- Email: [email protected]
Effect of different calcium phosphate scaffold ratios on odontogenic differentiation of human dental pulp cells
Materials Science and Engineering C 49 (2015) 225–233
Contents lists available at ScienceDirect
Materials Science and Engineering C journal homepage: www.elsevier.com/locate/msec
Effect of different calcium phosphate scaffold ratios on odontogenic differentiation of human dental pulp cells Sarah Talib AbdulQader a,d, Thirumulu Ponnuraj Kannan a,b,⁎, Ismail Ab Rahman a, Hanafi Ismail c, Zuliani Mahmood a a
School of Dental Sciences, Universiti Sains Malaysia, 16150 Kubang Kerian, Kelantan, Malaysia Human Genome Centre, School of Medical Sciences, Universiti Sains Malaysia, 16150 Kubang Kerian, Kelantan, Malaysia School of Materials and Minerals Resource Engineering, Universiti Sains Malaysia, 14300 Penang, Malaysia d Department of Pedodontic and Preventive Dentistry, College of Dentistry, University of Baghdad, Baghdad, Iraq b c
a r t i c l e
i n f o
Article history: Received 12 July 2014 Received in revised form 29 October 2014 Accepted 17 December 2014 Available online 20 December 2014 Keywords: Biphasic calcium phosphate Human dental pulp cells Calcium and phosphate ions Tissue regeneration Odontogenesis
a b s t r a c t Calcium phosphate (CaP) scaffolds have been widely and successfully used with osteoblast cells for bone tissue regeneration. However, it is necessary to investigate the effects of these scaffolds on odontoblast cells' proliferation and differentiation for dentin tissue regeneration. In this study, three different hydroxyapatite (HA) to beta tricalcium phosphate (β-TCP) ratios of biphasic calcium phosphate (BCP) scaffolds, BCP20, BCP50, and BCP80, with a mean pore size of 300 μm and 65% porosity were prepared from phosphoric acid (H2PO4) and calcium carbonate (CaCO3) sintered at 1000 °C for 2 h. The extracts of these scaffolds were assessed with regard to cell viability and differentiation of odontoblasts. The high alkalinity, more calcium, and phosphate ions released that were exhibited by BCP20 decreased the viability of human dental pulp cells (HDPCs) as compared to BCP50 and BCP80. However, the cells cultured with BCP20 extract expressed high alkaline phosphatase activity and high expression level of bone sialoprotein (BSP), dental matrix protein-1 (DMP-1), and dentin sialophosphoprotein (DSPP) genes as compared to that cultured with BCP50 and BCP80 extracts. The results highlighted the effect of different scaffold ratios on the cell microenvironment and demonstrated that BCP20 scaffold can support HDPC differentiation for dentin tissue regeneration. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Tissue engineering approaches have gained great importance to restore or replace tissues that were damaged or lost. This involves the regeneration of new functional healthy tissues using progenitor cells with scaffold in response to appropriate signals [1]. In clinical dentistry, dentin tissue regeneration is a desirable goal that allows the formation of new healthy dentin to be integrated with the pre-existing dentin to overcome the drawbacks of the conventional dental treatments [2–4]. Recent advances in stem-cell biology have revealed the possibility of dentin regeneration using human adult dental pulp cells (HDPCs) as progenitor cells that have the ability to form a dentin/pulp-like complex [5,6] and possess self-renewal [7] and multi-lineage differentiation capability [8–10]. These cells can be carried by scaffolds that play an important role in providing physical support for the cells and maintaining space for the regenerated tissue. These scaffolds guide new tissue growth before degradation and are eventually replaced by new tissues [11]. ⁎ Corresponding author at: School of Dental Sciences, Universiti Sains Malaysia, 16150 Kubang Kerian, Kelantan, Malaysia. E-mail address: [email protected] (T.P. Kannan).
http://dx.doi.org/10.1016/j.msec.2014.12.070 0928-4931/© 2014 Elsevier B.V. All rights reserved.
Scaffold biological properties depend primarily on the chemical composition of the biomaterial [12,13], and its porous structure: pore shape, pore size, porosity percentage and pore interconnection pathway [14,15]. Microporosity of diameter b10 μm permits body fluid circulation and macroporosity of diameter N100 μm is necessary for migration and proliferation of cells and tissue formation [16,17]. The total porosity of 65% is efficient for total protein production and alkaline phosphatase (ALP) activity that were taken as indicators of growth/matrix production and for the comparison of cell differentiation [18]. Among many types of scaffolds that have been used for the regeneration of hard tissues, biphasic calcium phosphate (BCP) scaffolds have shown to induce appropriate osteoblastic differentiation of stem/ progenitor cells in vitro and bone formation in vivo [19–21] due to their desirable properties including similarity in composition to the bone mineral, bioactivity and osteoconductivity [22,23]. They consist of an intimate mixture of hydroxyapatite (HA) [Ca10(PO4)6(OH)2], and beta-tricalcium phosphate (β-TCP) [Ca3(PO4)2] crystals of varying phase compositions (HA/β-TCP ratios) that have been reported to be more effective than pure HA or β-TCP alone [24–26]. They exhibit different biological behaviors of their components in which HA is bioactive and β-TCP is resorbable [27]. The effects of different HA/β-TCP ratios of several modified BCP scaffolds have been well studied for bone tissue
226
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
regeneration [15,28–31]. It was reported that BCP of 20/80 HA to β-TCP ratio stimulated the osteogenic differentiation of human mesenchymal stem cells (HMSCs) in vitro and fastest rate of bone induction in vivo as compared to 100% HA, HA/β-TCP (76/24, 63/37, 56/44), and 100% β-TCP [24]. However, these effects are not well known for odontogenic cell differentiation and dentin tissue regeneration. Although odontoblasts and osteoblasts show almost identical genetic profiles, there is a clear difference between the process of odontogenesis and osteogenesis and fundamental differences in cellular morphology between bone and dentin. These differences can affect the convenience and appropriateness of the scaffold used in tissue regeneration [32]. In spite of all improvements on the solubility and bioactivity obtained with BCPs, it is important to consider the possible impact of their composition on the processes linked to cell proliferation and differentiation, and to ensure the proportional amounts of each component do not impoverish the final composite properties. Hence, it would be of interest to evaluate the effect of different HA/β-TCP ratios of BCP scaffolds on HDPC odontogenesis for dentin tissue regeneration. The aim of the present study was to determine the optimum microenvironment and the ratio of HA to β-TCP that supported the viability and the differentiation of HDPCs. The influence of BCP characteristics of different ratios of HA to β-TCP and their degradation products on the microenvironment, viability, ALP activity, and gene expression of HDPCs has been investigated. The null hypothesis was that there would be no differences in the microenvironments and in HDPCs' odontogenesis due to the different ratios of HA to β-TCP. 2. Materials and methods 2.1. Scaffold preparation and characterization Calcium phosphate powders were prepared by a wet precipitation method using CaCO3 and H2PO4 as starting materials as mentioned previously [33]. Briefly, the reagents were mixed at Ca/P ratios of 1.517, 1.568, and 1.619 corresponding to BCP of 20/80 HA to β-TCP ratios (BCP20), 50/50 HA to β-TCP ratio (BCP50), and 80/20 HA to β-TCP ratio (BCP80) respectively. The precipitate was heated at 80 °C for an hour with stirring, aged for 48 h, washed, filtered, and dried at 100 °C. The ground powder was then added to polyethylene (PE) spherical particles as pore-former agents of 300–350 μm (Cospheric, USA) at a ratio of 4:2.5 v/v for total porosity of 65% and pore size range of 300 μm. The powder mixtures were compressed with a uniaxial press of 24 MPa in a 32 mm die to form 6 mm thickness pellets. Subsequently, pellets were submitted to thermal treatment of 400 °C for 2 h and sintered at 1000 °C for 2 h. Then, the pellets were crushed and sieved to obtain granules with the desired particle size range of 0.5–1 mm as shown in Fig. 1.
The phases of HA and β-TCP and the ratios of HA/β-TCP of the scaffold were analyzed qualitatively and quantitatively by X-ray diffraction (XRD) analysis using Eva and X'Pert HighScore Plus software, respectively. The registered patterns were compared with the International Centre for Diffraction Data (ICDD) powder diffraction file (PDF) to determine the crystalline phases. Field emission scanning electron microscopy (FESEM) (Zeiss Supra 55VP, Germany) was used to evaluate the surface morphology of the scaffolds. The scaffolds' surfaces were first sputter-coated with a layer of platinum alloy (150 Å thick). The total porosity of the scaffolds was assessed using Archimedes' method. 2.2. Preparation of scaffold extract The particles thus obtained were sterilized by autoclave before use. The extract of each scaffold sample (BCP20, BCP50, and BCP80) was prepared according to international standards for medical devices evaluation ISO standard (10993-12) [34]. After sterilization, 1 g of each scaffold material (in the form of particles) was incubated in 10 ml alpha minimum essential medium (Alpha-MEM, Invitrogen, USA) for 1, 3 and 7 days at 37 °C. After incubation, the extract was collected by filtration using 0.2 μm filter. 2.3. Measurements of calcium and phosphate ions, and pH values of the extracts The content of calcium and phosphate ions in each extract at each time was quantified colorimetrically using Calcium and Phosphate Colorimetric Assay Kits (BioVision, USA) according to the manufacturer's protocols. Briefly, for Ca2+ measurement, a chromogenic complex formed between Ca2+ and 0-cresolphthalein was measured at 575 nm using an ELISA reader (Sunrise, Tecan, Austria). For phosphate ion measurement, malachite green and ammonium molybdate that form a chromogenic complex with phosphate ions was measured at 405 nm. The intensity of the color was directly proportional to the concentration of ions in the samples. The difference between amounts of ions in the media exposed or not to particles, in the absence of cells, corresponded to the release of ions from the particles. Changes in the pH were measured at pre-determined time intervals as a function of time using a pH meter (Hanna 210, Japan). 2.4. Culture of HDPCs Immortalized human dental pulp cells (HDPCs) that was kindly provided by Professor Takashi Takata (Hiroshima University, Hiroshima, Japan) were used in this research [35]. Cells were cultured in AlphaMEM supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin, and 100 mg/ml streptomycin in a humidified atmosphere of 5% CO2 and 95% relative humidity at 37 °C. Culture medium was changed every three days until confluence. Cells were trypsinized, suspended in complete culture medium of Alpha-MEM, and subcultured at a cell density of 1 × 104 cells/well in 96-well culture plates (Corning, USA) with 200 μl of culture medium. After 24 h, the medium was removed from each well and replaced by 180 μl of each scaffold extract plus 20 μl of fetal bovine serum (final concentration of 10%). The extracts of BCP20, BCP50, and BCP80 scaffolds incubated in the medium for 3 days were chosen for the rest of the study's tests. 2.5. Cell viability assay
Fig. 1. Particle size of the prepared scaffolds after sieving as visualized by image analyzer.
The viability of HDPCs exposed to the different scaffold extracts was evaluated by MTT assay. After 1, 3, 5, and 7 days of incubation of cells with different scaffold extracts, the extracts were removed and the cells were tested for viability. The 3-(4,5-dimethyl-thiazoyl)-2,5diphenyl-tetrazolium bromide (MTT) (Sigma-Aldrich, USA) assay was used as previously described [36] to determine cell viability. This assay
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
is based on the reduction of the soluble yellow MTT tetrazolium salt to a blue insoluble formazan product by mitochondrial succinic dehydrogenase. After each incubation period, 30 μl of sterile-filtered MTT solution (5 mg/ml in PBS) was added into each well and re-incubated in 5% CO2 and 95% relative humidity at 37 °C for 4 h. Then, the culture medium with the MTT solution was aspirated and the formazan crystals were dissolved in 200 μl of dimethyl sulfoxide (DMSO). To extract and solubilize the formazan, the test plate was agitated by using a microplate shaker for several minutes. After confirmation of the homogeneity of the solutions, the absorbance of the solution was measured at 570 nm using an ELISA reader (Sunrise, Tecan, Austria). Cell viability was measured as a ratio of the optical density in the extract solution medium to the optical density in the fresh medium (control groups) using the following the formula: Cell viability% ¼ ½Absorbance of treated cells=Absorbance of control cells 100%: The absorbance was the average value measured from six wells in parallel for each sample.
