Physical Micro-Environment of Periodontal Ligament: Comparison
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Please note this is a comparison between Version 1 by Beatrice Masante and Version 3 by Jessie Wu.

In native periodontal ligament (PDL) development and homeostasis, mechanical loading plays a crucial role. Thus, any strategy aiming at periodontal tissue engineering should entail understanding and replicating in vitro the specific mechanical forces that guide the formation and remodeling of the PDL.

 

  • mechanical loading
  • mechanotransduction
  • periodontal ligament regeneration
  • tissue engineering
  • bioreactor
  • 3D environment

1. Periodontal DLigament Structure and Cells

The periodontium is a complex system composed of gingiva, periodontal ligament (PDL), cementum, and alveolar bone, featuring a hierarchically compartmentalized architecture [1]. The homeostasis of this system is maintained by the PDL, a specialized connective tissue, which is located between the cementum and alveolar bone and articulates (gomphosis) the teeth to the jaws [2][3]. From a histological perspective, PDL is an aligned fibrous network with a thickness ranging between 100 and 400 μm and is characterized by an extensive blood supply and a neural network [4]. PDL is constituted by a heterogeneous population of cells (namely PDL cells) that includes periodontal ligament fibroblasts (PDLFs), which represent by far the largest population and are responsible for the deposition and maintenance of the extracellular matrix (ECM) and periodontal ligament stem cells (PDLSCs), showing both osteogenic and tendo/ligamentogenic characteristics. Collagen type I and, in lesser amounts, type III constitute cross‐banded fibrils, named Sharpey’s fibers, which provide mechanical support and are usually classified as dentinogingival, transseptal, or alveolodental (forming the bulk of proper PDL fibers) [4]. In particular, fibers oblique or perpendicular to the long axis of the tooth are thought to play pivotal roles in eliciting adaptive responses during mastication and occlusion [4]. Among all the fibers, the horizontal ones withstand the greatest loads and exhibit the greatest strain under mastication [5], (Figure 1). The collagen fibers are generally aligned according to a periodic crimped pattern [6] that prevents ligament overextension [7][8]. Sharpey’s fibers anchor mostly to acellular cementum, a mineralized layer (50–300 μm thickness) covering the tooth dentin surface. PDL cells are arranged along PDL fibers so that the long cellular axis is parallel to the main fiber bundles of the PDL [9][10]. The presence of a particular type of elastic fibers named oxtytalan, made of fimbrillins, that form a network running parallel to cementum and are thought to interact with vessels and neural fibers is also noteworthy [11]. In a healthy subject, PDL covers the tooth root almost entirely, and a tight epithelial seal within the gingival sulcus prevents microorganisms from reaching the PDL. This delicate system is compromised by the onset of periodontal disease (PD), which affects in its severe form about 10% of adults, ranking sixth among the most prevalent diseases in the world [12]. PD starts as a localized and reversible inflammation of the gingiva (gingivitis) due to dental plaque, and, when untreated, it may become chronic periodontitis, which is characterized by the progressive destruction of the tooth‐supporting tissues, i.e., cementum, PDL, and bone [13][14].

In 2004, Kawaguchi et al. proposed the autograft of bone-marrow-derived mesenchymal stem cells (BMMSCs) to enhance the healing of periodontal defects, which proved successful in a dog model [15]. This approach highlighted the potential of cellular therapy and paved the way to other pre-clinical studies dealing with BMMSCs [16], adipose derived stem cells (ASCs) [17], and PDLSCs [18]. Among all the possible sources of mesenchymal stem cells (MSCs), PDLSCs (Figure 1) may be selected for PDL regeneration owing to their commitment capacity, as they express scleraxis, i.e., a tendon/ligament-specific transcription factor, more than BMMSCs or dental pulp stem cells (DPSCs) and have the potential to form cementum and PDL-like structures [19][20]. Indeed, the preservation of the PDL is essential to achieving proper regeneration of the periodontium and avoiding the ankylosis of the tooth, i.e., direct contact between the root and the alveolar bone.
From this perspective, the role of the transforming growth factor–β1 (TGF-β1) signaling becomes interesting since its activation enables the commitment of cementocytes, while its inhibition promotes fibroblastic differentiation of the ligament progenitors [21]. PDLSCs transplanted into a periodontal lesion in a rat model generated typical PDL-like structures in vivo by forming Sharpey’s-fiber-like collagen bundles that are connected to cementum-like structures [18]. New insights into the molecular regulation of periodontal attachment have been brought by Bai S et al., who investigated the regulatory mechanism of copine 7 (CPNE7) and cementum attachment protein (CAP) in coordination with cytoskeleton arrangement [22]. Regenerating cementum is regarded as key to promoting new fibrous tissue attachment, preserving the PDL, and avoiding tooth ankylosis [23].

