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Characterization of Dental Pulp Stem Cells of Human Tooth Germs
T. Takeda1,
Y. Tezuka2,
M. Horiuchi3,
K. Hosono2,
K. Iida1,
D. Hatakeyama1,
S. Miyaki3,
T. Kunisada2,
T. Shibata1 and
K. Tezuka2,4,*
1 Department of Oral and Maxillofacial Science and
2 Department of Tissue and Organ Development, Gifu University Graduate School of Medicine, 1-1 Yanagido, Gifu City, Gifu 501-1194, Japan;
3 Department of Regenerative Biology and Medicine, National Research Institute for Child Health and Development, 2-10-1 Ookura, Setagaya, Tokyo 157-8535, Japan; and
4 PRESTO, Japan Science and Technology Agency, 4-1-8 Honcho, Kawaguchi, Saitama 322-0012, Japan
Correspondence: * corresponding author, tezuka{at}gifu-u.ac.jp
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ABSTRACT
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In previous studies, human dental pulp stem cells (hDPSCs) were mainly isolated from adults. In this present study, we characterized hDPSCs isolated from an earlier developmental stage to evaluate the potential usage of these cells for tissue-regenerative therapy. hDPSCs isolated at the crown-completed stage showed a higher proliferation rate than those isolated at a later stage. When the cells from either group were cultured in medium promoting differentiation toward cells of the osteo/odontoblastic lineage, both became alkaline-phosphatase-positive, produced calcified matrix, and were also capable of forming dentin-like matrix on scaffolds in vivo. However, during long-term passage, these cells underwent a change in morphology and lost their differentiation ability. The results of a DNA array experiment showed that the expression of several genes, such as WNT16, was markedly changed with an increasing number of passages, which might have caused the loss of their characteristics as hDPSCs.
Key Words: human • dental pulp stem cells • tooth germs • gene expression • long-term culture
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INTRODUCTION
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Human dental pulp stem cells (hDPSCs) have been isolated from adult tooth pulps and characterized in terms of their self-renewal capability, multi-lineage differentiation, and clonogenic efficiency (Gronthos et al., 2000, 2002). Isolated pulp cells can be induced to differentiate into odontoblast-like cells and generate a dentin-like mineral structure in vitro (Tsukamoto et al., 1992; About et al., 2000). hDPSCs are multipotent cells known to form ectopic dentin and associated pulp tissue in vivo (Batouli et al., 2003). Tooth development is a complex process. A tooth germ starts as an aggregation of cells that forms a hard tissue called the crown. Tooth development then proceeds through the following stages: crown-completed, root-formative, and finally root-completed. Third molar development in humans proceeds with age as follows: crown-completed (12–16 yrs of age), root-formative (15–20 yrs), and root-completed (18–25 yrs) stages.
In previous studies, hDPSCs were mainly isolated from adults (older than 19 yrs) at the root-formative/-completed stage (Gronthos et al., 2000, 2002; Batouli et al., 2003; Papaccio et al., 2006; Yamada et al., 2006); however, detailed characterization of hDPSCs isolated from an earlier stage has not been well-documented. Therefore, the purpose of the present study was to characterize hDPSCs isolated from developing third molars at the crown-completed stage with respect to cell differentiation and gene expression profiles.
