Mechanical cues in orofacial tissue engineering and regenerative medicine Katrien M. Brouwer, PhD1,2; Ditte M. S. Lundvig, PhD1; Esther Middelkoop, PhD2,3; Frank A. D. T. G. Wagener, PhD1; Johannes W. Von den Hoff, PhD1 1. Department of Orthodontics and Craniofacial Biology, Radboud Institute for Molecular Life Sciences, Radboud university medical center, Nijmegen, The Netherlands, 2. Department of Plastic, Reconstructive and Hand Surgery, Research Institute MOVE, VU University Medical Center, Amsterdam, The Netherlands, 3. Association of Dutch Burn Centers, Beverwijk, The Netherlands

Reprint requests: Reprint requests: Johannes W. Von den Hoff, Department of Orthodontics and Craniofacial Biology, Radboud Institute for Molecular Life Sciences, Radboud university medical center, PO Box 9101, 6500 HB Nijmegen, The Netherlands. Tel: 10031 24 3614084; Fax: 10031 24 3540631; Email: [email protected] Manuscript received: October 30, 2014 Accepted in final form: March 11, 2015 DOI:10.1111/wrr.12283

ABSTRACT Cleft lip and palate patients suffer from functional, aesthetical, and psychosocial problems due to suboptimal regeneration of skin, mucosa, and skeletal muscle after restorative cleft surgery. The field of tissue engineering and regenerative medicine (TE/RM) aims to restore the normal physiology of tissues and organs in conditions such as birth defects or after injury. A crucial factor in cell differentiation, tissue formation, and tissue function is mechanical strain. Regardless of this, mechanical cues are not yet widely used in TE/RM. The effects of mechanical stimulation on cells are not straight-forward in vitro as cellular responses may differ with cell type and loading regime, complicating the translation to a therapeutic protocol. We here give an overview of the different types of mechanical strain that act on cells and tissues and discuss the effects on muscle, and skin and mucosa. We conclude that presently, sufficient knowledge is lacking to reproducibly implement external mechanical loading in TE/RM approaches. Mechanical cues can be applied in TE/RM by fine-tuning the stiffness and architecture of the constructs to guide the differentiation of the seeded cells or the invading surrounding cells. This may already improve the treatment of orofacial clefts and other disorders affecting soft tissues.

Orofacial clefts are the most common congenital abnormalities of the head, affecting about 1 in 600 children depending on ethnicity and geographical location.1 The etiology of orofacial clefts is not completely understood, but both genetic components as well as environmental factors are involved.2 Cleft lip and palate (CLP) patients need several surgical interventions to close the lip and restore palatal function (Figure 1). During wound repair diverse tissues are involved, including palatal skeletal muscle, skin, and mucosa. Unfortunately, CLP surgery leaves scars, which may interfere with normal development of the midface and dentition.3 Other complications include insufficient palatal muscle function which hampers swallowing, sucking, and speech.4 To prevent these complications, adjuvant therapies are necessary. Wounds normally heal by repair, which closes the defect but will lead to imperfect tissue structure. On the other end of the spectrum there is regeneration, which implicates perfect healing into the original tissue organization. Regeneration of wounds takes place in fetal tissues, but only during the early gestational period.5,6 The research field of tissue engineering and regenerative medicine (TE/RM) aims to stimulate regeneration of tissues and organs by either implanting biomaterials for in vivo regeneration, or by constructing substitutes in vitro.7,8 302

This may also be implemented in CLP treatment. A number of generally applicable requirements for a TE/RM construct must be met for optimal results: biocompatibility, biodegradability, a suitable architecture, and appropriate mechanical characteristics.9 Unfortunately, despite all the efforts, the success rate of clinical translation of TE/RM has been disappointing thus far. As tissue mechanics are crucial in the normal physiological behavior of cells, TE/RM approaches in orofacial clefting therapy may be more successful when mechanical strain is applied. In this review, we will first summarize the different types of mechanical forces normally experienced by tissues and cells. Thereafter, we will discuss studies that currently use mechanical stimulation in relevant TE/RM approaches with emphasis on muscle, skin, and mucosa as these tissues are central in CLP surgery. We will not discuss the application of mechanical strain in bone tissue engineering, as this topic has already been covered extensively elsewhere.10–12 Different methods may be required for specific tissues or organs, as the mechanical conditions in different tissues vary considerably. Applying mechanical cues in TE/RM approaches may greatly improve the outcome of surgical treatment of orofacial clefts as well as other tissue defects. C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

Brouwer et al.

Mechanical cues in orofacial tissue engineering and regenerative medicine

Figure 1. Cartoon of the soft tissues affected in CLP patients. Skin of the lip, mucosa, soft palate muscle, and bone are affected (latter not included in this review). Image courtesy of P. L. Carvajal Monroy, Department of Orthodontics and Craniofacial Biology, Dentistry, Radboudumc, Nijmegen, The Netherlands. [Color figure can be viewed in the online issue, which is available at]

TYPES OF MECHANICAL FORCES Cells constantly probe their mechanical environment and adapt their physiology accordingly. Each cell has its own mechanical niche. Depending on the tissue, cell type, and location, cells are subjected to different types and magnitudes of mechanical load (Figure 2, Table 1). Cells and tissues can be subjected to different types of external mechanical forces including (A) tensile forces, (B) compressive forces, and (C) (fluid) shear stress (Figure 2). Tensile forces increase the dimension of the tissue in the direction of the force. These are experienced by tissues such as tendons and muscles. Compressive forces reduce the dimension of the tissue in the direction of the force. For instance, body weight compresses the femoral bone. Shear stress is experienced by endothelial cells of the inner lining of the vasculature, but is also created by interstitial fluid flow in e.g., cartilage during loading. These different kinds of load cause cellular deformation or strain. Strain can differ between specific locations within the same tissue depending on the local microenvironment and composition of the extracellular matrix (ECM). The anatomical location, the macroenvironment, can also influence strain. For instance, in skin, the strain differs between locations of the body, e.g., on the scalp or the elbow. Specific lines of high tension exist (Langer’s lines) due to the orientation of the underlying collagen network.13 In addition to differences in direction, the magnitude of strain on cells also varies. For example, different muscles experience different strains, and need to produce different forces depending on their specific location and function.14 Strain is received and sensed both extra- and intracellularly. The ECM will receive most of the external strain, and depending on the interactions between the cell and the ECM, strain will be transferred to the cell. Cells can also directly pull on neighboring cells via connections such as adhesion molecules, tight junctions, gap junctions, adheC 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

rens junctions, or desmosomes. Besides strains induced by forces such as movement and growth, cells are also influenced by the mechanical properties of the surrounding ECM, and many cell types cannot survive when cultured in suspension. Stem cells have been shown to be sensitive to mechanical cues,15–25 and stem cell differentiation in vitro is influenced by the stiffness of the substrate.26 Cells probe the stiffness of the ECM and can remodel the ECM accordingly to maintain a suitable mechanical environment.27 Cells can remodel the ECM by compaction, contraction, and degradation of ECM components, but also by the deposition of new ECM. This changes matrix stiffness and composition, which also influences adjacent cells. The ECM architecture, such as shape and pore dimensions, can also modify the differentiation and responses of (stem) cells.15 In short, cell behavior is regulated by mechanical cues from the ECM (outside-in signaling), but cells also probe the mechanical properties of the ECM to remodel it to their needs (inside-out signaling). Strain is transduced intracellularly by a process called mechanotransduction, which is the translation of mechanical cues into biochemical signals. One of the key players herein are integrins, bidirectional signal transducers which deliver signals inside-out and outside-in.28 It is conceivable that TE/RM constructs should enable the binding of integrins through arginine–glycine–aspartate (RGD) peptides or other ligands to allow for signal transduction. Other key players in mechanotransduction are adhesion complexes, stretch-sensitive ion channels, and growth factor receptors.29–31 A number of excellent recent reviews have discussed this subject in depth.30,32