227
of forward primer, 3.0 μl of 0.6 μM of reverse primer, 2.0 μl of enzyme mix containing omniscript, sensiscript reverse transcriptases and HotStarTaq DNA polymerase enzymes, 25.0 μl of RNase-free water, and 5.0 μl of 100 ng of isolated RNA. HotStarTaq DNA Polymerase was activated to initiate PCR using Quick start mastercycler Pro (Eppendorf, Germany) at 95 °C for 15 min. cDNA amplification was carried out by initial denaturation at 95 °C for 30 s, annealing at 54–56 °C for 30 s, and extension at 72 °C for 60 s for 30 cycles; then, final extension at 72 °C for 10 min. The amplification reaction products were analyzed on 2% agarose gel, electrophoresed for 60 min at 75 V using Power Pac HC (Bio-Rad, USA) and visualized under ultraviolet light of Gel doc electrophoresis (Bio-Rad, USA). The genes selected in the current study were collagen 1 alpha 1 (COL1A1), bone sialoprotein (BSP), dental matrix protein-1 (DMP-1), and dentin sialophosphoprotein (DSPP), and GAPDH (glyceraldehyde3-phosphate dehydrogenase) as the reference housekeeping gene. The PCR primers of targeted genes were followed as reported earlier. The primers are listed in Table 1. 2.8. Statistical analysis
2.6. Assessment of alkaline phosphatase activity Quantitative ALP activity of HDPCs exposed to the different scaffold extracts was determined at 0–3, 3–6, 6–9, 9–12, and 12–15 days by the hydrolysis of p-nitrophenylphosphate to p-nitrophenol using an Alkaline Phosphatase Activity Colorimetric Assay Kit (BioVision, USA) according to the manufacturer's protocols. Cells were cultured at a cell density of 1 × 104 cells/well in 96-well culture plate (Corning, USA) with 200 μl of culture medium of 10% FBS until they reached 80% confluence. Cells were then exposed to different scaffold extracts with 50 μg/ml ascorbic acid (Sigma-Aldrich, USA), and 10 mmol/l β-glycerophosphate (Sigma-Aldrich, USA) for odontogenic induction and were cultured for 15 days. Culture extract medium was changed every three days. During each time of changing medium, 30 μl of supernatant was added to 50 μl of ALP assay buffer in 96 well plate. Then, 50 μl of 5 mM p-nitrophenylphosphate was added and the mixtures were incubated at 25 °C for 60 min. After incubation, 20 μl of stop solution was added to stop the enzymatic reaction and the absorbance was read at 405 nm using an ELISA reader (Sunrise, Tecan, Austria). ALP activity of the test samples was calculated using the following equation: ALP activity ðU=mlÞ ¼ A=V=T where A is the amount of pNP generated by the sample, V is the volume of the sample added in the assay well, and T is the reaction time (in minutes). 2.7. Total RNA extraction and reverse transcription-polymerase chain reaction analysis (RT-PCR) Total RNA of HDPCs cultured in 25 cm2 flask (n = 3 for each group) and exposed to BCP20, BCP50 and BCP80 extracts for 14, 21, and 28 days was extracted using an innuPREP RNA Mini Kit (analytikjena, Germany) according to the manufacturer's instructions. The concentration and purity of total RNA were determined by absorbance at 260/280 nm in a UV spectrophotometer (Biophotometer plus, Eppendorf, Germany), and the RNA quality was assessed by electrophoresis (Bio-Rad, USA) on a 2% agarose gel. The RNA was stored at − 80 °C until amplification by one-step RT-PCR. Both cDNA synthesis and amplification by RT-PCR were conducted using a QIAGEN OneStep RT-PCR Kit (QIAGEN, Germany). cDNA synthesis was carried out at 50 °C for 30 min in a final 50 μl volume of reaction mixture containing 10.0 μl of 5× buffer (12.5 mM MgCl2), 2.0 μl of deoxy nucleoside triphosphate (dNTP) (400 μM of each dNTP), 3.0 μl of 0.6 μM
Each experiment was performed in triplicate. Results are expressed as the mean and standard error. The statistical analysis of the data was performed by one-way analysis of variance followed by a multiplecomparison Tukey test, using SPSS 20.0 program. A value of P b 0.05 was considered statistically significant. 3. Results The qualitative XRD patterns for the synthesized BCP scaffolds of different ratios are shown in Fig. 2a. Two phases of BCP samples: β-TCP [ICDD 01-070-2065] and HA [ICDD 00-009-0432] only were present. The main peak was at 2θ angle of 31.04° for β-TCP and at 2θ angle of 31.86° for HA. Fig. 2b–d shows quantitative X-ray diffraction pattern of BCP scaffolds. Ca/P ratios of 1.517, 1.568, and 1.619 correspond to HA/β-TCP ratio of 20.1/79.9 (BCP20), 49.8/50.2 (BCP50), and 80.3/19.7 (BCP80), respectively. BCP20, BCP50, and BCP80 scaffolds possess uniform spheroid macropore shapes as shown in Fig. 3a–c that were fabricated using spherical particles of PE as pore-former agents. Statistical analysis of the macroporous structure of the scaffolds indicates that the mean pore size was 308 ± 12 μm. The scaffold micropores are shown in Fig. 3d–f. There is a decrease in micropore size with increase in grain growth as β-TCP ratio increased in the synthesized scaffold. The total porosity of the scaffolds as measured by Archimedes method was 64.62 ± 0.84%, 65.02 ± 0.72%, and 65.74 ± 0.52% for BCP20, BCP50, and BCP80, respectively that includes both micropore and macropore volume. There was no significant difference (P N 0.05) in the total porosity for these scaffolds of different ratios. The in vitro stability of the culture medium with and without BCP particles were quantitatively measured and are graphically represented in Fig. 4. The calcium and phosphate ions, and pH values of the extract without BCP particles were more stable throughout the experimental period. For each group, there was a significant difference (P b 0.05) in calcium and phosphate ions as well as in pH values between day 1 and day 3, while there was no significant difference (P N 0.05) in these values between day 3 and day 7. At each time point, there was a significant difference (P b 0.05) in calcium and phosphate ions as well as in pH values among study groups. Values of pH, calcium, and phosphate ions increased as HA/β-TCP ratio decreased. The extracts of different scaffold ratios incubated in the medium for 3 days were chosen for the rest of the study's tests. For cell viability assay, HDPCs were cultured with the extracts of BCP20, BCP50, and BCB80 for 1, 3, 5, and 7 days. The results showed
228
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
Table 1 List of primer sequences, melting temperature (Tm), GenBank accession no. and their respective product size. Target gene
Primer sequences
Tm (°C)
GenBank No.
Size
[References]
COL1A1
Forward: 5′-caccaatcacctgcgtacag-3′ Reverse: 5′-tggtttcttggtcggtgg-3′ Forward: 5′-gaaccacttccccacctttt-3′ Reverse: 5′-tctgaccatcatagccatcg-3′ Forward: 5′-caggagcacaggaaaaggag-3′ Reverse: 5′-ctggtggtatcttgggcact-3′ Forward: 5′-tgtcgctgttgtccaagaag-3′ Reverse: 5′-attctttggctgccattgtc-3′ Forward: 5′-cgaccactttgtcaagctca-3′ Reverse: 5′-aggggagattcagtgtggtg-3′
60. 2 60.5 60.2 59.6 60.0 60.0 60.0 60.1 60.0 60.0
NM_000088
216 bp
[37]
NM_004967
201 bp
[38]
NM_004407.3
213 bp
[39]
NM_014208.3
498 bp
[35]
BC001601
203 bp
[40]
BSP DMP-1 DSPP GAPDH
that the viability of cells increased significantly (P b 0.05) with the addition of BCP extracts as compared to control group as shown in Fig. 5a. The time-dependent result showed that there was a statistically significant increase (P b 0.05) in the cell viability of HDPCs for each study group. Day 7 showed the highest level of cell viability of HDPCs cultured with BCP20, BCP50, and BCB80 extracts as compared to day 1, day 3, and day 5. At day 7, the means of cell viability were 117.45 ± 8.29, 131.46 ± 6.89, and 145.26 ± 7.58 for HDPCs cultured with BCP20, BCP50, and BCB80 extracts, respectively. The results showed that HDPCs demonstrated statistically a significant increase (P b 0.05) in cell viability when cultured with BCP80 extract as compared to that cultured with BCP20, and BCP50 extracts. ALP activity of HDPCs cultured with BCP20 extract increased significantly (P b 0.05) as compared to that cultured with BCP50 and BCP80 at time periods of 3–6 days, 6–9 days, 9–12 days, and 12–15 days as shown in Fig. 5b. The time period of 12–15 days showed the highest level of ALP activity of HDPCs for all study groups. At time period of 12–15 days, the means of ALP activity were 174.21 ± 11.29, 137.37 ± 12.54, and 106.06 ± 9.82 for HDPCs cultured with BCP20, BCP50, and BCB80 extracts, respectively. A significant highest level of ALP activity was detected for HDPCs cultured with BCP20. For gene expression analysis (Fig. 6), HDPCs cultured with BCP extracts showed high expression level of COL1A1, BSP, DMP-1, and
DSPP genes as compared to that cultured without BCP extracts at 14, 21, and 28 days. Among study groups, there was a significant increase (P b 0.05) in the expression level of BSP, DMP-1, and DSPP genes in HDPCs cultured with BCP20 extract as compared to that cultured with BCP50 and BCP80 extracts, whereas there was no significant difference (P N 0.05) in the expression level of COL1A1 gene in HDPCs cultured with BCP20, BCP50 and BCP80 extracts. 4. Discussion The assessment of the microenvironment in the scaffold pores surrounding cells is important to study the cells' behavior for tissue regeneration. The microenvironment of the cells is affected by the degradation products of the scaffold during tissue regeneration. BCP scaffold releases ionic product during degradation process that primarily depended on its HA to β-TCP ratio. The aim of this study was to determine if the difference in the ratio of HA to β-TCP had an impact on viability and differentiation of HDPCs. In this study, all BCP scaffolds of different ratios were prepared using phosphoric acid and calcium carbonate sintered at 1000 °C for 2 h to eliminate the effect of different preparation methods on the scaffold properties. XRD analysis was carried out as shown in Fig. 2. The highest peak intensities for β-TCP and HA for all scaffolds were at 31.04° and
Fig. 2. (a) Qualitative X-ray diffraction pattern of BCP20, BCP50, and BCP80 scaffolds synthesized at pH 7.0 sintered at 1000 °C for 2 h. (b), (c), and (d) Quantitative X-ray diffraction patterns of BCP20, BCP50, and BCP80 corresponding to HA/β-TCP ratios of 20.1/79.9, 50.2/49.8 and 80.3/19.7, respectively.