2. Mimicking the Physical Micro-Environment of Periodontal DLigament

During normal oral functions, intermittent occlusal contacts accompanied by pressure from the tongue occur, and the PDL periodically undergoes different combinations of mechanical loading (i.e., compression, stretch, fluid-induced shear stress) that contribute to maintain the PDL homeostasis [24]. The mechanical response of PDL to experienced loading is determined by the combination of the oriented collagen fiber bundles and the distribution of the interstitial fluid, which makes the PDL acting as a shock absorber, increasing the tooth’s ability to withstand loading via the hydrostatic effect [25][26]. This is accompanied by a mechano-biological response of the PDL that, depending on the location under the tooth, changes structure and functions and induces remodeling in the surrounding tissues [4]. Cells detect and transduce the mechanical signals from their membrane to the nucleus through a molecular process named mechanotransduction [27]. Among the most relevant cell membrane mechanosensors, integrins play a fundamental role, mediating direct contact with the extracellular matrix (ECM). As transmembrane constituents of the focal adhesions (FA), integrins interact with scaffolding, docking, and signaling proteins linked to the actin cytoskeleton [28]. The variable composition of the FA core depends on ECM and mechanical stimuli. Cells modulate their own cytoskeletal architecture in response to applied forces [29] and remain in a sort of tensional homeostasis, i.e., a basal equilibrium stress state [30]. From a molecular point of view, the role of two actin-binding proteins associating with the cytoplasmic tail of the β1 integrin—talin and filamin A (FLNa) [31]—is noteworthy. By interacting with integrins, talin enhances cell adhesion to the ECM [32]. Without mechanical stimuli, talin is fully structured, but while under increasing force regimes, talin exposes progressively more vinculin binding sites (VBS), thus activating more vinculin proteins [33]. Vinculin may mediate reorganization of cell polarity, helping the cell to adapt to increased tensile forces [34]. FLNa competes with talin for binding to β1 integrin [35], and it is thought to antagonize integrin-mediated cell adhesion [36]. For example, Shifrin et al. showed that through the Rac/Pak/p38 signaling pathway, FLNa may prevent apoptosis in PDL in response to tensile forces [37].
Due to the fundamental role played by mechanical loading in vivo, a compelling strategy for directing cell commitment in periodontal tissue engineering may be represented by the reproduction in vitro of the dynamic environment in which PDL cells operate. Several groups investigated the sensitivity of PDL cells (i.e., cells harvested from the PDL, including not only PDLSCs but also more committed cells and even fibroblasts) to mechanical loading and their involvement in periodontal and bone remodeling in vitro. In the following paragraphs, investigations performed in the last decade on in vitro mechanically stimulated PDL cells and constructs are reported per type of force (i.e., compression, stretch, shear stress; Figure 1) and depending on the adopted in vitro mechanical loading method (Figure 2), highlighting the use of conventional two-dimensional (2D) or more physiological 3D cell culture techniques.
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Figure 1. Overview of the reviewed in vitro PDL mechanical loading studies. Schematic summary of the reviewed studies investigating the influence of mechanical loading on PDL cells based on the type of applied stimulation along with main outcomes.
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Figure 2. In vitro mechanical loading methods applied for PDL investigations. Schematic representation of the in vitro mechanical loading methods adopted in the reviewed studies for exposing PDL cells cultured in 2D layers or in 3D constructs to compression, stretch, and shear stress stimuli.

2.1. Compression

2.1.1. Weight Method

This method, based on the use of cover glasses or cylinders containing metal granules, which allows the application of tunable static compressive forces to the culture, was widely adopted to investigate in vitro how continuous compression can influence PDL cells. In 2011, Li et al. established a 3D model of PDL tissue based on hPDL cells seeded on a sheet of porous poly(lactic-co-glycolic acid) (PLGA) scaffold and exposed it to static compression (5–35 g/cm2 for 6–72 h), observing a significant induction of osteoclastogenic genes that did not occur when human gingival fibroblasts were used [38]. Moreover, a predominant upregulation of osteoclastogenesis inducers was observed at the early stage (6 h), while osteoclastogenesis inhibitor genes increased at the late stage (24–72 h), although cell proliferation was reduced [39]. In 2013, a comparison between hPDL cells cultured under compressive forces (2.0 g/cm2 for 2 or 48 h) in conventional 2D culture dishes or in 3D collagen gel highlighted significant alterations of the expression levels of several genes [40]. In particular, the number of activated integrin–focal adhesion kinase (FAK) was higher in 3D than in 2D culture, supporting that cellular attachment to ECM can strongly influence cellular responses to mechanical forces [40]. In 2016, it was demonstrated that compressive force (1 g/cm2 for 24 h) applied to hPDLSCs altered cell morphology and repressed collagen expression, which both recovered after force withdrawal [41]. Recently, continuous compression (0–1.5 g/cm2 for 12 h) on PDLSCs could reduce differentiation ability and increase macrophage migration, osteoclast differentiation, and proinflammatory factor expression. Moreover, a universal upregulation of the subfamily V member 4 of the transient receptor potential calcium channel (TRPV4), which regulated osteoclast differentiation by affecting the system receptor activator of nuclear factor kappa-B ligand (RANKL)/osteoprotegerin (OPG) via extracellular signal-regulated kinase (ERK) signaling (Figure 3a) [42], was shown. Brockhaus et al. reported that hPDLFs cultured under compression (2 g/cm2 for 24, 48, and 72 h) changed their morphology towards more unstructured, unsorted actin filaments, with a significant reduction of proliferation followed by recovery after 48 h, demonstrating that hPDLFs restore homeostasis and adapt to the compressive force through a lower cell division rate and a slowed cell cycle [43]. Moreover, Stemmler et al. reported that the inflammatory response of hPDLFs caused by periodontal pathogens combined with compressive load (2 g/cm2 for 6 h) was supported by the growth differentiation factor 15 (GDF15), which modulated the inflammatory response of PDLFs also regulating the levels of the key inflammatory molecule tumor necrosis factor α (TNFα) [44]. Recently, Jiang and colleagues showed a novel cellular mechanism. Indeed, continuous compressive force (0.5–2.5 g/cm2, 12 h) activated autophagy in hPDLSCs that induced M1 macrophage polarization via the inhibition of the AKT signaling pathway, contributing to the force-induced bone remodeling and tooth movement [45].

2.1.2. Hydrostatic Pressure Method

The hydrostatic pressure method exploits air pressure applied on the culture medium for imposing static or fluctuating compressive forces. Exerting a static compressive stress on PDLSCs (100 kPa, for 1, 6 or 12 h) [46] and on hPDL cells (1 MPa or 6 MPa for 10 or 60 min) [47], the expression of genes regulating osteoclastogenesis and osteoblastogenesis was induced. For mimicking the physiological loading during mastication, cyclic hydrostatic pressure was applied (1 MPa, 0.1 Hz, 3 h/day for 2 days) on hPDLFs, showing that the expression of several integrins, collagens, and metalloproteinases was significantly upregulated [48]. Recently, the inflammatory, osteogenic, and pro-osteoclastic effects of different cyclic compressive loading conditions (50–150 kPa, 0.1 Hz, for 1 h/day for 5 days) were investigated by stimulating hPDL cells in an inflammatory environment using a customized bioreactor [49]. According to the level of cyclic pressure, cells released different levels of inflammatory and pro-osteoclastic factors, modulating the downregulation (with150 kPa) or the upregulation (with 90 kPa) of osteogenic genes (alkaline phosphatase (ALP), collagen type I (COLL-1), RUNX2, OCN, osteopontin (OPN), and osterix (OSX)) [50].