Figure 2. Cartoon of different strains that cells can experience. (A) Tensile strain; (B) Compressive strain; (C) Shear strain. These strains can also occur in combination. ECM, extracellular matrix. [Color figure can be viewed in the online issue, which is available at]


Mechanical cues in orofacial tissue engineering and regenerative medicine

Brouwer et al.

Table 1. Definition of terms Term


Load Strain

All forces applied to an object Measure of deformation (change in shape or size of an object) due to an applied force Intensity of force, force per unit area Produced by applying forces from two opposing sides into an object, directed toward each other. Leading to negative strain Produced by two opposing forces that pull outward onto an object, directed in opposite directions away from each other. Leading to positive strain Displacement of layers of an object parallel to each other In one direction Identical in two directions In several directions

Stress (pressure) Compressive force Tensile force

Shear force Unidirectional Equibiaxial Multidirectional/multiaxial

Magnitude (Unit) F (N) e (unitless) r or P (N/m2 or Pa) F (N) F (N)

F (N) N/A N/A N/A

N/A, not applicable.

For TE/RM, several options for the application of load to constructs are available. Load can be applied in vitro in the form of stretch, fluid shear stress, and compression: high-tech bioreactors allow protocols with varying duration, interval, flow, and strain to match the specific needs of the target cells and tissues.33 In the following paragraphs, we will discuss how mechanical stimulation might be useful in TE/RM of skeletal muscle, and skin and mucosa. These soft tissues are all affected in CLP patients by cleft surgery, but also in other conditions such as trauma, tumour resection, or in other congenital defects. Specific mechanical cues may increase the quality of soft tissue constructs, and hence their clinical application.

SKELETAL MUSCLE AND MECHANICAL STRAIN Skeletal muscle is a clear example of a soft tissue that is influenced by mechanical forces. It consists of muscle fibers aligned parallel to the main lines of tension. Tension during muscle development causes an increase in myofiber length and myofibril number, and increases protein accumulation in later phases.34 In the adult situation, muscle cells turn hypertrophic in response to exercise, whereas atrophy occurs when muscle is not stimulated for longer periods of time. This indicates the importance of mechanical stimulation in the physiological function of muscle tissue and in the development of TE/RM muscle constructs. To prepare a functional muscle construct, muscle cells should proliferate and differentiate into aligned, cross-striated myofibers that produce physiological forces. In general, cells can respond to mechanical stimulation by changing their proliferation rate and differentiation status.35–40 Applying mechanical strain stimulates satellite cells—the normally quiescent muscle stem cells—to become activated.41 This might be caused by the release of hepatocyte growth factor from the ECM by stretching.42,43 The proliferation of avian myoblasts and murine C2C12 myoblasts is stimulated by mechanical strain.36,39,44,45 Muscle cell differentiation is also 304

enhanced by mechanical stimulation.39,46,47 In contrast to these findings, cyclic mechanical strain can also reduce the differentiation of C2C12 mouse myoblasts, adult bovine myoblasts, and murine muscle progenitor cells.36,44,45,48 These differences might be related to the applied strain regimen. Cells can be stretched unidirectionally or multi/(equi)biaxially. Stretching protocols can be used to create ECM alignment in constructs, which, vice versa, influences cell metabolism.49,50 This knowledge may be used to recreate the original ECM orientation of the tissue of interest. It should be kept in mind that cells can align differently depending on the applied strain. Cells have been shown to align parallel to the strain direction under static strain,39,51,52 and perpendicular under dynamic or cyclic strain.51,52 The latter is called stress shielding, and is hypothesised to be caused by the development of micro-ripples perpendicular to the strain.53,54 These observations are highly relevant for tissue regeneration as collagen is deposited primarily along the cells’ main axis in vivo, and constructs with an orientation like the original tissue may provide the best basis for regeneration. Uniaxial stretching of myoblasts increases the differentiation and alignment compared with equibiaxial stretching.55 Multiaxial stretch was shown to increase the phosphorylation of ribosomal S6 kinase, involved in muscle hypertrophy.56 The orientation, force and strength of strain can thus influence the differentiation of muscle cells. Another hurdle is that it is difficult to determine the exact strain experienced by muscle cells in vivo. To complicate matters further, the observed responses in vitro may also be caused by changes in nutrient diffusion47 and contact inhibition.39 These stretch-dependent outcomes underscore the need of well-established protocols before clinical translation.

MECHANICAL STRAIN IN TE/RM OF OROFACIAL MUSCLE In CLP patients, the major muscle of the soft palate (m. levator veli palatini, LVP) needs to be reconstructed for proper velopharyngeal functioning and normal speech development.57 A method to reconstruct this muscle is the C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

Brouwer et al.

