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. 2024 Aug 26;103(10):1008–1016. doi: 10.1177/00220345241264792

Engineered 3D Periodontal Ligament Model with Magnetic Tensile Loading

P Mulimani 1,2,3, NA Mazzawi 1,2,3,4, AJ Goldstein 2,3,5, AM Obenaus 2,3,6, SM Baggett 7, D Truong 2,3,8, TE Popowics 1,3,✉, NJ Sniadecki 2,3,5,6,9,✉
PMCID: PMC11465412  PMID: 39185630

Abstract

In vitro models are invaluable tools for deconstructing the biological complexity of the periodontal ligament (PDL). Model systems that closely reproduce the 3-dimensional (3D) configuration of cell–cell and cell–matrix interactions in native tissue can deliver physiologically relevant insights. However, 3D models of the PDL that incorporate mechanical loading are currently lacking. Hence, we developed a model where periodontal tissue constructs (PTCs) are made by casting PDL cells in a collagen gel suspended between a pair of slender, silicone posts for magnetic tensile loading. Specifically, one of the posts was rigid and the other was flexible with a magnet embedded in its tip so that PTCs could be subjected to tensile loading with an external magnet. Additionally, the deflection of the flexible post could be used to measure the contractile force of PDL cells in the PTCs. Prior to tensile loading, second harmonics generation analysis of collagen fibers in PTCs revealed that incorporation of PDL cells resulted in collagen remodeling. Biomechanical testing of PTCs by tensile loading revealed an elastic response at 4 h, permanent deformation by 1 d, and creep elongation by 1 wk. Subsequently, contractile forces of PDL cells were substantially lower for PTCs under tensile loading. Immunofluorescence analysis revealed that tensile loading caused PDL cells to increase in number, express higher levels of F-actin and α–smooth muscle actin, and become aligned to the tensile axis. Second harmonics generation analysis indicated that collagen fibers in PTCs progressively remodeled over time with tensile loading. Gene expression analysis also confirmed tension-mediated upregulation of the F-actin/Rho pathway and osteogenic genes. Our model is novel in demonstrating the mechanobiological behavior that results in cell-mediated remodeling of the PDL tissue in a 3D context. Hence, it can be a valuable tool to develop therapeutics for periodontitis, periodontal regeneration, and orthodontics.

Keywords: in vitro techniques, PDL, bioengineering, regeneration, mechanical stress, collagen

Introduction

The periodontal ligament (PDL) modulates its biomechanical properties and cellular functions in response to mechanical loads to ensure homeostasis, repair, or remodeling. Insights into its mechanobiology are needed for developing new therapeutic interventions for periodontitis, periodontal regeneration, and orthodontic tooth movement. As the PDL is a soft and extremely narrow band of tissue (i.e., 150 to 380 microns thick), studying the mechanobiology of the PDL in situ is difficult (Nanci 2012). As such, periodontal research has relied heavily on in vivo animal, in vitro, and in silico models (Meikle 2005). In particular, in vitro models can simplify biological complexity and thereby facilitate reductionist approaches. Mechanical in vitro models are indispensable to elucidate signaling pathways and molecular mechanisms that drive tissue behavior and cellular activity in response to mechanical loads (Yang et al. 2015; Aveic et al. 2021).

Two-dimensional (2D) models that deform monolayers of PDL cells have been widely used (Yang et al. 2015). However, findings obtained from 2D models can significantly differ from those in 3-dimensional (3D) models. Cell–cell and cell–extracellular matrix (ECM) connections present in 3D models are thought to be necessary for effective mechanotransduction (Kang et al. 2013). Similar to 3D ex vivo models, 3D in vitro models can enable studies of PDL biomechanical parameters such as stiffness, viscoelasticity, and strain (Fill et al. 2011). Tissue biomechanics can dictate cell structure, which in turn determines cell fate and function (Ruiz and Chen 2008; Yamamoto et al. 2018). Thus, 3D models are valuable for identifying therapeutic targets to regulate PDL cell differentiation and tissue remodeling.

