The Driving Force of Hierarchical Collagen Fiber Formation: A Review of Tendon,Ligament,and Meniscus Mechanobiology

Abstract

In tendons, ligaments, and menisci, collagen fibers running the length of the tissue are the primary source of strength and function. Cells assemble these fibers hierarchically from nanometer-wide fibrils into larger fibers and fascicles, increasing in size throughout development and with mechanical loading. These fibers largely do not regenerate after injury or with repair, limiting recovery options. Engineered replacements are a promising treatment option; however, it remains a challenge to produce the hierarchical collagen fibers essential to tissue strength, limiting their applications. To better repair, regenerate, and engineer these tissues, we must better understand how cells regulate hierarchical fiber formation and maintenance. It is well established that mechanical cues are critical for cell-driven hierarchical fiber formation, which cells sense through several mechanisms, such as integrin-mediated adhesions, cell-to-cell connections, mechanosensitive ion channels, primary cilia, and caveolae. These mechanisms of mechanosensation have been well studied at the fibril scale of collagen organization, but as tissues mature, the loading environment becomes more complex, with cells experiencing increasing secondary shear and compressive loads generated by the developing hierarchical structure. Mechanical cues in this environment are likely sensed through several pathways, each likely playing a role in tissue maturation and injury. There remains a clear gap in our understanding of the later stages of hierarchical fiber formation, which are crucial to better understand since large hierarchical fibers dominate the human musculoskeletal system. Here, we review the role of mechanobiology in hierarchical fiber development and maintenance, and highlight what still needs further research to better regenerate fibers in engineered replacements or in vivo after injury. A better understanding of the mechanisms by which cells form hierarchically organized collagen fibers could help to overcome the limitations of current tissue engineering techniques and help to create functional repairs and replacements.

Impact Statement

Here we review the various mechanisms by which cells within tendons, ligaments, and menisci sense mechanical stimuli and the role that each mechanosensor has been shown to play in hierarchical fiber formation. Furthermore, we highlight potential future research directions to better understand the role of mechanobiology in hierarchical fiber formation. Understanding the mechanisms by which hierarchical fibers are formed, and by which mature tissues are maintained, is crucial to regenerating fibers after injury or in tissue-engineered replacements, and will help to inform the pathophysiological effects of unloading and overloading.

Keywords

mechanobiology collagen tendon ligament meniscus tissue engineering

Introduction

Collagen fibers are the primary source of strength in tendons, ligaments, and menisci. In tendons and ligaments, these fibers translate load from muscle-to-bone and bone-to-bone, respectively.^1,2 In menisci, these fibers run circumferentially around the tissue, translating compressive loads into horizontal tensile hoop stresses and distributing loads across the knee.^3–5 In all of these tissues, cells organize these fibers hierarchically, assembling collagen molecules into aligned nanometer-wide fibrils, which they bundle into larger micron-wide fibers and millimeter-to-centimeter-wide fascicles, maturing with time and loading (Fig. 1).^1,4,6–19 Cells mature fibrils and fibers simultaneously throughout development,^1,10,20–22 with each hierarchical level contributing to the overall strength of the tissues.^10–13,23

FIG. 1.

Depiction of hierarchical fiber development and maturation, with relative diameters and localization of lysyl oxidase (LOX)-induced crosslinks and proteoglycans. Cells produce tropocollagen and assemble it into aligned fibrils, which they bundle together into larger fibers and fascicles. The exact timing of when these structures emerge and become mature is still largely unknown and will likely vary between species and tissues. However, current evidence suggests cells continue to mature fibrils and fibers simultaneously throughout early development, with each hierarchical level contributing to the overall strength of the tissue. The fibers are further reinforced by the accumulation of LOX crosslinks, SLRPs, and aggrecan throughout development. SLRPs, small leucine-rich proteoglycans.

Injuries that sever these fibers result in loss of function, pain, and decreased mobility.^3,5,24 These injuries have little-to-no healing, ultimately resulting in over 1,230,000 surgeries a year in the United States, including 850,000 meniscus,^3,25 130,000 anterior cruciate ligament,^26,27 and 250,000 rotator cuff²⁸ repairs annually. Small tears are often treated via rehabilitation, which typically produces unorganized scar tissue with inferior mechanics.^29,30 More severe tears are replaced by autograft or allograft transplants, which have limited availability, risk of immune response, donor-site morbidity, and high rerupture rates.^2,3,24,31,32 Collectively, current treatments for torn tendon, ligament, or meniscus, whether repaired or replaced, often fail to restore the native structure and function.^2,3,24,30,32 It remains a challenge to regenerate the large hierarchical fibers that dominate these tissues both in vivo and in engineered tissues, limiting repair options.³⁰

To regenerate hierarchical fibers, we must better understand how cells regulate their formation and maintenance.^10,33–35 The mechanisms by which cells mediate hierarchical formation are numerous and not thoroughly understood.^34,35 Integrin-mediated actin contractility, mechanosensitive ion channels, cell–cell connections, primary cilia, and caveolae are all known to have differential yet crucial roles.^34,35 Here we review the role of mechanobiology in hierarchical fiber development and maintenance, and highlight what still needs further research to better regenerate fibers in engineered replacements or in vivo after injury.

Hierarchical Fiber Development and Maintenance

Collagen fibrillogenesis begins during embryonic development when molecular signaling and cell–cell connections dominate.^1,22,36 While embryonic movements contribute to early fibril formation,^18,37–43 hierarchical fibers primarily mature during postnatal development and adolescence, when cell–matrix interactions and mechanical cues dominate.^{9,13,29,43–51} During development, cells deposit aligned collagen fibrils as nanometer-wide bundles between cells, which they grow longitudinally and then expand laterally, increasing individual fibril and bundle diameters to form mature fibrils, fibers, and fascicles.^{1,10,13–15,20–22,30,36,52,53} How collagen fibrils form and assemble into hierarchical formations is still largely debated,¹⁰ particularly at fascicle length scales, and will likely vary between species and tissues. However, current evidence suggests fibrils and fibers mature throughout development simultaneously,^{1,10,20–22,54} and with continued maturation, there is increased tissue strength (Fig. 1).^{10,13,19,30,55}

While fiber diameters tend to correlate with mechanical properties,¹⁹ it is not the sole determining factor of tissue strength.^10,13 Proteoglycans, enzymatic crosslinks, and different types of collagen play significant and complex roles in fiber maturation. Specifically, small leucine-rich proteoglycans (SLRPs), including decorin, biglycan, fibromodulin, and lumican, and larger proteoglycans, such as aggrecan, accumulate throughout development, with SLRPs influencing fibril length, fibril spacing, and fiber diameter, altering tissue mechanics through changes in collagen structure.^{1,13,33,48,52,56} Larger proteoglycans, like aggrecan, influence fiber spacing and load distribution between fibers and fascicles in hierarchical fibers (Fig. 1).^33,57 However, larger proteoglycans are at higher concentrations in the entheses of tendons, ligaments, and menisci and the inner region of menisci, where hierarchical fibers are less dominant. In these regions, the swelling properties of proteoglycans play a substantial role in the compressive properties.^{33,35,49,58–62} Enzymatic collagen crosslinks, induced by cellular expression of lysyl oxidase (LOX), also increase throughout development. Specifically, LOX reacts with lysine in collagen to form immature divalent crosslinks that spontaneously condense into mature trivalent crosslinks.^63–66 It has been suggested that mature crosslinks provide the primary strength in fibers.^{38,63,65,67,68} Furthermore, while type I collagen is the primary collagen in fibers, smaller amounts of minor collagens, including types III, V, XI, XII, and XIV, and structural proteins such as fibronectin also contribute, helping to regulate fiber diameter, spacing, length, and fibrillogenesis.^69–82 While it is largely unknown how cells regulate proteoglycans, LOX, or collagen types to develop fibers, inhibition of any results in altered organization and reduced tissue mechanics.^{1,38,56,66,69–74,76,78,79,82–84}

Additionally, fibers and fascicles exhibit wave-like morphologies called crimp. Crimp is fundamental in force transduction by responding first to load, forming the toe region of the stress–strain curve.^20,85,86 Crimp appears during early development^20,87 and increases in wavelength with time and load.^20,88–90 How crimp forms is largely unknown; however, it is suggested to be highly regulated by the mechanical environment.^87–89,91

Mechanical Regulation of Hierarchical Fiber Formation, Homeostasis, and Repair

Loading is essential for tendon, ligament, and meniscus development, homeostasis, and repair, but how loading influences cellular regulation of these tissues is poorly understood.^29,35,43,92 During development, cells convert mechanical stimuli into biochemical signals via mechanotransduction to develop hierarchical fibers. These mechanical cues are transduced to the cell nucleus via multiple signaling pathways, such as Hippo signaling through YAP/TAZ, Rho/ROCK signaling, and the LINC complex, resulting in nuclear and chromatin deformations to alter gene expression (these pathways are outside the scope of the article, but refer the reader to the following cited reviews for further information).^93–95 Generally, as loads increase throughout development, cells produce larger fibers and fascicles, increase crosslinking, and ultimately produce stronger tissues; however, the extent of hierarchical development varies between tissues.^{15,17,29,45,96} If mechanical loads are absent during development, tissues fail to mature, resulting in underdeveloped fibers and impaired expression of SLRPs, LOX, and collagens.^{9,13,18,29,35,43,45,51,75} Once fibers have formed, exercise can increase fiber maturation, producing stronger tissues, but too much exercise/load can lead to degeneration and injury.^9,29,35,48

The mechanisms by which cells sense mechanical cues are complex and likely change throughout hierarchical fiber formation (Fig. 2). It is established that cells initially form aligned fibrils largely through cell–cell connections and cellular contraction forces, highly regulated by integrins, focal adhesion kinase (FAK), and the actomyosin network,^{20,34,36,91,97–101} with cells remodeling the matrix to achieve a tensional homeostasis.^{34,98,102,103} Tensional homeostasis is a set point of tensile strain that cells attempt to maintain.^98,103 When this is lost, cells respond with an injurious, catabolic response.^88,98,103 However, stimulation that challenges tensional homeostasis can promote an anabolic response, increasing tissue maturation.^29,45,96,104 In fact, tensile loads, including slow growth and cyclic muscle activity, are thought to drive development by applying uniaxial stretch to fibers, which is translated to cells via focal adhesions (FAs), altering their tensional homeostasis and stimulating further development.^{9,29,43,45,51,105,106} However, while cells predominantly receive tensile strains transferred via cell–cell connections or aligned fibrils in early postnatal development, as tissues mature, this environment becomes more complex, with cells increasingly experiencing higher degrees of secondary shear and compressive loads generated by the developing hierarchical structure (Fig. 2).^44,107 Throughout development, cells organize themselves in rows, joined on either end by cell–cell adhesions, and form mature fibers around themselves.^100,108 Primary cilia align with tension, and cells alter the expression of mechanosensors such as mechanically gated ion channels and FA-related proteins.^104,109,110 Mechanical cues are sensed through several pathways, each likely playing a role in cellular response to load during fiber development and injury.

FIG. 2.

Depiction of major cell mechanosensation pathways and the evolving mechanical environment throughout hierarchical fiber development. (A) Initially, during embryonic development, cell–cell connections and cellular traction forces dominate as cells deposit aligned fibril bundles. (B) In early postnatal development, cells are oriented in channels between fibril bundles/immature fibers. Here, cells may predominantly receive tensile strains transferred via focal adhesions, based on the critical role of actin-cytoskeletal sensing in fibril formation (see Table 1). (C) As the tissues mature, this environment becomes more complex, with cells experiencing increasing amounts of secondary shear and compressive loads generated by the developing hierarchical structure. This changing loading environment may shift expression, activity, and roles of mechanosensors. Mechanosensors portrayed in the image are based on studies that have found their expression or reported on their activity at these stages of development and are not exhaustive of all sensors likely present.

Table 1.

Effects of Tensional Homeostasis/Cytoskeleton Disruption on Collagen Degradation, Fibril Formation, and Tissue Maturation

Treatment	Culture conditions	Cell type	Outcome
Disruption of actin polymerization	In vitro in 3D collagen gels⁹⁸	Rat tail tenocytes⁹⁸	Inhibited contraction, increased collagenase expression⁹⁸
Disruption of actin polymerization	Ex vivo in mouse tail tendon and chick metatarsal tendon³⁶	Mouse tail tenocytes, embryonic chick tenocytes³⁶	Reduced fibripositor function, reduced fibril alignment³⁶
Microtubule disruption via nocodazole and colchicine	Ex vivo in chick metatarsal tendon³⁶	Embryonic chick tenocytes³⁶	Reduced collagen secretion³⁶
Blebbistatin (NMMIIA inhibitor)	In vitro in 3D alginate gels¹⁰⁰ or de novo collagen gels²¹	Embryonic chick tendon progenitor cells^21,100	Reduced tissue elastic modulus,¹⁰⁰ reduced fibripositor function and fibril laydown²¹
	In vitro on an electrospun PCL scaffold¹²⁵	Mouse tendon fibroblasts¹²⁵	Increased catabolism, reduced YAP/TAZ signaling¹²⁵
	In ovo in chick flexor tendon¹⁰⁰	Embryonic chick tenocytes¹⁰⁰	Reduced tissue elastic modulus¹⁰⁰
FAK inhibition, FAK knockdown, or integrin deficiency	In vitro 2D culture^114,126	FAK-null or Y397F-FAK mouse embryonic fibroblasts,^114,126 mouse embryonic fibroblasts¹²⁶	Reduced cell motility and realignment in response to tractional force,¹¹⁴ increased MMP activity and reduced collagen alignment¹²⁶
	In vitro in 3D collagen-coated nylon meshes¹²⁶	Mouse embryonic fibroblasts¹²⁶	Increased MMP activity and reduced collagen alignment¹²⁶
	In vivo in α2β1 integrin-deficient mice¹²⁰	Mouse Achilles tenocytes¹²⁰	Increased MMP activity^120,126
Disruption of LINC complex function	In vivo in dnKASH mice⁹¹	Mouse Achilles and tail tenocytes⁹¹	Decreased Achilles tendon mechanics with increased cross-sectional area, decreased tail tendon cross-sectional area⁹¹
Loss of tension	In vitro in fibrin gels with or without TGF-β,¹²⁷ de novo collagen matrices,⁸⁹ or collagen gels^98,105	Human tenocytes,¹²⁷ embryonic chick metatarsal tenocytes,⁸⁹ rat tail tenocytes,⁹⁸ canine ACL fibroblasts¹⁰⁵	Rapid contraction of matrix,^89,127 abnormal cell phenotype, shift to an injurious or catabolic state,^98,105,127 decreased integrin expression and organization compared with tethered gels¹⁰⁵
	Ex vivo in rat tail tendon fascicles^{102,103,109,128,129}	Rat tail tenocytes^{102,103,109,128,129}	Rapid contraction of the matrix, abnormal cell phenotype, shift to an injurious state,^102,103,128 increased cell primary cilia length^109,129
	In vivo in rabbit lumbar IVD⁹⁹	Rabbit IVD fibroblasts⁹⁹	Rapid contraction of the matrix, abnormal cell phenotype, shift to an injurious state⁹⁹

ACL, anterior cruciate ligament; dnKASH, dominant negative expression of the Klarsicht, ACN-1, and Syne Homology domain; FAK, focal adhesion kinase; IVD, intervertebral disc; MMP, matrix metalloproteinase; NMMIIA, nonmuscle myosin II-A; PCL, polycaprolactone; TAZ, transcriptional coactivator of yes-associated protein with PDZ-binding motif; TGF-β, transforming growth factor-β; YAP, yes-associated protein.