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
229
Fig. 3. Field emission scanning electron microscope images (a), (b), (c) the macroporosity structure and (d), (e), (f) the microporosity structure of BCP80, BCP50, and BCP20, respectively. Blue arrow represents large grain size, red arrow represents small grain size, and yellow arrow represents micropores. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
31.86° corresponding to planes (0210) and (211), respectively, that were considered as the most appropriate peaks for XRD analysis [41]. The result confirmed that the prepared BCPs of Ca/P ratios of 1.517, 1.568, and 1.619 correspond to the HA/β-TCP ratio of 20.1/79.9 (BCP20), 49.8/50.2 (BCP50), and 80.3/19.7 (BCP80), respectively. These scaffolds consisted of crystalline phases as revealed by the narrow and sharp diffraction peaks. The scaffold macropores of 300 μm size were fabricated using PE particles as pore-former agents as shown in Fig. 3a–c. The size of macropores greatly affects the supply of nutrition, migration, cell attachment, and tissue in-growth as reported previously [16]. For dentin tissue regeneration, odontoblasts that are polarized cells align on the surface of existing dentin or matrix for dentin formation. It has been found that the porous scaffold provided appropriate environment for cell fate determination and induction of odontogenic differentiation as compared to non-porous scaffold. It was reported that a pore size of 300 μm was adequate for dental pulp-derived cells to align on the surface and to regenerate dentin-like tissue [31,32]. The prepared scaffolds exhibited an interconnected porous structure with open pores. The interconnectivity of the macropores of the biomaterial is important for the tissue in-growth into the material [14].
The microstructures of prepared BCP20, BCP50, and BCP80 scaffolds were observed by FESEM as shown in Fig. 3d–f. There were grains of two different structures that attributed to the β-TCP and HA phases. The grain size of β-TCP is usually larger than that of HA. This is due to the fact that β-TCP phase sintered before HA phase [42]. The difference in grain size became unobvious as HA/β-TCP ratio increased. This can be attributed to the presence of more HA phase scattered in the matrix that would act like an inhibitor of β-TCP grain growth and enlargement as confirmed previously [43]. Therefore, the micropore size and subsequently the total porosity increased as HA/β-TCP ratio increased. However, there was no significant difference in total porosity between the three prepared scaffolds. As mentioned previously, the chemical composition based on the different HA/β-TCP ratios affects the bioresorption properties of BCP scaffolds [44,45]. Bioresorption is an essential factor for the biomaterials intended to repair or regenerate defective hard tissues as it controls their bioactivity upon implantation. The ionic products that resulted from inorganic material dissolution are key to understand the behavior of these materials in vitro and in vivo, in the context of tissue engineering applications. Hence, in this study, the nature of bioresorbability of the prepared BCP scaffolds was evaluated by determining the amount
230
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
Fig. 4. Effect of scaffolds of different HA to β-TCP ratios on (a) the concentration of calcium ions in the culture medium, (b) the difference in color intensity that is proportional to the concentration of calcium ions in the medium at day 3, (c) the concentration of phosphate ions in the culture medium (d) the difference in color intensity that is proportional to the concentration of phosphate ions in the medium at day 3, (e) pH values of the culture medium. Data represent means of six samples in three separate experiments and the standard errors are shown as error bars (*P b 0.05). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
of calcium and phosphate ions released in the culture medium of pH 7.5 at 37 °C. The result showed that the release of calcium and phosphate ions increased with decrease in HA/β-TCP ratio of the prepared scaffolds as shown in Fig. 4a–d. The reason is that HA phase is more stable than β-TCP in which the resorbable nature of β-TCP has further stimulation for the release of calcium and phosphate ions from the scaffold. This release increased as β-TCP phase ratio increased in the prepared BCP. This result is consistent with those described that the presence of β-TCP, a soluble phase, in the composition of BCP avoided Ca2+ depletion in the medium caused by cell uptake [46,47]. The addition of BCP had more alkaline effect on the pH of the culture medium. This alkalinity increased with decrease in HA/β-TCP ratio as observed in Fig. 4e. The variation in the pH level depended on the amount of β-TCP present in the prepared BCP scaffold. This can be attributed to the increase in the dissociation rate of BCP as the soluble phase of β-TCP ratio increased in the synthesized BCP scaffolds. This dissolution resulted in increase in PO34 − in the surrounding medium that elevated the alkalinity of the medium according to the following reaction [42,48]: h i h i 2þ 3 3− 2 Ca3 ðPO4 Þ2 → Ca þ PO4 :
ð1Þ
There was a significant difference (P b 0.05) in the alkalinity as well as in the calcium and phosphate ions released from the prepared
scaffolds for each group between day 1 and day 3. However, there was no significant difference (P N 0.05) in these values between day 3 and day 7. This can be attributed to the formation of HA, a more stable phase, in a slightly alkaline environment as a precipitate layer that separated β-TCP phase from the surrounding medium which slowed down the β-TCP dissolution according to the following reaction [42]: 1−
5Ca3 ðPO4 Þ2 þ 3OH
→3Ca5 ðPO4 Þ3 OH þ PO4
3−
:
ð2Þ
This result is in agreement with those reported that a time-dependent depletion of calcium was demonstrated in the culture medium resulting from HA precipitation over the scaffold surface [12,49]. Therefore, the extracts of BCP20, BCP50, and BCP80 scaffolds that were incubated in the medium for 3 days were chosen for the rest of the study's tests. In this research, HDPCs that have been reported to have odontoblastic differentiation potential [35] were used as a cell model for studying the mechanism of proliferation and differentiation of odontoblasts. These cells were exposed to extracts prepared according to international standards, focusing on effects caused by the eventual release of substances in biological medium, which is especially relevant if we consider the difference in the solubility ability of the prepared BCPs based on the difference in the HA/β-TCP ratios. In spite of the possibility of the washing effect of the blood in the implant area that would remove the released ions in vivo, many studies reported that the continuous scaffold degradation would liberate a flux of calcium and phosphate
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
Fig. 5. Effect of extracts obtained from different scaffold ratios of HA and β-TCP on (a) cell viability of HDPCs, (b) ALP activity of HDPCs. Data represent means of six samples in three separate experiments and the standard errors are shown as error bars (*P b 0.05).
ions into the surroundings that causes a local supersaturation of the biologic fluid with these ions in the implant area lead to more hard tissue formation in vivo [24,50,51]. MTT test is commonly used to assess the cell viability and the cytotoxicity of the tissue regeneration materials. The results confirmed that no toxic effect existed to deter normal cellular growth and proliferation
231
in porous BCP extracts, but on the contrary, BCP extracts significantly increased the cell viability as compared to control group. Previous studies also showed that BCP had good biocompatibility and exhibited excellent cell viability with different types of cells [52–56]. The result of this study also showed that there was a significant difference in cell viability of HDPCs cultured with extracts of different scaffold ratios. The cell viability of HDPCs increased with increase in HA/β-TCP ratio in which BCP80 showed the highest cell viability of HDPCs as compared to BCP50 and BCP20. This result can be attributed to the microenvironment of alkalinity as well as calcium and phosphate ions provided by BCP80 that exhibited a positive effect on cell viability and proliferation of HDPCs. A good cytocompatibility of BCP scaffold with these important cells would, therefore, be one of the prerequisites prior to the application of porous BCP scaffold in the strategic delivery of odontogenic cells for faster tissue regeneration. ALP activity is one of the most important indicators of functional odontoblasts as differentiated odontoblasts showed much higher ALP activity than dental undifferentiated mesenchymal cells [57,58]. The activity of this enzyme is a prerequisite for pulp cells to differentiate into odontoblasts. This enzyme promotes type I collagen mineralization and prevents pyrophosphate ions from its inhibition effect of mineralization. It also provides inorganic phosphate for deposition of mineral crystals [59]. In this study, ALP activity of cells cultured with BCP extracts was assessed for up to 15 days, as shown in Fig. 5b. HDPCs cultured with BCP20 extract demonstrated a significant increase in ALP activity at time periods of 3–6 days, 6–9 days, 9–12 days, and 12–15 days as compared to that cultured with BCP50 and BCP80. This result is consistent with the result of Arinzeh et al. who quantified ALP activity of human mesenchymal stem cells cultured with BCP of 20/80 and 40/60 HA/β-TCP ratio at days 21 and 28. They found that the cells cultured with BCP of 20/80 HA/β-TCP ratio showed a significantly higher ALP activity than that cultured with control group at both days 21 and 28, and that cultured with BCP of 40/60 HA/β-TCP ratio at day 28 [24]. The ability of BCP20 to increase ALP activity of the cells can be attributed to the calcium and phosphate ion rich environment that resulted from BCP20 dissociation. It was found that these ions increased ALP activity of osteoblasts [60–64], and odontoblasts [65–68]. In addition, the alkaline pH condition that resulted from the dissociation of
Fig. 6. (a) Agarose gel electrophoresis analysis of gene expression of COL1A1, BSP, DMP-1, and DSPP in HDPCs cultured with BCP20, BCP50, and BCP80 extracts using RT-PCR, (b) fold change of COL1A1, BSP, DMP-1, and DSPP gene expressions versus control normalized to GAPDH at day 14, (c) fold change of COL1A1, BSP, DMP-1, and DSPP gene expressions versus control normalized to GAPDH at day 21, (d) fold change of COL1A1, BSP, DMP-1, and DSPP gene expressions versus control normalized to GAPDH at day 28. Data are expressed as means of three samples in three separate experiments and the standard errors are shown as error bars (*P b 0.05).
232
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
BCP in the extracellular fluid has been proven to increase ALP activity in HDPCs [69]. For gene expression analysis, HDPCs cultured with BCP extracts showed high COL1A1 gene expression as compared to that cultured without BCP extract. Among study groups, there was no significant difference in COL1A1 gene expression in cells cultured with BCP20, BCP50, and BCP80. The collagen matrix is essential as it provides a spatial template upon which the mineral crystals deposit [65]. The growth of these crystals is directed by the extracellular matrix proteins. In the current study, DMP-1, BSP, and DSPP genes that play positive roles in the process of odontoblast differentiation and maturation [67] were found to be significantly up-regulated in cells cultured with BCP20 extract as compared to that cultured with BCP50 and BCP80 extracts. These results suggested that alkaline, calcium, and phosphate rich environment provided by BCP20 promoted the odontoblastic differentiation of HDPCs. This result is consistent with other researchers who found that extracellular calcium ion [65,67], inorganic phosphate ion [68], and alkaline pH condition [69] were essential in the modulation and acceleration of the differentiation of HDPCs into odontoblasts. Our null hypothesis was rejected as differences were observed in the microenvironment, cell viability, ALP activity, and gene expression of HDPCs based on the different ratios of HA and β-TCP. This study demonstrated that BCP scaffold containing higher ratio of β-TCP promoting HDPC differentiation might be in part due to the rate of degradation, the degradation products, and the surface chemistry of BCP20 in relative to the other compositions. It has been elucidated that the high rate of degradation associated with BCP20 produced a localized alkaline, calcium and phosphate rich environment that might be more favorable for odontogenesis. 5. Conclusion In this study, different HA/β-TCP ratios of BCP scaffolds' extracts were compared for differentiation of HDPCs into odontoblasts. In spite of the differences in HA/β-TCP ratios of the prepared scaffolds being only 30%, it was found that 20/80 HA/β-TCP ratio provided more calcium and phosphate ion microenvironment with elevated pH level that restricted the viability of HDPCs as compared to other compositions. However, HDPCs cultured with the extract of 20/80 HA/β-TCP ratio scaffold expressed high ALP activity and up-regulated the expression level of BSP, DMP-1, and DSPP genes. The findings of this study demonstrated an optimal composition of HA/β-TCP as a scaffold for odontoblast differentiation. Acknowledgments The authors would like to thank Universiti Sains Malaysia for funding this research through the Research University Grant No. 1001/ PPSG/813073. The authors greatly acknowledge Professor Takashi Takata (Hiroshima University, Hiroshima, Japan) for providing Human Dental Pulp Cells to conduct this research. References [1] M. Nakashima, A. Akamine, J. Endod. 31 (2005) 711–718. [2] W. Zhang, X.F. Walboomers, G.J. VanOsch, J. VanDen Dolder, J.A. Jansen, Tissue Eng. A 14 (2008) 285–294. [3] X. Yang, X.F. Walboomers, J. Van Den Dolder, Tissue Eng. A 14 (2008) 71–81. [4] M. Miura, S. Gronthos, M. Zhao, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 5807–5812. [5] S. Gronthos, M. Mankani, J. Brahim, P.G. Robey, S. Shi, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 13625–13630. [6] K.M. Hargreaves, T. Giesler, M. Henry, Y. Wang, J. Endod. 34 (2008) 51–56. [7] S. Gronthos, J. Brahim, W. Li, J. Dent. Res. 8 (2002) 531–535. [8] G. Laino, R. D'Aquino, A. Graziano, J. Bone Miner. Res. 20 (2005) 1394–1402. [9] G. Laino, A. Graziano, R. D'Aquino, J. Cell. Physiol. 206 (2006) 693–701. [10] G. Papaccio, A. Graziano, R. D'Aquino, J. Cell. Physiol. 208 (2006) 319–325. [11] S.E. Saber, J. Oral Sci. 51 (2009) 495–507. [12] M. Fabio, A. Gutemberg, F. Gustavo, K. Bruno, J.R. Alexandre, G. Jose, Artif. Organs 36 (2012) 535–542.
