Skeletal muscle tissue engineering
D. Klumpp, R.E. Horch and J.P. Beier, University Hospital of Erlangen, Germany
Abstract:
Engineering functional skeletal muscle tissue is still a challenge, in particular in clinically relevant sizes. The development of a suitable scaffold for muscle tissue engineering in vivo remains a major obstacle. Materials and different architectures used for engineering skeletal muscle are presented here and experiences with electrospun scaffolds are highlighted. Characteristic demands of skeletal muscle tissue on a suitable scaffold including the necessary vascularization of implanted scaffolds and potential factors for myogenic differentiation are described and critically discussed. Finally, the options for future developments in the field of skeletal muscle tissue engineering are presented.
Key words
skeletal muscle tissue engineering; electrospinning; electrospun nanofibers; myoblasts; mesenchymal stem cells (MSCs)
17.1 Introduction
Engineering functional skeletal muscle tissue is still a challenge and clinically relevant sizes of functional skeletal muscle tissue have not yet been engineered. However, there are multiple options for clinical applications, so skeletal muscle tissue engineering is still a developing field. In the past, tissue engineering of skeletal muscle has had its first important phase in the late 1980s and early 1990s when Vandenburgh and Karlisch (1989) engineered contracting muscle cells in vitro for the first time. Thereafter, the myooids – muscle tissue engineered in vitro with a maximum thickness of 1 mm (Dennis et al., 2001) – introduced by Strohman et al. (1990), raised hopes that clinically relevant sizes of functional skeletal muscle may be engineered in the near future. Many problems still exist: one is to find the right scaffold for muscle tissue engineering in vivo. The materials and different architectures which have been used for functional skeletal muscle in the past will be discussed and experience with electrospun scaffolds will be highlighted in this chapter. Furthermore, the chapter will explain the characteristic demands of skeletal muscle tissue on a suitable scaffold including the necessary vascularization of implanted scaffolds in vivo and potential factors for myogenic differentiation. The options for future developments will be presented at the end of the chapter.
17.2 Clinical and scientific applications
17.2.1 Skeletal muscle for drug screening in vitro
Engineering of functional skeletal muscle in vitro offers the possibility of investigating potential side effects of newly developed drugs in general. Herein, the influence of certain drugs as well as the physiology of muscle tissue can be studied in detail without the necessity of laborious in vivo studies. For example, Kaji et al. (2010) have investigated the influence of exercise and insulin on glucose uptake in skeletal muscle tissue engineered and eletrically stimulated in vitro. Moreover, drug screening in vitro is an essential and economic tool for the development of orphan drugs for myopathies, e.g. Duchenne’s muscular dystrophy (DMD). Therefore, Vandenburgh and co-workers (2008) introduced a drug screening platform using dystrophic muscle tissue. Thus, costly animal studies are avoided which enables the screening of a variety of orphan drugs for musculoskeletal disorders. This shows the clinical relevance of engineered skeletal muscle tissue.
17.2.2 Clinical applications for engineered skeletal muscle tissue in vivo
In the past, tissue engineering of skeletal muscle has been tried in various studies for the treatment for muscle diseases, e.g. DMD. Though initially promising, the implantation of muscle precursor cells or engineered skeletal muscle tissue did not meet expectations. Herein, other options, e.g. gene therapy with antisense oligonucleotides (AONs), are more promising (Williams et al., 2008; Nelson et al., 2009).
The transfer of myocutaneous free flaps to cover soft tissue defects is one of the most frequent applications in a clinical setting. Unfortunately, this leads to a functional loss and loss of volume at the donor side, known as donor side morbidity. Moreover, the transferred tissue is supplied with blood by a microsurgically created anastomosis. This has the danger of flap failure due to technical problems at the anastomosis or reduced blood supply at the recipient site. Engineered flaps could be an alternative to cover soft tissue defects without the disadvantage of donor side morbidity. Even repeated coverages in case of flap failure or secondary defects could be mastered this way.