Actin-Cytoskeletal Tension and Integrin-Mediated Adhesions

While collagen remodeling is regulated by numerous signals, none are as well studied as actin-mediated mechanotransduction. Cells reach tensional homeostasis via regulation of the actin-cytoskeleton, largely through actin-to-extracellular matrix (ECM) connections called FAs.^{103,111–113} FAs are theorized to be one of the primary mechanisms by which cells in hierarchically organized tissues sense forces, though much of this work has been in 2D or ex vivo systems.^{103,105,111,114–120} These mechanosensitive macromolecular structures are composed of hundreds of proteins and act as a direct signaling pathway between the ECM and the nucleus.^111,121,122

FAs are primarily mediated by integrins, FAK, and the actin-cytoskeleton.^{112,116,118,123} FAs first form when integrins bind to ECM.^105,111 The cytoplasmic tail of integrins initiates a cascade of protein binding, including FAK, and the resulting complex of proteins attaches to the actin-cytoskeleton, providing a direct mechanical signal from ECM to nucleus.^111,112,124 Several components of this connection are known to have mediatory roles in fibril production and alignment (Table 1).³⁶ For example, nonmuscle myosin II-A (NMMIIA) of the actin-cytoskeleton contributes to collagen synthesis by forming cell extensions called fibripositors.^21,36 Fibripositors are extensions of the cell membrane that align with the axis of tension to lay down fibrils during embryonic development.^21,36 Disruption of these actin filaments with cytochalasin D (CytoD),³⁶ inhibition of NMMIIA via blebbistatin,¹⁰⁰ and inhibition of the LINC complex⁹¹ during embryonic development all result in reduced collagen organization and loss of tendon strength.

Cytoskeletal tension also plays a significant role in mature tissues. It is established that ex vivo untethered tendons, or tethered tendons treated with CytoD to disrupt actin polymerization, result in loss of cytoskeletal tension (Table 1).^{89,100,103,125,130–132} In both cases, cells lose their tensional homeostasis and respond injuriously, increasing catabolic gene expression and matrix degradation,^{98,99,103,127,128,130} resulting in reduced collagen secretion, alignment, and tissue strength.^{36,103,125,130–132} Treatment with blebbistatin also reduces YAP/TAZ activation, suggesting that actin-cytoskeletal tension may activate anabolic genes via this pathway.¹²⁵ This work demonstrates that the actin-cytoskeleton provides a mechanical cue to cells in the form of prestress to the ECM, but also a mechanism by which cells sense dynamic cues from the environment.^133,134

Harnessing actin-cytoskeletal tension in engineered tissues

The role of actin-cytoskeletal contraction in fibril formation is so critical that efforts to produce aligned collagen fibers in engineered tissues have primarily focused on harnessing cellular contraction. In these culture systems, cells are seeded into collagen or fibrin hydrogels, and boundary conditions are applied, often using pins or clamps to mimic the natural attachment points of these tissues (Table 2).^{40,105,133,134,136–148,150,151} As cells contract the gel through FAs to develop tensional homeostasis, they sense higher tension between the boundary constraints, prompting formation of aligned fibrils along the primary axis of stress.^{105,133,140,141,149,152} This actin-mediated ECM alignment and remodeling is notably similar to embryonic fibrillogenesis,^{40,105,136,137} with colocalization of collagen types I, III, XII, and XIV, a parallel fibronectin network, and fibripositors.⁴⁰ The boundary conditions allow cells to return to their tensile homeostatic set point.¹⁰³ However, these systems have primarily consisted of low-density collagen or fibrin gels (1–5 mg/mL), which have not been shown to support larger fibers and fascicle development. We have found that when using higher concentrations of collagen (10–20 mg/mL), cells form native-sized fibers and early fascicles.^{68,144,145,149} As collagen concentration increases, gross cellular contraction of the gel decreases, and tensile moduli significantly increase.^144,153,154 We hypothesize these attributes of high-density collagen gels provide the time and mechanical cues needed to support cellular development of larger hierarchical fibers.

Table 2.

Effects of Static Boundary Conditions on Collagen Fiber Formation and Maintenance In Vitro

Treatment	Culture conditions	Cell type	Outcome
Static boundary constraints	Ex vivo whole rat tail tendon^88,130,135	Rat tail tenocytes^88,130,135	Reduced MMP1 expression compared with stress-deprived tendons,¹³⁰ increased connexin 43 expression,¹³⁵ increased crimp length compared with untethered tendons⁸⁸
	In vitro in 3D fibrin gels^40,136–138	Human tenocytes,^40,137 mouse tendon and ligament progenitor cells,¹³⁶ embryonic chick tenocytes¹³⁸	Formation of an aligned collagen matrix, increased collagen fiber diameter and tissue mechanics,^40,136–138 improved tenogenic marker expression compared with collagen gels¹³⁶
	In vitro in 3D low-density collagen gels^{105,133,134,136,139–143}	Human fibroblasts,¹⁴⁰ mouse tendon and ligament progenitors,¹³⁶ rat tendon fibroblasts,¹³³ canine ACL fibroblasts,¹⁰⁵ mouse embryonic stem cells,¹⁴² rabbit mesenchymal stem cells,¹⁴³ human glioblastoma cells,¹³⁴ rat cardiac fibroblasts^133,141	Increased cell and collagen alignment compared with unclamped collagen,^{105,133,134,136,139–141,143} increased integrin α5 and β1 expression,¹⁰⁵ increased mechanics compared with unbound collagen,¹³⁶ increased mechanotransduction gene expression^136,142
	In vitro in 3D high-density collagen gels^{68,144,145,146–148,149}	Bovine meniscal fibrochondrocytes,^68,144–148 bovine ACL fibroblasts,^145,149 bovine flexor tenocytes,¹⁴⁵ bovine mesenchymal stem cells¹⁴⁸	Formation of hierarchically organized collagen fibers with fiber diameters that match native tissue, increased strength compared with unclamped constructs^{68,144,145,146–148,149}

MMP1, matrix metalloproteinase 1.

Increasing maturation with dynamic stimulation in engineered tissues

As discussed previously, dynamic stimulation is critical to fiber development in vivo and similarly in engineered tissues. A great deal of research has investigated the application of dynamic loading to drive fiber formation in vitro (Table 3).^{105,106,150,155,156,161,163–165,168–172} The most studied mechanical cues are those that mimic in situ cues, namely cyclic strain similar to what tendons, ligaments, and menisci undergo when walking or running. Cyclic strain upregulates integrin expression and cellular alignment, implying it is sensed through the integrin/FAK pathway.^{105,157,171,173} This change in integrin expression corresponds to changes in collagen organization at the micro- and macroscopic scale,¹⁰⁵ with cyclic load often driving collagen turnover and matrix maturation,^{105,155,156,163,164} resulting in increased fibril, fiber,^{136,161,164,165} and crimp formation.^164,165 Mechanotransduction has been less explored in the meniscus, and since the meniscus primarily experiences compressive loads that are translated into tensile hoop stresses, mechanosensors may have different roles in meniscal fibrochondrocytes. However, it has been shown in engineered menisci that dual compressive/tensile loading regimes, which convert compressive loads into tensile hoop stresses, further enhance fiber formation and zonal matrix formation over compressive loading alone, demonstrating the critical role of tensile cues.^{154,168,169,174}

Table 3.

Effects of Dynamic Load on Hierarchical Fiber Formation in Engineered Tissues and Fiber Maintenance in Mature Tissues

Treatment	Culture conditions	Cell type	Outcome
Cyclic tensile strain	In vitro 2D culture on flexible membranes^155–160	Human foreskin fibroblasts,¹⁵⁷ rabbit tenocytes,¹⁵⁶ human periodontal ligament cells,^155,158 human spinal ligament fibroblasts,¹⁵⁹ human semitendinosus tenocytes¹⁶⁰	Increased collagen expression,¹⁵⁹ Increased cell alignment,¹⁵⁷ increased MMP expression,^155,156 increased connexin 43 expression,^158,159 decreased primary cilia length¹⁶⁰
	In vitro in PCL scaffolds exposed to strain^161,162	Human mesenchymal stem cells,¹⁶¹ bovine meniscal fibrochondrocytes¹⁶²	Increased matrix maturation, collagen laydown, and tensile properties,¹⁶¹ increased Ca²⁺ flux¹⁶²
	In vitro in 3D collagen gels exposed to 2.5%^105,136 or 5%¹⁶³ cyclic strain, or fibrin gels exposed to 2.4% strain¹³⁶	Mouse tendon and ligament progenitor cells,¹³⁶ canine ACL fibroblasts,¹⁰⁵ human Achilles tenocytes¹⁶³	Increased expression of tenogenic markers and mechanics compared with static load,¹³⁶ increased integrin/FAK expression and activation,¹⁰⁵ increased cell spreading, increased matrix remodeling via MMPs¹⁶³
	In vitro in 3D high-density collagen gels with 5 or 10% cyclic load^164,165	Bovine ACL fibroblasts^164,165	Increased hierarchical fiber formation, crimp formation, collagen accumulation and tensile properties^164,165
	Ex vivo mouse patellar tendons loaded at 1 or 10% strain after injurious strain¹⁰⁴	Mouse patellar tenocytes¹⁰⁴	Increased actin and cell nucleus alignment with 10% strain¹⁰⁴
	Ex vivo rat tail tendons cyclically loaded at 1%, 3%, or 6% strain after stress-deprivation^102,129	Rat tail tenocytes^102,129	Cyclic load helps recover tensional homeostasis, reduced MMP-13 expression,¹⁰² and reduced cilia length compared with when stress-deprived¹²⁹
	Ex vivo bovine meniscus,¹⁶² ex vivo chicken flexor tendons¹⁶⁶	Bovine meniscal fibrochondrocytes,¹⁶² chicken flexor tenocytes¹⁶⁶	Increase Ca²⁺ flux,¹⁶² increased connexin 32 and 43 expression, altered vinculin expression¹⁶⁶
	In vivo in mouse supraspinatus tendon¹⁶⁷	Mouse supraspinatus tenocytes¹⁶⁷	Improved enthesis injury repair¹⁶⁷
Slow growth elongation	In vitro in pinned fibrin-based tendon constructs at 2 mm/day,¹⁰⁶ 3D high-density collagen gels with 0.1 mm/day of stretch¹⁶⁵	Embryonic chick metatarsal tenocytes,¹⁰⁶ bovine ACL fibroblasts¹⁶⁵	Increased collagen fibril diameter and tensile properties,^106,165 increased cell alignment, improved development of engineered enthesis¹⁶⁵
Dual cyclic compressive/tensile strain	In vitro in scaffold-free meniscal constructs,¹⁶⁸ or 3D high-density meniscus-shaped collagen gels¹⁶⁹	Bovine meniscal fibrochondrocytes^168,169	Increased matrix maturation and mechanics, increased fiber diameter, tensile properties, and zonal matrix formation^168,169

As discussed previously, while cells may predominantly receive tensile strains transferred via aligned collagen fibril bundles or cell–cell connections in early development, as tissues mature, this environment becomes more complex, with cells likely experiencing higher degrees of secondary shear and compressive loads generated by the developing hierarchical structure (Fig. 2).^150,175,176 Furthermore, as cells align collagen fibrils, they stiffen the tissue, activating further integrins and other mechanical signaling pathways such as mechanically gated ion channels.^126,142,162 In fact, in conditional FAK knockout mouse models, FAK inhibition during postnatal development did not abolish collagen organization in tendons and ligaments, and instead only reduced collagen alignment by 20% and collagen density by 40% in the periodontal ligament,¹²⁶ and reduced size and fibril diameters in flexor, Achilles, and tail tendons, but tensile properties were largely maintained.¹¹⁹ This demonstrates that while FAK- and integrin-mediated mechanotransduction play a significant role in collagen synthesis and alignment, other mechanisms beyond integrin-based contraction are needed during later stages of hierarchical fiber formation.

Cell–Cell Adhesions

Cell–cell adhesions are highly expressed in tendon, ligament, and meniscus cells and are crucial to tissue development and tensional homeostasis.^{158,177–184} In embryonic development, cells are tightly packed and communicate via cell–cell junctions mediated by cadherins and connexin-mediated gap junctions (Fig. 2).^{100,108,182,185} As tissues mature, cells are arranged in long rows joined together by cell–cell adhesions.^{100,108,166,182}

Gap junctions are crucial to fibril synthesis and alignment, with inhibition often increasing fibril dispersion and upregulating inflammatory genes.^{173,183,186,187} In particular, connexin (Cx)-32 and -43 are established to help regulate collagen secretion and to be mechanosensitive, but with distinct roles (Table 4).^{158,159,173,180} For example, in chick flexor tenocytes, signaling through Cx32 stimulates collagen secretion in 2D, while Cx43 signaling inhibits collagen secretion, both in static and dynamically loaded cultures.¹⁸⁰ Furthermore, while Cx32 and Cx43 are upregulated in response to load, strains beyond physiological levels (>8% strain) have been shown to suppress gap junction signaling, hampering intercellular communication.^135,188,189

Table 4.