Innovations in the fields of tissue engineering and biomaterials have been used to develop 3D models of the PDL (Aveic et al. 2021; Huang, Sanaei, et al. 2023). However, successful clinical translation requires 3D models that recapitulate physiological conditions. In particular, many of the current 3D models facilitate compressive loading but lack tensile loading (Yang et al. 2015). To address these problems, we conducted our research with 2 main objectives: first, to engineer a 3D model that recapitulates key elements of native periodontal architecture by embedding PDL cells in a collagen matrix hydrogel (i.e., periodontal tissue constructs [PTCs]) and, second, to design and test a magnet-based tensile stretching system on the model. Tensile stretching (interchangeably referred to as “tensile loading”), as during orthodontic force application, has been associated with higher cell numbers, collagen remodeling, elevated cellular F-actin and α–smooth muscle actin levels (αSMA), and osteogenic gene expression changes (Mabuchi et al. 2002; Martino et al. 2018; Kook et al. 2011; Huang et al. 2016; Yamamoto et al. 2018; Sun et al. 2021; Janjić et al. 2023). By presenting data on these parameters, our article demonstrates the feasibility of the PTC platform and validates its use in future studies as an in vitro 3D PDL model.

Materials and Methods

Isolating PDL Cells

PDL cells were harvested from the roots of a healthy donor’s premolars extracted for orthodontic purposes, using preestablished protocols (Somerman et al. 1988) (see Appendix).

Casting Tissues

PTCs were cast between pairs of magnetic posts by modifying a protocol for engineered heart tissues (Bremner et al. 2022). Specifically, an array of 6 pairs of posts—one post of the pair being rigid and the other flexible—was fabricated in polydimethylsiloxane (PDMS) (step i in Fig. 1A). Cubic neodymium magnets (1 mm3) were embedded in the tips of the flexible posts in each pair. Casting wells were made from PDMS in a 24-well plate. Each PTC was cast from a 100 μL cell-collagen mixture composed of 1:4 ratio of PDL cell solution (20 μL culture media containing 300,000 PDL cells) and collagen mixture [80 μL of 9 parts of collagen (4 mg/mL; Advanced Biomatrix) mixed with 1 part of neutralization solution (Advanced Biomatrix)] that was dispensed in each PDMS well (step ii in Fig. 1A). The post arrays were then inverted and placed into this mixture and transferred to an incubator at 37°C with 5% CO2 to solidify. After 48 h, the PTCs were transferred to a new plate with culture media (step iii in Fig. 1A) (see Appendix).

Figure 1.

Figure 1.

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Periodontal tissue constructs. (A) Casting procedure. (B) Periodontal tissue constructs (PTCs) in culture media in a 24-well plate. (C) PTCs on posts. (D) PTC observed under 1× brightfield microscope. Visible in the top post is the square cross section of an embedded magnet for tensile loading and in the bottom post is the round cross section of a glass capillary for rigid reinforcement. (E) PTC observed under 4× brightfield microscope.

Viability Assessment

Viability of PTCs was assessed using Nucleocounter-NC 200 machine (ChemoMetec) (see Appendix).

Calculation of PTC Contractile Force

Based on a modulus of elasticity of 2.5 MPa for PDMS, the bending stiffness (Kpost) of the flexible posts was calculated to be 0.95 µN/µm, as done previously (Bielawski et al. 2016). Contractile force was calculated by multiplying the bending stiffness by the deflection of the post (Δpost) (Fig. 2B).

Figure 2.

Figure 2.

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Cell-mediated contraction and extracellular matrix remodeling of periodontal tissue constructs (PTCs). (A) Contraction of PTCs is observed at 3 d after casting. (B) Calculation of contractile force. (C) Length, (D) force, and (E) viability of PTCs for (F) collagen constructs without cells (– PTC cells) and (G) PTC (+ PTC cells). Representative 2-photon images of PTC showing fibrillar collagen (green) in (G) constructs without cells and (I) PTCs. Insets show representative images of collagen fibers detected by Curve Align 5.0 beta in CT-FIRE fiber mode. Analysis of (J) fiber thickness, (K) length, (L) count, and (M) alignment in constructs without cells (n = 6) and PTCs (n = 9). For alignment, unity indicates all fibers are aligned in 1 direction while zero indicates a lack of alignment. Bar graphs and error bars depict mean ± SEM. Statistical significance using 2-tailed unpaired t test in panels C, D, and J–M or analysis of variance in panel E indicated by *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