In certain cases, the transferred muscle tissue is not used as coverage but as a functional substitute for paralysed, missing or denervated muscles. The most common situation for such a functional substitute is paralysis of the facial nerve. The transfer of innervated free flaps, e.g. the gracilis muscle of the thigh, can be used to reanimate the face (Terzis and Konofaos, 2008). Though the results are encouraging, this technique often does not produce satisfactory results (Terzis and Noah, 1997). Thus, in such special situations engineered muscle tissue could be designed more closely in size and shape to the original muscle. In this case, the engineered muscle tissue should also contain a motoric nerve of an adequate size to create a microsurgical anastomosis with a motoric nerve at the recipient site.
17.3 Characteristics of skeletal muscle
17.3.1 Architecture of mature skeletal muscle and extracellular matrix
The characteristic architecture of mature skeletal muscle lies in its highly orientated muscle fibers which are organized in bundles and the latter form muscles with a distinct function. The parallel orientation of the muscle fiber is a prerequisite to enable a longitudinal force generation which constitutes the exact function of the muscle.
Furthermore, muscle tissue – including skeletal as well as cardiac muscle – is one of the most vulnerable tissues regarding its tolerance of hypoxia. Thus, an inadequate vascularization rapidly leads to extended necrosis of muscle tissue and/or transformation into fibrous scar tissue, which is not contractile anymore. Therefore, vascularization plays an important role for tissue engineering of skeletal muscle in clinically relevant sizes. The vascularization as well as pro-angiogenic factors will be discussed in the following sections.
The characteristic parallel alignment of the muscle fibers is also found in the extracellular matrix (ECM) of skeletal muscle tissue. Natural ECM mainly consists of collagen type I fibers and elastin fibers with diameters in the range of 150 to 230 nm.
While stability is one of the main points of any ECM in general, the ECM of skeletal muscle tissue especially needs a certain elasticity to support and conduct the contraction of the muscle fibers. Thus, a potential scaffold for skeletal muscle tissue must balance stability and elasticity and enable the parallel alignment of muscle precursor cells. Boontheekul et al. (2007) showed that myogenic differentiation is clearly influenced by the stiffness of the scaffold.
17.3.2 Satellite cells in muscle regeneration and as a cell source for tissue engineering
In vivo, injured muscle tissue is regenerated by satellite cells (Snow, 1977). These cells were first described by Mauro (1961) and named satellite cells because of their localization beneath the basal lamina of muscle fibers. Today, the ‘satellite cell’ is identified by its expression of the transcription factor Pairedbox 7 (Pax7) (Seale et al., 2000). However, satellite cells enclose two sub-populations of cells depending on their co-expression of MyoD, a myogenic transcription factor. The main population of satellite cells (approximately 90%) also expresses MyoD (also named Myf5), which marks their commitment to the myogenic lineage (Weintraub et al., 1991). Along with this myogenic pre-differentiation goes the disadvantage of a decreased proliferation rate complicating the generation of clinically relevant sizes of muscle tissue in vitro. As an advantage, this commitment to the myogenic line also enables a safe application of satellite cells in a clinical setting without a significant risk of trans- or dedifferentiation. Therefore, satellite cells have been used mainly as cell source for skeletal muscle tissue engineering in the past (Otto et al., 2009).
Only a minor sub-population of approximately 10 per cent does not express the myogenic marker MyoD and shows in turn stem cell properties with the possibility of differentiation into multiple mesenchymal cell populations. This sub-population regenerates the Pax+/MyoD+-cell population in vivo through asymmetric self-renewal (Kuang et al., 2007). The regernerate potential of the Pax+/MyoD− sub-population is astonishing, since whole muscle bundles can be regenerated in vivo (Le Grand and Rudnicki, 2007). However, isolated satellite cells show a significant loss of their proliferative potential when cultured in vitro (Yaffe, 1968). This phenomenon has been explained with the loss of the stem cell niche, which is based on the cell contact to the basal lamina and the ECM (Boonen and Post, 2008). Thus, the generation of a suitable number of muscle-precursor cells for skeletal muscle tissue in vitro is still a challenge. Gilbert and his group could show that the satellite cell niche can be mimicked in vitro by cultivating isolated satellite cells on laminin cross-linked polyethylene glycol (PEG) hydrogels with an elasticity of 12 kPa, which exactly equals the elasticity of the basal lamina in skeletal muscle tissue (Gilbert et al., 2010).