2.1.3. Substrate Deformation Methods

Compression can also be achieved through several commercial and customized devices based on substrate deformation, in which an elastic membrane is deformed by force and the cells/constructs cultured on it are exposed to strain. In 2014, a 3D construct composed of hPDL cells seeded into a matrix of hyaluronan, gelatin, and COLL-1 was exposed to cyclic compression (340.6 g/cm2 for 1 s every 60 s for 6, 12, and 24 h) by using the commercial Flexercell FX-4000C Strain Unit (Flexcell International Corporation, Hillsborough, NC, USA). Compression increased cell death and the expression of several apoptosis-related genes. ECM genes were mostly upregulated after 6–12 h, but all were downregulated at 24 h, except for the three major ECM-degrading enzymes—MMPs1–3— and the connective tissue growth factor (CTGF), with upregulated matrix metalloproteinase-1 (MMP-1) and tissue inhibitor of metalloproteinases-1 (TIMP-1) protein levels without changes observed in RANKL, OPG, and basic fibroblast growth factor (FGF-2) expression [51]. By using the same device, Nettelhoff et al. exposed hPDLFs to compressive force (5 and 10% for 12 h). The 5% compression induced the highest ALP gene expression and the highest RANKL/OPG ratio, while 10% compression decreased cell viability without promoting apoptosis but resulting in tissue damage [52].
Thus, short-term static compression (almost for 1 h) can promote osteogenic differentiation of PDLSCs, while long-term static compression (for 12 h or longer) can alter the morphology of hPDL cells, may inhibit cell proliferation and the osteogenic differentiation of PDLSCs, and can promote the secretion of osteoclastogenesis-stimulating cytokines and ECM degradation.

2.2. Stretch

2.2.1. Substrate Deformation—Vacuum Approach

The vacuum approach stretches the cell-seeded membrane across a loading post by applying vacuum pressure, delivering a tunable biaxial or uniaxial tensile strain. This method is commonly applied by using commercial devices (e.g., Flexercell tension system (Flexcell International Corporation, Hillsborough, NC, USA). In 2013, Saminathan et al. embedded hPDLFs in an 80–100 μm thick 3D collagen membrane and cultured the construct under equibiaxial cyclic stretching (12%, 0.2 Hz, 5 s every 60 s for 6 h/day up to 21 days) to investigate the influence on ECM homeostasis. Mechanical loading did not affect the cell number, but it significantly upregulated the release of MMP-1 and TIMP-1 in the supernatants, suggesting that fibroblasts were remodeling the surrounding ECM [53]. This was confirmed by Chen et al. who exposed hPDLCs to equibiaxial cyclic stretching (12%, 0.1 Hz for 24 h), showing the upregulation of major periodontal ECM genes, such as COL1A1, COL3A1 and COL5A1 [54]. Some studies reported that hPDLSCs cultured in osteoinductive medium under cyclic stretching enhanced the osteogenic differentiation [55][56], both maintaining the same parameters stimulation and reducing the time of stimulation [57] or keeping the same strain magnitude but halving the frequency [58][59]. In particular, Xi et al. demonstrated that cyclic stretching (10%, 0.5 Hz, for up to 36 h) could increase the generation of reactive oxygen species (ROS), which may lead to the osteogenic differentiation of hPDLSCs (Figure 3b) [58]. In 2017, Liu et al., culturing healthy or pathological donor-derived PDLSCs under static strain (6–14% for 12 h), showed that PDLSCs from patients affected by periodontitis were more sensible to physical load than PDLSCs from healthy patients, likely due to the inflammatory milieu [60]. Recently, Salim et al. cultured human PDL cells under static strain (2.5, 5, and 10% for 24 h) and performed in vivo analyses on teeth with and without orthodontic tooth movement (OTM) [61]. Interestingly, they found that chaperone-assisted selective autophagy (CASA) machinery genes (chaperones HSPA8 and HSPB8, the cochaperones BAG3 and STUB1, and the molecule SYNPO2 interacting with BAG3 for autophagosome membrane formation) were inherently expressed in PDL cells and exhibited transcriptional induction upon in vitro mechanical strain and in vivo after OTM. The role of FLNa was also investigated, pointing out that it acts as a flexible actin crosslinker that is stretched under tension and degraded by CASA when damaged, which is consistent with previous works [62][63], further supporting the importance of the dynamic environment as a key factor of the homeostatic maintenance of PDL both in physiologic and treatment conditions [64].

2.2.2. Substrate Deformation—Pulling Approach

The substrate-pulling approach is based on a system that clamps the cell-seeded membrane and imposes uniaxial stretch by a controlled actuator. Adopting the commercial STB-140 STREX cell stretch system (Strex Co., Osaka, Japan), it was reported that a long-term cyclic stimulation (5%, 60 s/returns, resting time = 29 s for 7 days) could increase collagen mRNA and protein expression, suggesting that cyclic stretch on hPDLFs may contribute to the homeostasis of PDL fibers and to the ECM remodeling [65]. In 2012, a 3D construct based on a collagen film laden with rat PDLFs was exposed to uniaxial cyclic stretch (8%, 1 Hz (15 min stretch + 15 min rest) for 8 h/day for 5 days) using a customized device. After mechanical stimulation, the cells were perpendicularly oriented with respect to the stimulation direction and, analyzing several genes’ expression (COLL-1, RUNX2, c-fos, and Cox-2), the authors concluded that PDL cells under loading might tend to have bone-like and, at the same time, tendon-like behavior [66]. Applying the same stimulation for a shorter period (16 h) and in a cyclic or static manner, the cellular orientation could be reached, and three different pathways (ERK, p38 and JNK) were activated [67]. In 2021, Yu et al. demonstrated that exposing a hPDLSCs-laden 3D collagen membrane to uniaxial stretching (20% for 5 days) dramatically enhanced the bioactivity of PDLSC-derived exosomes [68].