Tensile Loading of PTCs

A tensile loading apparatus that held a bar magnet and a 24-well plate was 3D printed (Fig. 3A). For tensile loading, an array of PTCs was placed with flexible posts facing toward the bar magnet (Fig. 3B). At a separation of 1 cm, the attractive force between the bar magnet and embedded cube magnets in the flexible posts caused them to deflect. Prior to casting, post arrays were placed in the apparatus and the deflection of the posts was measured to be 430 ± 21 µm, which equates to 408.5 µN of force according to the bending stiffness. PTCs were loaded in the tensile apparatus 5 d after casting and subjected to tensile stretching for 4 h, 24 h, or 1 wk. Images of the stretched PTCs and unstretched controls were taken with brightfield microscopy (Nikon Ti-E) using a 2× objective and then processed using FIJI image analysis software to calculate PTC length. Based on PTC length, contractile force produced after tensile loading was calculated as mentioned in the section above.

Figure 3.

Figure 3.

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Tensile stretching of periodontal tissue constructs (PTCs). (A) Schematic of tensile stretching apparatus and setup. (B) Schematic of tensile stretching of PTCs as viewed from side. (C) Experiment timeline. Length changes of PTCs before, during, and after applying tensile stretch at (D) 4 h (n = 5), (E) 24 h (n = 6), and (F) 1 wk (n = 6). (G) Longitudinal change in PTC length with tensile stretching (for each time point: control, n = 27; tension, n = 24). (H) PTC contractile force in experimental (n = 41) and control groups (n = 39) before application of tensile stretch. (I) PTC contractile force after tensile stretching for 4 h (n = 30), 24 h (n = 28), and 1 wk (n = 30). Data are mean ± SEM, *P < 0.05, ##, **P < 0.01, ###, ***P < 0.001, ####, ****P < 0.0001 by repeated-measures 1-way analysis of variance (ANOVA) (D–G) or ordinary 1-way ANOVA (I) with Tukey’s multiple comparisons or 2-tailed unpaired t test (G, H). (G) Purple bars with asterisks depict intergroup differences among PTCs subjected to tension; # denotes intragroup difference between stretched and control PTCs at the same time point.

Immunofluorescence Imaging

Immunofluorescent staining of PTCs was carried out to detect nuclei, F-actin, and αSMA (see Appendix).

Collagen Structural Analysis

Collagen structure in PTCs was assessed using a multiphoton microscope (Olympus FV1000) to record the second harmonics generation by collagen fibers. Thickness, length, count, and alignment of the fibers were analyzed using CurveAlign 5.0 beta in CT-FIRE fiber mode (Liu et al. 2020) (see Appendix).

Gene Expression Analysis

PTCs tensile stretched for 4 and 24 h, and age-matched unstretched controls were harvested for bulk RNA sequencing and gene expression analysis (see Appendix).

Statistical Analysis

See Appendix.

Results

PTCs Exhibit Cell-Mediated Contractility and ECM Remodeling

We fabricated PTCs by casting patient-derived PDL cells within a collagen matrix surrounding 2 silicone posts (Fig. 1A, Appendix Fig. 1). In order to mimic the in vivo PDL, our PTCs were fabricated using a heterogenous PDL cell population. We have identified gene expression markers characteristic of fibroblasts and mesenchymal stem cells, suggesting the predominance of these cell types in our sample. In contrast, low gene expression was observed for markers for epithelial cells and gingival cells, suggesting the absence of these cell types (Appendix Table 1). Within 48 h, the cell–collagen mixture condensed into cell-ladened hydrogel suspended between the 2 posts (Fig. 1B–E). Three days after casting, PTCs reduced in length to 7,223 ± 37 µm, while collagen constructs without cells were 8,000 µm long (Fig. 2C). Based on the deflection of the posts, PTCs generated a contractile force of 739 ± 37 µN, while constructs without cells produced no force (Fig. 2D). Viability of PDL cells in the PTCs remained high over 2 wk after casting (Fig. 2E). We used second harmonic generation analysis to compare the structure of collagen fibers in constructs without cells (Fig. 2F, H) or PTCs (Fig. 2G, I). Collagen fibers in PTCs were significantly thicker (Fig. 2J) and shorter in length (Fig. 2K) as compared to constructs without cells. Fiber count was similar between constructs (Fig. 2L), but PTCs had fibers that were less aligned than constructs without cells (Fig. 2M).