Role of Cell–Cell Adhesions in Response to Load and in Collagen Maturation

Treatment	Culture conditions	Cell type	Outcome
Static load	Ex vivo rat tail tendon fascicle with 1N load¹³⁵	Rat tail tenocytes¹³⁵	Increased cellular diffusion with 10 min of load, decreased gap junction permeability and Cx43 expression with 1 h of load¹³⁵
	In vitro 2D culture on microgrooved device with 4% or 8% static strain^188,189	Rabbit Achilles tenocytes^188,189	Increased cellular diffusion through gap junctions and increased Cx43 gene expression at 4% strain, decreased diffusion and reduced Cx43 expression with 8% strain^188,189
Dynamic load	In vitro in 2D with 10% strain¹⁵⁹	Human longitudinal ligament fibroblasts¹⁵⁹	Increased Cx43 gene expression, increased gene expression related to ossification¹⁵⁹
	In vitro in 2D with 1%, 10%, or 20% strain¹⁵⁸	Human periodontal ligament fibroblasts¹⁵⁸	1% strain increased Cx43 gene and protein expression, 10% and 20% decreased Cx43 expression¹⁵⁸
	In vitro in 3D spheroid with 14% strain¹⁷³	Rabbit ACL fibroblasts¹⁷³	Increased Cx43 expression¹⁷³
Cx32 inhibition	In vitro 2D culture with or without mechanical strain¹⁸⁰	Chicken tenocytes¹⁸⁰	Reduced collagen synthesis and fibril laydown with Cx32 inhibition¹⁸⁰
Cx43 inhibition	In vivo in mice¹⁷⁹	Mouse supraspinatus and Achilles tenocytes¹⁷⁹	Impaired enthesis formation¹⁷⁹
	In vitro 2D culture with or without mechanical strain¹⁸⁰	Chicken tenocytes¹⁸⁰	Increased collagen synthesis and fibril laydown with Cx43 inhibition¹⁸⁰
	In vitro in 3D scaffold-free cultures¹⁸⁷	Human periodontal ligament fibroblasts¹⁸⁷	Reduced collagen alignment¹⁸⁷
Increased Cx43 expression	In vitro in 2D¹⁸³	Equine tenocytes¹⁸³	Increased collagen synthesis and intercellular communication¹⁸³
Cadherin 11 siRNA	In vitro 2D culture, ex vivo chick tendons¹⁹⁰	Embryonic chick tenocytes¹⁹⁰	Abnormal cell morphology, reduced collagen organization¹⁹⁰
	In vitro 2D culture with mechanical load¹⁹¹	Human periodontal ligament cells¹⁹¹	Abnormal cell morphology, reduced collagen synthesis¹⁹¹
Increased N-cadherin expression	In vitro in 2D¹⁹²	Rat periodontal ligament fibroblasts¹⁹²	Increased matrix mineralization¹⁹²

Cx32, connexin 32; Cx43, connexin 43.

Gap junctions are also critical in enthesis formation. The enthesis is the fibrocartilaginous transition region in tendons, ligaments, and menisci that translates loads between elastic ligamentous tissue and stiff bone.¹⁷⁹ In a murine model, deletion of Gja1, which encodes for Cx43, results in reduced cell alignment and mineralization in the enthesis compared with wildtype (WT) controls.¹⁷⁹ This correlates well with studies in periodontal ligament cells (PDLCs) and other ligament fibroblasts, where Cx43 was shown to have a significant role in ossification.^158,159,193

Another mechanism of force transduction in cell–cell adhesions is cadherins, transmembrane proteins that form cell–cell junctions. These proteins are highly expressed in musculoskeletal tissues, mechanosensitive, and have a role in regulating collagen secretion.^190,191 In particular, cadherin 11 is expressed in tendon and ligament fibroblasts and suppressed by load.^190,191 Furthermore, knockdown of cadherin 11 in these tissues reduces collagen synthesis¹⁹¹ and collagen organization.¹⁹⁰ N-cadherins have also been shown to be expressed in PDLCs and are more highly expressed when differentiated to form mineralized nodules, again suggesting that cell–cell connections have a role in ossification.¹⁹²

Mechanosensitive Ion Channels

Due to the large array of cellular functions regulated by calcium, stretch-activated calcium-permeable channels in the cell membrane are suggested to be significant regulators of musculoskeletal development.^162,194 Traditionally, these mechanically activated ion channels were thought to be primarily stimulated by external loads; however, recent work has reported that these channels have significant roles in regulating cellular contraction forces as well.¹⁹⁵ Cellular contraction or variations in matrix stiffness can activate ion channels, increasing intracellular calcium and altering cellular mechanosensitivity.^194–197 This allows cells to respond to a continuum of mechanical stimuli, often stimulating downstream events distinct from FAK-actin contractility.^198–200

While there are many ion channels, transient receptor potential vanilloid 4 (TRPV4), Piezo1 and 2, and voltage-gated channels are reported to have significant and independent roles in the development of many connective tissues^{194,198,201,202} and are found to be highly expressed in fibrous tissues, including tendons, ligaments, and menisci.^{175,198,203–206} These mechanosensitive channels respond to a wide range of mechanical cues, including passive stimulation from cellular contraction and dynamic stimulation from forces such as stretch, compression, shear flow, and osmotic stress.^{175,195–197,199,200,207,208} Furthermore, while all respond to a wide range of mechanical cues, they appear to differ in the strength of load needed for activation and often stimulate distinct downstream signaling (Table 5).^{198–201,207}

Table 5.

Role of Ion Channels in Cell Signaling, Collagen Fiber Formation, and Tissue Maturation

Treatment	Culture conditions	Cell type	Outcome
TRPV4
TRPV4 inhibitor	In vitro 2D culture¹⁹⁶	Mouse fibroblasts¹⁹⁶	Reduced integrin signaling, reduced FliI-NMMIIA interactions, reduced collagen remodeling¹⁹⁶
	In vitro on fibronectin-coated slides with oscillating fluid shear²⁰⁹	Human mesenchymal stem cells²⁰⁹	Lack of calcium signaling due to oscillating fluid shear²⁰⁹
	In vitro on aligned microphotopatterned substrates²¹⁰	Human mesenchymal stem cells²¹⁰	Reduced integrin signaling via altered vinculin tension, reduced collagen alignment²¹⁰
	In vitro in 3D agarose constructs²⁰⁷	Porcine articular chondrocytes²⁰⁷	Reduced Young’s modulus, disrupted fibril alignment, reduced collagen synthesis and remodeling¹³³
TRPV4 knockdown/knockout	In vitro in 2D on flexible collagen-coated wells¹⁹⁹	Mouse primary chondrocytes¹⁹⁹	Reduced Ca²⁺ flux, reduced cell response to cyclic strain¹⁹⁹
TRPV4 knockdown/knockout	In vitro in fibronectin-coated collagen gels²¹¹	Mouse and human gingival fibroblasts²¹¹	Reduced β1 integrin expression²¹¹
TRPV4 agonist/activation	In vitro on fibronectin-coated slides with oscillating fluid shear²⁰⁹	Human mesenchymal stem cells²⁰⁹	Increased expression of osteogenic markers²⁰⁹
TRPV4 agonist/activation	In vitro in fibronectin-coated collagen gels²¹¹	Mouse and human gingival fibroblasts²¹¹	Increased Ca²⁺ flux and TRPV4 mRNA expression²¹¹
Piezo1
Piezo1 inhibitor	In vitro 2D culture under static compressive load²¹²	Human periodontal ligament cells²¹²	Mechanical stimulation increased Piezo1 expression, Piezo1 inhibition suppressed NF-κB activation after load, and reduced mechano-response²¹²
Piezo1 knockdown	In vitro in 2D on flexible collagen-coated wells¹⁹⁹	Mouse primary chondrocytes¹⁹⁹	Reduced Ca²⁺ flux, reduced cellular response to cyclic strain¹⁹⁹
Conditional Piezo1 KO	In vitro in 2D¹⁷⁵	Human and rat tenocytes¹⁷⁵	Reduced Ca²⁺ response to shear stress¹⁷⁵
	Ex vivo rat tail tendon fascicles¹⁷⁵	Rat tail tenocytes¹⁷⁵	Reduced Ca²⁺ signaling, reduced tendon stiffness without alterations in diameter¹⁷⁵
	In vivo in mice¹⁷⁵	Mouse tendon¹⁷⁵	Decreased tissue stiffness¹⁷⁵
Piezo1 GOF mutation or overexpression	In vivo in mice^175,213 and in human patellar tendons²¹⁴	Mouse tenocytes,^175,213 Human patellar tenocytes²¹⁴	Increased tendon stiffness without alterations in diameter,^175,214 increased LOX crosslinking,¹⁷⁵ increased mouse jumping ability²¹³
Piezo1 agonist/activation	Ex vivo rat tail tendon fascicles¹⁷⁵	Mouse tail tenocytes¹⁷⁵	Increased LOX expression, increased tendon mechanics¹⁷⁵
Cav1.2
Cav1.2 GOF mutation	In vivo in mouse Achilles tendon²⁰⁶	Mouse tenocytes²⁰⁶	Increased elastic modulus, decreased cell density, increased collagen fibrillogenesis²⁰⁶
Cav1.2 blocker therapy	In vivo in humans with hypertension²⁰⁵	Human tenocytes²⁰⁵	Decreased incidence of tendinopathy²⁰⁵

FliI, flightless I; NMMIIA, nonmuscle myosin II A; LOX, lysyl oxidase; mRNA, messenger ribonucleic acid; TRPV4, transient receptor potential vanilloid 4.

TRPV4

Transient receptor potential (TRP) channels are a group of ion channels that are crucial in cell signal transduction, sensing various stimuli such as chemical, temperature, and mechanical cues.²⁰² Studies on 2D surfaces have found TRPV4, in comparison with several TRP family members, mediates the Ca²⁺ influx needed to develop cellular contraction and is required for collagen remodeling in fibroblasts and human mesenchymal stem cells (hMSCs).^194,196,210 Furthermore, TRPV4 has garnered interest since it has been shown to be highly expressed in musculoskeletal tissues,^{199,204,210,215} to play a significant role in fibril formation, and to respond to mechanical cues that tendons, ligaments, and menisci experience in situ (3–8% strain).^{199,201,207,216,217}

TRPV4 activation has been shown to regulate fibril laydown and collagen remodeling in several cell types (Table 5).^{194,196,210,211,218,219} More specifically, Ca²⁺ transfer through TRPV4 assists in a molecular chaperone function during protein synthesis,²²⁰ regulating early alignment of fibrils and the creation of fibripositors in dermal fibroblasts and stem cells.^196,210 In a microphotopatterned system, TRPV4 was critical during the initial stages of collagen matrix assembly in hMSCs, contributing to fibril synthesis and alignment.²¹⁰ However, its activity significantly decreased once aligned fibrils were produced,²¹⁰ suggesting its activity once fibrils are formed may be detrimental and it has less of a role during fiber and fascicle formation.

While TRPV4 mechanotransduction has a role in fibril formation independent of integrin-FAK-cytoskeletal signaling, it also helps regulate actin-cytoskeleton function, activating various proteins in the integrin signaling cascade.^{198,221–224} In particular, calcium flux through TRPV4 affects NMMIIA in cell extensions^194,196 and modulates tension across vinculin, significantly impacting FA formation and collagen remodeling.²¹⁰ Collectively, TRPV4 plays a significant role in FA function and cell–ECM interactions, including matrix synthesis, cellular contractility, MMP-1 activity, and cytokine production^194,218,225; however, this work has primarily been in 2D, and its role in organization beyond fibril alignment is largely unknown.

Piezo

Another family of ion channels that has received significant attention is the Piezo family, specifically Piezo1 and Piezo2, with Piezo1 suggested to be a critical regulator of LOX expression in mature tendons.¹⁷⁵ Similar to TRPV4, Piezo1 is activated by cellular contraction, with Piezo1 activation increasing as contraction forces increase,^{195,197,200,226} suggesting it is a higher-load sensor than TRPV4. Furthermore, Piezo channels respond to a wide range of mechanical cues, such as osmotic stress, compression, tension, and shear stress.^{197,227–231} A consensus on the mechanism of Piezo activation has yet to be reached, though it is well established that Piezo channels are activated by stretch and shear stress and are highly expressed in bone, cartilage, tendons, and ligaments (Table 5).^{198,212,232–235} In cartilage, Piezo1 and Piezo2 are believed to activate at different tensile strains, with Piezo2 activating at higher strains than Piezo1,^199,200 and have been shown to have synergistic effects.²⁰⁰ While this work was in chondrocytes, the same is likely true for cells in other musculoskeletal tissues where Piezo1 and 2 are both expressed.

In mature tendons, Piezo1 is reported to be a critical sensor of shear stress, responding to the level of stress generated by fiber sliding during physiological strains.¹⁷⁵ Recent work in mouse and human tenocytes has found that Piezo1 is associated with increases in tendon strength and LOX expression, without changes in tendon or fibril diameter, suggesting Piezo1 activation increases LOX crosslinking.^175,214,236 Furthermore, mice with a Piezo1 tendon-specific knock-in were found to have stiffer Achilles tendons, resulting in increased storage modulus, increased jumping ability, and faster running speeds compared with WT mice.²¹³ Humans carrying the Piezo1 GOF mutation, E756del, also had no differences in tendon size compared with healthy controls but had significantly increased jumping ability²¹³ and patellar tendon stiffness.²¹⁴

As already discussed, mechanosensitive ion channels are activated by external mechanical cues, with both TRPV4 and Piezo1 responding to stretch, compression, shear, and osmotic stress.^{175,195–197,199,200,207,208} In fact, mechanical stimulation of tendon, ligament, and meniscus significantly upregulates calcium signaling,^162,175 suggesting a role of these calcium stretch channels. Furthermore, TRPV4 and Piezo1 are both upregulated in response to load but respond to different ranges of load with differential gene expression.^{142,198,199,201} Specifically, in chondrocytes, it has been shown that TRPV4 responds to low strain, while Piezo1 responds to high strain, allowing cells to respond to a continuum of mechanical cues.^199,200,235

Voltage-gated channels

Voltage-gated calcium channels can also be activated as part of the cell’s response to mechanical stimulation. CaV1.2 is expressed in tendons and activates downstream of Piezo, as a result of the rapid influx of cations through Piezo1 and 2.²⁰⁶ It has been suggested that CaV1.2 helps regulate collagen fibrillogenesis and tendon mechanics, since mice with CaV1.2 GOF mutation have tendon hypertrophy, with increased tendon stiffness, ultimate tensile strength, and increased expression of ECM proteins (Table 5).^205,206

While the work to understand the role of mechanosensitive ion channels in tendons, ligaments, and menisci is promising, the scope of work is limited. There is still much to understand about the roles of these mechanosensitive ion channels in hierarchical fiber formation and maturation.

Other Mechanosensitive Cell Membrane Sensors

Primary cilia

The primary cilia of cells are microtubule-based organelles that sense force and transduce signals via the hedgehog pathway (Table 6). They are believed to be involved in the regulation of bone and cartilage^245,246 and are expressed in tenocytes, ligament fibroblasts, and meniscal fibrochondrocytes.^{184,238,247–251} Furthermore, they are found primarily in the outer region of the meniscus where hierarchical fibers dominate.¹⁸⁴ Primary cilia orient themselves parallel to the length of the collagen fibers and the direction of primary load,¹¹⁰ lengthening with reduced load to increase mechanosensation and shortening with cyclic loading.^109,129,160 Cilia length is also affected by hypoxia, shortening in hypoxic conditions, leading to a decreased response to mechanical strain.²³⁷ This suggests primary cilia have a smaller role in adult tissues, where tendons, ligaments, and menisci are in a more hypoxic environment with relatively constant cyclic load.^237,252 However, cilia may have a key role in wound healing and tissue generation, as stress-deprived tenocytes have been shown to lengthen their cilia as they attempt to re-establish tensional homeostasis both in 2D flexible membranes and ex vivo rat tail tendons.^109,129,160

Table 6.

