineralization: the origin and evolution of the secretory calcium-binding phosphoprotein family. J Exp Zool B Mol Dev Evol. 2006;306(3):295–316.[PubMed][Google Scholar]
63. Fisher LW, et al Flexible structures of SIBLING proteins, bone sialoprotein, and osteopontin. Biochem Biophys Res Commun. 2001;280(2):460–465.[PubMed][Google Scholar]
64. Chien YC, Hincke MT, McKee MDAvian eggshell structure and osteopontin. Cells Tissues Organs. 2009;189(1–4):38–43.[PubMed][Google Scholar]
65. Rittling SR, et al Mice lacking osteopontin show normal development and bone structure but display altered osteoclast formation in vitro. J Bone Miner Res. 1998;13(7):1101–1111.[PubMed][Google Scholar]
66. Chellaiah MA, et al Osteopontin deficiency produces osteoclast dysfunction due to reduced CD44 surface expression. Mol Biol Cell. 2003;14(1):173–189.[Google Scholar]
67. Steinfort J, van den Bos T, Beertsen WDifferences between enamel-related and cementum-related dentin in the rat incisor with special emphasis on the phosphoproteins. J Biol Chem. 1989;264(5):2840–2845.[PubMed][Google Scholar]
68. Steinfort J, Deblauwe B, Beertsen WThe inorganic components of cementum-and enamel-related dentin in the rat incisor. J Dent Res. 1990;69(6):1287–1292.[PubMed][Google Scholar]
69. Ohma N, Takagi Y, Takano YDistribution of non-collagenous dentin matrix proteins and proteoglycans, and their relation to calcium accumulation in bisphosphonate-affected rat incisors. Eur J Oral Sci. 2000;108(3):222–232.[PubMed][Google Scholar]
70. Salmon B, et al Abnormal osteopontin and matrix extracellular phosphoglycoprotein localization, and odontoblast differentiation, in X-linked hypophosphatemic teeth, Connect. Tissue Res. 2014;55(Suppl. 1):79–82.[PubMed][Google Scholar]
71. Barros NM, et al Proteolytic processing of osteopontin by PHEX and accumulation of osteopontin fragments in Hyp mouse bone, the murine model of X-linked hypophosphatemia. J Bone Miner Res. 2013;28(3):688–699.[PubMed][Google Scholar]
72. Boukpessi T, et al Osteopontin and the dento-osseous pathobiology of X-linked hypophosphatemia. Bone. 95(2017):151–161.[PubMed][Google Scholar]
73. Chung C, et al OPN deficiency suppresses appearance of odontoclastic cells and resorption of the tooth root induced by experimental force application. J Cell Physiol. 2008;214(3):614–620.[PubMed][Google Scholar]
74. Ishijima M, et al Enhancement of osteoclastic bone resorption and suppression of osteoblastic bone formation in response to reduced mechanical stress do not occur in the absence of osteopontin. J Exp Med. 2001;193(3):399–404.[Google Scholar]
75. Ishijima M, et al Resistance to unloading-induced three-dimensional bone loss in osteopontin-deficient mice. J Bone Miner Res. 2002;17(4):661–667.[PubMed][Google Scholar]
76. Rajachar R, Truong A, Giachelli CThe influence of surface mineral and osteopontin on the formation and function of murine bone marrow-derived osteoclasts. J Mater Sci Mater Med. 2008;19(10):3279–3285.[Google Scholar]
77. Chellaiah MA, Hruska KAThe integrin alpha(v)beta(3) and CD44 regulate the actions of osteopontin on osteoclast motility. Calcif Tissue Int. 2003;72(3):197–205.[PubMed][Google Scholar]
78. Fujihara S, et al Function and regulation of osteopontin in response to mechanical stress. J Bone Miner Res. 2006;21(6):956–964.[PubMed][Google Scholar]
79. Speer M, et al Smooth muscle cells deficient in osteopontin have enhanced susceptibility to calcification in vitro. Cardiovasc Res. 2005;66(2):324–333.[PubMed][Google Scholar]
80. Mo L, et al Renal calcinosis and stone formation in mice lacking osteopontin, Tamm-Horsfall protein, or both. Am J Physiol Ren Physiol. 2007;293(6):F1935–43.[PubMed][Google Scholar]
81. Fong H, et al Structure and mechanical properties of Ank/Ank mutant mouse dental tissues–an animal model for studying periodontal regeneration. Arch Oral Biol. 2009;54(6):570–576.[Google Scholar]
82. Fong H, et al Aberrant cementum phenotype associated with the hypophosphatemic hyp mouse. J Periodontol. 2009;80(8):1348–1354.[Google Scholar]
83. Chu E, et al Ablation of systemic phosphate-regulating gene fibroblast growth factor 23 (Fgf23) compromises the dentoalveolar complex. Anat Rec (Hoboken) 2010;293(7):1214–1226.[Google Scholar]
84. Kaipatur N, Murshed M, McKee MMatrix Gla protein inhibition of tooth mineralization. J Dent Res. 2008;87(9):839–844.[PubMed][Google Scholar]
85. Takano Y, et al Possible role of dentin matrix in region-specific deposition of cellular and acellular extrinsic fibre cementum. J Electron Microsc. 2003;52(6):573–580.[PubMed][Google Scholar]
86. Alatli-Kut I, Hultenby K, Hammarström LDisturbances of cementum formation induced by single injection of 1-hydroxyethylidene-1,1-bisphosphonate (HEBP) in rats: light and scanning electron microscopic studies. Scand J Dent Res. 1994;102(5):260–268.[PubMed][Google Scholar]
87. Ye L, et al Periodontal breakdown in the Dmp1 null mouse model of hypophosphatemic rickets. J Dent Res. 2008;87(7):624–629.[Google Scholar]
88. Soenjaya Y, et al Mechanical forces exacerbate periodontal defects in Bsp-null mice. J Dent Res. 2015;94(9):1276–1285.[Google Scholar]
89. Addison W, et al Phosphorylation-dependent inhibition of mineralization by osteopontin ASARM peptides is regulated by PHEX cleavage. J Bone Miner Res. 2010;25(4):695–705.[PubMed][Google Scholar]