[13] N. Thi-Phuong, L. Byong-Taek, J. Biomater. Appl. 27 (2012) 311–321. [14] F. Bai, J. Zhang, Z. Wang, J. Lu, J. Chang, J. Liu, G. Meng, X. Dong, Biomed. Mater. 6 (2011) 015007. [15] B.H. Fellah, N. Josselin, D. Chappard, P. Weiss, P. Layrolle, J. Mater. Sci. Mater. Med. 18 (2007) 287–294. [16] T. Abdurrahim, I. Sopyan, Recent Pat. Biomed. Eng. 1 (2008) 213–229. [17] T. Thamaraiselvi, S. Rajeswari, Artif. Organs 18 (2004) 9–17. [18] P. Kasten, I. Beyen, P. Niemeyer, R. Luginbu, M. Bohner, W. Richter, Acta Biomater. 4 (2008) 1904–1915. [19] R. Yang, Y. Feng, C. Li-jia, W. Jin-jing, L. Xiao-feng, S. Yu-jun, F. Hong-song, Z. Xing-dong, B. Hong, J. Zhejiang Univ. Sci. B 12 (2011) 582–590. [20] S. Rath, A. Leonie, A. Andreas, P. Justus, M. Anne‐Kathrin, G. Peter, H. Raymund, K. Ulrich, J. Cell. Mol. Med. 16 (2012) 2350–2361. [21] G. Song, H. Pamela, B. Chongyun, H. Jing, A. Clemens, Y. Huipin, C. Wenchuan, H. Hockin, Biomaterials 34 (2013) 2167–2176. [22] G. Daculsi, O. Laboux, O. Malard, P. Weiss, J. Mater. Sci. Mater. Med. 14 (2003) 195–200. [23] B.H. Fellah, P. Weiss, O. Gauthier, T. Rouillon, P. Pilet, G. Daculsi, P. Layrolle, J. Orthop. Res. 24 (2006) 628–653. [24] T.L. Arinzeh, T. Tran, J. Mcalary, G. Daculsi, Biomaterials 26 (2005) 3631–3638. [25] G. Daculsi, Biomaterials 19 (1998) 1473–1478. [26] V. Sanginario, M.P. Ginebra, K.E. Tanner, J.A. Planell, L. Amabrosio, J. Mater. Sci. Mater. Med. 17 (2006) 447–454. [27] S.C. Wu, H.C. Hsu, S.K. Hsu, W.H. Wang, W.F. Ho, Mater. Charact. 62 (2011) 526–534. [28] M. Alcaide, M.C. Serrano, R. Pagani, S. Sanchez-Salcedo, M. Vallet-Regi, M.T. Portoles, Biomaterials 30 (2009) 45–51. [29] M. Alcaide, M.C. Serrano, R. Pagani, J. Biomed. Mater. Res. A 89 (2009) 539–549. [30] M.C. Kruyt, C.E. Wilson, J.D. de Bruijn, C.A. van Blitterswijk, C.F. Oner, A.J. Verbout, W.J. Dhert, Biomaterials 27 (2006) 5099–5106. [31] H.S. Ryu, K.S. Hong, J.K. Lee, D.J. Kim, J.H. Lee, B.S. Chang, D.H. Lee, C.K. Lee, S.S. Chung, Biomaterials 25 (2004) 393–401. [32] M.J. MacDougall, J. Amjad, Bone Dev. (2010) 183–200. [33] S.T. AbdulQader, Ab.R. Ismail, I. Hanafi, T.P. Kannan, M. Zuliani, Ceram. Int. 39 (2013) 2375–2381. [34] ISO, 10993-12: Biological evaluation of medical devices - Part 12: sample preparation and reference materials, International Organization for Standardization 2007. [35] M. Kitagawa, H. Ueda, S. Iizuka, K. Sakamoto, H. Oka, Y. Kudo, I. Ogawa, M. Miyauchi, H. Tahara, T. Takata, Arch. Oral Biol. 52 (2007) 727–731. [36] T. Mossmann, J. Immunol. Methods 65 (1983) 55–63. [37] M.K. Sutherland, J.C. Geoghegan, C. Yu, D.G. Winkler, J.A. Latham, Bone 35 (2004) 448–454. [38] M. Kitagawa, H. Tahara, S. Kitagawa, H. Oka, Y. Kudo, S. Sato, I. Ogawa, M. Miyaichi, T. Takata, Bone 39 (2006) 1035–1042. [39] L. Casagrande, F.F. Demarco, Z. Zhang, F.B. Araujo, S. Shi, J.E. Nör, J. Dent. Res. 89 (2010) 603–608. [40] D. Iejima, Y. Sumita, H. Kagami, Y. Ando, M. Ueda, Arch. Oral Biol. 52 (2007) 924–931. [41] S. Raynaud, E. Champion, D. Bernache-Assollant, J.P. Laval, J. Am. Ceram. Soc. 84 (2001) 359–366. [42] A. Krajewski, A. Ravaglioli, R. Barbucci, Integr. Biomater. Sci. 189–254 (2002). [43] M. Descamps, J. Hornez, A. Leriche, J. Eur. Ceram. Soc. 27 (2007) 2401–2406. [44] P. Ducheyne, S. Radin, L. King, J. Biomed. Mater. Res. 27 (1993) 25–34. [45] R.Z. LeGeros, S. Lin, R. Rohanizadeh, D. Mijares, J.P. LeGeros, J. Mater. Sci. Mater. Med. 14 (2003) 201–209. [46] S. Sanchez-Salcedo, I. Izquierdo-Barba, D. Arcos, M. Vallet-Regı, Tissue Eng. 12 (2006) 279–290. [47] L. Saldaña, S. Sanchez-Salcedo, I. Izquierdo-Barba, F. Bensiamar, L. Munuera, M. Vallet-Regı, N. Vilaboa, Acta Biomater. 5 (2009) 1294–1305. [48] M. Aslanidou, V. Tiverios, M. Anastasios, T. Christos, Ceram. Int. 39 (2013) 539–546. [49] D. Arcos, S. Sanchez-Salcedo, I. Izquierdo-Barba, L. Ruiz, J. Gonzalez-Calbet, M. ValletRegi, J. Biomed. Mater. Res. A 78 (2006) 762–771. [50] S.S. Jensen, M.M. Bornstein, M. Dard, D.D. Bosshardt, D. Buser, J. Biomed. Mater. Res. B 90 (2009) 171–181. [51] N.M. Farina, F.M. Guzon, M.L. Pena, A.G. Cantalapiedra, J. Mater. Sci. Mater. Med. 19 (2008) 1565–1573. [52] A.l. Bignon, J. Chouteau, J.R. Chevalier, G. Fantozzi, J.P. Carret, P. Chavassieux, G. Boivin, M. Melin, D. Hartmann, J. Mater. Sci. Mater. Med. 14 (2003) 1089–1097. [53] W. Yang, G. Yin, D. Zhou, J. Gu, Y. Li, H. Zhang, J. Mater. Sci. Technol. 26 (2010) 754–758. [54] R. Li, W. Guo, B. Yang, L. Guo, L. Sheng, G. Chen, Y. Li, Q. Zou, D. Xie, X. An, Biomaterials 32 (2011) 4525–4538. [55] T.W. Kim, Y.M. Park, D.H. Kim, H.H. Jin, K.K. Shin, J.S. Jung, H.C. Park, S.Y. Yoon, Ceram. Int. 38 (2012) 1965–1974. [56] F. Mitri, G. Alves, G. Fernandes, B. König, A. Rossi, J. Granjeiro, Artif. Organs 36 (2012) 535–542. [57] E. Verron, J. Bouler, J. Guicheux, Acta Biomater. 8 (2012) 3541–3551. [58] M. Liu, S. Yao, L. Yang, Y. Mengtong, Z. Zhihui, H. Weiping, Molecules 17 (2012) 1219–1232. [59] C. Qin, O. Baba, W. Butler, Crit. Rev. Oral Biol. Med. 15 (2004) 126–136. [60] A. Suzuki, C. Ghayor, J.R. Guicheux, D. Magne, S. Quillard, A. Kakita, Y. Ono, Y. Miura, Y. Oiso, M. Itoh, J. Bone Miner. Res. 21 (2006) 674–683. [61] Y.K. Liu, Q.Z. Lu, R. Pei, H.J. Ji, G.S. Zhou, X.L. Zhao, R.K. Tang, M. Zhang, Biomed. Mater. 4 (2009) 025004. [62] L. Zhang, N. Hanagata, M. Maeda, T. Minowa, T. Ikoma, H. Fan, X. Zhang, Sci. Technol. Adv. Mater. 10 (2009) 025003.
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233 [63] S. Khoshniat, A. Bourgine, M. Julien, M. Petit, P. Pilet, T. Rouillon, M. Masson, M. Gatius, P. Weiss, J. Guicheux, Bone 48 (2011) 894–902. [64] S. Khoshniat, A. Bourgine, M. Julien, P. Weiss, J. Guicheux, L. Beck, Cell. Mol. Life Sci. 68 (2011) 205–218. [65] F. Rashid, S. Hideki, M. Noriyoshi, M. Yoshihiro, F. Tsuyoshi, S. Hiroaki, O. Tetsuji, K. Hiroyuki, K. Hidemi, J. Endod. 29 (2003) 104–107. [66] B.L. Foster, K.A. Tompkins, R.B. Rutherford, H. Zhang, E.Y. Chu, H. Fong, M.J. Somerman, Birth Defects Res. C. Embryo Today 84 (2008) 281–314.
233
[67] H. Tada, E. Nemoto, S. Kanaya, N. Hamaji, H. Sato, H. Shimauchi, Biochem. Biophys. Res. Commun. 394 (2010) 1093–1097. [68] H. Tada, E. Nemoto, B.L. Foster, M.J. Somerman, H. Shimauchi, Bone 48 (2011) 1409–1416. [69] T. Okabe, M. Sakamoto, H. Takeuchi, K. Matsushima, J. Endod. 32 (2006) 198–201.