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MATERIALS & METHODS
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Isolation and Culture of Human Dental Pulp Stem Cells
Normal human third molars were extracted at Gifu University Medical Hospital after informed consent had been obtained from each person (age range, 12–25 yrs), according to a protocol approved by the Institutional Review Board. hDPSCs and human marrow stromal cells (hMSCs, from a 20-year-old individual) were isolated and cultured as previously described (Tanaka et al., 1995; Kuznetsov et al., 1997; Gronthos et al., 2000). The developmental stage of each tooth was estimated by morphology. The isolated hDPSCs were cultured in MSCGM medium (Cambrex, Walkersville, MD, USA). For in vitro differentiation experiments, we used at least 2 independent cell lines for each group. For the induction of osteo/odontogenesis, hDPSCs were cultured in MSCGM medium supplemented with 0.1 µM dexamethasone, 50 µg/mL ascorbic acid (Wako, Tokyo, Japan), and 0.1% β-glycerophosphate (Sigma, St. Louis, MO, USA). hDPSCs were plated at 1.0 x 105 cells/well in 12-well plates. After a culture period of 15 days, alkaline phosphatase (ALP) analysis was performed with an ALP staining kit (Muto, Tokyo, Japan), and von Kossa staining was conducted as described previously (Tezuka et al., 2002). For the induction of adipogenesis, an adipogenic induction medium kit (Cambrex) was used according to the manufacturer’s instructions. We assessed the accumulation of neutral lipid-containing vacuoles by staining with Oil red O. For neural differentiation, cells attached to 0.1% gelatin-coated dishes were incubated in Neurobasal A medium containing B27 supplement (GIBCO/Invitrogen, Carlsbad, CA, USA), 1% penicillin, 20 ng/mL epidermal growth factor, and 40 ng/mL fibroblast growth factor (R&D Systems, Minneapolis, MN, USA) for 18 days (Miura et al., 2003).
To determine the number of cumulative population doublings, we seeded hDPSCs into 10-cm dishes. Every 3–5 days, when the cells had become confluent, they were trypsinized and re-seeded at a density of 1.0 x 106 cells/dish. This procedure was repeated for every passage (P). Cells were counted at each passage with a hemocytometer.
Transplantation and Preparation of Histological Sections
Approximately 1.0 x 106 hDPSCs were seeded onto a Calcium Phosphate Scaffold (BD Biosciences, Bedford, MA, USA), cultured for 7 days, and then transplanted subcutaneously into the dorsal surfaces of 8-week-old immunocompromised mice (NOD/Shiscid; CLEA, Tokyo, Japan), as previously described (Krebsbach et al., 1997). Our animal-use protocols were reviewed and approved by the Gifu University Institutional Review Board. The transplants were removed at 8 and 15 wks post-transplantation, and preparation of frozen non-decalcified sections was made with a cryofilm-transfer kit (FINETEC Co. Ltd., Tokyo, Japan; Kawamoto, 2003). Briefly, the transplants were immediately frozen in dry ice-hexane. Each sample was embedded in 4% carboxymethyl cellulose gel, completely frozen, and then attached to the sample stage of a cryomicrotome (CM3050S; Leica Instruments, Heidelberg, Germany). Thereafter, the surface of each sample was covered with adhesive film (FINETEC Co. Ltd.), and the sample was sagittally sectioned along with the film at a thickness of 7 µm by the use of a disposable tungsten carbide blade (Leica Instruments). Each section was then stained with hematoxylin and eosin or von Kossa stain.
cDNA Microarray Analysis
Total RNA was isolated from the cells with an RNeasy Plus Mini Kit (Qiagen, Valencia, CA, USA). Human gene expression was examined by means of a Human Genome U133 Plus 2.0 probe array (GeneChip; Affymetrix, Santa Clara, CA, USA), which contains approximately 54,000 genes, and a Hewlett-Packard Gene Array Scanner (Palo Alto, CA, USA). Each sample was analyzed once. The fluorescence intensity of each probe was quantified by GeneChip Analysis Suite 5.0 (Affymetrix). The level of gene expression was determined as the average difference, obtained by the GeneChip software. Micro-array data compliant with the MIAME guidelines (http://www.mged.org/Workgroups/MIAME/miame.html) and experimental details have been submitted to GEO (http://www.ncbi.nlm.nih.gov/geo/) under series accession number GSE10444. Gene functional classification analyses were performed with DAVID software (Dennis et al., 2003).
Semi-quantitative Reverse-transcriptase/Polymerase Chain-reaction (RT-PCR)
First-strand cDNA mixture was used for PCR with Taq polymerase (Takara, Shiga, Japan), and the PCR was performed as described previously (Miura et al., 2003). The primers used for the PCR are shown in Appendix Table 1.