Role of Mechanosensitive Cell Membrane Sensors in Cell Signaling, Collagen Fiber Formation, and Tissue Maturation

Treatment	Culture conditions	Cell type	Outcome
Primary cilia
Stress deprivation	Ex vivo rat tail tendon fascicles^109,129,160	Rat tail tenocytes^109,129,160	Increased ciliary length over the course of 24 h^109,129,160
Cyclic load	In vitro in 2D on a flexible membrane with up to 3% strain¹⁶⁰	Human semitendinosus tenocytes¹⁶⁰	Ciliary disassembly¹⁶⁰
Cyclic load	Ex vivo rat tail tendon fascicles at 3% strain^129,160	Rat tail tenocytes^129,160	Reduction of ciliary length compared with stress-deprived tendons, ciliary disassembly^129,160
Hypoxia	In vitro in 2D²³⁷	Rat tail tenocytes²³⁷	Reduced cilia elongation compared with normoxic conditions²³⁷
Reduced cilia function	In vitro in 2D culture with a scratch wound^133,238	Human periodontal ligament cells²³⁸	Reduced collagen alignment and cell proliferation²³⁸
	In vitro in 2D culture with siRNA targeting IFT88 + TRPV4 activation²⁰⁹	Human mesenchymal stem cells²⁰⁹	Reduced cell response to TRPV4 activation²⁰⁹
	In vivo in mouse supraspinatus tendon^167,239	Mouse supraspinatus tenocytes^167,239	Abnormal enthesis formation with reduced mineralization,²³⁹ impaired healing after injury¹⁶⁷
DDR1
Increased DDR1 expression	In vitro 2D culture²¹⁹	Mouse embryonic fibroblasts²¹⁹	Increased TRPV4 localization to FAs, increased NMMIIA activity, increased Ca²⁺ flux through TRPV4²¹⁹
Reduced DDR1 function	In vitro 2D culture²⁴⁰	Mouse NIH3T3 cells, mouse embryonic fibroblasts²⁴⁰	Reduced cell spreading and migration²⁴⁰
Reduced DDR1 function	In vivo in DDR1 KO mice^241,242	Mouse periodontal ligament,²⁴¹ spiral ligament fibroblasts²⁴²	Decreased collagen content and cellular contraction,²⁴¹ altered cell morphology, reduced auditory signal transduction²⁴²
Caveolins
Compressive load	In vitro in 2D culture on collagen-coated glass in a pressurized incubator²⁴³	Mouse chondrocytes²⁴³	Increased caveolin-1 phosphorylation, ERK1/2 activation²⁴³
Cav-1 siRNA + Cyclic tensile load + IL-1β treatment	In vitro 2D culture under cyclic tensile strains at 2%, 5%, or 12%²⁴⁴	Rat annulus fibrosis cells²⁴⁴	Decreased inflammation due to IL-1β with siRNA treatment, decreased integrin β1 expression²⁴⁴

DDR1, discoidin domain receptor 1; ERK1/2, extracellular signal-related kinase 1/2; IFT88, intraflagellar transport protein 88 homolog; IL-1β, interleukin 1β; KO, knockout; siRNA, small interfering ribonucleic acid; TRPV4, transient receptor potential vanilloid 4.

Primary cilia also play a major role in the enthesis.^167,238,239 Cilia are present in more cells in the tendon enthesis than the midsection and coincide with hedgehog signaling and enthesis mineralization.^167,238,239 Furthermore, primary cilia are critical to enthesis mechano-responsiveness in developing and adult tissue, with loading driving assembly and disassembly of cilia and removal of cilia causing significant structural and compositional deficits in the enthesis.²³⁹

Interestingly, primary cilia may also have crosstalk with TRPV4. It has been reported that TRPV4 localizes to the primary cilium of MSCs, and the effects of TRPV4 activation were reduced without functional primary cilia.²⁰⁹ Collectively, while explored in the enthesis, the role of primary cilia in hierarchical fiber formation is less established. More work is necessary to understand how it directly, or in combination with other mechanosensors, influences hierarchical fiber formation.

Discoidin domain receptors

Discoidin domain receptors (DDRs) are mechanosensitive collagen-binding tyrosine kinases with a possible role in fibril alignment (Table 6).²⁵³ DDR1 is a cell surface receptor that associates with NMMIIA upon binding to collagen, promoting cell migration and inhibiting spreading.²⁴⁰ This suggests interplay between DDR binding and actin-cytoskeletal pathways, aligning with research that indicates both have a role in collagen fibril alignment.^36,219,241 DDR1 also colocalizes with TRPV4 in mouse embryonic fibroblasts, and DDR1-collagen binding can induce TRPV4-mediated Ca²⁺ signaling.²¹⁹ This results in enhanced collagen binding and remodeling mediated by DDR1, which is lost if TRPV4 is inhibited.²¹⁹ The mechanosensitivity of DDR1 has been shown in the spiral ligament of the ear canal, where deletion of the DDR1 gene in mice altered outer hair cell shape.²⁴² The altered cell morphology was observed in the areas where DDR1 and NMMIIA colocalized in healthy mice, suggesting that altered DDR1 function results in alterations in actomyosin-mediated tensional homeostasis.²⁴²

Caveolae

Caveolae are mechanosensitive indentations in the cell membrane of several cell types (Table 6).^254,255 These indentations associate with actin and contain proteins such as cavin-1 and EH-containing domain 2 (EHD2), which are distributed into the cytoplasm and localize to the nucleus when caveolae are flattened due to stretching of the cell membrane.^172,254 Caveolae have been implicated in altering actin-cytoskeletal dynamics, as well as activation of downstream signaling pathways such as YAP/TAZ and RhoA.²⁵⁵ Caveolae are expressed in meniscal fibrochondrocytes and are again more numerous in the outer region of the meniscus where hierarchical fibers dominate, suggesting a role in responding to tension and regulation of collagen fibers.²⁵⁶ While the role of caveolae in meniscal mechanobiology has not been thoroughly explored, similar tissues have shown that caveolins such as Cav1 respond to compressive and cyclic tensile load and may contribute to cellular tensional homeostasis.^243,244 Despite the known association of caveolae and the actin-cytoskeleton, little is still known about their role in hierarchical fiber development or in tendons and ligaments, making them an enticing target for future research.

Conclusion

Our understanding of cell-driven hierarchical fiber formation is still rapidly evolving. There remain several research gaps and unanswered questions that are crucial to further our understanding, such as investigating how mechanical forces contribute to later stages of hierarchical fiber development, which mechanotransduction pathways contribute to tissue maturation via increased synthesis of different types of collagen, SLRPs, and LOX, and how various mechanosensation pathways interact with and alter one another. Most work has focused on the roles of actin-cytoskeletal mechanosensation in fiber development. While it is clear that actin-cytoskeletal-based mechanotransduction is a key mediator of fibril synthesis and tissue growth,^{119,126,157,257} it is becoming clearer that other mechanosensors are crucial to fiber maturation. For example, ion channels are now well established to play a role in modulating collagen synthesis and LOX crosslinking, both key to fiber maturation.^{196,199,210,213,214,219,258} Yet other mechanosensors such as primary cilia, DDRs, and caveolae are just beginning to be explored. All of these mechanosensation pathways, how they interact with each other, and how they influence downstream signaling pathways are exciting future research directions. Furthermore, as hierarchical fibers form, from embryonic development through adolescence, the loading environment, including the type, rate, and magnitude of load to which cells are exposed, will shift. We suspect this shifts the activity, role, or quantity of these mechanosensors and may suggest a need for adaptive stimulation or stage-targeted therapies to fully regenerate hierarchical fibers. In particular, we suspect that while actin-cytoskeletal sensing is critical to initial fibril and fiber formation, this sensing pathway will be less critical as fibers and fascicles mature and the cell experiences more secondary shear and compression. Here we may find a more dominant role of ion channels or other mechanotransduction pathways yet to be discovered.

Paramount to furthering our understanding of hierarchical fiber formation is understanding the mechanisms by which cells organize larger fibers and fascicles, which dominate the human musculoskeletal system. The majority of studies evaluating fiber formation are in 2D or rodent models. While mouse transgenic models are extremely powerful for exploring mechanisms, rodents largely lack the larger hierarchical organization of human musculoskeletal tissues,^17,49,259 limiting our ability to understand how cells regulate fascicle formation. Thus, there is a need to use larger animal models for in vivo and ex vivo analysis of fiber regulation and to develop engineered systems that can reliably reproduce cellular environments within these tissues to better understand the role of mechanotransduction in fascicle formation. Little is understood about the mechanisms by which cells mediate later stages of hierarchical development, and cells largely do not regenerate these fibers after injury or in engineered replacements. Understanding the mechanisms by which hierarchical fiber formation is mediated and maintained is crucial to regenerating fibers after injury or in engineered replacements, and will help to inform the pathophysiological effects of unloading and overloading, and will help inform the benefits of controlled loading during rehabilitation.^9,29,34

Authors’ Contributions

All authors have made substantial contributions to drafting and critical revision of the article and have read and approved the final submitted article.

Footnotes

Author Disclosure Statement

The authors declare they have no known competing financial interests or personal relationships that influenced the work reported in this article.

Funding Information

This work was supported in part by the National Institute of Arthritis and Musculoskeletal and Skin Disease of the NIH (R01 AR082365).

References

1. Zhang

, Young

, Ezura

, et al. Development of tendon structure and function: Regulation of collagen fibrillogenesis. J Musculoskelet Neuronal Interact 2005;5(1):5–21.

2. Rodrigues

, Reis

, Gomes

. Engineering tendon and ligament tissues: Present developments towards successful clinical products. J Tissue Eng Regen Med 2013;7(9):673–686; doi: 10.1002/TERM.1459

3. Hasan

, Fisher

, Ingham

. Current strategies in meniscal regeneration. J Biomed Mater Res B Appl Biomater 2014;102(3):619–634; doi: 10.1002/JBM.B.33030

4. Petersen

, Tillmann

. Collagenous fibril texture of the human knee joint menisci. Anat Embryol (Berl) 1998;197(4):317–324; doi: 10.1007/S004290050141

5. Irwin

, Brown

, Koff

, et al. Generating new meniscus therapies via recent breakthroughs in development, model systems, and clinical diagnostics. J Orthop Res 2025;43(6):1073–1089; doi: 10.1002/JOR.26066

6. Gautieri

, Vesentini

, Redaelli

, et al. Hierarchical structure and nanomechanics of collagen microfibrils from the atomistic scale up. Nano Lett 2011;11(2):757–766; doi: 10.1021/nl103943u

7. Buehler

. Nature designs tough collagen: Explaining the nanostructure of collagen fibrils. Proc Natl Acad Sci USA 2006;103(33):12285–12290; doi: 10.1073/pnas.0603216103

8. Li

, Qu

, Han

, et al. Micromechanical anisotropy and heterogeneity of the meniscus extracellular matrix. Acta Biomater 2017;54:356–366; doi: 10.1016/j.actbio.2017.02.043

9. Killian

, Cavinatto

, Galatz

, et al. The role of mechanobiology in tendon healing. J Shoulder Elbow Surg 2012;21(2):228–237; doi: 10.1016/J.JSE.2011.11.002

10.

10. Siadat

, Zamboulis

, Thorpe

, et al. Tendon extracellular matrix assembly, maintenance and dysregulation throughout life. Adv Exp Med Biol 2021;1348:45–103; doi: 10.1007/978-3-030-80614-9_3

11.

11. Svensson

, Smith

, Moyer

, et al. Effects of maturation and advanced glycation on tensile mechanics of collagen fibrils from rat tail and Achilles tendons. Acta Biomater 2018;70:270–280; doi: 10.1016/J.ACTBIO.2018.02.005

12.

12. Gohl

, Listrat

, Béchet

. Hierarchical mechanics of connective tissues: Integrating insights from nano to macroscopic studies. J Biomed Nanotechnol 2014;10(10):2464–2507.

13.

13. Connizzo

, Yannascoli

, Soslowsky

. Structure-function relationships of postnatal tendon development: A parallel to healing. Matrix Biol 2013;32(2):106–116; doi: 10.1016/J.MATBIO.2013.01.007

14.

14. Clark

, Ogden

. Prenatal and postnatal development of human knee joint menisci. Iowa Orthop J 1981;1:20–27.

15.

15. Clark

, Ogden

. Development of the menisci of the human knee joint. Morphological changes and their potential role in childhood meniscal injury. J Bone Joint Surg Am 1983;65(4):538–547.

16.

16. Clark

, Sidles

. The interrelation of fiber bundles in the anterior cruciate ligament. J Orthop Res 1990;8(2):180–188; doi: 10.1002/JOR.1100080205

17.

17. Lee

, Elliott

. Comparative multi-scale hierarchical structure of the tail, plantaris, and Achilles tendons in the rat. J Anat 2019;234(2):252–262; doi: 10.1111/JOA.12913

18.

18. Tsinman

, Huang

, Ahmed

, et al. Lack of skeletal muscle contraction disrupts fibrous tissue morphogenesis in the developing murine knee. J Orthop Res 2023;41(10):2305–2314; doi: 10.1002/JOR.25659

19.

19. Parry

DAD

, Barnes

GRG

, Craig

. A comparison of the size distribution of collagen fibrils in connective tissues as a function of age and a possible relation between fibril size distribution and mechanical properties. Proc R Soc Lond B Biol Sci 1978;203(1152):305–321; doi: 10.1098/RSPB.1978.0107

20.

20. Kalson

, Lu

, Taylor

, et al. A structure-based extracellular matrix expansion mechanism of fibrous tissue growth. Elife 2015;4:e05958; doi: 10.7554/ELIFE.05958

21.

21. Kalson

, Starborg

, Lu

, et al. Nonmuscle myosin II powered transport of newly formed collagen fibrils at the plasma membrane. Proc Natl Acad Sci U S A 2013;110(49):E4743–E4752; doi: 10.1073/pnas.1314348110

22.

22. Holmes

, Lu

, Starborg

, et al. Collagen Fibril Assembly and Function. Curr Top Dev Biol 2018;130:107–142; doi: 10.1016/BS.CTDB.2018.02.004

23.

23. Ryan

CNM

, Sorushanova

, Lomas

, et al. Glycosaminoglycans in tendon physiology, pathophysiology, and therapy. Bioconjug Chem 2015;26(7):1237–1251; doi: 10.1021/ACS.BIOCONJCHEM.5B00091

24.

24. Nau

, Teuschl

. Regeneration of the anterior cruciate ligament: Current strategies in tissue engineering. World J Orthop 2015;6(1):127–136; doi: 10.5312/wjo.v6.i1.127

25.