Mechanical cues in orofacial tissue engineering and regenerative medicine

implantation of a (cellularized) scaffold in the palatum. Cells may experience different levels of strain when cultured in vitro in a two-dimensional or in a more biomimetic three-dimensional (3D) model.33 An often used 3D model for tissue-engineered muscle is the production of myofibers in ECM-based hydrogels. These constructs allow assessment of the effects of strain on muscle cell proliferation and differentiation. The molecular responses to movement/exercise in such a model58 were comparable to the in vivo situation in terms of growth factor synthesis and force production.47,59–61 The stiffness of a substrate can determine whether cells will proliferate or differentiate.62 For in vivo use, hydrogels may be less suitable due to their limited mechanical stiffness and force production.63 The mechanical stiffness of hydrogels can be improved by chemical crosslinking,64 but the maximum force they can produce is still about seven times lower than native adult skeletal muscle.63 Using unstrained decellularized muscle scaffolds seeded with C2C12 mouse myoblasts, in vitro, only low forces were produced.65 Comparable constructs seeded with human satellite cells and cyclically strained in a bioreactor, gave a contractile response in vivo of 1% of that of native muscle. In contrast, statically cultured muscle constructs did not generate any contractions.66 This indicates that mechanical preconditioning can increase the produced force. However, the implantation of these constructs in a rat limb muscle defect increased the functional capacity in only about 50% of the cases.67 This may be related to a stronger immune reaction in the “negative responders.”68 Another issue is the limited myofiber survival after implantation of muscle constructs.68,69 These experiments show that in vivo implantation does not readily lead to long-lasting improvement in muscle function. Moreover, several studies have shown a decrease in regenerative capacity in vivo after in vitro culture of muscle cells,70–72 which might be related to dedifferentiation.73 An alternative approach would be to skip the in vitro culturing step and to apply in vivo TE. Then, the cells are isolated, seeded into an appropriate scaffold, and implanted directly into the defect.74 This construct will then be exposed to a physiological level of mechanical force, avoiding the time-consuming culture step and complicated regulatory issues. However, obtaining enough cells from biopsies without culturing may be impossible. A totally different view on the effect of mechanical load on muscle regeneration emerged from a study by Corona et al.75 They observed that decellularized muscle ECM was unable to induce muscle regeneration, but still muscle function had improved significantly. A possible explanation for this could be that the construct protected the remaining muscle from overload.75–77 Although TE/RM research has focused rather intensively on skeletal muscle, this mainly concerns muscles of the limbs and trunk. There are, however, significant differences between these muscles and the muscles of the head (reviewed in Carvajal Monroy et al.4, probably related to their different embryonic origin,78,79 which gives rise to architectural and cellular differences. Craniofacial muscles regenerate slower, and seem to produce more fibrotic tissue after injury than limb muscles.4 In vitro, the satellite cells of the masseter muscle (one of the muscles of mastication) proliferate more and differentiate later.4 Muscle fiber types also differ between muscles, and the LVP in C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

healthy individuals contains a larger proportion of slow fibers, whereas that of CLP patients has a higher proportion of fast fibers.80–82 Stretching seems to guide the type of muscle fibers formed in culture, as MyHC-2b, a fasttwitch subtype, is up-regulated after rapid stretching of C2C12 cells, whereas slow stretching induces the slowtwitch MyHC-2a and MyHC-1.83 Fast muscle fibers generally contain less satellite cells,84,85 and these cells proliferate less in vitro.78 Moreover, mechanical strain stimulates protein synthesis and fiber growth in glycolytic fast fibers less than both slow and fast oxidative fibers from rat soleus muscle.86 During exercise, protein synthesis first decreases, to increase again after a lag phase.87–90 This is specifically the case in muscles with a high proportion of fast glycolytic fibers,87 and thus of particular relevance for CLP patients. These data indicate that specialized protocols are needed for the stimulation of proliferation and differentiation for orofacial muscle cells. Muscle regeneration in the soft palate of CLP patients may be limited due to fibrosis, reduced differentiation of myoblasts, and reduced protein synthesis. All muscles have a different force production, which may ask for “customized” protocols. For instance, jaw muscle needs to be able to produce more force, whereas muscles in the soft palate, which are used during speech, sucking, and swallowing, need to have a better endurance. A slow stretch protocol stimulating slow, enduring muscle fiber formation may, therefore, be most appropriate for CLP muscle TE/RM. However, the application of TE/RM in the orofacial region is still in its infancy, and specific animal models such as a recently developed palatal wound model91 are required to evaluate TE/RM approaches for the muscles in the orofacial region.

SKIN AND MUCOSA AND MECHANICAL STRAIN Mechanical factors are involved in the physiology of normal skin and mucosa. Its elasticity, for instance, protects skin from rupture during contact. This elasticity is derived from the basket-weave orientation of the ECM fibers collagen and elastin.92,93 During wound healing, several mechanical issues are involved, including the wounding itself, but also the pulling by myofibroblasts during wound contraction. Mechanical properties are also altered by granulation tissue formation and edema.94 Suturing of a wound leads to mechanical strain, especially when a larger tissue defect is present. Moreover, wounds over the joints heal with more scar formation,29 showing that mechanical tension increases scarring.95 Normal skin and scar tissue have different mechanical characteristics.96–99 In scars, collagen type I has a more parallel orientation than in normal skin, which reduces compliance at low loads and resistance to failure.97 Thus, constructs should guide proper collagen deposition as tissue stiffness may compromise function. As strain contributes to (hypertrophic) scar formation, methods have been developed to reduce strain on wounds. Stress-shielding devices that reduce the mechanical strain have been developed to diminish scarring. In studies on porcine skin wounds, the scar area was strongly reduced in stress-shielded wounds compared with control and highly stressed wounds.100 The beneficial effect of stress305

Mechanical cues in orofacial tissue engineering and regenerative medicine

shielding was also confirmed in a human study.100 Another method is the use of negative-pressure wound therapy, in which a vacuum is created over the wound. This decreases wound surface area and enhances healing rates,101–103 possibly through enhanced granulation tissue formation, angiogenesis, cellular proliferation, and myofibroblast differentiation.101,104 In contrast, pressure therapy is also used to reduce scar formation. Pressure garments have been found to improve scars with respect to maturation and hypertrophy. They seem to reduce collagen synthesis by limiting the blood supply, and to help realignment of collagen fiber bundles (reviewed in Macintyre and Baird105). Scar formation can thus be reduced by an initial reduction of (tensile) stress on the wound (stress-shielding devices), whereas later on pressure therapy limits hypertrophic scarring. Comparing these two methods, the strain direction seems to be different, as stress shielding reduces tensile strain parallel to the skin surface, whereas negativepressure therapy and pressure garments strain the skin perpendicular to the surface.

MECHANICAL STRAIN IN TE/RM OF OROFACIAL SKIN AND MUCOSA The structure and composition, but also the mechanics of skin and mucosa vary for different locations in the body. This may have implications for TE/RM approaches of specific locations such as the orofacial region in CLP patients. Skin has a different mechanical orientation (Langer’s lines) all over the body, and it differs locally in thickness and structure. Skin in the lip region, for instance, is rather thin, as opposed to skin on the palms of the hands. Skin around the lips is also well perfused, which may be beneficial for healing. The healing of oral mucosa is generally faster and leads to less scar formation.106 This may be caused by healing-promoting factors in the saliva,107 but also by an inherent difference in phenotype between oral and skin tissue and fibroblasts, with respect to resolution of inflammation, ECM remodeling and ECM expression.108,109 This latter is especially important, as a dermal construct should stimulate fibroblast proliferation and differentiation, but conversely, the balance between cell number and ECM (production) is crucial in skin regeneration as hypertrophic scarring involves too many myofibroblasts and too much ECM. Mechanical loading of wounds causes hypertrophic scarring in mice,110 and both in vitro and in vivo studies have shown that static as well as cyclic strain increase myofibroblast formation,111 proliferation,37,112,113 and differentiation (the expression of several ECM genes including a smooth muscle actin [aSMA]).37,112–114 Strain was also found to reduce myofibroblast apoptosis,110,115 whereas its release increased apoptosis.116 Others have shown that aSMA is down-regulated after cyclic equibiaxial strain in vitro,117 and these differences may result from different strain profiles.118 Strain direction and donor age also affect responses regarding cellular morphology, proliferation, and collagen production.119,120 Finally, in lung fibroblasts differentiation is reduced after cyclic strain,121 in contrast to dermal fibroblasts that generally showed increased differentiation.37,112,113 In gingival fibroblasts, mechanical strain leads to proliferative and antiapoptotic stimuli,122 whereas others confirmed the induction of pro306