Tensile Loading Causes Biomechanical Creep Response

To assess tissue behavior in response to mechanical stretching, we applied tensile strain on PTCs using a magnetic apparatus (Fig. 3A, B). PTCs were magnetically stretched to generate an initial strain of 4% to 7% and held in a stretched position for 4 h, 24 h, or 1 wk (Fig. 3C). Releasing PTCs from their stretched length after 4 h resulted in partial elastic recovery and mild permanent deformation as compared to their prestretched length (Fig. 3D). Stretching for 24 h caused significant permanent deformation and little elastic recovery, as indicated by the similarity in length of PTCs to their stretched value (Fig. 3E). Similarly, stretching for 1 wk caused significant permanent deformation of PTCs such that their length exceeded their initial stretched length, indicating a creep response in the tissue (Fig. 3F).

Overall, the creep response in PTCs resulted in significant elongation at 1 wk with tensile loading as compared to 4 and 24 h (Fig. 3G). Consequently, PTCs stretched for 1 wk generated weaker contractile forces as compared to PTCs stretched for 4 or 24 h (Fig. 3I). Prior to magnetic stretching, there was not a significant difference in contractile force or tissue length between control and experimental groups (Fig. 3H, Appendix Fig. 2A). These results indicate that tensile loading of PTCs results in increased length and reduced contractile force generation as compared to controls (Fig. 3G, I, Appendix Fig. 2B–K).

Tensile Loading Triggers Mechanobiological Responses

To assess the effects of tensile stretching on the biological response of PDL cells, we performed immunofluorescent staining (Fig. 4A–C). We found that cell count (Fig. 4D), F-actin levels (Fig. 4E), and αSMA expression (Fig. 4F) were higher for PDL cells in PTCs with 1 wk of tensile loading as compared to those in unloaded controls (Appendix Fig. 3). However, F-actin and αSMA production was similar on a per-cell basis for tensile-loaded PTCs versus controls (Appendix Fig. 4). These results indicate that elevated F-actin and αSMA levels observed at 1 wk of tensile loading are mainly due to an increase in cell count, induced by mechanical loading. Additionally, directionality of the cells increased significantly with time, but by 1 wk, PTCs under tensile loading were more aligned as compared to unloaded controls (Fig. 4G, Appendix Fig. 5). Together, these results indicate there is a mechanobiological response in PDL cells due to tensile mechanical loading.

Figure 4.

Figure 4.

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Periodontal tissue construct (PTC) structural analysis and tensile loading effects. (A–C) Representative images of immunofluorescent stained PTCs 3 d after casting for nuclei (blue), α–smooth muscle actin levels (αSMA) (green), and F-actin (red). (A) Whole-mounted representative composite image of a PTC. (B, C) Networked appearance of stellate and spindle-shaped PDL cells interspersed with αSMA-positive green myofibroblast cells. (D) Quantification of immunofluorescent data on tensile stretching for 4 h, 24 h, and 1 wk for cell count. Quantification of (E) F-actin and (F) αSMA levels normalized by area after tensile stretching for 1 wk. (G) Quantification of immunofluorescent data on tensile stretching for 4 h, 24 h, and 1 wk for tissue directionality. Quantification of collagen fiber (H) thickness, (I) length, (J) count, and (K) alignment on tensile-stretching PTCs for 4 h, 24 h, and 1 wk, using Curve Align 5.0 beta in CT-FIRE fiber mode. Data are mean ± SEM, #, *P < 0.05, **P < 0.01, ***P < 0.001, ####, ****P < 0.0001 by 1-way analysis of variance with Tukey’s multiple comparisons (D, G, H–K) or 2-tailed unpaired t test (D–K). (H–K) Purple bars with asterisks depict intergroup differences among PTCs subjected to tension, and # denotes intragroup difference between stretched and control PTCs at the same time point (n = 9–12 PTCs per group from 3 independent experiments; 30–50 images per group were taken for analysis).