25. Kahan

, Burroughs

, Petit

, et al. Rates of subsequent surgeries after meniscus repair with and without concurrent anterior cruciate ligament reconstruction. PLoS One 2023;18(11):e0294964; doi: 10.1371/JOURNAL.PONE.0294964

26.

26. Sugimoto

, LeBlanc

, Wooley

, et al. The effectiveness of a functional knee brace on joint-position sense in anterior cruciate ligament–reconstructed individuals. J Sport Rehabil 2016;25(2):190–194; doi: 10.1123/JSR.2014-0226

27.

27. Buller

, Best

, Baraga

, et al. Trends in anterior cruciate ligament reconstruction in the United States. Orthop J Sports Med 2015;3(1):2325967114563664; doi: 10.1177/2325967114563664

28.

28. Jain

, Higgins

, Losina

, et al. Epidemiology of musculoskeletal upper extremity ambulatory surgery in the United States. BMC Musculoskelet Disord 2014;15(4):1–7; doi: 10.1186/1471-2474-15-4

29.

29. Magnusson

, Kjaer

, Petersen

, et al. The impact of loading, unloading, ageing and injury on the human tendon. J Physiol 2019;597(5):1283–1298; doi: 10.1113/JP275450

30.

30. Yang

, Rothrauff

, Tuan

. Tendon and ligament regeneration and repair: Clinical relevance and developmental paradigm. Birth Defects Res C Embryo Today 2013;99(3):203–222; doi: 10.1002/BDRC.21041

31.

31. Patel

, Caldwell

, Doty

, et al. Integrating soft and hard tissues via interface tissue engineering. J Orthop Res 2018;36(4):1069–1077; doi: 10.1002/jor.23810

32.

32. Bramson

MTK

, Van Houten

, Corr

. Mechanobiology in tendon, ligament, and skeletal muscle tissue engineering. J Biomech Eng 2021;143(7):e070801; doi: 10.1115/1.4050035

33.

33. Lopez

, Bonassar

. The role of SLRPs and large aggregating proteoglycans in collagen fibrillogenesis, extracellular matrix assembly, and mechanical function of fibrocartilage. Connect Tissue Res 2022;63(3):269–286; doi: 10.1080/03008207.2021.1903887

34.

34. Lavagnino

, Wall

, Little

, et al. Tendon mechanobiology: Current knowledge and future research opportunities. J Orthop Res 2015;33(6):813–822; doi: 10.1002/JOR.22871

35.

35. McNulty

, Guilak

. Mechanobiology of the meniscus. J Biomech 2015;48(8):1469–1478; doi: 10.1016/J.JBIOMECH.2015.02.008

36.

36. Canty

, Starborg

, Lu

, et al. Actin filaments are required for fibripositor-mediated collagen fibril alignment in tendon. J Biol Chem 2006;281(50):38592–38598; doi: 10.1074/jbc.M607581200

37.

37. Birk

, Nurminskaya

, Zycband

. Collagen fibrillogenesis in situ: Fibril segments undergo post-depositional modifications resulting in linear and lateral growth during matrix development. Dev Dyn 1995;202(3):229–243; doi: 10.1002/AJA.1002020303

38.

38. Pan

, Li

, Brown

, et al. Embryo movements regulate tendon mechanical property development. Philos Trans R Soc Lond B Biol Sci 2018;373(1759):20170325; doi: 10.1098/RSTB.2017.0325

39.

39. Schiele

, Marturano

, Kuo

. Mechanical factors in embryonic tendon development: Potential cues for stem cell tenogenesis. Curr Opin Biotechnol 2013;24(5):834–840; doi: 10.1016/J.COPBIO.2013.07.003

40.

40. Bayer

, Yeung

CYC

, Kadler

, et al. The initiation of embryonic-like collagen fibrillogenesis by adult human tendon fibroblasts when cultured under tension. Biomaterials 2010;31(18):4889–4897; doi: 10.1016/J.BIOMATERIALS.2010.02.062

41.

41. Maeda

, Sakabe

, Sunaga

, et al. Conversion of mechanical force into TGF-β-mediated biochemical signals. Curr Biol 2011;21(11):933–941; doi: 10.1016/J.CUB.2011.04.007

42.

42. Schweitzer

, Zelzer

, Volk

. Connecting muscles to tendons: Tendons and musculoskeletal development in flies and vertebrates. Development 2010;137(17):2807–2817; doi: 10.1242/DEV.047498

43.

43. Mikic

, Johnson

, Chhabra

, et al. Differential effects of embryonic immobilization on the development of fibrocartilaginous skeletal elements. J Rehabil Res Dev 2000;37(2):127–133.

44.

44. Wang

, Chen

, Zheng

, et al. In vitro loading models for tendon mechanobiology. J Orthop Res 2018;36(2):566–575; doi: 10.1002/jor.23752

45.

45. Galloway

, Lalley

, Shearn

. The role of mechanical loading in tendon development, maintenance, injury, and repair. J Bone Joint Surg Am 2013;95(17):1620–1628; doi: 10.2106/JBJS.L.01004

46.

46. Cone

, Barnes

, Howe

, et al. Age- and sex-specific differences in ACL and ACL bundle size during adolescent growth. J Orthop Res 2022;40(7):1613–1620; doi: 10.1002/JOR.25198

47.

47. Cone

, Piercy

, Lambeth

, et al. Tissue-specific changes in size and shape of the ligaments and tendons of the porcine knee during post-natal growth. PLoS One 2019;14(10):e0219637; doi: 10.1371/JOURNAL.PONE.0219637

48.

48. Pedrini‐Mille

, Pedrini

, Maynard

, et al. Response of immature chicken meniscus to strenuous exercise: Biochemical studies of proteoglycan and collagen. J Orthopaedic Res 1988;6(2):196–204; doi: 10.1002/JOR.1100060206

49.

49. Tsinman

, Jiang

, Han

, et al. Intrinsic and growth-mediated cell and matrix specialization during murine meniscus tissue assembly. FASEB J 2021;35(8):e21779; doi: 10.1096/FJ.202100499R

50.

50. Usami

, Jiang

, Dyment

, et al. Limb motility and ambulation as mechanical cues in postnatal murine tendon development. Dev Biol 2025;525:130–138; doi: 10.1016/J.YDBIO.2025.05.020

51.

51. Schwartz

, Lipner

, Pasteris

, et al. Muscle loading is necessary for the formation of a functional tendon enthesis. Bone 2013;55(1):44–51; doi: 10.1016/J.BONE.2013.03.010

52.

52. Ansorge

, Adams

, Birk

, et al. Mechanical, compositional, and structural properties of the post-natal mouse Achilles tendon. Ann Biomed Eng 2011;39(7):1904–1913; doi: 10.1007/s10439-011-0299-0

53.

53. Fukazawa

, Hatta

, Uchio

, et al. Development of the meniscus of the knee joint in human fetuses. Congenit Anom (Kyoto) 2009;49(1):27–32; doi: 10.1111/J.1741-4520.2008.00216.X

54.

54. Canty

, Lu

, Meadows

, et al. Coalignment of plasma membrane channels and protrusions (fibripositors) specifies the parallelism of tendon. J Cell Biol 2004;165(4):553–563; doi: 10.1083/JCB.200312071

55.

55. Ansorge

, Adams

, Jawad

, et al. Mechanical property changes during neonatal development and healing using a multiple regression model. J Biomech 2012;45(7):1288–1292; doi: 10.1016/J.JBIOMECH.2012.01.030

56.

56. Chen

, Birk

. The regulatory roles of small leucine-rich proteoglycans in extracellular assembly. FEBS J 2013;280(10):2120–2137; doi: 10.1111/FEBS.12136

57.

57. Han

, Heo

, Driscoll

, et al. Macro- to microscale strain transfer in fibrous tissues is heterogeneous and tissue-specific. Biophys J 2013;105(3):807–817; doi: 10.1016/J.BPJ.2013.06.023

58.

58. Qu

, Subramony

, Boskey

, et al. Compositional mapping of the mature anterior cruciate ligament-to-bone insertion. J Orthop Res 2017;35(11):2513–2523; doi: 10.1002/JOR.23539

59.

59. Vogel

, Ördög

, Pogány

, et al. Proteoglycans in the compressed region of human tibialis posterior tendon and in ligaments. J Orthop Res 1993;11(1):68–77; doi: 10.1002/JOR.1100110109

60.

60. Wang

INE

, Mitroo

, Chen

, et al. Age-dependent changes in matrix composition and organization at the ligament-to-bone insertion. J Orthop Res 2006;24(8):1745–1755; doi: 10.1002/JOR.20149

61.

61. Benjamin

, Toumi

, Ralphs

, et al. Where tendons and ligaments meet bone: Attachment sites (‘entheses’) in relation to exercise and/or mechanical load. J Anat 2006;208(4):471–490; doi: 10.1111/J.1469-7580.2006.00540.X

62.

62. Melrose

, Smith

, Cake

, et al. Comparative spatial and temporal localisation of perlecan, aggrecan and type I, II and IV collagen in the ovine meniscus: An ageing study. Histochem Cell Biol 2005;124(3–4):225–235; doi: 10.1007/S00418-005-0005-0

63.

63. Eyre

, Paz

, Gallop

. Cross-Linking in Collagen and Elastin. Annu Rev Biochem 1984;53(1):717–748; doi: 10.1146/annurev.bi.53.070184.003441

64.

64. Depalle

, Qin

, Shefelbine

, et al. Influence of cross-link structure, density and mechanical properties in the mesoscale deformation mechanisms of collagen fibrils. J Mech Behav Biomed Mater 2015;52:1–13; doi: 10.1016/j.jmbbm.2014.07.008

65.

65. Reiser

, Mccormick

, Rucker

. Enzymatic and nonensymatic cross-linking of collagen and elastin. Faseb J 1992;6(7):2439–2449; doi: 10.1096/fasebj.6.7.1348714

66.

66. Ellingson

, Pancheri

, Schiele

. Regulators of collagen crosslinking in developing and adult tendons. Eur Cell Mater 2022;43:130–152; doi: 10.22203/eCM.v043a11

67.

67. Marturano

, Xylas

, Sridharan

, et al. Lysyl oxidase-mediated collagen crosslinks may be assessed as markers of functional properties of tendon tissue formation. Acta Biomater 2014;10(3):1370–1379; doi: 10.1016/j.actbio.2013.11.024

68.

68. Bates

, Troop

, Brown

, et al. Temporal application of lysyl oxidase during hierarchical collagen fiber formation differentially effects tissue mechanics. Acta Biomater 2023;160:98–111; doi: 10.1016/J.ACTBIO.2023.02.024

69.

69. Leiphart

, Weiss

, DiStefano

, et al. Collagen V deficiency during murine tendon healing results in distinct healing outcomes based on knockdown severity. J Biomech 2022;144:111315; doi: 10.1016/J.JBIOMECH.2022.111315

70.

70. Izu

, Adams

, Connizzo

, et al. Collagen XII mediated cellular and extracellular mechanisms regulate establishment of tendon structure and function. Matrix Biol 2021;95:52–67; doi: 10.1016/J.MATBIO.2020.10.004

71.

71. Leiphart

, Pham

, Harvey

, et al. Coordinate roles for collagen VI and biglycan in regulating tendon collagen fibril structure and function. Matrix Biol Plus 2022;13:100099; doi: 10.1016/J.MBPLUS.2021.100099

72.

72. Sun

, Luo

, Adams

, et al. Collagen XI regulates the acquisition of collagen fibril structure, organization and functional properties in tendon. Matrix Biol 2020;94:77–94; doi: 10.1016/J.MATBIO.2020.09.001

73.

73. Izu

, Ansorge

, Zhang

, et al. Dysfunctional tendon collagen fibrillogenesis in collagen VI null mice. Matrix Biol 2011;30(1):53–61; doi: 10.1016/J.MATBIO.2010.10.001

74.

74. Carlson

, Shetye

, Sun

, et al. Collagen V haploinsufficiency in female murine patellar tendons results in altered matrix engagement and cellular density, demonstrating decreased healing. J Orthop Res 2024;42(5):950–960; doi: 10.1002/JOR.25740

75.

75. Connizzo

, Freedman

, Fried

, et al. Regulatory role of collagen V in establishing mechanical properties of tendons and ligaments is tissue-dependent. J Orthop Res 2015;33(6):882–888; doi: 10.1002/JOR.22893

76.

76. Connizzo

, Han

, Birk

, et al. Collagen V-heterozygous and -null supraspinatus tendons exhibit altered dynamic mechanical behaviour at multiple hierarchical scales. Interface Focus 2016;6(1):20150043; doi: 10.1098/RSFS.2015.0043

77.

77. Buckley

, Evans

, Matuszewski

, et al. Distributions of Types I, II and III collagen by region in the human supraspinatus tendon. Connect Tissue Res 2013;54(6):374–379; doi: 10.3109/03008207.2013.847096

78.

78. Cohen

, Fung

, Stein

, et al. Tendon-targeted knockout of collagen XI disrupts patellar and Achilles tendon structure and mechanical properties during murine postnatal development. Connect Tissue Res 2024;65(6):497–510; doi: 10.1080/03008207.2024.2432324

79.

79. Ye

, Shetye

, Birk

, et al. Regulatory role of collagen Xi in the establishment of mechanical properties of tendons and ligaments in mice is tissue dependent. J Biomech Eng 2025;147(1):e011003; doi: 10.1115/1.4066570

80.

80. Johnston

, Connizzo

, Shetye

, et al. Collagen V haploinsufficiency in a murine model of classic Ehlers-Danlos syndrome is associated with deficient structural and mechanical healing in tendons. J Orthop Res 2017;35(12):2707–2715; doi: 10.1002/JOR.23571

81.

81. Ansorge

, Meng

, Zhang

, et al. Type XIV collagen regulates fibrillogenesis: Premature collagen fibril growth and tissue dysfunction in null mice. J Biol Chem 2009;284(13):8427–8438; doi: 10.1074/JBC.M805582200

82.

82. Fung

, Sun

, Soslowsky

, et al. Targeted conditional collagen XII deletion alters tendon function. Matrix Biol Plus 2022;16:100123; doi: 10.1016/J.MBPLUS.2022.100123

83.

83. Herchenhan

, Uhlenbrock

, Eliasson

, et al. Lysyl oxidase activity is required for ordered collagen fibrillogenesis by tendon cells. J Biol Chem 2015;290(26):16440–16450; doi: 10.1074/jbc.M115.641670

84.

84. Wang

, Fan

, Heo

, et al. Structure, mechanics, and mechanobiology of fibrocartilage pericellular matrix mediated by type V collagen. Adv Sci (Weinh) 2025;12(32):e14750; doi: 10.1002/ADVS.202414750

85.

85. Franchi

, Trirè

, Quaranta

, et al. Collagen structure of tendon relates to function. ScientificWorldJournal 2007;7:404–420; doi: 10.1100/TSW.2007.92

86.