Brouwer et al.

liferation but did not observe increased expression of myofibroblast markers.123 Cell type is thus also important in responses to mechanical strain, and results should, therefore, specifically be defined for cells from the orofacial region. In the epidermis, the specific architecture of the rete ridges provides a mechanical niche for keratinocytes. This knowledge may be used in TE/RM, as in constructs with an architecture comparable to this rete ridge structure the epithelialization rate was higher than in constructs with a flat surface, and the expression of genes related to proliferation and migration was increased in keratinocytes.124–126 Mechanical strain has further been found to increase the proliferation of keratinocytes by both static and cyclic stretch,38,40 whereas differentiation was down-regulated by static stretch.40 Keratinocyte proliferation may be beneficial in the closure of wounds but can also cause a thickened epidermis as observed in hypertrophic scars and psoriasis.127 Normal skin and mucosal ECM fibers are present in a random, basket-weave orientation, although alignment of collagen fibers has been observed in specific locations.128 Scar tissue usually has more parallel collagen fibers than normal tissue. In clinical wound closure, skin is sometimes stretched to close larger defects, leading to a more parallel alignment of collagen and elastin bundles.129 The effects on the quality of the resulting skin are not exactly known. Mechanical strain can thus influence the orientation and density of (newly deposited) fibers within a construct, and therewith skin quality. This may in turn modulate the cells, as for instance the orientation or nanotopography of the substrate has been found to influence fibroblast migration.128 The effects of a unidirectional orientation or a more random orientation of ECM fibers both in vitro as well as in vivo should, therefore, be investigated. In vivo, faster vascularization and myofibroblast ingrowth was observed in collagen scaffolds implanted in the mucosa covering the hard palate compared with scaffolds implanted in the skin of rats. Scaffolds implanted in the loose skin on the back of the rats showed more inflammation compared with scaffolds in the palate (which lies over bone) or in the skin on the skull. This might be caused by the higher level of mechanical stimuli in the back skin.130 These differences indicate the importance to perform studies specifically for the orofacial region. Another method to implement strain in TE/RM for skin is by controlling scaffold stiffness. Higher scaffold stiffness accelerates angiogenesis, decreases wound contraction, and improves dermal regeneration.131 As the application of mechanical strain during in vitro culture can also improve the (experimental or surgical) handling properties of the constructs by increasing ECM production,37,132 these are all beneficial effects of a higher construct strength.

CONCLUSION AND FUTURE DIRECTIONS The surgical treatment of CLP is still suboptimal, and TE/ RM strategies could be useful to improve regeneration of the tissues involved. As mechanical strain is important for the development and maintenance of healthy tissues, TE/ RM constructs may also be enhanced by applying strain. C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

Brouwer et al.

Mechanical cues in orofacial tissue engineering and regenerative medicine

First, it is essential to define the desired aim of mechanical stimulation for regeneration. During production of a skeletal muscle construct, the aim is to stimulate cellular proliferation and differentiation, whereas when implanted the construct should experience and produce physiological strains. Although the reported optimal strain parameters for well-differentiated muscle constructs vary, the application of strain by stretching is probably the most physiological form of mechanical stimulation. For CLP muscle TE/ RM, unidirectional strain using a slow stretch protocol may be most appropriate as this will stimulate the development of slow twitch, (endurance) muscle fibers, which are present in normal palatal muscle. For use in vivo, the stiffness of constructs used for muscle TE/RM should be increased by the use of other materials than hydrogels. Vascularization and innervation should be stimulated to optimize myofiber survival and force production,73 using for instance growth factors. Although in vitro mechanical strain can improve the orientation and strength of muscle constructs, the results are not yet optimal. In vivo TE/RM by the implantation of an acellular scaffold with chemotactic agents for (stem) cells seems to be promising. Alternatively, cell-seeded scaffolds could be implanted after which the cells are stimulated by the physiological load on the construct. In vivo TE/RM with acellular constructs would also ease the extensive regulatory issues and high costs associated with culturing cells for clinical application. Mechanical factors are crucial for the normal physiology of skin and mucosa, but forces on a wound may also increase the risk of (hypertrophic) scarring. Scar formation can be diminished by limiting these forces, which is already clinically applied in pressure therapy. For CLP patients, the most obvious method is to prevent tight suturing and to use constructs that can withstand the forces on the wound. The response of skin cells to mechanical strain in vitro varies considerably depending on strain type, strain direction and cell type and origin, which as yet precludes the establishment of solid protocols. It is evident, however, that the application of mechanical strain in vitro can strengthen constructs before implantation. Moreover, constructs with an appropriate ECM composition and geometry, in combination with a suitable stiffness, may already improve regeneration without external mechanical stimulation. Concluding, there are several opportunities to implement mechanical cues into TE/RM approaches for CLP and other soft tissue disorders. Currently, the most feasible approach to provide mechanical cues to cells is by optimizing scaffold architecture and stiffness, and not by in vitro mechanical stimulation. The effects of dynamic vs. static mechanical strain on cellular differentiation and orientation should be further elucidated as the present results are contradictory. We also conclude that more research should be performed to optimize the parameters for in vitro loading of soft tissue constructs to further improve translation to the patient. Source of Funding: This work was supported by the Dutch Burns Foundation project 09.110, TASENE (NWO/ SIDA/COSTECH W02.29.101), and the European Commission (FP7-HEALTH-2011-1, Grant Agreement no. 279024), project EuroSkinGraft. The authors declare no conflicts of interest for this work. C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