Contents lists available at ScienceDirect
Materials Science and Engineering C journal homepage: www.elsevier.com/locate/msec
Effect of different calcium phosphate scaffold ratios on odontogenic differentiation of human dental pulp cells Sarah Talib AbdulQader a,d, Thirumulu Ponnuraj Kannan a,b,⁎, Ismail Ab Rahman a, Hanafi Ismail c, Zuliani Mahmood a a
School of Dental Sciences, Universiti Sains Malaysia, 16150 Kubang Kerian, Kelantan, Malaysia Human Genome Centre, School of Medical Sciences, Universiti Sains Malaysia, 16150 Kubang Kerian, Kelantan, Malaysia School of Materials and Minerals Resource Engineering, Universiti Sains Malaysia, 14300 Penang, Malaysia d Department of Pedodontic and Preventive Dentistry, College of Dentistry, University of Baghdad, Baghdad, Iraq b c
a r t i c l e
i n f o
Article history: Received 12 July 2014 Received in revised form 29 October 2014 Accepted 17 December 2014 Available online 20 December 2014 Keywords: Biphasic calcium phosphate Human dental pulp cells Calcium and phosphate ions Tissue regeneration Odontogenesis
a b s t r a c t Calcium phosphate (CaP) scaffolds have been widely and successfully used with osteoblast cells for bone tissue regeneration. However, it is necessary to investigate the effects of these scaffolds on odontoblast cells' proliferation and differentiation for dentin tissue regeneration. In this study, three different hydroxyapatite (HA) to beta tricalcium phosphate (β-TCP) ratios of biphasic calcium phosphate (BCP) scaffolds, BCP20, BCP50, and BCP80, with a mean pore size of 300 μm and 65% porosity were prepared from phosphoric acid (H2PO4) and calcium carbonate (CaCO3) sintered at 1000 °C for 2 h. The extracts of these scaffolds were assessed with regard to cell viability and differentiation of odontoblasts. The high alkalinity, more calcium, and phosphate ions released that were exhibited by BCP20 decreased the viability of human dental pulp cells (HDPCs) as compared to BCP50 and BCP80. However, the cells cultured with BCP20 extract expressed high alkaline phosphatase activity and high expression level of bone sialoprotein (BSP), dental matrix protein-1 (DMP-1), and dentin sialophosphoprotein (DSPP) genes as compared to that cultured with BCP50 and BCP80 extracts. The results highlighted the effect of different scaffold ratios on the cell microenvironment and demonstrated that BCP20 scaffold can support HDPC differentiation for dentin tissue regeneration. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Tissue engineering approaches have gained great importance to restore or replace tissues that were damaged or lost. This involves the regeneration of new functional healthy tissues using progenitor cells with scaffold in response to appropriate signals [1]. In clinical dentistry, dentin tissue regeneration is a desirable goal that allows the formation of new healthy dentin to be integrated with the pre-existing dentin to overcome the drawbacks of the conventional dental treatments [2–4]. Recent advances in stem-cell biology have revealed the possibility of dentin regeneration using human adult dental pulp cells (HDPCs) as progenitor cells that have the ability to form a dentin/pulp-like complex [5,6] and possess self-renewal [7] and multi-lineage differentiation capability [8–10]. These cells can be carried by scaffolds that play an important role in providing physical support for the cells and maintaining space for the regenerated tissue. These scaffolds guide new tissue growth before degradation and are eventually replaced by new tissues [11]. ⁎ Corresponding author at: School of Dental Sciences, Universiti Sains Malaysia, 16150 Kubang Kerian, Kelantan, Malaysia. E-mail address: [email protected] (T.P. Kannan).
http://dx.doi.org/10.1016/j.msec.2014.12.070 0928-4931/© 2014 Elsevier B.V. All rights reserved.
Scaffold biological properties depend primarily on the chemical composition of the biomaterial [12,13], and its porous structure: pore shape, pore size, porosity percentage and pore interconnection pathway [14,15]. Microporosity of diameter b10 μm permits body fluid circulation and macroporosity of diameter N100 μm is necessary for migration and proliferation of cells and tissue formation [16,17]. The total porosity of 65% is efficient for total protein production and alkaline phosphatase (ALP) activity that were taken as indicators of growth/matrix production and for the comparison of cell differentiation [18]. Among many types of scaffolds that have been used for the regeneration of hard tissues, biphasic calcium phosphate (BCP) scaffolds have shown to induce appropriate osteoblastic differentiation of stem/ progenitor cells in vitro and bone formation in vivo [19–21] due to their desirable properties including similarity in composition to the bone mineral, bioactivity and osteoconductivity [22,23]. They consist of an intimate mixture of hydroxyapatite (HA) [Ca10(PO4)6(OH)2], and beta-tricalcium phosphate (β-TCP) [Ca3(PO4)2] crystals of varying phase compositions (HA/β-TCP ratios) that have been reported to be more effective than pure HA or β-TCP alone [24–26]. They exhibit different biological behaviors of their components in which HA is bioactive and β-TCP is resorbable [27]. The effects of different HA/β-TCP ratios of several modified BCP scaffolds have been well studied for bone tissue
226
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
regeneration [15,28–31]. It was reported that BCP of 20/80 HA to β-TCP ratio stimulated the osteogenic differentiation of human mesenchymal stem cells (HMSCs) in vitro and fastest rate of bone induction in vivo as compared to 100% HA, HA/β-TCP (76/24, 63/37, 56/44), and 100% β-TCP [24]. However, these effects are not well known for odontogenic cell differentiation and dentin tissue regeneration. Although odontoblasts and osteoblasts show almost identical genetic profiles, there is a clear difference between the process of odontogenesis and osteogenesis and fundamental differences in cellular morphology between bone and dentin. These differences can affect the convenience and appropriateness of the scaffold used in tissue regeneration [32]. In spite of all improvements on the solubility and bioactivity obtained with BCPs, it is important to consider the possible impact of their composition on the processes linked to cell proliferation and differentiation, and to ensure the proportional amounts of each component do not impoverish the final composite properties. Hence, it would be of interest to evaluate the effect of different HA/β-TCP ratios of BCP scaffolds on HDPC odontogenesis for dentin tissue regeneration. The aim of the present study was to determine the optimum microenvironment and the ratio of HA to β-TCP that supported the viability and the differentiation of HDPCs. The influence of BCP characteristics of different ratios of HA to β-TCP and their degradation products on the microenvironment, viability, ALP activity, and gene expression of HDPCs has been investigated. The null hypothesis was that there would be no differences in the microenvironments and in HDPCs' odontogenesis due to the different ratios of HA to β-TCP. 2. Materials and methods 2.1. Scaffold preparation and characterization Calcium phosphate powders were prepared by a wet precipitation method using CaCO3 and H2PO4 as starting materials as mentioned previously [33]. Briefly, the reagents were mixed at Ca/P ratios of 1.517, 1.568, and 1.619 corresponding to BCP of 20/80 HA to β-TCP ratios (BCP20), 50/50 HA to β-TCP ratio (BCP50), and 80/20 HA to β-TCP ratio (BCP80) respectively. The precipitate was heated at 80 °C for an hour with stirring, aged for 48 h, washed, filtered, and dried at 100 °C. The ground powder was then added to polyethylene (PE) spherical particles as pore-former agents of 300–350 μm (Cospheric, USA) at a ratio of 4:2.5 v/v for total porosity of 65% and pore size range of 300 μm. The powder mixtures were compressed with a uniaxial press of 24 MPa in a 32 mm die to form 6 mm thickness pellets. Subsequently, pellets were submitted to thermal treatment of 400 °C for 2 h and sintered at 1000 °C for 2 h. Then, the pellets were crushed and sieved to obtain granules with the desired particle size range of 0.5–1 mm as shown in Fig. 1.
The phases of HA and β-TCP and the ratios of HA/β-TCP of the scaffold were analyzed qualitatively and quantitatively by X-ray diffraction (XRD) analysis using Eva and X'Pert HighScore Plus software, respectively. The registered patterns were compared with the International Centre for Diffraction Data (ICDD) powder diffraction file (PDF) to determine the crystalline phases. Field emission scanning electron microscopy (FESEM) (Zeiss Supra 55VP, Germany) was used to evaluate the surface morphology of the scaffolds. The scaffolds' surfaces were first sputter-coated with a layer of platinum alloy (150 Å thick). The total porosity of the scaffolds was assessed using Archimedes' method. 2.2. Preparation of scaffold extract The particles thus obtained were sterilized by autoclave before use. The extract of each scaffold sample (BCP20, BCP50, and BCP80) was prepared according to international standards for medical devices evaluation ISO standard (10993-12) [34]. After sterilization, 1 g of each scaffold material (in the form of particles) was incubated in 10 ml alpha minimum essential medium (Alpha-MEM, Invitrogen, USA) for 1, 3 and 7 days at 37 °C. After incubation, the extract was collected by filtration using 0.2 μm filter. 2.3. Measurements of calcium and phosphate ions, and pH values of the extracts The content of calcium and phosphate ions in each extract at each time was quantified colorimetrically using Calcium and Phosphate Colorimetric Assay Kits (BioVision, USA) according to the manufacturer's protocols. Briefly, for Ca2+ measurement, a chromogenic complex formed between Ca2+ and 0-cresolphthalein was measured at 575 nm using an ELISA reader (Sunrise, Tecan, Austria). For phosphate ion measurement, malachite green and ammonium molybdate that form a chromogenic complex with phosphate ions was measured at 405 nm. The intensity of the color was directly proportional to the concentration of ions in the samples. The difference between amounts of ions in the media exposed or not to particles, in the absence of cells, corresponded to the release of ions from the particles. Changes in the pH were measured at pre-determined time intervals as a function of time using a pH meter (Hanna 210, Japan). 2.4. Culture of HDPCs Immortalized human dental pulp cells (HDPCs) that was kindly provided by Professor Takashi Takata (Hiroshima University, Hiroshima, Japan) were used in this research [35]. Cells were cultured in AlphaMEM supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin, and 100 mg/ml streptomycin in a humidified atmosphere of 5% CO2 and 95% relative humidity at 37 °C. Culture medium was changed every three days until confluence. Cells were trypsinized, suspended in complete culture medium of Alpha-MEM, and subcultured at a cell density of 1 × 104 cells/well in 96-well culture plates (Corning, USA) with 200 μl of culture medium. After 24 h, the medium was removed from each well and replaced by 180 μl of each scaffold extract plus 20 μl of fetal bovine serum (final concentration of 10%). The extracts of BCP20, BCP50, and BCP80 scaffolds incubated in the medium for 3 days were chosen for the rest of the study's tests. 2.5. Cell viability assay
Fig. 1. Particle size of the prepared scaffolds after sieving as visualized by image analyzer.
The viability of HDPCs exposed to the different scaffold extracts was evaluated by MTT assay. After 1, 3, 5, and 7 days of incubation of cells with different scaffold extracts, the extracts were removed and the cells were tested for viability. The 3-(4,5-dimethyl-thiazoyl)-2,5diphenyl-tetrazolium bromide (MTT) (Sigma-Aldrich, USA) assay was used as previously described [36] to determine cell viability. This assay
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
is based on the reduction of the soluble yellow MTT tetrazolium salt to a blue insoluble formazan product by mitochondrial succinic dehydrogenase. After each incubation period, 30 μl of sterile-filtered MTT solution (5 mg/ml in PBS) was added into each well and re-incubated in 5% CO2 and 95% relative humidity at 37 °C for 4 h. Then, the culture medium with the MTT solution was aspirated and the formazan crystals were dissolved in 200 μl of dimethyl sulfoxide (DMSO). To extract and solubilize the formazan, the test plate was agitated by using a microplate shaker for several minutes. After confirmation of the homogeneity of the solutions, the absorbance of the solution was measured at 570 nm using an ELISA reader (Sunrise, Tecan, Austria). Cell viability was measured as a ratio of the optical density in the extract solution medium to the optical density in the fresh medium (control groups) using the following the formula: Cell viability% ¼ ½Absorbance of treated cells=Absorbance of control cells 100%: The absorbance was the average value measured from six wells in parallel for each sample.