Another cell population in skeletal muscle tissue has been characterized by Popescu and coworkers: the telocyte, a Pax negative cell population with typical prolongations, called ‘telopodes’ (Popescu and Faussone-Pellegrini, 2010). The telopodes are located near vessels, nerves and muscle progenitor cells in mature cardiac and skeletal muscle tissue (Gherghiceanu and Popescu, 2010). Therefore, telocytes are believed to regulate myogenesis and muscle tissue regeneration. This cell type expresses c-kit and caveolin-1 but is also known to secrete vascular endothelial growth factor (VEGF) which could be an interesting property in muscle tissue engineering in vivo (Suciu et al., 2010). Thus, the functions of muscle-precursor cells including Pax+/MyoD+-, Pax+/MyoD− cells and telocytes are exactly balanced for muscle regeneration in vivo. Therefore, these muscle precursor cells isolated from mature muscle tissue seem to be the most suitable cell source for skeletal muscle tissue engineering. However, the isolation of an adequate number of precursor cells and the preservation of the proliferation rate in vitro are still obstacles that must be overcome before a clinical application can be realized.
The use of mesenchymal stem cells (MSCs) has often been proposed because of their higher proliferation rate in vitro (Deans and Elisseeff, 2009). The use of adipose-derived mesenchymal stem cells (ADMSCs) offers an easy, accessible and interesting possibility for their application in a clinical setting (Zhu et al., 2008). However, myogenic differentiation of MSCs is challenging in vitro as well as in vivo. Brazelton et al. (2003) have reported a poor incorporation rate of 5–10% of transplanted MSCs in skeletal muscle tissue in vivo. Though the majority of transplanted MSCs shows no myogenic differentiation in vivo, the transplanted MSCs contribute to myogenic regeneration in injured or dystrophic muscle tissue through paracrine effects (Satija et al., 2009). The secretion of anti-inflammatory, anti-apoptotic and angiogenic factors by transplanted MSCs constitutes this paracrine effects which support the local regeneration of injured skeletal muscle tissue (Meirelles Lda and Nardi, 2009). The pro-angiogenic effect is also seen when the secretome of MSCs is added in vivo (Estrada et al., 2009). However, MSCs also fuse with co-cultured with myoblasts in vitro (Beier et al., 2011) which underlines their versatile contribution to muscle regeneration. As an interesting feature, MSCs can be transplanted allogenically due to their low immunogenicity (García-Castro et al., 2008; Rossignol et al., 2009).
Besides MSCs, induced pluripotent stem cells (iPSCs) offer an even higher proliferation rate but also a seriously augmented risk of dedifferentiation and tumorigenicity in vivo (Klumpp et al., 2010). Therefore, this cell source has only rarely been studied for tissue engineering applications in vivo, as yet.