2.2.3. Substrate Deformation—Inflation and Bending Approaches

Studies based on the substrate inflation approach were inspired by Howard et al. [69]. A cell-seeded membrane is clamped and deflected by hydrostatic pressure applied to the underside, providing uniform biaxial stretch. In 2012, Xu and colleagues showed that hPDL cells exposed to cyclic stretching (1–20%, 0.1 Hz up to 24 h) appeared aligned perpendicularly to the stretching direction, and the expression of the membrane connexin 43 (Cx43) protein could be modulated in a time- and magnitude-dependent manner via cyclic stretching [70]. By adopting the substrate-bending approach, the osteogenic differentiation of hPDLSCs cultured under tensile stress (3000 μstrain, 0.5 Hz) was reached after 24 h of stimulation, demonstrating upregulation of different osteogenic markers [71].
In summary, the aforementioned studies demonstrated that tensile stress could promote osteogenic differentiation. In particular, cyclic stretch can upregulate the protein and mRNA expression of osteogenic genes and the synthesis of osteoclastogenesis-inhibitory molecules, complemented by an increased expression of the major periodontal ECM genes, leading to the homeostasis and organization of the PDL fibers and to ECM remodeling.

2.3. Shear Stress

The effect of shear stress on PDL cells has been poorly investigated in the literature, even though this stimulus is an important cue in the physiological environment. The most adopted method for investigating the effect of shear stress on PDL behavior is based on tangential fluid provided by parallel plate flow chambers. In 2014, Tang and colleagues cultured hPDL cells in osteogenic medium under steady fluid shear stress (12 dyn/cm2 for 2 h), showing an early morphologic change and rearrangement of filamentous actin with significant increases in ALP activity and mRNA levels of osteogenic genes and osteoid nodules [72]. Similarly, Zheng et al. exposed hPDL cells to fluid shear stress (6 dyn/cm2 up to 12 h), reporting a rearranged cell alignment, an inhibited cell proliferation and migration, and osteogenic differentiation [73]. Very recently, Shi et al. observed that fluid shear stress (6 dyn/cm2 for 4 h) promoted cell proliferation by activating mechanotransduction pathways involving the p38 mitogen-activated protein kinases, angiomotin (AMOT), and Yes-associated protein (YAP) (Figure 3c) [74]. Adopting the sliding plate method where the 3D construct is housed between two parallel plates with one of them connected to an actuator that imposes controlled sliding motion and consequent shear stress on the construct, a static shear stress was applied to a construct composed of hPDL cells embedded in a collagen gel. After 24 h of stimulation, cells and collagen fibers aligned in the direction of the principal strain vector [75]. Recently, a model of PDL regeneration based on a fiber-guiding scaffold seeded with PDL cells and subjected to shear stress in a laminar flow-based bioreactor (6 dynes/cm2 for 1–4 h) showed increased viability, adhesion, and cytoskeleton arrangement compared to cells in the absence of load [76].

The above-mentioned findings highlight how shear stress can represent an effective approach for guiding cell and fiber alignment and for promoting osteogenic differentiation.

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Figure 3. Schematic representation of some mechanotransduction pathways activated on PDL cells by the reported physical stimuli (i.e., compression, stretch, and shear stress). (a) External compression applied on rat primary PDLSCs can induce the activation of TRPV4 and consequently modulate bone remodeling [42]; (b) external cyclic stretch can promote the osteogenic differentiation of hPDLSCs by activating the Nrf2 [58]; (c) shear stress applied to hPDL cells can activate p38, which regulates the nuclear translocation YAP, promoting cell proliferation [74].
Overall, since mechanical stimulation can be essential for promoting specific PDL cellular and tissue processes, the biomimetic approach demands the use of technological devices for supporting the in vitro maturation of the PDL tissue to be grafted, particularly bioreactors providing controlled physical stimuli to reproduce the actual combined mechanical loading experienced in vivo [77][78][79]. 