227
of forward primer, 3.0 μl of 0.6 μM of reverse primer, 2.0 μl of enzyme mix containing omniscript, sensiscript reverse transcriptases and HotStarTaq DNA polymerase enzymes, 25.0 μl of RNase-free water, and 5.0 μl of 100 ng of isolated RNA. HotStarTaq DNA Polymerase was activated to initiate PCR using Quick start mastercycler Pro (Eppendorf, Germany) at 95 °C for 15 min. cDNA amplification was carried out by initial denaturation at 95 °C for 30 s, annealing at 54–56 °C for 30 s, and extension at 72 °C for 60 s for 30 cycles; then, final extension at 72 °C for 10 min. The amplification reaction products were analyzed on 2% agarose gel, electrophoresed for 60 min at 75 V using Power Pac HC (Bio-Rad, USA) and visualized under ultraviolet light of Gel doc electrophoresis (Bio-Rad, USA). The genes selected in the current study were collagen 1 alpha 1 (COL1A1), bone sialoprotein (BSP), dental matrix protein-1 (DMP-1), and dentin sialophosphoprotein (DSPP), and GAPDH (glyceraldehyde3-phosphate dehydrogenase) as the reference housekeeping gene. The PCR primers of targeted genes were followed as reported earlier. The primers are listed in Table 1. 2.8. Statistical analysis
2.6. Assessment of alkaline phosphatase activity Quantitative ALP activity of HDPCs exposed to the different scaffold extracts was determined at 0–3, 3–6, 6–9, 9–12, and 12–15 days by the hydrolysis of p-nitrophenylphosphate to p-nitrophenol using an Alkaline Phosphatase Activity Colorimetric Assay Kit (BioVision, USA) according to the manufacturer's protocols. Cells were cultured at a cell density of 1 × 104 cells/well in 96-well culture plate (Corning, USA) with 200 μl of culture medium of 10% FBS until they reached 80% confluence. Cells were then exposed to different scaffold extracts with 50 μg/ml ascorbic acid (Sigma-Aldrich, USA), and 10 mmol/l β-glycerophosphate (Sigma-Aldrich, USA) for odontogenic induction and were cultured for 15 days. Culture extract medium was changed every three days. During each time of changing medium, 30 μl of supernatant was added to 50 μl of ALP assay buffer in 96 well plate. Then, 50 μl of 5 mM p-nitrophenylphosphate was added and the mixtures were incubated at 25 °C for 60 min. After incubation, 20 μl of stop solution was added to stop the enzymatic reaction and the absorbance was read at 405 nm using an ELISA reader (Sunrise, Tecan, Austria). ALP activity of the test samples was calculated using the following equation: ALP activity ðU=mlÞ ¼ A=V=T where A is the amount of pNP generated by the sample, V is the volume of the sample added in the assay well, and T is the reaction time (in minutes). 2.7. Total RNA extraction and reverse transcription-polymerase chain reaction analysis (RT-PCR) Total RNA of HDPCs cultured in 25 cm2 flask (n = 3 for each group) and exposed to BCP20, BCP50 and BCP80 extracts for 14, 21, and 28 days was extracted using an innuPREP RNA Mini Kit (analytikjena, Germany) according to the manufacturer's instructions. The concentration and purity of total RNA were determined by absorbance at 260/280 nm in a UV spectrophotometer (Biophotometer plus, Eppendorf, Germany), and the RNA quality was assessed by electrophoresis (Bio-Rad, USA) on a 2% agarose gel. The RNA was stored at − 80 °C until amplification by one-step RT-PCR. Both cDNA synthesis and amplification by RT-PCR were conducted using a QIAGEN OneStep RT-PCR Kit (QIAGEN, Germany). cDNA synthesis was carried out at 50 °C for 30 min in a final 50 μl volume of reaction mixture containing 10.0 μl of 5× buffer (12.5 mM MgCl2), 2.0 μl of deoxy nucleoside triphosphate (dNTP) (400 μM of each dNTP), 3.0 μl of 0.6 μM
Each experiment was performed in triplicate. Results are expressed as the mean and standard error. The statistical analysis of the data was performed by one-way analysis of variance followed by a multiplecomparison Tukey test, using SPSS 20.0 program. A value of P b 0.05 was considered statistically significant. 3. Results The qualitative XRD patterns for the synthesized BCP scaffolds of different ratios are shown in Fig. 2a. Two phases of BCP samples: β-TCP [ICDD 01-070-2065] and HA [ICDD 00-009-0432] only were present. The main peak was at 2θ angle of 31.04° for β-TCP and at 2θ angle of 31.86° for HA. Fig. 2b–d shows quantitative X-ray diffraction pattern of BCP scaffolds. Ca/P ratios of 1.517, 1.568, and 1.619 correspond to HA/β-TCP ratio of 20.1/79.9 (BCP20), 49.8/50.2 (BCP50), and 80.3/19.7 (BCP80), respectively. BCP20, BCP50, and BCP80 scaffolds possess uniform spheroid macropore shapes as shown in Fig. 3a–c that were fabricated using spherical particles of PE as pore-former agents. Statistical analysis of the macroporous structure of the scaffolds indicates that the mean pore size was 308 ± 12 μm. The scaffold micropores are shown in Fig. 3d–f. There is a decrease in micropore size with increase in grain growth as β-TCP ratio increased in the synthesized scaffold. The total porosity of the scaffolds as measured by Archimedes method was 64.62 ± 0.84%, 65.02 ± 0.72%, and 65.74 ± 0.52% for BCP20, BCP50, and BCP80, respectively that includes both micropore and macropore volume. There was no significant difference (P N 0.05) in the total porosity for these scaffolds of different ratios. The in vitro stability of the culture medium with and without BCP particles were quantitatively measured and are graphically represented in Fig. 4. The calcium and phosphate ions, and pH values of the extract without BCP particles were more stable throughout the experimental period. For each group, there was a significant difference (P b 0.05) in calcium and phosphate ions as well as in pH values between day 1 and day 3, while there was no significant difference (P N 0.05) in these values between day 3 and day 7. At each time point, there was a significant difference (P b 0.05) in calcium and phosphate ions as well as in pH values among study groups. Values of pH, calcium, and phosphate ions increased as HA/β-TCP ratio decreased. The extracts of different scaffold ratios incubated in the medium for 3 days were chosen for the rest of the study's tests. For cell viability assay, HDPCs were cultured with the extracts of BCP20, BCP50, and BCB80 for 1, 3, 5, and 7 days. The results showed
228
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
Table 1 List of primer sequences, melting temperature (Tm), GenBank accession no. and their respective product size. Target gene
Primer sequences
Tm (°C)
GenBank No.
Size
[References]
COL1A1
Forward: 5′-caccaatcacctgcgtacag-3′ Reverse: 5′-tggtttcttggtcggtgg-3′ Forward: 5′-gaaccacttccccacctttt-3′ Reverse: 5′-tctgaccatcatagccatcg-3′ Forward: 5′-caggagcacaggaaaaggag-3′ Reverse: 5′-ctggtggtatcttgggcact-3′ Forward: 5′-tgtcgctgttgtccaagaag-3′ Reverse: 5′-attctttggctgccattgtc-3′ Forward: 5′-cgaccactttgtcaagctca-3′ Reverse: 5′-aggggagattcagtgtggtg-3′
60. 2 60.5 60.2 59.6 60.0 60.0 60.0 60.1 60.0 60.0
NM_000088
216 bp
[37]
NM_004967
201 bp
[38]
NM_004407.3
213 bp
[39]
NM_014208.3
498 bp
[35]
BC001601
203 bp
[40]
BSP DMP-1 DSPP GAPDH
that the viability of cells increased significantly (P b 0.05) with the addition of BCP extracts as compared to control group as shown in Fig. 5a. The time-dependent result showed that there was a statistically significant increase (P b 0.05) in the cell viability of HDPCs for each study group. Day 7 showed the highest level of cell viability of HDPCs cultured with BCP20, BCP50, and BCB80 extracts as compared to day 1, day 3, and day 5. At day 7, the means of cell viability were 117.45 ± 8.29, 131.46 ± 6.89, and 145.26 ± 7.58 for HDPCs cultured with BCP20, BCP50, and BCB80 extracts, respectively. The results showed that HDPCs demonstrated statistically a significant increase (P b 0.05) in cell viability when cultured with BCP80 extract as compared to that cultured with BCP20, and BCP50 extracts. ALP activity of HDPCs cultured with BCP20 extract increased significantly (P b 0.05) as compared to that cultured with BCP50 and BCP80 at time periods of 3–6 days, 6–9 days, 9–12 days, and 12–15 days as shown in Fig. 5b. The time period of 12–15 days showed the highest level of ALP activity of HDPCs for all study groups. At time period of 12–15 days, the means of ALP activity were 174.21 ± 11.29, 137.37 ± 12.54, and 106.06 ± 9.82 for HDPCs cultured with BCP20, BCP50, and BCB80 extracts, respectively. A significant highest level of ALP activity was detected for HDPCs cultured with BCP20. For gene expression analysis (Fig. 6), HDPCs cultured with BCP extracts showed high expression level of COL1A1, BSP, DMP-1, and
DSPP genes as compared to that cultured without BCP extracts at 14, 21, and 28 days. Among study groups, there was a significant increase (P b 0.05) in the expression level of BSP, DMP-1, and DSPP genes in HDPCs cultured with BCP20 extract as compared to that cultured with BCP50 and BCP80 extracts, whereas there was no significant difference (P N 0.05) in the expression level of COL1A1 gene in HDPCs cultured with BCP20, BCP50 and BCP80 extracts. 4. Discussion The assessment of the microenvironment in the scaffold pores surrounding cells is important to study the cells' behavior for tissue regeneration. The microenvironment of the cells is affected by the degradation products of the scaffold during tissue regeneration. BCP scaffold releases ionic product during degradation process that primarily depended on its HA to β-TCP ratio. The aim of this study was to determine if the difference in the ratio of HA to β-TCP had an impact on viability and differentiation of HDPCs. In this study, all BCP scaffolds of different ratios were prepared using phosphoric acid and calcium carbonate sintered at 1000 °C for 2 h to eliminate the effect of different preparation methods on the scaffold properties. XRD analysis was carried out as shown in Fig. 2. The highest peak intensities for β-TCP and HA for all scaffolds were at 31.04° and
Fig. 2. (a) Qualitative X-ray diffraction pattern of BCP20, BCP50, and BCP80 scaffolds synthesized at pH 7.0 sintered at 1000 °C for 2 h. (b), (c), and (d) Quantitative X-ray diffraction patterns of BCP20, BCP50, and BCP80 corresponding to HA/β-TCP ratios of 20.1/79.9, 50.2/49.8 and 80.3/19.7, respectively.