PDL Cells Remodel Collagen upon Tensile Loading

To identify ECM structural changes underlying PTC behavior on tensile loading, we performed second harmonic generation analysis of collagen fibers. By 1 wk, tensile stretching significantly decreased fiber thickness, increased fiber length, reduced fiber count, and increased fiber alignment as compared to stretching for either 4 or 24 h (Fig. 4H–K, Appendix Fig. 6). Tensile loading for 4 h significantly increased fiber thickness and reduced fiber length compared to controls (Fig. 4H, I), whereas loading for 24 h significantly increased fiber thickness alone compared to unloaded controls (Fig. 4H). After loading for 1 wk, PTCs had significantly longer fibers, lower fiber count, and higher fiber alignment compared to the controls (Fig. 4H, I). These results indicate significant collagen remodeling after 1 wk of tensile stretching reflected as permanent deformation and creep of the PTC longitudinal dimension.

Tensile Stretch Alters Gene Pathways for Tissue Remodeling

To identify early transcriptional changes from tensile loading, we performed bulk RNA sequencing of PTCs stretched for 4 and 24 h. Tensile loading resulted in a higher number of differentially expressed genes (DEGs) over time (Fig. 5A). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis indicated that signal transduction pathways had the most DEGs after 4 and 24 h of tensile loading (Fig. 5B). At 24 h, DEGs for cell growth and death, as well as replication and repair pathways, were among the highest. Genes related to collagen fibrillogenesis and ECM organization (COL1A1, PLOD1, PLOD2, COMP), collagen breakdown and tissue turnover (ADAMTS3, MMP1, MMP3, MMP8, TIMP3) (Fig. 5C), and cell cycle and cell death (CDK1, CDCA2, CCNB1, CENPF) (Fig. 5D) were downregulated at 4 h and upregulated at 24 h (Appendix Fig. 7). A similar response of downregulation followed by upregulation was observed for genes related to cytoskeletal organization and the Rho pathway (ACTN1, FLNA, RHOA, ARPC3, LIMK1) (Fig. 5E), αSMA production and transforming growth factor β (TGF-β) pathway (ACTA2, BMP8A, SMAD7, TGFB1) (Fig. 5F), and osteoblast differentiation and mineralized matrix production (BMP8B, IBSP, SPP1, DMP1) (Fig. 5G, Appendix Fig. 8). Overall, these results indicate a mechanobiological response and concur with previously reported gene expression changes due to tensile stretching (Meikle 2006; Wise and King 2008; Yamamoto et al. 2018; Sun et al. 2021; Janjić et al. 2023).

Figure 5.

Figure 5.

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Gene expression analysis at 4 h and 24 h after tensile stretching (for each time point, n = 1). (A) Bar chart demonstrates split-up of upregulated and downregulated genes among genes differentially expressed at 4 h and 24 h, compared to control and to each other. (B) Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis depicting biological pathways that differentially expressed genes (DEGs) belong to at 4 h and 24 h compared to controls. Heatmaps showing log2 fold change in subsets of DEGs related to (C) collagen remodeling, (D) cell cycle and division, (E) F-actin and Rho pathway, (F) α–smooth muscle actin levels (αSMA) and transforming growth factor β (TGF-β) pathway, and (G) mineralization.

Discussion

Our study has provided the proof of principle for a novel in vitro 3D model that emulates some key structural and functional features of the human PDL. We find that PDL cells in our model remodel collagen fibers in PTCs suspended between 2 silicone posts (Fig. 1C, D). This design resembles cells enmeshed within principal collagen fiber bundles spanning alveolar bone and cementum (Nanci 2012). Spindle-shaped and stellate PDL cells and collagen fibers in PTCs are organized in parallel and are aligned along the tensile axis (Appendix Fig. 5, 6) resembling in vivo observations (Beertsen et al. 1997; Hirashima et al. 2016). The thickness of collagen fibers in PTCs is within the physiological range for collagen fiber bundles in PDL tissue (i.e., 1 to 4 µm) (Berkovitz 1990; Pini et al. 2004). Additionally, PDL cells in our model were distributed uniformly throughout the construct, as opposed to other 3D models where cells accumulate at the periphery (Yang et al. 2015).