REFERENCES 1. Vanderas AP. Incidence of cleft lip, cleft palate, and cleft lip and palate among races: a review. Cleft Palate J 1987; 24: 216–25. 2. Watkins SE, Meyer RE, Strauss RP, Aylsworth AS. Classification, epidemiology, and genetics of orofacial clefts. Clin Plast Surg 2014; 41: 149–63. 3. Shetye PR. Facial growth of adults with unoperated clefts. Clin Plast Surg 2004; 31: 361–71. 4. Carvajal Monroy PL, Grefte S, Kuijpers-Jagtman AM, Wagener FA, Von den Hoff JW. Strategies to improve regeneration of the soft palate muscles after cleft palate repair. Tissue Eng Part B Rev 2012; 18: 468–77. 5. Bullard KM, Longaker MT, Lorenz HP. Fetal wound healing: current biology. World J Surg 2003; 27: 54–61. 6. Cass DL, Bullard KM, Sylvester KG, Yang EY, Longaker MT, Adzick NS. Wound size and gestational age modulate scar formation in fetal wound repair. J Pediatr Surg 1997; 32: 411–5. 7. Langer R, Vacanti JP. Tissue engineering. Science 1993; 260: 920–6. 8. Mason C, Dunnill P. A brief definition of regenerative medicine. Regen Med 2008; 3: 1–5. 9. O’Brien FJ. Biomaterials and scaffolds for tissue engineering. Mater Today 2011; 14: 88–95. 10. McCoy RJ, O’Brien FJ. Influence of shear stress in perfusion bioreactor cultures for the development of threedimensional bone tissue constructs: a review. Tissue Eng Part B Rev 2010; 16: 587–601. 11. Sladkova M, de Peppo G. Bioreactor systems for human bone tissue engineering. Processes 2014; 2: 494–525. 12. Rauh J, Milan F, Gunther KP, Stiehler M. Bioreactor systems for bone tissue engineering. Tissue Eng Part B Rev 2011; 17: 263–80. 13. Langer K. On the anatomy and physiology of the skin. Br J Plast Surg 1978; 31: 93–106. 14. Lieber RL, Ward SR. Skeletal muscle design to meet functional demands. Philos Trans R Soc Lond B Biol Sci 2011; 366: 1466–76. 15. Qu X, Zhu W, Huang S, Li YS, Chien S, Zhang K, et al. Relative impact of uniaxial alignment vs. form-induced stress on differentiation of human adipose derived stem cells. Biomaterials 2013; 34: 9812–8. 16. Delaine-Smith RM, Reilly GC. Mesenchymal stem cell responses to mechanical stimuli. Muscles Ligaments Tendons J 2012; 2: 169–80. 17. Discher DE, Mooney DJ, Zandstra PW. Growth factors, matrices, and forces combine and control stem cells. Science 2009; 324: 1673–7. 18. Evans ND, Oreffo RO, Healy E, Thurner PJ, Man YH. Epithelial mechanobiology, skin wound healing, and the stem cell niche. J Mech Behav Biomed Mater 2013; 28: 397–409. 19. Kurpinski K, Chu J, Hashi C, Li S. Anisotropic mechanosensing by mesenchymal stem cells. Proc Natl Acad Sci USA 2006; 103: 16095–100. 20. Morita Y, Watanabe S, Ju Y, Xu B. Determination of optimal cyclic uniaxial stretches for stem cell-to-tenocyte differentiation under a wide range of mechanical stretch conditions by evaluating gene expression and protein synthesis levels. Acta Bioeng Biomech 2013; 15: 71–9. 21. Pan F, Zhang M, Wu G, Lai Y, Greber B, Scholer HR, Chi L. Topographic effect on human induced pluripotent stem


Mechanical cues in orofacial tissue engineering and regenerative medicine







28. 29.






35. 36.





cells differentiation towards neuronal lineage. Biomaterials 2013; 34: 8131–9. Rehfeldt F, Engler AJ, Eckhardt A, Ahmed F, Discher DE. Cell responses to the mechanochemical microenvironmentimplications for regenerative medicine and drug delivery. Adv Drug Deliv Rev 2007; 59: 1329–39. Saha K, Keung AJ, Irwin EF, Li Y, Little L, Schaffer DV, et al. Substrate modulus directs neural stem cell behavior. Biophys J 2008; 95: 4426–38. Teh TK, Toh SL, Goh JC. Aligned fibrous scaffolds for enhanced mechanoresponse and tenogenesis of mesenchymal stem cells. Tissue Eng Part A 2013; 19: 1360–72. Dado-Rosenfeld D, Tzchori I, Fine A, Chen-Konak L, Levenberg S. Tensile forces applied on a cell-embedded three-dimensional scaffold can direct early differentiation of embryonic stem cells toward the mesoderm germ layer. Tissue engineering Part A 2015; 21: 124–33. Engler AJ, Sen S, Sweeney HL, Discher DE. Matrix elasticity directs stem cell lineage specification. Cell 2006; 126: 677–89. Brown RA, Prajapati R, McGrouther DA, Yannas IV, Eastwood M. Tensional homeostasis in dermal fibroblasts: mechanical responses to mechanical loading in threedimensional substrates. J Cell Physiol 1998; 175: 323–32. Dedhar S. Integrins and signal transduction. Curr Opin Hematol 1999; 6: 37–43. Wong VW, Longaker MT, Gurtner GC. Soft tissue mechanotransduction in wound healing and fibrosis. Semin Cell Dev Biol 2012; 23: 981–6. DuFort CC, Paszek MJ, Weaver VM. Balancing forces: architectural control of mechanotransduction. Nat Rev Mol Cell Biol 2011; 12: 308–19. Janmey PA, Wells RG, Assoian RK, McCulloch CA. From tissue mechanics to transcription factors. Differentiation 2013; 86: 112–20. Mammoto A, Mammoto T, Ingber DE. Mechanosensitive mechanisms in transcriptional regulation. J Cell Sci 2012; 125: 3061–73. Riehl BD, Park JH, Kwon IK, Lim JY. Mechanical stretching for tissue engineering: two-dimensional and threedimensional constructs. Tissue Eng Part B Rev 2012; 18: 288–300. Vandenburgh HH. Motion into mass: how does tension stimulate muscle growth? Med Sci Sports Exerc 1987; 19: S142–9. Goldspink G. Gene expression in muscle in response to exercise. J Muscle Res Cell Motil 2003; 24: 121–6. Kumar A, Murphy R, Robinson P, Wei L, Boriek AM. Cyclic mechanical strain inhibits skeletal myogenesis through activation of focal adhesion kinase, Rac-1 GTPase, and NF-kappaB transcription factor. FASEB J 2004; 18: 1524–35. Powell HM, McFarland KL, Butler DL, Supp DM, Boyce ST. Uniaxial strain regulates morphogenesis, gene expression, and tissue strength in engineered skin. Tissue Eng Part A 2010; 16: 1083–92. Takei T, Han O, Ikeda M, Male P, Mills I, Sumpio BE. Cyclic strain stimulates isoform-specific PKC activation and translocation in cultured human keratinocytes. J Cell Biochem 1997; 67: 327–37. Vandenburgh HH, Karlisch P. Longitudinal growth of skeletal myotubes in vitro in a new horizontal mechanical cell stimulator. In Vitro Cell Dev Biol 1989; 25: 607–16.

Brouwer et al.