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
229
Fig. 3. Field emission scanning electron microscope images (a), (b), (c) the macroporosity structure and (d), (e), (f) the microporosity structure of BCP80, BCP50, and BCP20, respectively. Blue arrow represents large grain size, red arrow represents small grain size, and yellow arrow represents micropores. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
31.86° corresponding to planes (0210) and (211), respectively, that were considered as the most appropriate peaks for XRD analysis [41]. The result confirmed that the prepared BCPs of Ca/P ratios of 1.517, 1.568, and 1.619 correspond to the HA/β-TCP ratio of 20.1/79.9 (BCP20), 49.8/50.2 (BCP50), and 80.3/19.7 (BCP80), respectively. These scaffolds consisted of crystalline phases as revealed by the narrow and sharp diffraction peaks. The scaffold macropores of 300 μm size were fabricated using PE particles as pore-former agents as shown in Fig. 3a–c. The size of macropores greatly affects the supply of nutrition, migration, cell attachment, and tissue in-growth as reported previously [16]. For dentin tissue regeneration, odontoblasts that are polarized cells align on the surface of existing dentin or matrix for dentin formation. It has been found that the porous scaffold provided appropriate environment for cell fate determination and induction of odontogenic differentiation as compared to non-porous scaffold. It was reported that a pore size of 300 μm was adequate for dental pulp-derived cells to align on the surface and to regenerate dentin-like tissue [31,32]. The prepared scaffolds exhibited an interconnected porous structure with open pores. The interconnectivity of the macropores of the biomaterial is important for the tissue in-growth into the material [14].
The microstructures of prepared BCP20, BCP50, and BCP80 scaffolds were observed by FESEM as shown in Fig. 3d–f. There were grains of two different structures that attributed to the β-TCP and HA phases. The grain size of β-TCP is usually larger than that of HA. This is due to the fact that β-TCP phase sintered before HA phase [42]. The difference in grain size became unobvious as HA/β-TCP ratio increased. This can be attributed to the presence of more HA phase scattered in the matrix that would act like an inhibitor of β-TCP grain growth and enlargement as confirmed previously [43]. Therefore, the micropore size and subsequently the total porosity increased as HA/β-TCP ratio increased. However, there was no significant difference in total porosity between the three prepared scaffolds. As mentioned previously, the chemical composition based on the different HA/β-TCP ratios affects the bioresorption properties of BCP scaffolds [44,45]. Bioresorption is an essential factor for the biomaterials intended to repair or regenerate defective hard tissues as it controls their bioactivity upon implantation. The ionic products that resulted from inorganic material dissolution are key to understand the behavior of these materials in vitro and in vivo, in the context of tissue engineering applications. Hence, in this study, the nature of bioresorbability of the prepared BCP scaffolds was evaluated by determining the amount
230
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
Fig. 4. Effect of scaffolds of different HA to β-TCP ratios on (a) the concentration of calcium ions in the culture medium, (b) the difference in color intensity that is proportional to the concentration of calcium ions in the medium at day 3, (c) the concentration of phosphate ions in the culture medium (d) the difference in color intensity that is proportional to the concentration of phosphate ions in the medium at day 3, (e) pH values of the culture medium. Data represent means of six samples in three separate experiments and the standard errors are shown as error bars (*P b 0.05). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)
of calcium and phosphate ions released in the culture medium of pH 7.5 at 37 °C. The result showed that the release of calcium and phosphate ions increased with decrease in HA/β-TCP ratio of the prepared scaffolds as shown in Fig. 4a–d. The reason is that HA phase is more stable than β-TCP in which the resorbable nature of β-TCP has further stimulation for the release of calcium and phosphate ions from the scaffold. This release increased as β-TCP phase ratio increased in the prepared BCP. This result is consistent with those described that the presence of β-TCP, a soluble phase, in the composition of BCP avoided Ca2+ depletion in the medium caused by cell uptake [46,47]. The addition of BCP had more alkaline effect on the pH of the culture medium. This alkalinity increased with decrease in HA/β-TCP ratio as observed in Fig. 4e. The variation in the pH level depended on the amount of β-TCP present in the prepared BCP scaffold. This can be attributed to the increase in the dissociation rate of BCP as the soluble phase of β-TCP ratio increased in the synthesized BCP scaffolds. This dissolution resulted in increase in PO34 − in the surrounding medium that elevated the alkalinity of the medium according to the following reaction [42,48]: h i h i 2þ 3 3− 2 Ca3 ðPO4 Þ2 → Ca þ PO4 :
ð1Þ
There was a significant difference (P b 0.05) in the alkalinity as well as in the calcium and phosphate ions released from the prepared
scaffolds for each group between day 1 and day 3. However, there was no significant difference (P N 0.05) in these values between day 3 and day 7. This can be attributed to the formation of HA, a more stable phase, in a slightly alkaline environment as a precipitate layer that separated β-TCP phase from the surrounding medium which slowed down the β-TCP dissolution according to the following reaction [42]: 1−
5Ca3 ðPO4 Þ2 þ 3OH
→3Ca5 ðPO4 Þ3 OH þ PO4
3−
:
ð2Þ
This result is in agreement with those reported that a time-dependent depletion of calcium was demonstrated in the culture medium resulting from HA precipitation over the scaffold surface [12,49]. Therefore, the extracts of BCP20, BCP50, and BCP80 scaffolds that were incubated in the medium for 3 days were chosen for the rest of the study's tests. In this research, HDPCs that have been reported to have odontoblastic differentiation potential [35] were used as a cell model for studying the mechanism of proliferation and differentiation of odontoblasts. These cells were exposed to extracts prepared according to international standards, focusing on effects caused by the eventual release of substances in biological medium, which is especially relevant if we consider the difference in the solubility ability of the prepared BCPs based on the difference in the HA/β-TCP ratios. In spite of the possibility of the washing effect of the blood in the implant area that would remove the released ions in vivo, many studies reported that the continuous scaffold degradation would liberate a flux of calcium and phosphate
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
Fig. 5. Effect of extracts obtained from different scaffold ratios of HA and β-TCP on (a) cell viability of HDPCs, (b) ALP activity of HDPCs. Data represent means of six samples in three separate experiments and the standard errors are shown as error bars (*P b 0.05).
ions into the surroundings that causes a local supersaturation of the biologic fluid with these ions in the implant area lead to more hard tissue formation in vivo [24,50,51]. MTT test is commonly used to assess the cell viability and the cytotoxicity of the tissue regeneration materials. The results confirmed that no toxic effect existed to deter normal cellular growth and proliferation
231
in porous BCP extracts, but on the contrary, BCP extracts significantly increased the cell viability as compared to control group. Previous studies also showed that BCP had good biocompatibility and exhibited excellent cell viability with different types of cells [52–56]. The result of this study also showed that there was a significant difference in cell viability of HDPCs cultured with extracts of different scaffold ratios. The cell viability of HDPCs increased with increase in HA/β-TCP ratio in which BCP80 showed the highest cell viability of HDPCs as compared to BCP50 and BCP20. This result can be attributed to the microenvironment of alkalinity as well as calcium and phosphate ions provided by BCP80 that exhibited a positive effect on cell viability and proliferation of HDPCs. A good cytocompatibility of BCP scaffold with these important cells would, therefore, be one of the prerequisites prior to the application of porous BCP scaffold in the strategic delivery of odontogenic cells for faster tissue regeneration. ALP activity is one of the most important indicators of functional odontoblasts as differentiated odontoblasts showed much higher ALP activity than dental undifferentiated mesenchymal cells [57,58]. The activity of this enzyme is a prerequisite for pulp cells to differentiate into odontoblasts. This enzyme promotes type I collagen mineralization and prevents pyrophosphate ions from its inhibition effect of mineralization. It also provides inorganic phosphate for deposition of mineral crystals [59]. In this study, ALP activity of cells cultured with BCP extracts was assessed for up to 15 days, as shown in Fig. 5b. HDPCs cultured with BCP20 extract demonstrated a significant increase in ALP activity at time periods of 3–6 days, 6–9 days, 9–12 days, and 12–15 days as compared to that cultured with BCP50 and BCP80. This result is consistent with the result of Arinzeh et al. who quantified ALP activity of human mesenchymal stem cells cultured with BCP of 20/80 and 40/60 HA/β-TCP ratio at days 21 and 28. They found that the cells cultured with BCP of 20/80 HA/β-TCP ratio showed a significantly higher ALP activity than that cultured with control group at both days 21 and 28, and that cultured with BCP of 40/60 HA/β-TCP ratio at day 28 [24]. The ability of BCP20 to increase ALP activity of the cells can be attributed to the calcium and phosphate ion rich environment that resulted from BCP20 dissociation. It was found that these ions increased ALP activity of osteoblasts [60–64], and odontoblasts [65–68]. In addition, the alkaline pH condition that resulted from the dissociation of
Fig. 6. (a) Agarose gel electrophoresis analysis of gene expression of COL1A1, BSP, DMP-1, and DSPP in HDPCs cultured with BCP20, BCP50, and BCP80 extracts using RT-PCR, (b) fold change of COL1A1, BSP, DMP-1, and DSPP gene expressions versus control normalized to GAPDH at day 14, (c) fold change of COL1A1, BSP, DMP-1, and DSPP gene expressions versus control normalized to GAPDH at day 21, (d) fold change of COL1A1, BSP, DMP-1, and DSPP gene expressions versus control normalized to GAPDH at day 28. Data are expressed as means of three samples in three separate experiments and the standard errors are shown as error bars (*P b 0.05).
232
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233
BCP in the extracellular fluid has been proven to increase ALP activity in HDPCs [69]. For gene expression analysis, HDPCs cultured with BCP extracts showed high COL1A1 gene expression as compared to that cultured without BCP extract. Among study groups, there was no significant difference in COL1A1 gene expression in cells cultured with BCP20, BCP50, and BCP80. The collagen matrix is essential as it provides a spatial template upon which the mineral crystals deposit [65]. The growth of these crystals is directed by the extracellular matrix proteins. In the current study, DMP-1, BSP, and DSPP genes that play positive roles in the process of odontoblast differentiation and maturation [67] were found to be significantly up-regulated in cells cultured with BCP20 extract as compared to that cultured with BCP50 and BCP80 extracts. These results suggested that alkaline, calcium, and phosphate rich environment provided by BCP20 promoted the odontoblastic differentiation of HDPCs. This result is consistent with other researchers who found that extracellular calcium ion [65,67], inorganic phosphate ion [68], and alkaline pH condition [69] were essential in the modulation and acceleration of the differentiation of HDPCs into odontoblasts. Our null hypothesis was rejected as differences were observed in the microenvironment, cell viability, ALP activity, and gene expression of HDPCs based on the different ratios of HA and β-TCP. This study demonstrated that BCP scaffold containing higher ratio of β-TCP promoting HDPC differentiation might be in part due to the rate of degradation, the degradation products, and the surface chemistry of BCP20 in relative to the other compositions. It has been elucidated that the high rate of degradation associated with BCP20 produced a localized alkaline, calcium and phosphate rich environment that might be more favorable for odontogenesis. 5. Conclusion In this study, different HA/β-TCP ratios of BCP scaffolds' extracts were compared for differentiation of HDPCs into odontoblasts. In spite of the differences in HA/β-TCP ratios of the prepared scaffolds being only 30%, it was found that 20/80 HA/β-TCP ratio provided more calcium and phosphate ion microenvironment with elevated pH level that restricted the viability of HDPCs as compared to other compositions. However, HDPCs cultured with the extract of 20/80 HA/β-TCP ratio scaffold expressed high ALP activity and up-regulated the expression level of BSP, DMP-1, and DSPP genes. The findings of this study demonstrated an optimal composition of HA/β-TCP as a scaffold for odontoblast differentiation. Acknowledgments The authors would like to thank Universiti Sains Malaysia for funding this research through the Research University Grant No. 1001/ PPSG/813073. The authors greatly acknowledge Professor Takashi Takata (Hiroshima University, Hiroshima, Japan) for providing Human Dental Pulp Cells to conduct this research. References [1] M. Nakashima, A. Akamine, J. Endod. 31 (2005) 711–718. [2] W. Zhang, X.F. Walboomers, G.J. VanOsch, J. VanDen Dolder, J.A. Jansen, Tissue Eng. A 14 (2008) 285–294. [3] X. Yang, X.F. Walboomers, J. Van Den Dolder, Tissue Eng. A 14 (2008) 71–81. [4] M. Miura, S. Gronthos, M. Zhao, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 5807–5812. [5] S. Gronthos, M. Mankani, J. Brahim, P.G. Robey, S. Shi, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 13625–13630. [6] K.M. Hargreaves, T. Giesler, M. Henry, Y. Wang, J. Endod. 34 (2008) 51–56. [7] S. Gronthos, J. Brahim, W. Li, J. Dent. Res. 8 (2002) 531–535. [8] G. Laino, R. D'Aquino, A. Graziano, J. Bone Miner. Res. 20 (2005) 1394–1402. [9] G. Laino, A. Graziano, R. D'Aquino, J. Cell. Physiol. 206 (2006) 693–701. [10] G. Papaccio, A. Graziano, R. D'Aquino, J. Cell. Physiol. 208 (2006) 319–325. [11] S.E. Saber, J. Oral Sci. 51 (2009) 495–507. [12] M. Fabio, A. Gutemberg, F. Gustavo, K. Bruno, J.R. Alexandre, G. Jose, Artif. Organs 36 (2012) 535–542.