Quantitative Real-time PCR
PCR amplification of cDNA was performed by SYBR Premix Ex Taq (Takara). The primers for the real-time PCR are listed in Appendix Table 1. After an initial denaturation at 95°C for 10 min, a two-step cycle procedure was used (denaturation at 95°C for 5 sec, annealing and extension at 60°C for 30 sec) for 40 cycles in a Thermal Cycler Dice (Takara). Gene expression levels were normalized according to the level of GAPDH expression. Relative amounts of WNT16 and GAPDH mRNA in each sample were calculated from standard curves obtained by sequential dilution of total RNA prepared from hDPSCs at passage 20 (P20); and those of tumor protein D52 (TPD52), inositol 1,4,5-trisphosphate receptor (ITPR1), Toll-like receptor 4 (TLR4), and GAPDH mRNA, from the standard curves obtained for hDPSCs at P4.
Statistical Analysis
Data are presented as means ± standard deviation. The differences in mean values of doubling times were evaluated by the t test after evaluation of variances (Microsoft Excel, Windows XP). For real-time PCR, the mean and standard deviation of expression coefficient were calculated by Thermal Cycler Dice Real Time System Software Ver. 1.03 (Takara).
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RESULTS
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Isolation and Characterization of hDPSCs
Dental pulp tissues were isolated from the third molars of 100 persons (DP 1-100), and hDPSCs were successfully isolated from 91 of them (Appendix Table 2). Nine samples were lost due to bacterial and/or fungal contamination. After collagenase digestion, 2.2 ± 0.7 x 105 and 1.2 ± 0.7 x 105 live nucleated cells were isolated from each third molar at crown-completed and root-formative/-completed stages, respectively. When seeded in culture dishes, these cells formed colonies with a scattered morphology after 5 days’ incubation (Fig. 1A ). We calculated that 48 ± 30 and 40 ± 35 colonies/104 cells were formed from randomly selected crown-completed (N = 3) and root-formative/-completed (N = 4) teeth, respectively. Cells isolated from either group were then cultured in medium promoting osteo/odontoblastic differentiation. After 15 days in culture, they became ALP-positive and produced calcified matrix, as judged from the results of von Kossa staining (Figs. 1B, 1C). When the cells were grown for 4 wks in medium conducive to adipogenic differentiation, small lipid-containing vesicles were observed in cells of either crown-completed or root-formative/-completed cultures (Appendix Fig. 1A). hMSCs isolated from bone marrow formed numerous adipocytes containing larger lipid vesicles under the same culture conditions (Appendix Fig. 1B). Peroxisome proliferator-activated receptor- 2 and lipoprotein lipase were not detected in hDPSCs, even after adipogenic induction (Appendix Fig. 1C). Next, we found that hDPSCs cultured in neural differentiation medium expressed neural cell markers such as nestin (Appendix Fig. 1D) and βIII-tubulin (Appendix Fig. 1E). Control antibody showed no immunoreactivity (Appendix Fig. 1F). These results suggest that hDPSCs isolated from developing third molars had differentiation potential comparable with that previously reported for later stages.
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Figure 1. Differentiation of hDPSCs. (A–C) In vitro differentiation of hDPSCs. hDPSCs collected at the crown- (DP1) and root-completed (DP75) stages were cultured and then tested for their ability to differentiate into cells of multiple lineages. Cell morphology of DP1 cells after 5 days in culture is shown in (A). Osteogenesis was assessed by ALP-activity staining (B) and with von Kossa stain (C). Scale bar = 200 µm (A–C). (D–I) Characterization of hDPSCs (crown-completed stage [DP1] and root-formative stage [DP27]) transplanted into NOD-SCID mice for assessment of differentiation in vivo. DP1 (D–F) and DP27 (G–I) cells were seeded onto calcium-phosphate scaffolds and transplanted subcutaneously into NOD-SCID mice. Tissues were isolated at 15 wks post-transplantation and stained with hematoxylin and eosin (D,E,G,H) or with von Kossa (F,I) stain. Polarized light demonstrates alignment of the collagen fibers on the forming surfaces (E,H). In the hDPSC transplants, the scaffolds (Sc) are lined with a dentin-like matrix (d) surrounding connective tissue (CT). Arrows show the calcified matrix (F,I). Scale bar = 500 µm (D–I).