40. Yano S, Komine M, Fujimoto M, Okochi H, Tamaki K. Mechanical stretching in vitro regulates signal transduction pathways and cellular proliferation in human epidermal keratinocytes. J Invest Dermatol 2004; 122: 783–90. 41. Tatsumi R, Sheehan SM, Iwasaki H, Hattori A, Allen RE. Mechanical stretch induces activation of skeletal muscle satellite cells in vitro. Exp Cell Res 2001; 267: 107–14. 42. Tatsumi R. Mechano-biology of skeletal muscle hypertrophy and regeneration: possible mechanism of stretch-induced activation of resident myogenic stem cells. Anim Sci J 2010; 81: 11–20. 43. Grefte S, Kuijpers-Jagtman AM, Torensma R, Von den Hoff JW. Skeletal muscle development and regeneration. Stem Cells Dev 2007; 16: 857–68. 44. Kook SH, Lee HJ, Chung WT, Hwang IH, Lee SA, Kim BS, et al. Cyclic mechanical stretch stimulates the proliferation of C2C12 myoblasts and inhibits their differentiation via prolonged activation of p38 MAPK. Mol Cells 2008; 25: 479–86. 45. Kook SH, Son YO, Choi KC, Lee HJ, Chung WT, Hwang IH, et al. Cyclic mechanical stress suppresses myogenic differentiation of adult bovine satellite cells through activation of extracellular signal-regulated kinase. Mol Cell Biochem 2008; 309: 133–41. 46. Engler AJ, Griffin MA, Sen S, Bonnemann CG, Sweeney HL, Discher DE. Myotubes differentiate optimally on substrates with tissue-like stiffness: pathological implications for soft or stiff microenvironments. J Cell Biol 2004; 166: 877–87. 47. Powell CA, Smiley BL, Mills J, Vandenburgh HH. Mechanical stimulation improves tissue-engineered human skeletal muscle. Am J Physiol Cell Physiol 2002; 283: C1557–65. 48. Boonen KJ, Langelaan ML, Polak RB, van der Schaft DW, Baaijens FP, Post MJ. Effects of a combined mechanical stimulation protocol: value for skeletal muscle tissue engineering. J Biomech 2010; 43: 1514–21. 49. Hu JJ, Humphrey JD, Yeh AT. Characterization of engineered tissue development under biaxial stretch using nonlinear optical microscopy. Tissue Eng Part A 2009; 15: 1553–64. 50. Nguyen TD, Liang R, Woo SL, Burton SD, Wu C, Almarza A, et al. Effects of cell seeding and cyclic stretch on the fiber remodeling in an extracellular matrix-derived bioscaffold. Tissue Eng Part A 2009; 15: 957–63. 51. de Jonge N, Kanters FM, Baaijens FP, Bouten CV. Straininduced collagen organization at the micro-level in fibrinbased engineered tissue constructs. Ann Biomed Eng 2013; 41: 763–74. 52. Vandenburgh HH. A computerized mechanical cell stimulator for tissue culture: effects on skeletal muscle organogenesis. In Vitro Cell Dev Biol 1988; 24: 609–19. 53. Ostrovidov S, Hosseini V, Ahadian S, Fujie T, Parthiban SP, Ramalingam M, et al. Skeletal muscle tissue engineering: methods to form skeletal myotubes and their applications. Tissue Eng Part B Rev 2014; 20: 403–36. 54. Lin P, Yang S. Spontaneous formation of one-dimensional ripples in transit to highly ordered two-dimensional herringbone structures through sequential and unequal biaxial mechenical stretching. Appl Phys Lett 2007; 90: 241903. 55. Pennisi CP, Olesen CG, de Zee M, Rasmussen J, Zachar V. Uniaxial cyclic strain drives assembly and differentiation of skeletal myocytes. Tissue Eng Part A 2011; 17: 2543–50. C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

Brouwer et al.

Mechanical cues in orofacial tissue engineering and regenerative medicine

56. Hornberger TA, Armstrong DD, Koh TJ, Burkholder TJ, Esser KA. Intracellular signaling specificity in response to uniaxial vs. multiaxial stretch: implications for mechanotransduction. Am J Physiol Cell Physiol 2005; 288: C185– 94. 57. Braithwaite F, Maurice DG. The importance of the levator palati muscle in cleft palate closure. Br J Plast Surg 1968; 21: 60–2. 58. Khodabukus A, Paxton JZ, Donnelly K, Baar K. Engineered muscle: a tool for studying muscle physiology and function. Exerc Sport Sci Rev 2007; 35: 186–91. 59. Baar K, Torgan CE, Kraus WE, Esser K. Autocrine phosphorylation of p70(S6k) in response to acute stretch in myotubes. Mol Cell Biol Res Commun 2000; 4: 76–80. 60. Cheema U, Brown R, Mudera V, Yang SY, McGrouther G, Goldspink G. Mechanical signals and IGF-I gene splicing in vitro in relation to development of skeletal muscle. J Cell Physiol 2005; 202: 67–75. 61. Vandenburgh H, Kaufman S. In vitro model for stretchinduced hypertrophy of skeletal muscle. Science 1979; 203: 265–8. 62. Breuls RG, Jiya TU, Smit TH. Scaffold stiffness influences cell behavior: opportunities for skeletal tissue engineering. Open Orthop J 2008; 2: 103–9. 63. Huang YC, Dennis RG, Larkin L, Baar K. Rapid formation of functional muscle in vitro using fibrin gels. J Appl Physiol 2005; 98: 706–13. 64. Zhu J, Marchant RE. Design properties of hydrogel tissueengineering scaffolds. Expert Rev Med Devices 2011; 8: 607–26. 65. Borschel GH, Dennis RG, Kuzon WM, Jr. Contractile skeletal muscle tissue-engineered on an acellular scaffold. Plast Reconstr Surg 2004; 113: 595–602; discussion 3–4. 66. Moon du G, Christ G, Stitzel JD, Atala A, Yoo JJ. Cyclic mechanical preconditioning improves engineered muscle contraction. Tissue Eng Part A 2008; 14: 473–82. 67. Corona BT, Ward CL, Baker HB, Walters TJ, Christ GJ. Implantation of in vitro tissue engineered muscle repair constructs and bladder acellular matrices partially restore in vivo skeletal muscle function in a rat model of volumetric muscle loss injury. Tissue Eng Part A 2014; 20: 705–15. 68. Thorrez L, Shansky J, Wang L, Fast L, VandenDriessche T, Chuah M, et al. Growth, differentiation, transplantation and survival of human skeletal myofibers on biodegradable scaffolds. Biomaterials 2008; 29: 75–84. 69. Thorrez L, Vandenburgh H, Callewaert N, Mertens N, Shansky J, Wang L, et al. Angiogenesis enhances factor IX delivery and persistence from retrievable human bioengineered muscle implants. Mol Ther 2006; 14: 442–51. 70. Cerletti M, Jurga S, Witczak CA, Hirshman MF, Shadrach JL, Goodyear LJ, et al. Highly efficient, functional engraftment of skeletal muscle stem cells in dystrophic muscles. Cell 2008; 134: 37–47. 71. Montarras D, Morgan J, Collins C, Relaix F, Zaffran S, Cumano A, et al. Direct isolation of satellite cells for skeletal muscle regeneration. Science 2005; 309: 2064–7. 72. Sacco A, Doyonnas R, Kraft P, Vitorovic S, Blau HM. Selfrenewal and expansion of single transplanted muscle stem cells. Nature 2008; 456: 502–6. 73. Bilodeau K, Mantovani D. Bioreactors for tissue engineering: focus on mechanical constraints. A comparative review. Tissue Eng 2006; 12: 2367–83. C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