[13] N. Thi-Phuong, L. Byong-Taek, J. Biomater. Appl. 27 (2012) 311–321. [14] F. Bai, J. Zhang, Z. Wang, J. Lu, J. Chang, J. Liu, G. Meng, X. Dong, Biomed. Mater. 6 (2011) 015007. [15] B.H. Fellah, N. Josselin, D. Chappard, P. Weiss, P. Layrolle, J. Mater. Sci. Mater. Med. 18 (2007) 287–294. [16] T. Abdurrahim, I. Sopyan, Recent Pat. Biomed. Eng. 1 (2008) 213–229. [17] T. Thamaraiselvi, S. Rajeswari, Artif. Organs 18 (2004) 9–17. [18] P. Kasten, I. Beyen, P. Niemeyer, R. Luginbu, M. Bohner, W. Richter, Acta Biomater. 4 (2008) 1904–1915. [19] R. Yang, Y. Feng, C. Li-jia, W. Jin-jing, L. Xiao-feng, S. Yu-jun, F. Hong-song, Z. Xing-dong, B. Hong, J. Zhejiang Univ. Sci. B 12 (2011) 582–590. [20] S. Rath, A. Leonie, A. Andreas, P. Justus, M. Anne‐Kathrin, G. Peter, H. Raymund, K. Ulrich, J. Cell. Mol. Med. 16 (2012) 2350–2361. [21] G. Song, H. Pamela, B. Chongyun, H. Jing, A. Clemens, Y. Huipin, C. Wenchuan, H. Hockin, Biomaterials 34 (2013) 2167–2176. [22] G. Daculsi, O. Laboux, O. Malard, P. Weiss, J. Mater. Sci. Mater. Med. 14 (2003) 195–200. [23] B.H. Fellah, P. Weiss, O. Gauthier, T. Rouillon, P. Pilet, G. Daculsi, P. Layrolle, J. Orthop. Res. 24 (2006) 628–653. [24] T.L. Arinzeh, T. Tran, J. Mcalary, G. Daculsi, Biomaterials 26 (2005) 3631–3638. [25] G. Daculsi, Biomaterials 19 (1998) 1473–1478. [26] V. Sanginario, M.P. Ginebra, K.E. Tanner, J.A. Planell, L. Amabrosio, J. Mater. Sci. Mater. Med. 17 (2006) 447–454. [27] S.C. Wu, H.C. Hsu, S.K. Hsu, W.H. Wang, W.F. Ho, Mater. Charact. 62 (2011) 526–534. [28] M. Alcaide, M.C. Serrano, R. Pagani, S. Sanchez-Salcedo, M. Vallet-Regi, M.T. Portoles, Biomaterials 30 (2009) 45–51. [29] M. Alcaide, M.C. Serrano, R. Pagani, J. Biomed. Mater. Res. A 89 (2009) 539–549. [30] M.C. Kruyt, C.E. Wilson, J.D. de Bruijn, C.A. van Blitterswijk, C.F. Oner, A.J. Verbout, W.J. Dhert, Biomaterials 27 (2006) 5099–5106. [31] H.S. Ryu, K.S. Hong, J.K. Lee, D.J. Kim, J.H. Lee, B.S. Chang, D.H. Lee, C.K. Lee, S.S. Chung, Biomaterials 25 (2004) 393–401. [32] M.J. MacDougall, J. Amjad, Bone Dev. (2010) 183–200. [33] S.T. AbdulQader, Ab.R. Ismail, I. Hanafi, T.P. Kannan, M. Zuliani, Ceram. Int. 39 (2013) 2375–2381. [34] ISO, 10993-12: Biological evaluation of medical devices - Part 12: sample preparation and reference materials, International Organization for Standardization 2007. [35] M. Kitagawa, H. Ueda, S. Iizuka, K. Sakamoto, H. Oka, Y. Kudo, I. Ogawa, M. Miyauchi, H. Tahara, T. Takata, Arch. Oral Biol. 52 (2007) 727–731. [36] T. Mossmann, J. Immunol. Methods 65 (1983) 55–63. [37] M.K. Sutherland, J.C. Geoghegan, C. Yu, D.G. Winkler, J.A. Latham, Bone 35 (2004) 448–454. [38] M. Kitagawa, H. Tahara, S. Kitagawa, H. Oka, Y. Kudo, S. Sato, I. Ogawa, M. Miyaichi, T. Takata, Bone 39 (2006) 1035–1042. [39] L. Casagrande, F.F. Demarco, Z. Zhang, F.B. Araujo, S. Shi, J.E. Nör, J. Dent. Res. 89 (2010) 603–608. [40] D. Iejima, Y. Sumita, H. Kagami, Y. Ando, M. Ueda, Arch. Oral Biol. 52 (2007) 924–931. [41] S. Raynaud, E. Champion, D. Bernache-Assollant, J.P. Laval, J. Am. Ceram. Soc. 84 (2001) 359–366. [42] A. Krajewski, A. Ravaglioli, R. Barbucci, Integr. Biomater. Sci. 189–254 (2002). [43] M. Descamps, J. Hornez, A. Leriche, J. Eur. Ceram. Soc. 27 (2007) 2401–2406. [44] P. Ducheyne, S. Radin, L. King, J. Biomed. Mater. Res. 27 (1993) 25–34. [45] R.Z. LeGeros, S. Lin, R. Rohanizadeh, D. Mijares, J.P. LeGeros, J. Mater. Sci. Mater. Med. 14 (2003) 201–209. [46] S. Sanchez-Salcedo, I. Izquierdo-Barba, D. Arcos, M. Vallet-Regı, Tissue Eng. 12 (2006) 279–290. [47] L. Saldaña, S. Sanchez-Salcedo, I. Izquierdo-Barba, F. Bensiamar, L. Munuera, M. Vallet-Regı, N. Vilaboa, Acta Biomater. 5 (2009) 1294–1305. [48] M. Aslanidou, V. Tiverios, M. Anastasios, T. Christos, Ceram. Int. 39 (2013) 539–546. [49] D. Arcos, S. Sanchez-Salcedo, I. Izquierdo-Barba, L. Ruiz, J. Gonzalez-Calbet, M. ValletRegi, J. Biomed. Mater. Res. A 78 (2006) 762–771. [50] S.S. Jensen, M.M. Bornstein, M. Dard, D.D. Bosshardt, D. Buser, J. Biomed. Mater. Res. B 90 (2009) 171–181. [51] N.M. Farina, F.M. Guzon, M.L. Pena, A.G. Cantalapiedra, J. Mater. Sci. Mater. Med. 19 (2008) 1565–1573. [52] A.l. Bignon, J. Chouteau, J.R. Chevalier, G. Fantozzi, J.P. Carret, P. Chavassieux, G. Boivin, M. Melin, D. Hartmann, J. Mater. Sci. Mater. Med. 14 (2003) 1089–1097. [53] W. Yang, G. Yin, D. Zhou, J. Gu, Y. Li, H. Zhang, J. Mater. Sci. Technol. 26 (2010) 754–758. [54] R. Li, W. Guo, B. Yang, L. Guo, L. Sheng, G. Chen, Y. Li, Q. Zou, D. Xie, X. An, Biomaterials 32 (2011) 4525–4538. [55] T.W. Kim, Y.M. Park, D.H. Kim, H.H. Jin, K.K. Shin, J.S. Jung, H.C. Park, S.Y. Yoon, Ceram. Int. 38 (2012) 1965–1974. [56] F. Mitri, G. Alves, G. Fernandes, B. König, A. Rossi, J. Granjeiro, Artif. Organs 36 (2012) 535–542. [57] E. Verron, J. Bouler, J. Guicheux, Acta Biomater. 8 (2012) 3541–3551. [58] M. Liu, S. Yao, L. Yang, Y. Mengtong, Z. Zhihui, H. Weiping, Molecules 17 (2012) 1219–1232. [59] C. Qin, O. Baba, W. Butler, Crit. Rev. Oral Biol. Med. 15 (2004) 126–136. [60] A. Suzuki, C. Ghayor, J.R. Guicheux, D. Magne, S. Quillard, A. Kakita, Y. Ono, Y. Miura, Y. Oiso, M. Itoh, J. Bone Miner. Res. 21 (2006) 674–683. [61] Y.K. Liu, Q.Z. Lu, R. Pei, H.J. Ji, G.S. Zhou, X.L. Zhao, R.K. Tang, M. Zhang, Biomed. Mater. 4 (2009) 025004. [62] L. Zhang, N. Hanagata, M. Maeda, T. Minowa, T. Ikoma, H. Fan, X. Zhang, Sci. Technol. Adv. Mater. 10 (2009) 025003.
S.T. AbdulQader et al. / Materials Science and Engineering C 49 (2015) 225–233 [63] S. Khoshniat, A. Bourgine, M. Julien, M. Petit, P. Pilet, T. Rouillon, M. Masson, M. Gatius, P. Weiss, J. Guicheux, Bone 48 (2011) 894–902. [64] S. Khoshniat, A. Bourgine, M. Julien, P. Weiss, J. Guicheux, L. Beck, Cell. Mol. Life Sci. 68 (2011) 205–218. [65] F. Rashid, S. Hideki, M. Noriyoshi, M. Yoshihiro, F. Tsuyoshi, S. Hiroaki, O. Tetsuji, K. Hiroyuki, K. Hidemi, J. Endod. 29 (2003) 104–107. [66] B.L. Foster, K.A. Tompkins, R.B. Rutherford, H. Zhang, E.Y. Chu, H. Fong, M.J. Somerman, Birth Defects Res. C. Embryo Today 84 (2008) 281–314.
233
[67] H. Tada, E. Nemoto, S. Kanaya, N. Hamaji, H. Sato, H. Shimauchi, Biochem. Biophys. Res. Commun. 394 (2010) 1093–1097. [68] H. Tada, E. Nemoto, B.L. Foster, M.J. Somerman, H. Shimauchi, Bone 48 (2011) 1409–1416. [69] T. Okabe, M. Sakamoto, H. Takeuchi, K. Matsushima, J. Endod. 32 (2006) 198–201.