86. Stouffer

, Butler

, Hosny

. The relationship between crimp pattern and mechanical response of human patellar tendon-bone units. J Biomech Eng 1985;107(2):158–165; doi: 10.1115/1.3138536

87.

87. Shah

, Palacios

. Development of crimp morphology and cellular changes in chick tendons. Dev Biol 1982;94(2):499–504; doi: 10.1016/0012-1606(82)90366-9

88.

88. Lavagnino

, Brooks

, Oslapas

, et al. Crimp length decreases in lax tendons due to cytoskeletal tension, but is restored with tensional homeostasis. J Orthop Res 2017;35(3):573–579; doi: 10.1002/JOR.23489

89.

89. Herchenhan

, Kalson

, Holmes

, et al. Tenocyte contraction induces crimp formation in tendon-like tissue. Biomech Model Mechanobiol 2012;11(3–4):449–459; doi: 10.1007/S10237-011-0324-0

90.

90. Järvinen

TAH

, Jozsa

, Kannus

, et al. Mechanical loading regulates tenascin-C expression in the osteotendinous junction. J Cell Sci 1999;112 Pt 18(18):3157–3166; doi: 10.1242/JCS.112.18.3157

91.

91. Pancheri

, Daw

, Ditton

, et al. The LINC complex regulates tendon elastic modulus, collagen crimp, and lateral expansion during early postnatal development. J Orthop Res 2025;43(6):1090–1100; doi: 10.1002/JOR.26069

92.

92. Andarawis-Puri

, Flatow

, Soslowsky

. Tendon basic science: Development, repair, regeneration, and healing. J Orthop Res 2015;33(6):780–784; doi: 10.1002/jor.22869

93.

93. Maurer

, Lammerding

. The Driving Force: Nuclear mechanotransduction in cellular function, fate, and disease. Annu Rev Biomed Eng 2019;21:443–468; doi: 10.1146/ANNUREV-BIOENG-060418-052139

94.

94. Dupont

, Morsut

, Aragona

, et al. Role of YAP/TAZ in mechanotransduction. Nature 2011;474(7350):179–183; doi: 10.1038/NATURE10137

95.

95. Amano

, Nakayama

, Kaibuchi

. Rho-Kinase/ROCK: A Key Regulator of the Cytoskeleton and Cell Polarity. Cytoskeleton (Hoboken) 2010;67(9):545–554; doi: 10.1002/CM.20472

96.

96. Banes

, Horesovsky

, Larson

, et al. Mechanical load stimulates expression of novel genes in vivo and in vitro in avian flexor tendon cells. Osteoarthritis Cartilage 1999;7(1):141–153; doi: 10.1053/JOCA.1998.0169

97.

97. Holmes

, Yeung

CYC

, Garva

, et al. Synchronized mechanical oscillations at the cell–matrix interface in the formation of tensile tissue. Proc Natl Acad Sci U S A 2018;115(40):E9288–E9297; doi: 10.1073/pnas.1801759115

98.

98. Lavagnino

, Arnoczky

. In vitro alterations in cytoskeletal tensional homeostasis control gene expression in tendon cells. J Orthop Res 2005;23(5):1211–1218; doi: 10.1016/J.ORTHRES.2005.04.001

99.

99. Bonnevie

, Gullbrand

, Ashinsky

, et al. Aberrant mechanosensing in injured intervertebral discs as a result of boundary-constraint disruption and residual-strain loss. Nat Biomed Eng 2019;3(12):998–1008; doi: 10.1038/s41551-019-0458-4

100.

100. Schiele

, Von Flotow

, Tochka

, et al. Actin cytoskeleton contributes to the elastic modulus of embryonic tendon during early development. J Orthop Res 2015;33(6):874–881; doi: 10.1002/JOR.22880

101.

101. Sakar

, Eyckmans

, Pieters

, et al. Cellular forces and matrix assembly coordinate fibrous tissue repair. Nat Commun 2016;7(1):11036–11038; doi: 10.1038/ncomms11036

102.

102. Arnoczky

, Lavagnino

, Egerbacher

, et al. Loss of homeostatic strain alters mechanostat “set point” of tendon cells in vitro. Clin Orthop Relat Res 2008;466(7):1583–1591; doi: 10.1007/S11999-008-0264-X

103.

103. Gardner

, Lavagnino

, Egerbacher

, et al. Re-establishment of cytoskeletal tensional homeostasis in lax tendons occurs through an actin-mediated cellular contraction of the extracellular matrix. J Orthop Res 2012;30(11):1695–1701; doi: 10.1002/JOR.22131

104.

104. Freedman

, Rodriguez

, Leiphart

, et al. Dynamic Loading and Tendon Healing Affect Multiscale Tendon Properties and ECM Stress Transmission. Sci Rep 2018;8(1):10854; doi: 10.1038/S41598-018-29060-Y

105.

105. Henshaw

, Attia

, Bhargava

, et al. Canine ACL fibroblast integrin expression and cell alignment in response to cyclic tensile strain in three-dimensional collagen gels. J Orthop Res 2006;24(3):481–490; doi: 10.1002/JOR.20050

106.

106. Kalson

, Holmes

, Herchenhan

, et al. Slow stretching that mimics embryonic growth rate stimulates structural and mechanical development of tendon-like tissue in vitro. Dev Dyn 2011;240(11):2520–2528; doi: 10.1002/DVDY.22760

107.

107. Screen

HRC

, Lee

, Bader

, et al. An investigation into the effects of the hierarchical structure of tendon fascicles on micromechanical properties. Proc Inst Mech Eng H 2004;218(2):109–119; doi: 10.1243/095441104322984004

108.

108. Theodossiou

, Murray

, Schiele

. Cell-cell junctions in developing and adult tendons. Tissue Barriers 2020;8(1):1695491; doi: 10.1080/21688370.2019.1695491

109.

109. Lavagnino

, Gardner

, Sedlak

, et al. Tendon cell ciliary length as a biomarker of in situ cytoskeletal tensional homeostasis. Muscles Ligaments Tendons J 2013;3(3):118–121.

110.

110. Donnelly

, Ascenzi

, Farnum

. Primary cilia are highly oriented with respect to collagen direction and long axis of extensor tendon. J Orthop Res 2010;28(1):77–82; doi: 10.1002/JOR.20946

111.

111. Sun

, Guo

, Fässler

. Integrin-mediated mechanotransduction. J Cell Biol 2016;215(4):445–456; doi: 10.1083/JCB.201609037

112.

112. Cary

, Guan

. Focal adhesion kinase in integrin-mediated signaling. Front Biosci 1999;4(5):D102–D113; doi: 10.2741/CARY

113.

113. Kuo

. Mechanotransduction at focal adhesions: Integrating cytoskeletal mechanics in migrating cells. J Cell Mol Med 2013;17(6):704–712; doi: 10.1111/JCMM.12054

114.

114. Wang

, Dembo

, Hanks

, New Collective Author. et al.; Focal adhesion kinase is involved in mechanosensing during fibroblast migration. Proc Natl Acad Sci U S A 2001;98(20):11295–11300; doi: 10.1073/PNAS.201201198

115.

115. Wang

, Butler

, Ingber

, New Collective Author. Mechanotransduction across the cell surface and through the cytoskeleton. Science 1993;260(5111):1124–1127; doi: 10.1126/SCIENCE.7684161

116.

116. Wolfenson

, Bershadsky

, Henis

, et al. Actomyosin-generated tension controls the molecular kinetics of focal adhesions. J Cell Sci 2011;124(Pt 9):1425–1432; doi: 10.1242/JCS.077388

117.

117. Wolfenson

, Henis

, Geiger

, et al. The heel and toe of the cell’s foot: A multifaceted approach for understanding the structure and dynamics of focal adhesions. Cell Motil Cytoskeleton 2009;66(11):1017–1029; doi: 10.1002/CM.20410

118.

118. Geiger

, Spatz

, Bershadsky

. Environmental sensing through focal adhesions. Nat Rev Mol Cell Biol 2009;10(1):21–33; doi: 10.1038/NRM2593

119.

119. Leahy

, Chenna

, Soslowsky

, et al. Focal adhesion kinase regulates tendon cell mechanoresponse and physiological tendon development. FASEB J 2024;38(17):e70050; doi: 10.1096/FJ.202400151R

120.

120. Kronenberg

, Michel

, Hochstrat

, et al. Increased collagen turnover impairs tendon microstructure and stability in integrin α2β1-deficient mice. Int J Mol Sci 2020;21(8):2835–2815; doi: 10.3390/IJMS21082835

121.

121. Wang

, Liu

, Yu

, et al. Inhibition of integrin αvβ6 activation of TGF-β attenuates tendinopathy. Adv Sci (Weinh) 2022;9(11):e2104469; doi: 10.1002/ADVS.202104469

122.

122. Galbraith

, Yamada

, Sheetz

. The relationship between force and focal complex development. J Cell Biol 2002;159(4):695–705; doi: 10.1083/JCB.200204153

123.

123. Mitra

, Hanson

, Schlaepfer

. Focal adhesion kinase: In command and control of cell motility. Nat Rev Mol Cell Biol 2005;6(1):56–68; doi: 10.1038/NRM1549

124.

124. Moser

, Legate

, Zent

, et al. The tail of integrins, talin, and kindlins. Science 2009;324(5929):895–899; doi: 10.1126/SCIENCE.1163865

125.

125. Jones

, Hallström

, Jiang

, et al. Mechanoepigenetic regulation of extracellular matrix homeostasis via Yap and Taz. Proc Natl Acad Sci U S A 2023;120(22):e2211947120; doi: 10.1073/pnas.2211947120

126.

126. Rajshankar

, Wang

, Worndl

, et al. Focal adhesion kinase regulates tractional collagen remodeling, matrix metalloproteinase expression, and collagen structure, which in turn affects matrix-induced signaling. J Cell Physiol 2020;235(3):3096–3111; doi: 10.1002/JCP.29215

127.

127. Bayer

, Schjerling

, Herchenhan

, et al. Release of tensile strain on engineered human tendon tissue disturbs cell adhesions, changes matrix architecture, and induces an inflammatory phenotype. PLoS One 2014;9(1):e86078; doi: 10.1371/JOURNAL.PONE.0086078

128.

128. Lavagnino

, Arnoczky

, Egerbacher

, et al. Isolated fibrillar damage in tendons stimulates local collagenase mRNA expression and protein synthesis. J Biomech 2006;39(13):2355–2362; doi: 10.1016/J.JBIOMECH.2005.08.008

129.

129. Gardner

, Arnoczky

, Lavagnino

. Effect of in vitro stress-deprivation and cyclic loading on the length of tendon cell cilia in situ. J Orthop Res 2011;29(4):582–587; doi: 10.1002/JOR.21271

130.

130. Arnoczky

, Tian

, Lavagnino

, et al. Ex vivo static tensile loading inhibits MMP-1 expression in rat tail tendon cells through a cytoskeletally based mechanotransduction mechanism. J Orthop Res 2004;22(2):328–333; doi: 10.1016/S0736-0266(03)00185-2

131.

131. Connizzo

, Grodzinsky

. Lose-Dose administration of dexamethasone is beneficial in preventing secondary tendon damage in a stress-deprived joint injury explant model. J Orthop Res 2020;38(1):139–149; doi: 10.1002/JOR.24451

132.

132. Connizzo

, Piet

, Shefelbine

, et al. Age-associated changes in the response of tendon explants to stress deprivation is sex-dependent. Connect Tissue Res 2020;61(1):48–62; doi: 10.1080/03008207.2019.1648444

133.

133. Thomopoulos

, Fomovsky

, Holmes

. The development of structural and mechanical anisotropy in fibroblast populated collagen gels. J Biomech Eng 2005;127(5):742–750; doi: 10.1115/1.1992525

134.

134. Vader

, Kabla

, Weitz

, et al. Strain-induced alignment in collagen gels. PLoS One 2009;4(6):e5902; doi: 10.1371/JOURNAL.PONE.0005902

135.

135. Maeda

, Ye

, Wang

, et al. Gap junction permeability between tenocytes within tendon fascicles is suppressed by tensile loading. Biomech Model Mechanobiol 2012;11(3–4):439–447; doi: 10.1007/S10237-011-0323-1

136.

136. Breidenbach

, Dyment

, Lu

, et al. Fibrin gels exhibit improved biological, structural, and mechanical properties compared with collagen gels in cell-based tendon tissue-engineered constructs. Tissue Eng Part A 2015;21(3–4):438–450; doi: 10.1089/TEN.TEA.2013.0768

137.

137. Herchenhan

, Bayer

, Svensson

, et al. In vitro tendon tissue development from human fibroblasts demonstrates collagen fibril diameter growth associated with a rise in mechanical strength. Dev Dyn 2013;242(1):2–8; doi: 10.1002/DVDY.23896

138.

138. Paxton

, Grover

, Baar

. Engineering an in vitro model of a functional ligament from bone to bone. Tissue Eng Part A 2010;16(11):3515–3525; doi: 10.1089/TEN.TEA.2010.0039

139.

139. Bowles

, Williams

, Zipfel

, et al. Self-assembly of aligned tissue-engineered annulus fibrosus and intervertebral disc composite via collagen gel contraction. Tissue Eng Part A 2010;16(4):1339–1348; doi: 10.1089/TEN.TEA.2009.0442

140.

140. Bell

, Ivarsson

, Merrill

. Production of a tissue-like structure by contraction of collagen lattices by human fibroblasts of different proliferative potential in vitro. Proc Natl Acad Sci U S A 1979;76(3):1274–1278; doi: 10.1073/PNAS.76.3.1274

141.

141. Chen

, Vigliotti

, Bacca

, et al. Role of boundary conditions in determining cell alignment in response to stretch. Proc Natl Acad Sci U S A 2018;115(5):986–991; doi: 10.1073/pnas.1715059115

142.

142. Kaji

, Montero

, Patel

, et al. Transcriptional profiling of mESC-derived tendon and fibrocartilage cell fate switch. Nat Commun 2021;12(1):4208; doi: 10.1038/S41467-021-24535-5

143.

143. Nirmalanandhan

, Rao

, Sacks

, et al. Effect of length of the engineered tendon construct on its structure–function relationships in culture. J Biomech 2007;40(11):2523–2529; doi: 10.1016/J.JBIOMECH.2006.11.016

144.

144. Puetzer

, Koo

, Bonassar

. Induction of fiber alignment and mechanical anisotropy in tissue engineered menisci with mechanical anchoring. J Biomech 2015;48(8):1436–1443; doi: 10.1016/J.JBIOMECH.2015.02.033

145.

145. Puetzer

, Ma

, Sallent

, et al. Driving hierarchical collagen fiber formation for functional tendon, ligament, and meniscus replacement. Biomaterials 2021;269:120527; doi: 10.1016/J.BIOMATERIALS.2020.120527

146.