17.4 Potential scaffolds for skeletal muscle tissue engineering
17.4.1 Materials and scaffolds
Though a variety of materials exist, only a few meet the special demands of skeletal muscle tissue. In the first place, biocompatibility and the absence of tumorigenicity are vital features for application in vivo.Therefore, certain materials which have been widely used for tissue engineering are not suitable for in vivo studies. Matrigel™ for example, a hydrogel extracted from Engelbreth-Holm-Swarm (EHS) mouse sarcoma cells and containing a variety of growth factors, shows good results in vitro but cannot be used in a clinical setting (Vukicevic et al., 1992). Components of the natural ECM of skeletal muscle, e.g. collagen I and elastin, are the most suitable candidates for tissue engineering of skeletal muscle in vivo. Their use in vivo is non-hazardous and bovine collagen I shows a very low immunogenicity in xenogenous models in vivo (Peng et al., 2010). However, their disadvantage lies in their fast degradation in vivo. The stability of fibrin, elastin as well as collagen is completely lost after several weeks in vivo (Arkudas et al., 2009a). Since vascularization, neurotization and myogenic differentiation of implanted myoblasts into mature muscle fibers takes several months, these materials are not suitable as the only components in clinical settings. As an exception, silk fibroin shows a long-term stability over one year in vivo at a concentration of 17% (solved in hexafluoro-iso-propanol, HFIP) (Zhao et al., 2003). Even when an all-aqueous dissolution instead of the cell-toxic HFIP is used, fibroin with a concentration of 10% is stable up to 6 months in vivo (Wang et al., 2008). The second component of silk, i.e. sericin, has been identified as the reason for the initially high immunogenicity of silk hydrogels (Panilaitis et al., 2003). The biocompatibility of pure fibroin equals that of materials like collagen (Meinel et al., 2005). Therefore, silk fibroin extracted from silk worms has been analyzed in several studies in vitro (Mandal and Kundu, 2009) and in vivo (MacIntosh et al., 2008; Unger et al., 2010) as hydrogels or sponge-like scaffolds. The disadvantage of the highly hydrophobic fibroin lies in its low cell attachment and low elasticity which renders the material not suitable for engineering skeletal muscle.
Concerning stability, biodegradable synthetic materials are a cost-effective and easy to handle alternative. Materials like poly-l-lactic acid (PLLA) or poly-ε-caprolyctone (PCL) are stable over approximately one year in vivo (Gunatillake and Adhikari, 2003; Bolgen et al., 2005). Though both materials are biocompatible, the acidic degradation products of PLLA can lead to cell toxic effects (Ignatius and Claes, 1996). Furthermore, the disadvantages of PCL are mostly its high hydrophobicity and low elasticity. Though these features equal silk fibroin, PCL is easier to handle and can be used in combination with other biomaterials. Herein, PCL can be coated or blended with materials like collagen (Schnell et al., 2007; Klumpp et al., 2012) or gelatine (Kim et al., 2010) to enhance cell attachment.
17.4.2 Three-dimensional architecture
The challenge of designing scaffolds for tissue engineering lies in the generation of a three-dimensional architecture mimicking the natural ECM with its biologic as well as mechanic properties. Therefore, potential scaffolds for the tissue engineering of skeletal muscle should reflect the parallel alignment of native myotubes and myofibrils. Following the cell guidance theory described by Curtis and Wilkinson (1997), the myogenic differentiation and parallel alignment of myogenic cells can thus be enhanced (Gingras et al., 2009). Hence matrices like fibrin or other hydrogels with random orientation may not render best results. A possible way to gain parallel alignment inside a scaffold is the unidirectional freeze-drying of hydrogels. Thus, ice crystals form in a spatially orientated pattern leading to orientated pores afterwards. This method has successfully been used for materials like collagen and silk fibroin (Madaghiele et al., 2008) and the pore size can be controlled by the freezing temperature (Schoof et al., 2001). The disadvantages are that the alignment of the pores remains spatial and the scaffold itself shows a random architecture, though.