74. Rossi CA, Flaibani M, Blaauw B, Pozzobon M, Figallo E, Reggiani C, et al. In vivo tissue engineering of functional skeletal muscle by freshly isolated satellite cells embedded in a photopolymerizable hydrogel. FASEB J 2011; 25: 2296–304. 75. Corona BT, Wu X, Ward CL, McDaniel JS, Rathbone CR, Walters TJ. The promotion of a functional fibrosis in skeletal muscle with volumetric muscle loss injury following the transplantation of muscle-ECM. Biomaterials 2013; 34: 3324–35. 76. Warren GL, Hayes DA, Lowe DA, Armstrong RB. Mechanical factors in the initiation of eccentric contraction-induced injury in rat soleus muscle. J Physiol 1993; 464: 457–75. 77. Warren GL, Hayes DA, Lowe DA, Prior BM, Armstrong RB. Materials fatigue initiates eccentric contraction-induced injury in rat soleus muscle. J Physiol 1993; 464: 477–89. 78. Ono Y, Boldrin L, Knopp P, Morgan JE, Zammit PS. Muscle satellite cells are a functionally heterogeneous population in both somite-derived and branchiomeric muscles. Dev Biol 2010; 337: 29–41. 79. Pavlath GK, Thaloor D, Rando TA, Cheong M, English AW, Zheng B. Heterogeneity among muscle precursor cells in adult skeletal muscles with differing regenerative capacities. Dev Dyn 1998; 212: 495–508. 80. Hanes MC, Weinzweig J, Kuzon WM, Panter KE, Buchman SR, Faulkner JA, et al. Contractile properties of single permeabilized muscle fibers from congenital cleft palates and normal palates of Spanish goats. Plast Reconstr Surg 2007; 119: 1685–94. 81. Lindman R, Paulin G, Stal PS. Morphological characterization of the levator veli palatini muscle in children born with cleft palates. Cleft Palate Craniofac J 2001; 38: 438–48. 82. Rader EP, Cederna PS, Weinzweig J, Panter KE, Yu D, Buchman SR, et al. Contraction-induced injury to single permeabilized muscle fibers from normal and congenitallyclefted goat palates. Cleft Palate Craniofac J 2007; 44: 216–22. 83. Kurokawa K, Abe S, Sakiyama K, Takeda T, Ide Y, Ishigami K. Effects of stretching stimulation with different rates on the expression of MyHC mRNA in mouse cultured myoblasts. Biomed Res 2007; 28: 25–31. 84. Gibson MC, Schultz E. The distribution of satellite cells and their relationship to specific fiber types in soleus and extensor digitorum longus muscles. Anat Rec 1982; 202: 329–37. 85. Schmalbruch H, Hellhammer U. The number of nuclei in adult rat muscles with special reference to satellite cells. Anat Rec 1977; 189: 169–75. 86. Goodman CA, Kotecki JA, Jacobs BL, Hornberger TA. Muscle fiber type-dependent differences in the regulation of protein synthesis. PLoS One 2012; 7: e37890. 87. Bylund-Fellenius AC, Ojamaa KM, Flaim KE, Li JB, Wassner SJ, Jefferson LS. Protein synthesis versus energy state in contracting muscles of perfused rat hindlimb. Am J Physiol 1984; 246: E297–305. 88. Dreyer HC, Fujita S, Cadenas JG, Chinkes DL, Volpi E, Rasmussen BB. Resistance exercise increases AMPK activity and reduces 4E-BP1 phosphorylation and protein synthesis in human skeletal muscle. J Physiol 2006; 576: 613–24. 89. Tipton KD, Elliott TA, Ferrando AA, Aarsland AA, Wolfe RR. Stimulation of muscle anabolism by resistance exercise and ingestion of leucine plus protein. Appl Physiol Nutr Metab 2009; 34: 151–61.


Mechanical cues in orofacial tissue engineering and regenerative medicine

90. Passey S, Martin N, Player D, Lewis MP. Stretching skeletal muscle in vitro: does it replicate in vivo physiology? Biotechnol Lett 2011; 33: 1513–21. 91. Carvajal Monroy PL, Grefte S, Kuijpers-Jagtman AM, Helmich MPAC, Ulrich DJO, Von den Hoff JW, et al. A rat model for muscle regeneration in the soft palate. PLoS One 2013; 8: 1–8. 92. de Vries HJ, Enomoto DN, van Marle J, van Zuijlen PP, Mekkes JR, Bos JD. Dermal organization in scleroderma: the fast Fourier transform and the laser scatter method objectify fibrosis in nonlesional as well as lesional skin. Lab Invest 2000; 80: 1281–9. 93. Smitha B, Donoghue M. Clinical and histopathological evaluation of collagen fiber orientation in patients with oral submucous fibrosis. J Oral Maxillofac Pathol 2011; 15: 154– 60. 94. Ogawa R. Mechanobiology of scarring. Wound Repair Regen 2011; 19 (Suppl 1): s2–9. 95. Wray RC. Force required for wound closure and scar appearance. Plast Reconstr Surg 1983; 72: 380–2. 96. Agache PG, Monneur C, Leveque JL, De Rigal J. Mechanical properties and Young’s modulus of human skin in vivo. Arch Dermatol Res 1980; 269: 221–32. 97. Corr DT, Gallant-Behm CL, Shrive NG, Hart DA. Biomechanical behavior of scar tissue and uninjured skin in a porcine model. Wound Repair Regen 2009; 17: 250–9. 98. Clark JA, Cheng JC, Leung KS. Mechanical properties of normal skin and hypertrophic scars. Burns 1996; 22: 443–6. 99. Ogawa R, Okai K, Tokumura F, Mori K, Ohmori Y, Huang C, et al. The relationship between skin stretching/contraction and pathologic scarring: the important role of mechanical forces in keloid generation. Wound Repair Regen 2012; 20: 149–57. 100. Gurtner GC, Dauskardt RH, Wong VW, Bhatt KA, Wu K, Vial IN, et al. Improving cutaneous scar formation by controlling the mechanical environment: large animal and phase I studies. Ann Surg 2011; 254: 217–25. 101. Daigle P, Despatis MA, Grenier G. How mechanical deformations contribute to the effectiveness of negative-pressure wound therapy. Wound Repair Regen 2013; 21: 498–502. 102. Azzopardi EA, Boyce DE, Dickson WA, Azzopardi E, Laing JH, Whitaker IS, et al. Application of topical negative pressure (vacuum-assisted closure) to split-thickness skin grafts: a structured evidence-based review. Ann Plast Surg 2013; 70: 23–9. 103. Bloemen MC, van der Wal MB, Verhaegen PD, Nieuwenhuis MK, van Baar ME, van Zuijlen PP, et al. Clinical effectiveness of dermal substitution in burns by topical negative pressure: a multicenter randomized controlled trial. Wound Repair Regen 2012; 20: 797–805. 104. Lancerotto L, Bayer LR, Orgill DP. Mechanisms of action of microdeformational wound therapy. Semin Cell Dev Biol 2012; 23: 987–92. 105. Macintyre L, Baird M. Pressure garments for use in the treatment of hypertrophic scars—a review of the problems associated with their use. Burns 2006; 32: 10–5. 106. Schor SL, Ellis I, Irwin CR, Banyard J, Seneviratne K, Dolman C, et al. Subpopulations of fetal-like gingival fibroblasts: characterisation and potential significance for wound healing and the progression of periodontal disease. Oral Dis 1996; 2: 155–66. 107. Hakkinen L, Uitto VJ, Larjava H. Cell biology of gingival wound healing. Periodontol 2000 2000; 24: 127–52.