Our model demonstrates a link between tissue behavior and cellular remodeling of ECM. PDL cells caused the PTCs to contract between the pairs of posts and reduce in length (Fig. 2A–D). In contrast, there was no compaction in collagen constructs that lacked cells. This demonstrates the role played by cells in generating contractile force to compact the tissue. Further, PDL cells contributed to collagen remodeling as shown by increased fiber thickness, reduced fiber length, and reduced fiber alignment, supporting the contribution of PDL cells to ECM turnover (Fig. 2J, K, M) (Beertsen et al. 1997).

The PDL in vivo is likened to a complex, fiber-reinforced substance that responds to force in a viscoelastic, nonlinear, and time-dependent manner (Jónsdóttir et al. 2006). Tensile loads are more predominant than compressive ones in the PDL under both physiologic and orthodontic tooth movement situations yet remain understudied in a 3D in vitro model (Beertsen et al. 1997; Cattaneo et al. 2005; Yang et al. 2015). We find that under tensile loading, PTCs exhibited permanent deformation that increased over time. This indicates a creep response (Fig. 3F) presumably due to longer and thinner collagen fibers resulting from stretch-induced deformation (Fig. 4H, I) (Lanir et al. 1988; van Driel et al. 2000, Gauthier et al. 2021). Alternatively, one could attribute the creep response to enzymatic degradation as indicated by the reduction in fiber count in PTCs (Fig. 4J). Additionally, proliferating cells can contribute to increased enzymatic degradation (Green et al. 1990). Correspondingly, PTCs did have a higher cell count at 1 wk of tensile loading as compared to controls (Fig. 4D). In vivo models of tooth movement have shown increased cellularity due to a 6-fold increase in cell proliferation and upregulation of metalloproteinases in PDL tissues under tensile loading (Mabuchi et al. 2002; Kook et al. 2011).

The time-dependent PTC response to tensile load resulted in cellular changes that may be applicable in periodontal tissue regeneration. Enhanced creep, stress relaxation, and higher loss of modulus of elasticity of collagen networks promote spreading of adherent cells such as myofibroblasts, fibroblasts, and mesenchymal stem cells (Chaudhuri et al. 2020; Huang, Li, et al. 2023). In line with this observation, PTCs stretched for 1 wk displayed αSMA-positive PDL cells, which indicates differentiation into myofibroblasts (Appendix Fig. 5D, H). Previously, αSMA- positive perivascular cells and myofibroblasts in the PDL were found to be osteoblast progenitors (Roguljic et al. 2013; Feng et al. 2016). Thus, our platform can be used in future investigation of the relationship between modulating collagen fibrillogenesis and ECM biomechanical properties and differentiation of periodontal cells.

Elevated F-actin and αSMA production generally correlates with increased cellular contractility (Wise and King 2008; Yamamoto et al. 2018). However, our PTCs exhibited reduced levels of contractile force despite higher F-actin and αSMA production after tensile stretching (Fig. 3I, Fig. 4E, F, Appendix Fig. 2D). This discrepancy may be attributed to the creep response in PTC, which elongated under tension, presumably due to the remodeling of collagen, and thereby increased the distance between the silicone posts used to measure tissue-level force. It is unknown if the contractile forces of individual cells were affected by tensile loading in our study. However, F-actin and αSMA levels on a per-cell basis were similar between tensile-loaded and unloaded controls (Appendix Fig. 4), so it is plausible that cellular forces were similar.

Our limited gene expression analysis found tension to stimulate a higher number of DEGs at 24 h (Fig. 5D–H) compared to 4 h. The large number of cell cycle and cell death genes upregulated in the 24-h tension group (Fig. 5B, E) corroborates the anabolic and cell-proliferative effects of mechanical tension (Huang et al. 2016). Mechanical stretching is known to activate the Rho mechanotransduction pathway and upregulate F-actin formation, along with TGF-β pathway activation and αSMA production (Meng et al. 2010; Martino et al. 2018). We see an upregulation of all these genes in the 24-h tension group, but upregulation of F-actin and αSMA at a protein level is only seen in the 1-wk tension group, presumably because protein production trails messenger RNA synthesis. Furthermore, our finding of upregulation of osteogenic genes due to tensile stretching is consistent with other studies showing that tension drives osteogenesis (Meikle 2005; Sun et al. 2021; Janjić et al. 2023). However, further studies are needed to validate these findings with proteomic analysis.