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hDPSCs can Generate a Dentin/Pulp-like Structure in vivo
At 8 wks post-transplantation, cells from both crown-completed stage (DP1) and root-formative stage (DP27) samples started to generate a matrix on the surface of the implanted scaffolds, and by 15 wks post-transplantation, they had formed dentin- and pulp-like connective tissues (Figs. 1D, 1G ). As shown previously (Gronthos et al., 2000), a highly ordered collagenous matrix, deposited perpendicular to the odontoblast-like layer, was visualized by illumination with polarized light (Figs. 1E, 1H). Evidence of calcification of this dentin-like matrix was obtained by von Kossa staining (Figs. 1F, 1I).
Changes in Cell Morphology and Differentiation Capability during Long-term Culture
Our study compared the long-term growth curves of crown-completed (DP1, 2, 11, 28, and 31) and root-formative/-completed (DP27, 29, 33, 34, 54, and 75; Fig. 2A) groups. The former maintained a high growth rate for more than 2 mos, a rate higher than that of the latter. The average doubling time was 42 ± 2.8 hrs for the crown-completed group and 65 ± 6.5 hrs for the root-formative/-completed one, a difference that was statistically significant (P = 0.0001). The growth rate of the cells obviously decreased with an increase in the number of passages. In addition, the morphology of crown-completed stage (DP1) cells changed rapidly during long-term culture (Fig. 1A, Appendix Fig. 2). These cells changed to a spindle-like shape and then became flattened. More importantly, they gradually lost their cell differentiation phenotype after long-term culture (Fig. 2B). Less apparent, but similar, changes in morphology and ALP activity were also observed with root-formative stage (DP27) cells (Fig. 2B and Appendix Fig. 2). At 8 wks post-transplantation, we also confirmed that crown-completed stage (DP1) cells could not produce dentin-like tissue in vivo after 10 passages (Fig. 2C, right); whereas P4 transplants showed a layer of dentin-like matrix covered with an interface layer of odontoblast-like cells (Fig. 2C, left and middle).
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Figure 2. Growth curves and change in differentiation potential of hDPSCs during long-term culture. (A) Curves for long-term growth obtained from 11 cell lines isolated from individuals of various stages (crown-completed stage, DP1, 2, 11, 28, and 31 [red]; root-formative stage, DP27 [black]; root-completed stage, DP29, 33, 34, 54, and 75 [black]). Cells isolated from the crown-completed stage maintained a high growth rate for more than 2 mos, whereas those from later stages showed a lower growth rate. (B) ALP staining of crown-completed stage (DP1 [top]) and root-formative stage (DP27 [bottom]) cell cultures at P3, 10, 20, and 30, respectively. DP1 and DP27 cells lost their differentiation capability induced by confluence after long-term culture. (C) In vivo differentiation of crown-completed stage (DP1) cells at P4 and P10. Transplants were removed at 8 wks post-transplantation, sectioned, and stained with hematoxylin and eosin. High-magnification view of the boxed area in the left photo is shown in the middle one. In P4 transplants, the scaffolds (Sc) were lined with a layer of dentin-like matrix (d, middle) surrounding connective tissue (CT) and an interface layer of odontoblast-like cells (od, middle); whereas no dentin-like matrix was observed in P10 transplants. Scale bar = 200 µm (left and right), 100 µm (middle).
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Comparison of Gene Expression Profiles
To identify genes modulated by passage, we performed cDNA microarray analysis on 3 independent samples at the crown-completed stage (DP2, 28, and 31) that had undergone 4 (P4) or 10 passages (P10). The number of genes down-regulated more than two-fold at P10, compared with their expression at P4, was 719. In contrast, the number of genes up-regulated more than two-fold at P10, compared with their expression at P4, was 642. These genes were classified into 14 (P4>P10) and 11 (P4<P10) functional categories by DAVID (Appendix Table 3). Genes up-regulated or down-regulated more than five-fold in all 3 samples, when P4 was compared with P10, were selected and listed (Table and Appendix Table 4).
To evaluate the reliability of microarray analysis and to confirm the passage-dependent increase or decrease in mRNA expression level, we performed real-time PCR. As expected, the expression of WNT16 mRNA increased significantly with an increase in passage number in all the samples tested (Fig. 3A). In contrast, expression of TPD52, ITPR1, and TLR4 mRNA decreased with passage number (Figs. 3B–3D). This decrease in TLR4 mRNA expression tended to be slower for root-completed- than crown-completed-stage hDPSCs (Fig. 3D).