Brouwer et al.

108. Mah W, Jiang G, Olver D, Cheung G, Kim B, Larjava H, et al. Human gingival fibroblasts display a non-fibrotic phenotype distinct from skin fibroblasts in three-dimensional cultures. PLoS One 2014; 9: e90715. 109. Glim JE, Everts V, Niessen FB, Ulrich MM, Beelen RH. Extracellular matrix components of oral mucosa differ from skin and resemble that of foetal skin. Arch Oral Biol 2014; 59: 1048–55. 110. Aarabi S, Bhatt KA, Shi Y, Paterno J, Chang EI, Loh SA, et al. Mechanical load initiates hypertrophic scar formation through decreased cellular apoptosis. FASEB J 2007; 21: 3250–61. 111. Junker JP, Kratz C, Tollback A, Kratz G. Mechanical tension stimulates the transdifferentiation of fibroblasts into myofibroblasts in human burn scars. Burns 2008; 34: 942–6. 112. Chiquet M, Sarasa-Renedo A, Tunc-Civelek V. Induction of tenascin-C by cyclic tensile strain versus growth factors: distinct contributions by Rho/ROCK and MAPK signaling pathways. Biochim Biophys Acta 2004; 1693: 193–204. 113. Gilbert TW, Stewart-Akers AM, Sydeski J, Nguyen TD, Badylak SF, Woo SL. Gene expression by fibroblasts seeded on small intestinal submucosa and subjected to cyclic stretching. Tissue Eng 2007; 13: 1313–23. 114. Hinz B, Mastrangelo D, Iselin CE, Chaponnier C, Gabbiani G. Mechanical tension controls granulation tissue contractile activity and myofibroblast differentiation. Am J Pathol 2001; 159: 1009–20. 115. Derderian CA, Bastidas N, Lerman OZ, Bhatt KA, Lin SE, Voss J, et al. Mechanical strain alters gene expression in an in vitro model of hypertrophic scarring. Ann Plast Surg 2005; 55: 69–75; discussion. 116. Grinnell F, Zhu M, Carlson MA, Abrams JM. Release of mechanical tension triggers apoptosis of human fibroblasts in a model of regressing granulation tissue. Exp Cell Res 1999; 248: 608–19. 117. Peters AS, Brunner G, Blumbach K, Abraham DJ, Krieg T, Eckes B. Cyclic mechanical stress downregulates endothelin-1 and its responsive genes independently of TGFbeta1 in dermal fibroblasts. Exp Dermatol 2012; 21: 765–70. 118. Kessler D, Dethlefsen S, Haase I, Plomann M, Hirche F, Krieg T, et al. Fibroblasts in mechanically stressed collagen lattices assume a “synthetic” phenotype. J Biol Chem 2001; 276: 36575–85. 119. Xie KY, Yang L, Chen K, Li Q. In vitro study of the effect of cyclic strains on the dermal fibroblast (GM3384) morphology—mapping of cell responses to strain field. Med Eng Phys 2012; 34: 826–31. 120. Berry CC, Cacou C, Lee DA, Bader DL, Shelton JC. Dermal fibroblasts respond to mechanical conditioning in a strain profile dependent manner. Biorheology 2003; 40: 337–45. 121. Blaauboer ME, Smit TH, Hanemaaijer R, Stoop R, Everts V. Cyclic mechanical stretch reduces myofibroblast differentiation of primary lung fibroblasts. Biochem Biophys Res Commun 2011; 404: 23–7. 122. Danciu TE, Gagari E, Adam RM, Damoulis PD, Freeman MR. Mechanical strain delivers anti-apoptotic and proliferative signals to gingival fibroblasts. J Dent Res 2004; 83: 596–601. 123. Guo F, Carter DE, Leask A. Mechanical tension increases CCN2/CTGF expression and proliferation in gingival fibroblasts C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

Brouwer et al.

Mechanical cues in orofacial tissue engineering and regenerative medicine

via a TGFbeta-dependent mechanism. PLoS One 2011; 6: e19756. 124. Bush KA, Pins GD. Development of microfabricated dermal epidermal regenerative matrices to evaluate the role of cellular microenvironments on epidermal morphogenesis. Tissue Eng Part A 2012; 18: 2343–53. 125. Clement AL, Moutinho TJ, Jr., Pins GD. Micropatterned dermal-epidermal regeneration matrices create functional niches that enhance epidermal morphogenesis. Acta Biomater 2013; 9: 9474–84. 126. Lammers G, Roth G, Heck M, Zengerle R, Tjabringa GS, Versteeg EM, et al. Construction of a microstructured collagen membrane mimicking the papillary dermis architecture and guiding keratinocyte morphology and gene expression. Macromol Biosci 2012; 12: 675–91. 127. Raut AS, Prabhu RH, Patravale VB. Psoriasis clinical implications and treatment: a review. Crit Rev Ther Drug Carrier Syst 2013; 30: 183–216.

C 2015 by the Wound Healing Society Wound Rep Reg (2015) 23 302–311 V

128. Kim HN, Hong Y, Kim MS, Kim SM, Suh KY. Effect of orientation and density of nanotopography in dermal wound healing. Biomaterials 2012; 33: 8782–92. 129. Verhaegen PD, Schouten HJ, Tigchelaar-Gutter W, van Marle J, van Noorden CJ, Middelkoop E, et al. Adaptation of the dermal collagen structure of human skin and scar tissue in response to stretch: an experimental study. Wound Repair Regen 2012; 20: 658–66. 130. Jansen RG, van Kuppevelt TH, Daamen WF, KuijpersJagtman AM, Von den Hoff JW. Tissue reactions to collagen scaffolds in the oral mucosa and skin of rats: environmental and mechanical factors. Arch Oral Biol 2008; 53: 376–87. 131. Wang X, You C, Hu X, Zheng Y, Li Q, Feng Z, et al. The roles of knitted mesh-reinforced collagen-chitosan hybrid scaffold in the one-step repair of full-thickness skin defects in rats. Acta Biomater 2013; 9: 7822–32. 132. Balestrini JL, Billiar KL. Equibiaxial cyclic stretch stimulates fibroblasts to rapidly remodel fibrin. J Biomech 2006; 39: 2983–90.


Mechanical cues in orofacial tissue engineering and regenerative medicine.

Cleft lip and palate patients suffer from functional, aesthetical, and psychosocial problems due to suboptimal regeneration of skin, mucosa, and skele...
194KB Sizes 1 Downloads 10 Views