146. McCorry

, Mansfield

, Sha

, et al. A model system for developing a tissue engineered meniscal enthesis. Acta Biomater 2017;56:110–117; doi: 10.1016/J.ACTBIO.2016.10.040

147.

147. Kim

, Boys

, Estroff

, et al. Combining TGF-β1 and mechanical anchoring to enhance collagen fiber formation and alignment in tissue-engineered menisci. ACS Biomater Sci Eng 2021;7(4):1608–1620; doi: 10.1021/ACSBIOMATERIALS.0C01791

148.

148. McCorry

, Bonassar

. Fiber development and matrix production in tissue engineered menisci using bovine mesenchymal stem cells and fibrochondrocytes. Connect Tissue Res 2017;58(3–4):329–341; doi: 10.1080/03008207.2016.1267152

149.

149. Brown

, Puetzer

. Driving native-like zonal enthesis formation in engineered ligaments using mechanical boundary conditions and β-tricalcium phosphate. Acta Biomater 2022;140:700–716; doi: 10.1016/J.ACTBIO.2021.12.020

150.

150. Wang

, Pearson

, Kutys

, et al. Extracellular matrix alignment dictates the organization of focal adhesions and directs uniaxial cell migration. APL Bioeng 2018;2(4):e046107; doi: 10.1063/1.5052239

151.

151. Costa

, Lee

, Holmes

. Creating alignment and anisotropy in engineered heart tissue: Role of boundary conditions in a model three-dimensional culture system. Tissue Eng 2003;9(4):567–577; doi: 10.1089/107632703768247278

152.

152. Loerakker

, Obbink-Huizer

, Baaijens

FPT

. A physically motivated constitutive model for cell-mediated compaction and collagen remodeling in soft tissues. Biomech Model Mechanobiol 2014;13(5):985–1001; doi: 10.1007/s10237-013-0549-1

153.

153. Cross

, Zheng

, Won Choi

, et al. Dense type I collagen matrices that support cellular remodeling and microfabrication for studies of tumor angiogenesis and vasculogenesis in vitro. Biomaterials 2010;31(33):8596–8607; doi: 10.1016/J.BIOMATERIALS.2010.07.072

154.

154. Puetzer

, Bonassar

. High density type I collagen gels for tissue engineering of whole menisci. Acta Biomater 2013;9(8):7787–7795; doi: 10.1016/J.ACTBIO.2013.05.002

155.

155. Ziegler

, Alonso

, Steinberg

, et al. Mechano-transduction in periodontal ligament cells identifies activated states of MAP-kinases p42/44 and p38-stress kinase as a mechanism for MMP-13 expression. BMC Cell Biol 2010;11:10–14; doi: 10.1186/1471-2121-11-10

156.

156. Archambault

, Tsuzaki

, Herzog

, et al. Stretch and interleukin-1beta induce matrix metalloproteinases in rabbit tendon cells in vitro. J Orthop Res 2002;20(1):36–39; doi: 10.1016/S0736-0266(01)00075-4

157.

157. Wen

, Blume

, Sumpio

. Role of integrins and focal adhesion kinase in the orientation of dermal fibroblasts exposed to cyclic strain. Int Wound J 2009;6(2):149–158; doi: 10.1111/J.1742-481X.2009.00591.X

158.

158. Xu

, Fan

, Shan

, et al. Cyclic stretch influenced expression of membrane connexin 43 in human periodontal ligament cell. Arch Oral Biol 2012;57(12):1602–1608; doi: 10.1016/J.ARCHORALBIO.2012.07.002

159.

159. Yang

, Lu

, Chen

, et al. Mechanical strain induces Cx43 expression in spinal ligament fibroblasts derived from patients presenting ossification of the posterior longitudinal ligament. Eur Spine J 2011;20(9):1459–1465; doi: 10.1007/S00586-011-1767-9

160.

160. Rowson

, Shelton

, Screen

HRC

, et al. Mechanical loading induces primary cilia disassembly in tendon cells via TGFβ and HDAC6. Sci Rep 2018;8(1):11107; doi: 10.1038/S41598-018-29502-7

161.

161. Baker

, Shah

, Huang

, et al. Dynamic tensile loading improves the functional properties of mesenchymal stem cell-laden nanofiber-based fibrocartilage. Tissue Eng Part A 2011;17(9–10):1445–1455; doi: 10.1089/TEN.TEA.2010.0535

162.

162. Han

, Heo

, Driscoll

, et al. Impact of cellular microenvironment and mechanical perturbation on calcium signalling in meniscus fibrochondrocytes. Eur Cell Mater 2014;27:321–331; doi: 10.22203/ECM.V027A23

163.

163. Jones

, Jones

, Legerlotz

, et al. Cyclical strain modulates metalloprotease and matrix gene expression in human tenocytes via activation of TGFβ. Biochim Biophys Acta 2013;1833(12):2596–2607; doi: 10.1016/J.BBAMCR.2013.06.019

164.

164. Troop

, Puetzer

. Intermittent cyclic stretch of engineered ligaments drives hierarchical collagen fiber maturation in a dose- and organizational-dependent manner. Acta Biomater 2024;185:296–311; doi: 10.1016/J.ACTBIO.2024.07.025

165.

165. Brown

, Puetzer

. Enthesis maturation in engineered ligaments is differentially driven by loads that mimic slow growth elongation and rapid cyclic muscle movement. Acta Biomater 2023;172:106–122; doi: 10.1016/J.ACTBIO.2023.10.012

166.

166. Ralphs

, Waggett

, Benjamin

. Actin stress fibres and cell-cell adhesion molecules in tendons: Organisation in vivo and response to mechanical loading of tendon cells in vitro. Matrix Biol 2002;21(1):67–74; doi: 10.1016/S0945-053X(01)00179-2

167.

167. Xiao

, Zhang

, Li

, et al. Mechanical stimulation promotes enthesis injury repair by mobilizing Prrx1+ cells via ciliary TGF-β signaling. Elife 2022;11:e73614–e23; doi: 10.7554/ELIFE.73614

168.

168. Huey

, Athanasiou

. Tension-compression loading with chemical stimulation results in additive increases to functional properties of anatomic meniscal constructs. PLoS One 2011;6(11):e27857; doi: 10.1371/JOURNAL.PONE.0027857

169.

169. Puetzer

, Bonassar

. Physiologically distributed loading patterns drive the formation of zonally organized collagen structures in tissue-engineered meniscus. Tissue Eng Part A 2016;22(13–14):907–916; doi: 10.1089/ten.tea.2015.0519

170.

170. Dyment

, Barrett

, Awad

, et al. A brief history of tendon and ligament bioreactors: Impact and future prospects. J Orthop Res 2020;38(11):2318–2330; doi: 10.1002/JOR.24784

171.

171. Vanderploeg

, Imler

, Brodkin

, et al. Oscillatory tension differentially modulates matrix metabolism and cytoskeletal organization in chondrocytes and fibrochondrocytes. J Biomech 2004;37(12):1941–1952; doi: 10.1016/J.JBIOMECH.2004.02.048

172.

172. Ma

, Vyhlidal

, Li

, et al. Mechano-bioengineering of the knee meniscus. Am J Physiol Cell Physiol 2022;323(6):C1652–C1663; doi: 10.1152/ajpcell.00336.2022

173.

173. Gögele

, Hoffmann

, Konrad

, et al. Cyclically stretched ACL fibroblasts emigrating from spheroids adapt their cytoskeleton and ligament-related expression profile. Cell Tissue Res 2021;384(3):675–690; doi: 10.1007/S00441-021-03416-9

174.

174. Zhang

, Chen

, Wang

, et al. Orchestrated biomechanical, structural, and biochemical stimuli for engineering anisotropic meniscus. Sci Transl Med 2019;11(487):eaao0750; doi: 10.1126/SCITRANSLMED.AAO0750

175.

175. Passini

, Jaeger

, Saab

, et al. Shear-stress sensing by PIEZO1 regulates tendon stiffness in rodents and influences jumping performance in humans. Nat Biomed Eng 2021;5(12):1457–1471; doi: 10.1038/S41551-021-00716-X

176.

176. Screen

HRC

, Bader

, Lee

, et al. Local strain measurement within tendon. Strain 2004;40(4):157–163; doi: 10.1111/J.1475-1305.2004.00164.X

177.

177. Yamaoka

, Sawa

, Ebata

, et al. Cultured periodontal ligament fibroblasts express diverse connexins. Tissue Cell 2002;34(6):375–380; doi: 10.1016/S0040816602000381

178.

178. Willecke

, Eiberger

, Degen

, et al. Structural and functional diversity of connexin genes in the mouse and human genome. Biol Chem 2002;383(5):725–737; doi: 10.1515/BC.2002.076

179.

179. Shen

, Schwartz

, Civitelli

, et al. Connexin 43 is necessary for murine tendon enthesis formation and response to loading. J Bone Miner Res 2020;35(8):1494–1503; doi: 10.1002/JBMR.4018

180.

180. Waggett

, Benjamin

, Ralphs

. Connexin 32 and 43 gap junctions differentially modulate tenocyte response to cyclic mechanical load. Eur J Cell Biol 2006;85(11):1145–1154; doi: 10.1016/J.EJCB.2006.06.002

181.

181. Kato

, Ishihara

, Kawanabe

, et al. Gap-junction-mediated communication in human periodontal ligament cells. J Dent Res 2013;92(7):635–640; doi: 10.1177/0022034513489992

182.

182. Mcneilly

, Banes

, Benjamin

, et al. Tendon cells in vivo form a three dimensional network of cell processes linked by gap junctions. J Anat 1996;189(Pt 3):593.

183.

183. Young

, Becker

, Fleck

, et al. Maturational alterations in gap junction expression and associated collagen synthesis in response to tendon function. Matrix Biol 2009;28(6):311–323; doi: 10.1016/J.MATBIO.2009.05.002

184.

184. Hellio Le Graverand

, Ou

, Schield-Yee

, et al. The cells of the rabbit meniscus: Their arrangement, interrelationship, morphological variations and cytoarchitecture. J Anat 2001;198(Pt 5):525–535; doi: 10.1046/J.1469-7580.2000.19850525.X

185.

185. Chi

, Rattner

, Sciore

, et al. Gap junctions of the medial collateral ligament: Structure, distribution, associations and function. J Anat 2005;207(2):145–154; doi: 10.1111/J.1469-7580.2005.00440.X

186.

186. Maeda

, Kimura

, Yamada

, et al. Enhanced gap junction intercellular communication inhibits catabolic and pro-inflammatory responses in tenocytes against heat stress. J Cell Commun Signal 2017;11(4):369–380; doi: 10.1007/S12079-017-0397-3

187.

187. Hirashima

, Ohta

, Togo

, et al. Mesoscopic structural analysis via deep learning processing, with a special reference to in vitro alteration in collagen fibre induced by a gap junction inhibitor. Microscopy (Oxf) 2023;72(1):18–26; doi: 10.1093/JMICRO/DFAC044

188.

188. Maeda

, Pian

, Ohashi

. Temporal regulation of gap junctional communication between tenocytes subjected to static tensile strain with physiological and non-physiological amplitudes. Biochem Biophys Res Commun 2017;482(4):1170–1175; doi: 10.1016/J.BBRC.2016.12.007

189.

189. Maeda

, Ohashi

. Mechano-regulation of gap junction communications between tendon cells is dependent on the magnitude of tensile strain. Biochem Biophys Res Commun 2015;465(2):281–286; doi: 10.1016/J.BBRC.2015.08.021

190.

190. Richardson

, Starborg

, Lu

, et al. Tendon development requires regulation of cell condensation and cell shape via cadherin-11-mediated cell-cell junctions. Mol Cell Biol 2007;27(17):6218–6228; doi: 10.1128/MCB.00261-07

191.

191. Feng

, Zhang

, Kou

, et al. Cadherin-11 modulates cell morphology and collagen synthesis in periodontal ligament cells under mechanical stress. Angle Orthod 2017;87(2):193–199; doi: 10.2319/020716-107.1

192.

192. Lin

, Chien

, Cho

. N-cadherin expression during periodontal ligament cell differentiation in vitro. J Periodontol 1999;70(9):1039–1045; doi: 10.1902/JOP.1999.70.9.1039

193.

193. Dieterle

, Husari

, Steinberg

, et al. Role of Mechanotransduction in Periodontal Homeostasis and Disease. J Dent Res 2021;100(11):1210–1219; doi: 10.1177/00220345211007855

194.

194. Ji

, McCulloch

. TRPV4 integrates matrix mechanosensing with Ca2+ signaling to regulate extracellular matrix remodeling. Febs J 2021;288(20):5867–5887; doi: 10.1111/FEBS.15665

195.

195. Ellefsen

, Holt

, Chang

, et al. Myosin-II mediated traction forces evoke localized Piezo1-dependent Ca2+ flickers. Commun Biol 2019;2(1):298–213; doi: 10.1038/s42003-019-0514-3

196.

196. Arora

, Di Gregorio

, He

, et al. TRPV4 mediates the Ca2+ influx required for the interaction between flightless-1 and non-muscle myosin, and collagen remodeling. J Cell Sci 2017;130(13):2196–2208; doi: 10.1242/JCS.201665

197.

197. Nourse

, Pathak

. How cells channel their stress: Interplay between Piezo1 and the cytoskeleton. Semin Cell Dev Biol 2017;71:3–12; doi: 10.1016/J.SEMCDB.2017.06.018

198.

198. Shen

, Pan

, Guo

, et al. The roles of mechanosensitive ion channels and associated downstream MAPK signaling pathways in PDLC mechanotransduction. Mol Med Rep 2020;21(5):2113–2122; doi: 10.3892/MMR.2020.11006

199.

199. Du

, Li

, Zhang

, et al. Roles of TRPV4 and piezo channels in stretch-evoked Ca2+ response in chondrocytes. Exp Biol Med (Maywood) 2020;245(3):180–189; doi: 10.1177/1535370219892601

200.

200. Lee

, Leddy

, Chen

, et al. Synergy between Piezo1 and Piezo2 channels confers high-strain mechanosensitivity to articular cartilage. Proc Natl Acad Sci U S A 2014;111(47):E5114–E5122; doi: 10.1073/PNAS.1414298111

201.

201. Zhang

, Meng

, Wang

, et al. TRPV4 and PIEZO channels mediate the mechanosensing of chondrocytes to the biomechanical microenvironment. Membranes (Basel) 2022;12(2):237; doi: 10.3390/MEMBRANES12020237

202.

202. Guilak

, Leddy

, Liedtke

. Transient receptor potential vanilloid 4: The sixth sense of the musculoskeletal system? Ann N Y Acad Sci 2010;1192:404–409; doi: 10.1111/J.1749-6632.2010.05389.X

203.