90. Bronckers A, et al Studies of osteocalcin function in dentin formation in rodent teeth. Eur J Oral Sci. 1998;106(3):795–807.[PubMed][Google Scholar]
91. D’Errico J, et al Expression of bone associated markers by tooth root lining cells, in situ and in vitro. Bone. 1997;20(2):117–126.[PubMed][Google Scholar]
92. Kagayama M, et al Expression of osteocalcin in cementoblasts forming acellular cementum. J Periodontal Res. 1997;32(3):273–278.[PubMed][Google Scholar]
93. Hunter GKRole of osteopontin in modulation of hydroxyapatite formation, Calcif. Tissue Int. 2013;93(4):348–354.[PubMed][Google Scholar]
94. Gericke A, et al Importance of phosphorylation for osteopontin regulation of biomineralization, Calcif. Tissue Int. 2005;77(1):45–54.[Google Scholar]
95. Prince CW, et al Isolation, characterization, and biosynthesis of a phosphorylated glycoprotein from rat bone. J Biol Chem. 1987;262(6):2900–2907.[PubMed][Google Scholar]
96. Neame PJ, Butler WTPosttranslational modification in rat bone osteopontin, Connect. Tissue Res. 1996;35(1–4):145–150.[PubMed][Google Scholar]
97. Salih E, Zhou HY, Glimcher MJPhosphorylation of purified bovine bone sialoprotein and osteopontin by protein kinases. J Biol Chem. 1996;271(28):16897–16905.[PubMed][Google Scholar]
98. Millán JL Medicine and Biotechnology. Vol. 337. Wiley-VCH; Weinheim, Germany: 2006. Mammalian Alkaline Phosphatases: From Biology to Applications. [PubMed][Google Scholar]
99. Millan JL, Whyte MPAlkaline phosphatase and hypophosphatasia, Calcif. Tissue Int. 2016;98(4):398–416.[Google Scholar]
100. Addison W, et al Pyrophosphate inhibits mineralization of osteoblast cultures by binding to mineral, up-regulating osteopontin, and inhibiting alkaline phosphatase activity. J Biol Chem. 2007;282(21):15872–15883.[PubMed][Google Scholar]
101. McKee MD, et al Compounded PHOSPHO1/ALPL deficiencies reduce dentin mineralization. J Dent Res. 2013;92(8):721–727.[Google Scholar]
102. van den Bos T, Beertsen WAlkaline phosphatase activity in human periodontal ligament: age effect and relation to cementum growth rate. J Periodontal Res. 1999;34(1):1–6.[PubMed][Google Scholar]
103. Groeneveld M, Everts V, Beertsen WAlkaline phosphatase activity in the periodontal ligament and gingiva of the rat molar: its relation to cementum formation. J Dent Res. 1995;74(7):1374–1381.[PubMed][Google Scholar]
104. Groeneveld M, Everts V, Beertsen WA quantitative enzyme histochemical analysis of the distribution of alkaline phosphatase activity in the periodontal ligament of the rat incisor. J Dent Res. 1993;72(9):1344–1350.[PubMed][Google Scholar]
105. Christensen B, et al C-terminal modification of osteopontin inhibits interaction with the αVβ3-integrin. J Biol Chem. 2012;287(6):3788–3797.[Google Scholar]
106. VandenBos T, et al Blood circulation as source for osteopontin in acellular extrinsic fiber cementum and other mineralizing tissues. J Dent Res. 1999;78(11):1688–1695.[PubMed][Google Scholar]
107. Liaw L, et al Altered wound healing in mice lacking a functional osteopontin gene (spp1) J Clin Invest. 101(1998):1468–1478.[Google Scholar]
108. Ishii T, et al Osteopontin as a positive regulator in the osteoclastogenesis of arthritis. Biochem Biophys Res Commun. 2004;316(3):809–815.[PubMed][Google Scholar]
109. Yuan Q, et al Increased osteopontin contributes to inhibition of bone mineralization in FGF23-deficient mice. J Bone Miner Res. 2014;29(3):693–704.[Google Scholar]
110. Doetschman TInfluence of genetic background on genetically engineered mouse phenotypes. Methods Mol Biol. 530(2009):423–433.[Google Scholar]
111. Sanford LP, et al Influence of genetic background on knockout mouse phenotypes. Methods Mol Biol. 158(2001):217–225.[PubMed][Google Scholar]
112. Sanford LP, et al TGFbeta2 knockout mice have multiple developmental defects that are non-overlapping with other TGFbeta knockout phenotypes. Development. 1997;124(13):2659–2670.[Google Scholar]
113. Schiviz A, et al Influence of genetic background on bleeding phenotype in the tail-tip bleeding model and recommendations for standardization: communication from the SSC of the ISTH. J Thromb Haemost. 2014;12(11):1940–1942.[PubMed][Google Scholar]
114. Carleton SM, et al Role of genetic background in determining phenotypic severity throughout postnatal development and at peak bone mass in Col1a2 deficient mice (oim) Bone. 2008;42(4):681–694.[Google Scholar]
115. Pirog KA, et al Abnormal chondrocyte apoptosis in the cartilage growth plate is influenced by genetic background and deletion of CHOP in a targeted mouse model of pseudoachondroplasia. PLoS One. 2014;9(2):e85145.[Google Scholar]
116. Ge C, et al Discoidin receptor 2 controls bone formation and marrow adipogenesis. J Bone Miner Res. 2016;31(12):2193–2203.[Google Scholar]
117. Li Y, et al Mouse genetic background influences the dental phenotype. Cells Tissues Organs. 2013;198(6):448–456.[Google Scholar]
118. Bonyadi M, et al Mapping of a major genetic modifier of embryonic lethality in TGF beta 1 knockout mice. Nat Genet. 1997;15(2):207–211.[PubMed][Google Scholar]
119. Coley WD, et al Effect of genetic background on the dystrophic phenotype in mdx mice. Hum Mol Genet. 2016;25(1):130–145.[Google Scholar]
120. Eshraghi M, et al Effect of genetic background on the phenotype of the Smn2B/− mouse model of spinal muscular atrophy. Hum Mol Genet. 2016;25(20):4494–4506.[Google Scholar]