References

  1. Menicanin, D.; Hynes, K.; Han, J.; Gronthos, S.; Bartold, P.M.; Cementum and Periodontal Ligament Regeneration. Adv. Exp. Med. Biol. 2015, 881, 207–236, 10.1007/978-3-319-22345-2_12.
  2. Socransky, S.S.; Haffajee, A.D.; Periodontal microbial ecology. Periodontology 2000 2005, 38, 135–187, 10.1111/j.1600-0757.2005.00107.x.
  3. Beertsen, W.; McCulloch, C.A.; Sodek, J.; The periodontal ligament: A unique, multifunctional connective tissue. Periodontology 2000 1997, 13, 20–40, 10.1111/j.1600-0757.1997.tb00094.x.
  4. de Jong, T.; Bakker, A.D.; Everts, V.; Smit, T.H.; The intricate anatomy of the periodontal ligament and its development: Lessons for periodontal regeneration.. J. Periodontal Res 2017, 52, 965–974, 10.1111/jre.12477.
  5. Ortun-Terrazas, J.; Cegonino, J.; Santana-Penin, U.; Santana-Mora, U.; Perez Del Palomar, A.; Approach towards the porous fibrous structure of the periodontal ligament using micro-computerized tomography and finite element analysis. J. Mech. Behav. Biomed. Mater. 2018, 79, 135–149, 10.1016/j.jmbbm.2017.12.022.
  6. Gathercole, L.J.; Keller, A.; Crimp morphology in the fibre-forming collagens. Matrix 1991, 11, 214–234, 0.1016/s0934-8832(11)80161-7.
  7. Maceri, F.; Marino, M.; Vairo, G.; A unified multiscale mechanical model for soft collagenous tissues with regular fiber arrangement.. J. Biomech. 2010, 43, 355–363, 10.1016/j.jbiomech.2009.07.040.
  8. Szczesny, S.E.; Driscoll, T.P.; Tseng, H.Y.; Liu, P.C.; Heo, S.J.; Mauck, R.L.; Chao, P.G.; Crimped Nanofibrous Biomaterials Mimic Microstructure and Mechanics of Native Tissue and Alter Strain Transfer to Cells. ACS Biomater. Sci. Eng. 2017, 3, 2869–2876, 10.1021/acsbiomaterials.6b00646.
  9. Cho, M.I.; Garant, P.R.; Development and general structure of the periodontium.. Periodontology 2000 2000, 24, 9–27, 10.1034/j.1600-0757.2000.2240102.x.
  10. Bosshardt, D.D.; Bergomi, M.; Vaglio, G.; Wiskott, A.; Regional structural characteristics of bovine periodontal ligament samples and their suitability for biomechanical tests. J. Anat. 2008, 212, 319–329, 10.1111/j.1469-7580.2008.00856.x.
  11. Tsuruga, E.; Irie, K.; Sakakura, Y.; Yajima, T.; Expression of fibrillins and tropoelastin by human gingival and periodontal ligament fibroblasts in vitro. J. Periodontal Res. 2002, 37, 23–28, 10.1034/j.1600-0765.2002.00662.x.
  12. Peres, M.A.; Macpherson, L.M.D.; Weyant, R.J.; Daly, B.; Venturelli, R.; Mathur, M.R.; Listl, S.; Celeste, R.K.; Guarnizo-Herreno, C.C.; Kearns, C.; et al.et al. Oral diseases: A global public health challenge.. Lancet 2019, 394, 249–260, 10.1016/S0140-6736(19)31146-8.
  13. Lourenco, T.G.; Heller, D.; Silva-Boghossian, C.M.; Cotton, S.L.; Paster, B.J.; Colombo, A.P.; Microbial signature profiles of periodontally healthy and diseased patients. J. Clin. Periodontol. 2014, 41, 1027–1036, 10.1111/jcpe.12302.
  14. Feres, M.; Teles, F.; Teles, R.; Figueiredo, L.C.; Faveri, M.; The subgingival periodontal microbiota of the aging mouth. Periodontology 2000 2016, 72, 30–53, 10.1111/prd.12136.
  15. Kawaguchi, H.; Hirachi, A.; Hasegawa, N.; Iwata, T.; Hamaguchi, H.; Shiba, H.; Takata, T.; Kato, Y.; Kurihara, H.; Enhancement of periodontal tissue regeneration by transplantation of bone marrow mesenchymal stem cells. J. Periodontol. 2004, 75, 1281–1287, 10.1902/jop.2004.75.9.1281.
  16. Du, J.; Shan, Z.; Ma, P.; Wang, S.; Fan, Z.; Allogeneic bone marrow mesenchymal stem cell transplantation for periodontal regeneration.. J. Dent. Res. 2014, 93, 183–188, 10.1177/0022034513513026.
  17. Mohammed, E.; Khalil, E.; Sabry, D.; Effect of Adipose-Derived Stem Cells and Their Exo as Adjunctive Therapy to Nonsurgical Periodontal Treatment: A Histologic and Histomorphometric Study in Rats. Biomolecules 2018, 8, 167, 10.3390/biom8040167.
  18. Seo, B.M.; Miura, M.; Gronthos, S.; Bartold, P.M.; Batouli, S.; Brahim, J.; Young, M.; Robey, P.G.; Wang, C.Y.; Shi, S.; et al. Investigation of multipotent postnatal stem cells from human periodontal ligament.. Lancet 2004, 364, 149–155, 10.1016/S0140-6736(04)16627-0.
  19. Bartold, P.M.; Gronthos, S.; Ivanovski, S.; Fisher, A.; Hutmacher, D.W.; Tissue engineered periodontal products. J. Periodontal. Res. 2016, 51, 1–15, 10.1111/jre.12275.
  20. Iwata, T.; Yamato, M.; Zhang, Z.; Mukobata, S.; Washio, K.; Ando, T.; Feijen, J.; Okano, T.; Ishikawa, I; Validation of human periodontal ligament-derived cells as a reliable source for cytotherapeutic use.. J. Clin. Periodontol. 2010, 37, 1088–1099, 10.1111/j.1600-051X.2010.01597.x.
  21. Lim, J.C.; Bae, S.H.; Lee, G.; Ryu, C.J.; Jang, Y.J. Activation of beta-catenin by TGF-beta1 promotes ligament-fibroblastic differentiation and inhibits cementoblastic differentiation of human periodontal ligament cells. Stem Cells 2020. Online ahead of print.10.1002/stem.3275
  22. Bai, S.; Lee, J.H.; Son, C.; Lee, D.S.; Park, J.C; CPNE7 regenerates periodontal ligament via TAU-mediated alignment and cementum attachment protein-mediated attachment.. J. Clin. Periodontol. 2022, 49, 609–620, 10.1111/jcpe.13621.