203. Lee

, Nims

, Savadipour

, et al. Inflammatory signaling sensitizes Piezo1 mechanotransduction in articular chondrocytes as a pathogenic feed-forward mechanism in osteoarthritis. Proc Natl Acad Sci U S A 2021;118(13):e2001611118; doi: 10.1073/PNAS.2001611118

204.

204. Ma

, Li

, Kunze

, et al. Engineered human meniscus in modeling sex differences of knee osteoarthritis in vitro. Front Bioeng Biotechnol 2022;10:823679; doi: 10.3389/FBIOE.2022.823679

205.

205. Ciufo

, Cao

, Mendias

, et al. clinical association of achilles tendinopathy and hypertension mediated by CaV1.2 voltage-gated Ca2+ Channel. Foot Ankle Orthop 2023;8(4):1–2; doi: 10.1177/2473011423S00084

206.

206. Li

, Korcari

, Ciufo

, et al. Increased Ca2+ signaling through CaV1.2 induces tendon hypertrophy with increased collagen fibrillogenesis and biomechanical properties. Faseb J 2023;37(7):e23007; doi: 10.1096/FJ.202300607R

207.

207. O’Conor

, Leddy

, Benefield

, et al. TRPV4-mediated mechanotransduction regulates the metabolic response of chondrocytes to dynamic loading. Proc Natl Acad Sci USA 2014;111(4):1316–1321; doi: 10.1073/PNAS.1319569111

208.

208. Savadipour

, Nims

, Katz

, et al. Regulation of chondrocyte biosynthetic activity by dynamic hydrostatic pressure: The role of TRP channels. Connect Tissue Res 2022;63(1):69–81; doi: 10.1080/03008207.2020.1871475

209.

209. Corrigan

, Johnson

, Stavenschi

, et al. TRPV4-mediates oscillatory fluid shear mechanotransduction in mesenchymal stem cells in part via the primary cilium. Sci Rep 2018;8(1):3824–3813; doi: 10.1038/s41598-018-22174-3

210.

210. Gilchrist

, Leddy

, Kaye

, et al. TRPV4-mediated calcium signaling in mesenchymal stem cells regulates aligned collagen matrix formation and vinculin tension. Proc Natl Acad Sci U S A 2019;116(6):1992–1997; doi: 10.1073/PNAS.1811095116

211.

211. Ji

, Wang

, et al. TRPV4 regulates β1 integrin-mediated cell–matrix adhesions and collagen remodeling. Faseb J 2023;37(6):e22946; doi: 10.1096/FJ.202300222R

212.

212. Jin

, Li

, Wang

, et al. Functional role of mechanosensitive ion channel Piezo1 in human periodontal ligament cells. Angle Orthod 2015;85(1):87–94; doi: 10.2319/123113-955.1

213.

213. Nakamichi

, Ma

, Nonoyama

, et al. The mechanosensitive ion channel PIEZO1 is expressed in tendons and regulates physical performance. Sci Transl Med 2022;14(647):eabj5557; doi: 10.1126/SCITRANSLMED.ABJ5557

214.

214. Götschi

, Held

, Klucker

, et al. PIEZO1 gain-of-function gene variant is associated with elevated tendon stiffness in humans. J Appl Physiol (1985) 2023;135(1):165–173; doi: 10.1152/JAPPLPHYSIOL.00573.2022():

215.

215. Muramatsu

, Wakabayashi

, Ohno

, et al. Functional gene screening system identified TRPV4 as a regulator of chondrogenic differentiation. J Biol Chem 2007;282(44):32158–32167; doi: 10.1074/JBC.M706158200

216.

216. Irwin

, Puranam

, Hoffman

, et al. Differential response of inner and outer zone meniscal cells to tensile load under non-inflammatory and inflammatory conditions. Osteoarthritis Cartilage 2021;29:S3–S4; doi: 10.1016/j.joca.2021.05.011

217.

217. Grace

, Bonvini

, Belvisi

, et al. Modulation of the TRPV4 ion channel as a therapeutic target for disease. Pharmacol Ther 2017;177:9–22; doi: 10.1016/J.PHARMTHERA.2017.02.019

218.

218. Sharma

, Goswami

, Zhang

, et al. TRPV4 regulates matrix stiffness and TGFβ1-induced epithelial-mesenchymal transition. J Cell Mol Med 2019;23(2):761–774; doi: 10.1111/JCMM.13972

219.

219. Wang

, Coelho

, Arora

, et al. DDR1 associates with TRPV4 in cell-matrix adhesions to enable calcium-regulated myosin activity and collagen compaction. J Cell Physiol 2022;237(5):2451–2468; doi: 10.1002/JCP.30696

220.

220. Wang

, Ji

, Smith

, et al. Impact of TRP channels on extracellular matrix remodeling: Focus on trpv4 and collagen. Int J Mol Sci 2024;25(7):3566; doi: 10.3390/IJMS25073566

221.

221. Jin

, He

, Wang

, et al. Mechanical force modulates periodontal ligament stem cell characteristics during bone remodelling via TRPV4. Cell Prolif 2020;53(10):e12912; doi: 10.1111/CPR.12912

222.

222. Lukacs

, Rennekampff

, Tenenhaus

, et al. The periodontal ligament, temperature-sensitive ion channels TRPV1-4, and the mechanosensitive ion channels Piezo1 and 2: A Nobel connection. J Periodontal Res 2023;58(4):687–696; doi: 10.1111/JRE.13137

223.

223. Sharma

, Goswami

, Merth

, et al. TRPV4 ion channel is a novel regulator of dermal myofibroblast differentiation. Am J Physiol Cell Physiol 2017;312(5):C562–C572; doi: 10.1152/AJPCELL.00187.2016

224.

224. Yoneda

, Suzuki

, Hatano

, et al. PIEZO1 and TRPV4, which are distinct mechano-sensors in the osteoblastic MC3T3-E1 cells, modify cell-proliferation. Int J Mol Sci 2019;20(19):4960; doi: 10.3390/IJMS20194960

225.

225. Zhao

, Sullivan

, Chase

, et al. Calcineurin/nuclear factor of activated T cells-coupled vanilliod transient receptor potential channel 4 Ca2+ sparklets stimulate airway smooth muscle cell proliferation. Am J Respir Cell Mol Biol 2014;50(6):1064–1075; doi: 10.1165/RCMB.2013-0416OC

226.

226. Atcha

, Jairaman

, Holt

, et al. Mechanically activated ion channel Piezo1 modulates macrophage polarization and stiffness sensing. Nat Commun 2021;12(1):3256–3214; doi: 10.1038/s41467-021-23482-5

227.

227. Haliloglu

, Becker

, Temucin

, et al. Recessive PIEZO2 stop mutation causes distal arthrogryposis with distal muscle weakness, scoliosis and proprioception defects. J Hum Genet 2017;62(4):497–501; doi: 10.1038/JHG.2016.153

228.

228. Grotle

, Huo

, Harrison

, et al. GsMTx-4 normalizes the exercise pressor reflex evoked by intermittent muscle contraction in early stage type 1 diabetic rats. Am J Physiol Heart Circ Physiol 2021;320(4):H1738–H1748; doi: 10.1152/AJPHEART.00794.2020

229.

229. Bartoli

, Evans

, Blythe

, et al. Global PIEZO1 Gain-of-Function Mutation Causes Cardiac Hypertrophy and Fibrosis in Mice. Cells 2022;11(7):1199–1115; doi: 10.3390/CELLS11071199

230.

230. Swain

, Romac

JMJ

, Vigna

, et al. Piezo1-mediated stellate cell activation causes pressure-induced pancreatic fibrosis in mice. JCI Insight 2022;7(8):e158288–e15; doi: 10.1172/JCI.INSIGHT.158288

231.

231. Copp

, Kim

, Ruiz-Velasco

, et al. The mechano-gated channel inhibitor GsMTx4 reduces the exercise pressor reflex in decerebrate rats. J Physiol 2016;594(3):641–655; doi: 10.1113/JP271714

232.

232. Savadipour

, Palmer

, Ely

, et al. The role of PIEZO ion channels in the musculoskeletal system. Am J Physiol Cell Physiol 2023;324(3):C728–C740; doi: 10.1152/AJPCELL.00544.2022

233.

233. Gaite

, Solé-Magdalena

, García-Mesa

, et al. Immunolocalization of the mechanogated ion channels PIEZO1 and PIEZO2 in human and mouse dental pulp and periodontal ligament. Anat Rec (Hoboken) 2024;307(5):1960–1968; doi: 10.1002/AR.25351

234.

234. Zhou

, Gao

, Fan

, et al. Piezo1/2 mediate mechanotransduction essential for bone formation through concerted activation of NFAT-YAP1-ß-catenin. Elife 2020;9:e52779–e38; doi: 10.7554/ELIFE.52779

235.

235. Lee

, Guilak

, Liedtke

. Role of Piezo Channels in Joint Health and Injury. Curr Top Membr 2017;79:263–273; doi: 10.1016/BS.CTM.2016.10.003

236.

236. Emig

, Knodt

, Krussig

, et al. Piezo1 Channels Contribute to the Regulation of Human Atrial Fibroblast Mechanical Properties and Matrix Stiffness Sensing. Cells 2021;10(3):663–621; doi: 10.3390/CELLS10030663

237.

237. Lavagnino

, Oslapas

, Gardner

, et al. Hypoxia inhibits primary cilia formation and reduces cell-mediated contraction in stress-deprived rat tail tendon fascicles. Muscles Ligaments Tendons J 2016;6(2):193–197; doi: 10.11138/MLTJ/2016.6.2.193

238.

238. Chang

, Li

, Kim

, et al. BBS7-SHH Signaling Activity Regulates Primary Cilia for Periodontal Homeostasis. Front Cell Dev Biol 2021;9:796274–796212; doi: 10.3389/FCELL.2021.796274

239.

239. Fang

, Schwartz

, Moore

, et al. Primary cilia as the nexus of biophysical and hedgehog signaling at the tendon enthesis. Sci Adv 2020;6(44):eabc1799; doi: 10.1126/SCIADV.ABC1799

240.

240. Huang

, Arora

, McCulloch

, et al. The collagen receptor DDR1 regulates cell spreading and motility by associating with myosin IIA. J Cell Sci 2009;122(Pt 10):1637–1646; doi: 10.1242/JCS.046219

241.

241. Coelho

, Arora

, van Putten

, et al. Discoidin domain receptor 1 mediates myosin-dependent collagen contraction. Cell Rep 2017;18(7):1774–1790; doi: 10.1016/J.CELREP.2017.01.061

242.

242. Meyer Zum Gottesberge

, Hansen

. The collagen receptor DDR1 co-localizes with the non-muscle myosin IIA in mice inner ear and contributes to the cytoarchitecture and stability of motile cells. Cell Tissue Res 2014;358(3):729–736; doi: 10.1007/S00441-014-2009-3

243.

243. Ren

, Tang

, Jiang

, et al. Periodic mechanical stress stimulates Cav-1-dependent IGF-1R mitogenic signals in rat chondrocytes through ERK1/2. Cell Physiol Biochem 2018;48(4):1652–1663; doi: 10.1159/000492288

244.

244. Zhang

, Wang

, Yuan

, et al. Moderate mechanical stimulation rescues degenerative annulus fibrosus by suppressing caveolin-1 mediated pro-inflammatory signaling pathway. Int J Biol Sci 2021;17(5):1395–1412; doi: 10.7150/IJBS.57774

245.

245. Li

, Guo

, Su

, et al. Role of Primary Cilia in Skeletal Disorders. Stem Cells Int 2022;2022:6063423–6063412; doi: 10.1155/2022/6063423

246.

246. Barsch

, Niedermair

, Mamilos

, et al. Physiological and pathophysiological aspects of primary cilia—a literature review with view on functional and structural relationships in cartilage. Int J Mol Sci 2020;21(14):4959; doi: 10.3390/IJMS21144959

247.

247. Martínez

, Smith

, Rodriguez

, et al. Sonic hedgehog stimulates proliferation of human periodontal ligament stem cells. J Dent Res 2011;90(4):483–488; doi: 10.1177/0022034510391797

248.

248. Hisamoto

, Goto

, Muto

, et al. Developmental changes in primary cilia in the mouse tooth germ and oral cavity. Biomed Res 2016;37(3):207–214; doi: 10.2220/BIOMEDRES.37.207

249.

249. Beertsen

, Everts

, Houtkooper

. Frequency of occurrence and position of cilia in fibroblasts of the periodontal ligament of the mouse incisor. Cell Tissue Res 1975;163(4):415–431; doi: 10.1007/BF00218489

250.

250. Grevenstein

, Oppermann

, Winter

, et al. First detection of primary cilia in injured human anterior cruciate ligament: A pilot study with pathophysiological reflections. Pathol Res Pract 2022;237:154036; doi: 10.1016/J.PRP.2022.154036

251.

251. Donnelly

, Williams

, Farnum

. The primary cilium of connective tissue cells: Imaging by multiphoton microscopy. Anat Rec (Hoboken) 2008;291(9):1062–1073; doi: 10.1002/AR.20665

252.

252. Rhatomy

, Utomo

, Prakoeswa

CRS

, et al. Ligament/tendon culture under hypoxic conditions: A systematic review. Adv Pharm Bull 2021;11(4):595–600; doi: 10.34172/APB.2021.069

253.

253. Vogel

, Gish

, Alves

, et al. The discoidin domain receptor tyrosine kinases are activated by collagen. Mol Cell 1997;1(1):13–23; doi: 10.1016/S1097-2765(00)80003-9

254.

254. Vyhlidal

, Adesida

. Mechanotransduction in meniscus fibrochondrocytes: What about caveolae? J Cell Physiol 2022;237(2):1171–1181; doi: 10.1002/JCP.30616

255.

255. Echarri

, Del Pozo

. Caveolae - mechanosensitive membrane invaginations linked to actin filaments. J Cell Sci 2015;128(15):2747–2758; doi: 10.1242/jcs.153940

256.

256. Schwab

, Hempel

, Funk

RHW

, et al. Ultrastructural identification of caveolae and immunocytochemical as well as biochemical detection of caveolin in chondrocytes. Histochem J 1999;31(5):315–320; doi: 10.1023/A:1003718002088

257.

257. Chen

, He

, Zhong

, et al. Roles of focal adhesion proteins in skeleton and diseases. Acta Pharm Sin B 2023;13(3):998–1013; doi: 10.1016/J.APSB.2022.09.020

258.

258. Rashidi

, Harasymowicz

, Savadipour

, et al. PIEZO1-mediated mechanotransduction regulates collagen synthesis on nanostructured 2D and 3D models of fibrosis. Acta Biomater 2025;193:242–254; doi: 10.1016/J.ACTBIO.2024.12.034

259.

259. Gomez-Florit

, Labrador-Rached

, Domingues

RMA

, et al. The tendon microenvironment: Engineered in vitro models to study cellular crosstalk. Adv Drug Deliv Rev 2022;185:114299; doi: 10.1016/J.ADDR.2022.114299