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Figure 3. Change in expression levels of WNT16, TPD52, ITPR1, and TLR4 mRNA in long-term cultures. Real-time PCR analysis was performed with crown-completed stage (DP2, DP28, and DP31) and root-completed stage (DP33, DP54, and DP75) cells at P4, P10, and P20. The relative amounts of WNT16 (A), TPD52 (B), ITPR1 (C), and TLR4 (D) mRNA were divided by that of GAPDH and used to calculate expression coefficients. Error bars indicate the standard deviation obtained from duplicate analyses.
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DISCUSSION
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In this study, we successfully isolated hDPSCs from developing third molars, extracted before eruption. We observed that the hDPSCs isolated from the crown-completed stage showed higher proliferation potential compared with those from later stages. Cells isolated from any stage formed dentin-like structures when transplanted into immunodeficient mice. Combined with previous data (Batouli et al., 2003), these facts suggest that the dental pulp tissue contained stem cells capable of generating dentin-like structures, even when isolated at the early stage of tooth development.
Observation of the cells in long-term culture revealed that, both in vitro and in vivo, they lost differentiation capability; and by microarray analysis, we showed that WNT16 expression markedly increased with passage number. We also found that WNT1 expression also increased in P10 by 4.9-fold. The WNT family is a group of signaling molecules shown to control a diverse range of developmental processes, including fate specification, proliferation, polarity, and migration of cells (Nusse and Varmus, 1982; Wodarz and Nusse, 1998), as well as tooth renewal (Jarvinen et al., 2006). In contrast, expression of TPD52 mRNA decreased with an increase in passage number. TPD52 has been implicated in cell proliferation, apoptosis, and vesicle trafficking (Lewis et al., 2007). Therefore, these may be key molecules suppressing cell differentiation and/or maintaining proliferation of hDPSCs in long-term culture.
Among the genes expressed at higher levels at P4 than at P10, Rho GTPase activating protein 8 (ARHGAP8) specifically binds to RhoA and is involved in the regulation of actin cytoskeleton organization, membrane trafficking, gene expression, and cell proliferation (Shang et al., 2003). High expression of this molecule along with that of other GTP-binding proteins may be related to the morphology of hDPSCs freshly isolated from the crown-completed stage, and to the morphological change observed in these cells at later passages. Another gene down-regulated after P4, ITPR1 acts as a channel to release Ca2+ from internal stores (Takei et al., 1998; Mikoshiba, 2006). TLR4 is expressed in dendritic and other cells in dental pulp (Okiji et al., 1997; Sakurai et al., 1999), and has been immunolocalized in human odontoblasts (Veerayutthwilai et al., 2007). These signaling molecules may be involved in the differentiation capability and/or proliferation of hDPSCs; however, there is too little information for an in-depth discussion of the possible roles of these molecules.
In this study, we characterized hDPSCs isolated from tooth germs at the crown-completed stage and found that these cells were highly proliferative and had the potential to generate a dentin-like matrix in vivo. However, these characteristics were lost in long-term culture, with a change in their gene expression profile. These results will provide fundamental information to prepare a large number of cells for use in tissue-regeneration therapy without loss of differentiation capability.
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ACKNOWLEDGMENTS
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We thank the members of the Department of Tissue and Organ Development and the Department of Oral and Maxillofacial Sciences of Gifu University Graduate School of Medicine for their technical help and fruitful discussions. This work was supported by grants from the Ministry of Education, Science, and Culture of Japan and the Japan Science and Technology Agency.
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FOOTNOTES
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A supplemental appendix to this article is published electronically only at http://jdr.iadrjournals.org/cgi/content/full/87/7/676/DC1.
Received for publication September 3, 2007.
Revision received February 15, 2008.
Accepted for publication March 4, 2008.
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Journal of Dental Research, Vol. 87, No. 7,
676-681 (2008)
DOI: 10.1177/154405910808700716
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ABSTRACT
INTRODUCTION