  23. Park, C.H.; Oh, J.H.; Jung, H.M.; Choi, Y.; Rahman, S.U.; Kim, S.; Kim, T.I.; Shin, H.I.; Lee, Y.S.; Yu, F.H.; et al.et al. Effects of the incorporation of epsilon-aminocaproic acid/chitosan particles to fibrin on cementoblast differentiation and cementum regeneration.. Acta Biomater. 2017, 61, 134–143, 10.1016/j.actbio.2017.07.039.
  24. Gauthier, R.; Jeannin, C.; Attik, N.; Trunfio-Sfarghiu, A.M.; Gritsch, K.; Grosgogeat, B.; Tissue Engineering for Periodontal Ligament Regeneration: Biomechanical Specifications.. J. Biomech. Eng. 2021, 143, 030801, 10.1115/1.4048810.
  25. Yang, L.; Yang, Y.; Wang, S.; Li, Y.; Zhao, Z.; In vitro mechanical loading models for periodontal ligament cells: From two-dimensional to three-dimensional models.. Arch. Oral Biol. 2015, 60, 416–424, 10.1016/j.archoralbio.2014.11.012.
  26. Natali, A.N.; Pavan, P.G.; Scarpa, C.; Numerical analysis of tooth mobility: Formulation of a non-linear constitutive law for the periodontal ligament. Dent. Mater 2004, 20, 623–629, 10.1016/j.dental.2003.08.003.
  27. Martino, F.; Perestrelo, A.R.; Vinarsky, V.; Pagliari, S.; Forte, G.; Cellular Mechanotransduction: From Tension to Function.. Front. Physiol. 2018, 9, 824, 10.3389/fphys.2018.00824.
  28. Schiller, H.B.; Fassler, R.; Mechanosensitivity and compositional dynamics of cell-matrix adhesions.. EMBO Rep 2013, 14, 509–519, 10.1038/embor.2013.49.
  29. Webster, K.D.; Ng, W.P.; Fletcher, D.A.; Tensional homeostasis in single fibroblasts.. Biophys. J. 2014, 107, 146–155, 10.1016/j.bpj.2014.04.051.
  30. Brown, R.A.; Prajapati, R.; McGrouther, D.A.; Yannas, I.V.; Eastwood, M.; Tensional homeostasis in dermal fibroblasts: Mechanical responses to mechanical loading in three-dimensional substrates.. J. Cell. Physiol 1998, 175, 323–332, 10.1002/(SICI)1097-4652(199806)175:3<323::AID-JCP10>3.0.CO;2-6.
  31. Kim, C.; Ye, F.; Ginsberg, M.H.; Regulation of integrin activation. Annu. Rev. Cell. Dev. Biol. 2011, 27, 321–345, 10.1146/annurev-cellbio-100109-104104.
  32. Calderwood, D.A.; Zent, R.; Grant, R.; Rees, D.J.; Hynes, R.O.; Ginsberg, M.H.; The Talin head domain binds to integrin beta subunit cytoplasmic tails and regulates integrin activation.. J. Biol. Chem. 1999, 274, 28071–28074, 10.1074/jbc.274.40.28071.
  33. Haining, A.W.; von Essen, M.; Attwood, S.J.; Hytonen, V.P.; Del Rio Hernandez, A.; All Subdomains of the Talin Rod Are Mechanically Vulnerable and May Contribute to Cellular Mechanosensing.. ACS Nano 2016, 10, 6648–6658, 10.1021/acsnano.6b01658.
  34. Carisey, A.; Tsang, R.; Greiner, A.M.; Nijenhuis, N.; Heath, N.; Nazgiewicz, A.; Kemkemer, R.; Derby, B.; Spatz, J.; Ballestrem, C.; et al. Vinculin regulates the recruitment and release of core focal adhesion proteins in a force-dependent manner.. Curr. Biol. 2013, 23, 271–281, 10.1016/j.cub.2013.01.009.
  35. Kiema, T.; Lad, Y.; Jiang, P.; Oxley, C.L.; Baldassarre, M.; Wegener, K.L.; Campbell, I.D.; Ylanne, J.; Calderwood, D.A.; The molecular basis of filamin binding to integrins and competition with talin.. Mol. Cell 2006, 21, 337–347, 10.1016/j.molcel.2006.01.011.
  36. Das, M.; Ithychanda, S.S.; Qin, J.; Plow, E.F.; Migfilin and filamin as regulators of integrin activation in endothelial cells and neutrophils. PLoS ONE 2011, 6, e26355, 10.1371/journal.pone.0026355.
  37. Shifrin, Y.; Pinto, V.I.; Hassanali, A.; Arora, P.D.; McCulloch, C.A.; Force-induced apoptosis mediated by the Rac/Pak/p38 signalling pathway is regulated by filamin A.. Biochem. J. 2012, 445, 57–67, 10.1042/BJ20112119.
  38. Li, Y.; Zheng, W.; Liu, J.S.; Wang, J.; Yang, P.; Li, M.L.; Zhao, Z.H.; Expression of osteoclastogenesis inducers in a tissue model of periodontal ligament under compression. J. Dent. Res. 2011, 90, 115–120, 10.1177/0022034510385237.
  39. Li, Y.; Li, M.; Tan, L.; Huang, S.; Zhao, L.; Tang, T.; Liu, J.; Zhao, Z.; Analysis of time-course gene expression profiles of a periodontal ligament tissue model under compression.. Arch. Oral Biol. 2013, 58, 511–522, 10.1016/j.archoralbio.2012.10.006.
  40. Kang, K.L.; Lee, S.W.; Ahn, Y.S.; Kim, S.H.; Kang, Y.G.; Bioinformatic analysis of responsive genes in two-dimension and three-dimension cultured human periodontal ligament cells subjected to compressive stress. J. Periodontal Res. 2013, 48, 87–97, 10.1111/j.1600-0765.2012.01507.x.
  41. Feng, L.; Yang, R.; Liu, D.; Wang, X.; Song, Y.; Cao, H.; He, D.; Gan, Y.; Kou, X.; Zhou, Y.; et al. PDL Progenitor-Mediated PDL Recovery Contributes to Orthodontic Relapse. J. Dent. Res. 2016, 95, 1049–1056, 10.1177/0022034516648604.
  42. Jin, S.S.; He, D.Q.; Wang, Y.; Zhang, T.; Yu, H.J.; Li, Z.X.; Zhu, L.S.; Zhou, Y.H.; Liu, Y.; Mechanical force modulates periodontal ligament stem cell characteristics during bone remodelling via TRPV4.. Cell Prolif. 2020, 53, e12912, 10.1111/cpr.12912.