Key limitations of our study are that the model size and scale of forces are not the same as in vivo. Our model is relatively unconstrained compared to in vivo tissue, which is enclosed circumferentially by alveolar bone and tooth root. The modulus of elasticity of our PTC model has not been measured, so its resemblance to in vivo PDL is unknown. Matching the in vitro modulus to that in vivo is also problematic because of the lack of consensus, which has been reported to range from 0.01 to 1750 MPa (Fill et al. 2011). Additionally, the gene expression analysis herein was limited only to early changes due to tensile loading. Future analyses with PTCs should examine osteogenesis and mineralization of PTCs in response to tension on a longer time scale. Finally, our study does not investigate the molecular mechanisms that underlie the changes induced by tensile loading.

To our knowledge, we have fabricated a first of its kind 3D PDL model that facilitates exploration of links between macroscale tissue behavior and microscale structural changes. PTCs may be produced at a larger scale and lower cost than animal models and could be used as an in vitro alternative in drug development, testing biological agents, elucidating molecular mechanisms, and developing periodontal grafts for transplantation. Moreover, with magnetic tensile loading, our PTC model fills a critical gap for investigating periodontal mechanobiology. A future goal would also be to develop a PTC model that would be able to apply compressive and cyclic loading along with tensile loading at various magnitudes, to recapitulate the orthodontic tooth movement processes more comprehensively. These applications have significant translational potential for periodontitis treatment, periodontal regeneration, and orthodontic treatment and retention.

Author Contributions

P. Mulimani, contributed to conception, design, data acquisition, analysis, and interpretation, drafted and critically revised the manuscript; N.A. Mazzawi, contributed to data acquisition and analysis, critically revised the manuscript; A.J. Goldstein, A.M. Obenaus, S.M. Baggett, D. Truong, contributed to conception, design, data acquisition, critically revised the manuscript; T.E. Popowics, N.J. Sniadecki, contributed to conception, design, data interpretation, critically revised the manuscript. All authors have their final approval and agree to be accountable for all aspects of work.

Supplemental Material

sj-docx-1-jdr-10.1177_00220345241264792 – Supplemental material for Engineered 3D Periodontal Ligament Model with Magnetic Tensile Loading
sj-docx-1-jdr-10.1177_00220345241264792.docx (6.9MB, docx)

Supplemental material, sj-docx-1-jdr-10.1177_00220345241264792 for Engineered 3D Periodontal Ligament Model with Magnetic Tensile Loading by P. Mulimani, N.A. Mazzawi, A.J. Goldstein, A.M. Obenaus, S.M. Baggett, D. Truong, T.E. Popowics and N.J. Sniadecki in Journal of Dental Research

Footnotes

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding: The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: We would like to gratefully acknowledge grant support for P. Mulimani from Ruth L. Kirschstein Interdisciplinary Research Training Award (R90) Grant #R90DE023059, support for N.A. Mazzawi through the Mary Gates Research Scholarship, and funding for the project from University of Washington, School of Dentistry’s Dr. Douglas L. Morell Dentistry Research Fund and National Institutes of Health/National Institute of Dental and Craniofacial Research NIH/NIDCR R03 Grant # 5R03DE0 29827-02.

A supplemental appendix to this article is available online.

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Associated Data

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Supplementary Materials

sj-docx-1-jdr-10.1177_00220345241264792 – Supplemental material for Engineered 3D Periodontal Ligament Model with Magnetic Tensile Loading
sj-docx-1-jdr-10.1177_00220345241264792.docx (6.9MB, docx)

Supplemental material, sj-docx-1-jdr-10.1177_00220345241264792 for Engineered 3D Periodontal Ligament Model with Magnetic Tensile Loading by P. Mulimani, N.A. Mazzawi, A.J. Goldstein, A.M. Obenaus, S.M. Baggett, D. Truong, T.E. Popowics and N.J. Sniadecki in Journal of Dental Research

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