Osteopontin regulates dentin and alveolar bone development and mineralization

1. Introduction

2. Materials and methods

2.1. Mice

2.2. Laser capture microdissection and protein extraction

2.3. LC-MS/MS and bioinformatics analysis

2.4. Microcomputed tomography

2.5. Histology, immunohistochemistry, and in situ hybridization

2.6. Histomorphometry

2.7. Quantitative real-time polymerase chain reaction

2.8. Statistical analysis

2.1. Mice

2.2. Laser capture microdissection and protein extraction

2.3. LC-MS/MS and bioinformatics analysis

2.4. Microcomputed tomography

2.5. Histology, immunohistochemistry, and in situ hybridization

2.6. Histomorphometry

2.7. Quantitative real-time polymerase chain reaction

2.8. Statistical analysis

3. Results

3.1. Proteomic analysis of PDL in Ank versus control mice

3.2. Osteopontin is expressed in the periodontium during tooth root development

3.3. Loss of OPN affects dental and periodontal development and mineralization

3.4. Altered gene expression in periodontal tissues of Spp1 mice

3.5. OPN does not regulate cementogenesis in Ank mice

3.1. Proteomic analysis of PDL in Ank versus control mice

3.2. Osteopontin is expressed in the periodontium during tooth root development

3.3. Loss of OPN affects dental and periodontal development and mineralization

3.4. Altered gene expression in periodontal tissues of Spp1 mice

3.5. OPN does not regulate cementogenesis in Ank mice

4. Discussion

4.1. Role of OPN in regulation of dentoalveolar development and mineralization

4.2. Osteopontin and cementum formation

4.1. Role of OPN in regulation of dentoalveolar development and mineralization

4.2. Osteopontin and cementum formation

Supplementary Material

1

2

3

4

5

S1

S10

S2

S3

S4

S5

S6

S7

S8

S9

1

2

3

4

5

S1

S10

S2

S3

S4

S5

S6

S7

S8

S9

Acknowledgments

Abstract

Footnotes

References

3.1. Proteomic analysis of PDL in Ank^{versus control mice}

3.4. Altered gene expression in periodontal tissues of Spp1^mice

3.5. OPN does not regulate cementogenesis in Ank^mice

3.1. Proteomic analysis of PDL in Ank^{versus control mice}

3.4. Altered gene expression in periodontal tissues of Spp1^mice

3.5. OPN does not regulate cementogenesis in Ank^mice