  43. Brockhaus, J.; Craveiro, R.B.; Azraq, I.; Niederau, C.; Schroder, S.K.; Weiskirchen, R.; Jankowski, J.; Wolf, M.; In Vitro Compression Model for Orthodontic Tooth Movement Modulates Human Periodontal Ligament Fibroblast Proliferation, Apoptosis and Cell Cycle. Biomolecules 2021, 11, 932, 10.3390/biom11070932.
  44. Stemmler, A.; Symmank, J.; Steinmetz, J.; von Brandenstein, K.; Hennig, C.L.; Jacobs, C.; GDF15 Supports the Inflammatory Response of PdL Fibroblasts Stimulated by P. gingivalis LPS and Concurrent Compression.. Int. J. Mol. Sci. 2021, 22, 13608, 10.3390/ijms222413608.
  45. Jiang, N.; He, D.; Ma, Y.; Su, J.; Wu, X.; Cui, S.; Li, Z.; Zhou, Y.; Yu, H.; Liu, Y.; et al. Force-Induced Autophagy in Periodontal Ligament Stem Cells Modulates M1 Macrophage Polarization via AKT Signaling. Front. Cell Dev. Biol. 2021, 9, 666631, 10.3389/fcell.2021.666631.
  46. Zhang, L.; Liu, W.; Zhao, J.; Ma, X.; Shen, L.; Zhang, Y.; Jin, F.; Jin, Y.; Mechanical stress regulates osteogenic differentiation and RANKL/OPG ratio in periodontal ligament stem cells by the Wnt/beta-catenin pathway. Biochim. Biophys. Acta 2016, 1860, 2211–2219, 10.1016/j.bbagen.2016.05.003.
  47. Yamamoto, T.; Kita, M.; Kimura, I.; Oseko, F.; Terauchi, R.; Takahashi, K.; Kubo, T.; Kanamura, N.; Mechanical stress induces expression of cytokines in human periodontal ligament cells.. Oral Dis. 2006, 12, 171–175, 10.1111/j.1601-0825.2005.01179.x.
  48. Wenger, K.H.; El-Awady, A.R.; Messer, R.L.; Sharawy, M.M.; White, G.; Lapp, C.A.; Pneumatic pressure bioreactor for cyclic hydrostatic stress application: Mechanobiology effects on periodontal ligament cells.. J. Appl. Physiol. (1985) 2011, 111, 1072–1079, 10.1152/japplphysiol.01175.2010.
  49. Yousefian, J.; Firouzian, F.; Shanfeld, J.; Ngan, P.; Lanese, R.; Davidovitch, Z.; A new experimental model for studying the response of periodontal ligament cells to hydrostatic pressure.. Am. J. Orthod. Dentofac. Orthop. 1995, 108, 402–409, 10.1016/s0889-5406(95)70038-2.
  50. Jia, R.; Yi, Y.; Liu, J.; Pei, D.; Hu, B.; Hao, H.; Wu, L.; Wang, Z.; Luo, X.; Lu, Y.; et al. Cyclic compression emerged dual effects on the osteogenic and osteoclastic status of LPS-induced inflammatory human periodontal ligament cells according to loading force.. BMC Oral Health. 2020, 20, 7, 10.1186/s12903-019-0987-y.
  51. Saminathan, A.; Sriram, G.; Vinoth, J.K.; Cao, T.; Meikle, M.C.; Engineering the periodontal ligament in hyaluronan-gelatin-type I collagen constructs: Upregulation of apoptosis and alterations in gene expression by cyclic compressive strain.. Tissue Eng. Part A 2015, 21, 518–529, 10.1089/ten.TEA.2014.0221.
  52. Nettelhoff, L.; Grimm, S.; Jacobs, C.; Walter, C.; Pabst, A.M.; Goldschmitt, J.; Wehrbein, H.; Influence of mechanical compression on human periodontal ligament fibroblasts and osteoblasts. Clin. Oral Investig. 2016, 20, 621–629, 10.1007/s00784-015-1542-0.
  53. Saminathan, A.; Vinoth, K.J.; Low, H.H.; Cao, T.; Meikle, M.C.; Engineering three-dimensional constructs of the periodontal ligament in hyaluronan-gelatin hydrogel films and a mechanically active environment.. J. Periodontal Res. 2013, 48, 790–801, 10.1111/jre.12072.
  54. Chen, Y.; Mohammed, A.; Oubaidin, M.; Evans, C.A.; Zhou, X.; Luan, X.; Diekwisch, T.G.; Atsawasuwan, P.; Cyclic stretch and compression forces alter microRNA-29 expression of human periodontal ligament cells. Gene 2015, 566, 13–17, 10.1016/j.gene.2015.03.055.
  55. Wei, F.; Liu, D.; Feng, C.; Zhang, F.; Yang, S.; Hu, Y.; Ding, G.; Wang, S.; microRNA-21 mediates stretch-induced osteogenic differentiation in human periodontal ligament stem cells.. Stem Cells Dev. 2015, 24, 312–319, 10.1089/scd.2014.0191.
  56. Shen, T.; Qiu, L.; Chang, H.; Yang, Y.; Jian, C.; Xiong, J.; Zhou, J.; Dong, S.; Cyclic tension promotes osteogenic differentiation in human periodontal ligament stem cells.. Int. J. Clin. Exp. Pathol. 2014, 7, 7872–7880, PMC4270604.
  57. Wang, H.; Feng, C.; Jin, Y.; Tan, W.; Wei, F.; Identification and characterization of circular RNAs involved in mechanical force-induced periodontal ligament stem cells.. J. Cell. Physiol. 2019, 234, 10166–10177, 10.1002/jcp.27686.
  58. Xi, X.; Zhao, Y.; Liu, H.; Li, Z.; Chen, S.; Liu, D.; Nrf2 activation is involved in osteogenic differentiation of periodontal ligament stem cells under cyclic mechanical stretch.. Exp. Cell Res. 2021, 403, 112598, 10.1016/j.yexcr.2021.112598.
  59. Meng, X.; Wang, W.; Wang, X.; MicroRNA-34a and microRNA-146a target CELF3 and suppress the osteogenic differentiation of periodontal ligament stem cells under cyclic mechanical stretch. J. Dent. Sci. 2022, 17, 1281–1291, 10.1016/j.jds.2021.11.011.
  60. Liu, J.; Li, Q.; Liu, S.; Gao, J.; Qin, W.; Song, Y.; Jin, Z.; Periodontal Ligament Stem Cells in the Periodontitis Microenvironment Are Sensitive to Static Mechanical Strain.. Stem Cells Int. 2017, 2017, 1380851, 10.1155/2017/1380851.
  61. Salim, C.; Muders, H.; Jager, A.; Konermann, A.; Role of chaperone-assisted selective autophagy (CASA) in mechanical stress protection of periodontal ligament cells.. J. Orofac. Orthop. 2022, 83, 1–12, 10.1007/s00056-021-00358-3.
  62. Ulbricht, A.; Eppler, F.J.; Tapia, V.E.; van der Ven, P.F.; Hampe, N.; Hersch, N.; Vakeel, P.; Stadel, D.; Haas, A.; Saftig, P.; et al.et al. Cellular mechanotransduction relies on tension-induced and chaperone-assisted autophagy. Curr. Biol. 2013, 23, 430–435, 10.1016/j.cub.2013.01.064.
  63. Ehrlicher, A.J.; Nakamura, F.; Hartwig, J.H.; Weitz, D.A.; Stossel, T.P.; Mechanical strain in actin networks regulates FilGAP and integrin binding to filamin A.. Nature 2011, 478, 260–263, 10.1038/nature10430.
  64. Huang, H.; Yang, R.; Zhou, Y.H.; Mechanobiology of Periodontal Ligament Stem Cells in Orthodontic Tooth Movement.. Stem Cells Int. 2018, 2018, 6531216, 10.1155/2018/6531216.
  65. Nemoto, T.; Kajiya, H.; Tsuzuki, T.; Takahashi, Y.; Okabe, K.; Differential induction of collagens by mechanical stress in human periodontal ligament cells. Arch. Oral Biol. 2010, 55, 981–987, 10.1016/j.archoralbio.2010.08.004.
  66. Oortgiesen, D.A.; Yu, N.; Bronckers, A.L.; Yang, F.; Walboomers, X.F.; Jansen, J.A.; A three-dimensional cell culture model to study the mechano-biological behavior in periodontal ligament regeneration.. Tissue Eng. Part C Methods 2012, 18, 81–89, 10.1089/ten.TEC.2011.0367.
  67. Papadopoulou, A.; Iliadi, A.; Eliades, T.; Kletsas, D.; Early responses of human periodontal ligament fibroblasts to cyclic and static mechanical stretching. Eur. J. Orthod 2017, 39, 258–263, 10.1093/ejo/cjw075.
  68. Yu, W.; Su, X.; LI, M.; Wan, W.; Li, A.; Zhou, H.; Xu, F; Three-dimensional mechanical microenvironment enhanced osteogenic activity of mesenchymal stem cells-derived exosomes. Chem. Eng. J. 2021, 417, 128040, 10.1016/j.cej.2020.128040.
  69. Howard, P.S.; Kucich, U.; Taliwal, R.; Korostoff, J.M.; Mechanical forces alter extracellular matrix synthesis by human periodontal ligament fibroblasts. J. Periodontal Res. 1998, 33, 500–508, 10.1111/j.1600-0765.1998.tb02350.x.
  70. Xu, C.; Fan, Z.; Shan, W.; Hao, Y.; Ma, J.; Huang, Q.; Zhang, F.; Cyclic stretch influenced expression of membrane connexin 43 in human periodontal ligament cell. Arch. Oral Biol. 2012, 57, 1602–1608, 10.1016/j.archoralbio.2012.07.002.
  71. Tang, N.; Zhao, Z.; Zhang, L.; Yu, Q.; Li, J.; Xu, Z.; Li, X.; Up-regulated osteogenic transcription factors during early response of human periodontal ligament stem cells to cyclic tensile strain. Arch. Med. Sci. 2012, 8, 422–430, 10.5114/aoms.2012.28810.
  72. Tang, M.; Peng, Z.; Mai, Z.; Chen, L.; Mao, Q.; Chen, Z.; Chen, Q.; Liu, L.; Wang, Y.; Ai, H.; et al. Fluid shear stress stimulates osteogenic differentiation of human periodontal ligament cells via the extracellular signal-regulated kinase 1/2 and p38 mitogen-activated protein kinase signaling pathways. J. Periodontol. 2014, 85, 1806–1813, 10.1902/jop.2014.140244.
  73. Zheng, L.; Chen, L.; Chen, Y.; Gui, J.; Li, Q.; Huang, Y.; Liu, M.; Jia, X.; Song, W.; Ji, J.; et al.et al. The effects of fluid shear stress on proliferation and osteogenesis of human periodontal ligament cells. J. Biomech. 2016, 49, 572–579, 10.1016/j.jbiomech.2016.01.034.
  74. Shi, Q.; Zheng, L.; Na, J.; Li, X.; Yang, Z.; Chen, X.; Song, Y.; Li, C.; Zhou, L.; Fan, Y.; et al. Fluid shear stress promotes periodontal ligament cells proliferation via p38-AMOT-YAP. Cell. Mol. Life Sci. 2022, 79, 551, 10.1007/s00018-022-04591-w.
  75. Kim, S.G.; Kim, S.G.; Viechnicki, B.; Kim, S.; Nah, H.D.; Engineering of a periodontal ligament construct: Cell and fibre alignment induced by shear stress. J. Clin. Periodontol. 2011, 38, 1130–1136, 10.1111/j.1600-051X.2011.01790.x.
  76. Lin, H.H.; Chao, P.G.; Tai, W.C.; Chang, P.C.; 3D-Printed Collagen-Based Waveform Microfibrous Scaffold for Periodontal Ligament Reconstruction.. Int. J. Mol. Sci. 2021, 22, 7725, 10.3390/ijms22147725.
  77. Putame, G.; Gabetti, S.; Carbonaro, D.; Meglio, F.D.; Romano, V.; Sacco, A.M.; Belviso, I.; Serino, G.; Bignardi, C.; Morbiducci, U.; et al.et al. Compact and tunable stretch bioreactor advancing tissue engineering implementation. Application to engineered cardiac constructs. Med. Eng. Phys. 2020, 84, 1–9, 10.1016/j.medengphy.2020.07.018.
  78. Lim, D.; Renteria, E.S.; Sime, D.S.; Ju, Y.M.; Kim, J.H.; Criswell, T.; Shupe, T.D.; Atala, A.; Marini, F.C.; Gurcan, M.N.; et al.et al. Bioreactor design and validation for manufacturing strategies in tissue engineering. Bio-Des. Manuf. 2022, 5, 43–63, 10.1007/s42242-021-00154-3.
  79. Gabetti, S.; Masante, B.; Cochis, A.; Putame, G.; Sanginario, A.; Armando, I.; Fiume, E.; Scalia, A.C.; Daou, F.; Baino, F.; et al.et al. An automated 3D-printed perfusion bioreactor combinable with pulsed electromagnetic field stimulators for bone tissue investigations. . Sci. Rep. 2022, 12, 13859, 10.1038/s41598-022-18075-1.
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