The most suitable method to achieve strict parallel alignment is the electrospinning technique (Ayres et al., 2006). The formation of fibers by electrical voltage is a complex method and the multiple parameters like concentration of the solution, flow rate and viscosity as well as the voltage and distance to the counter-electrode enables the adjustment of the resulting scaffold’s properties in a wide range (Boudriot et al., 2006). Thus, a plethora of biomaterials can be spun to nano- or microfibers like hyaluronic acid, collagen I, elastin as well as synthetic polymers (Sell et al., 2009). As discussed previously, the use of synthetic polymers result in stable and slow-degrading scaffolds but with low cell attachment due to their hydrophobicity. Therefore, post-spinning modifications like coating (Riboldi et al., 2005) or plasma treatment (Martins et al., 2010) can enhance cell attachment in vitro and in vivo. However, these methods are limited to a few fiber layers. For three-dimensional nanofibrous scaffolds synthetic polymers can be blended with biopolymers like collagen, hyaluron acid or elastin (Schnell et al., 2007). Furthermore, two different polymer solutions can be spun separately into one fiber. The surface of the resulting fibers of this core–shell spinning method is formed solely by the surrounding polymer. Zhang et al. (2005) showed that the core–shell spinning technique using PCL as core and collagen I as shell is superior to a post-spinning coating of PCL fibers with collagen. Regarding the control of the pore size or interspaces between the fibers, this is still a challenge and the most prominent disadvantage of the electrospinning method in general. In scaffolds with parallel fiber alignment especially, the fibers are densly packed after spinning and the absence of adequate interspaces hinders cell migration and – more vital for in vivo application – also ingrowth of vessels into the scaffold (Telemeco et al., 2005). Baker and co-workers have addressed this point by co-spinning a water-soluble polymer solution, e.g. poly(ethylene-oxide) (PEO). The resulting fibers – named sacrificial fibers – can be solved after spinning and leave behind interspaces for enhanced cell migration and vascularization (Baker et al., 2008). Thus, the properties of PCL–collagen blend nanofibers meet the demand of muscle tissue, especially when adequate interspaces can be generated.
17.5 Smart matrices
The challenge to meet every demand of the tissue engineered within a certain matrix has led to the development of smart matrices which mimic the natural ECM more closely. Fibrin gel, for example, shows binding sites for the pro-angiogenic factor VEGF. Thus, fibrin hydrogels naturally enhance vascularization in vivo (Arkudas et al., 2009b). In hydrogels in general, different factors can be mixed into the matrix easily. Another option is to bind VEGF and other factors to nanoparticles which shows a therapeutic effect for example in ischemic muscle (Kim et al., 2011). Since the application of VEGF is expensive, the restriction of the pro-angiogenic effect to the ischemic site only is economically important. Therefore, Ye et al. (2011) introduced a hypoxia-regulated system releasing VEGF selectively at the ischemic sites in myocardial repair.
The electrospinning technique holds multiple options for the design of smart matrices as well. Herein, the method of core–shell spinning can also be used to create a drug delivery system with a controlled release of different drugs (Jiang et al., 2005). Even enclosing pro-angiogenic factors into the shell and different factors into the core is technically possible. The time of release is also controllable through the degradation time of the material and the way of binding the factor to certain materials. Thus, smart matrices could facilitate the ingrowth of vessels through an early release of proangiogenic factors like VEGF (Yang et al., 2010) as well as the prolonged release of factors enhancing differentiation of the implanted cells inside the matrix.
Different factors exist to either enhance cell proliferation or myogenic differentiation. Among these insuline-like growth factor (IGF-1) is one of the best known factors, increasing the proliferation of muscle precursor cells as well as promoting myogenic differentiation (Allen and Boxhorn, 1989). In vivo, overexpression of IGF-1 accelerates muscle regeneration after injury with less fibrosis (Menetrey et al., 2000; Sato et al., 2003) and even results in muscular hypertrophy in normal muscle tissue (Adams and McCue, 1998). When muscle precursor cells are implanted in vivo, the release or overexpression of IGF-1 enhances survival of the implanted cells (Wang et al., 2009). Even without the implantation of muscle precursor cells, a local release IGF-1 is able to attract stem cells, e.g. MSCs in vivo (Haider et al., 2008). These properties are the reason for the extensive use of IGF-1 in vitro and in vivo for muscle tissue engineering.
The effect of attracting autologous stem cells and muscle precursor cells to the implanted matrix or site of injury in musle tissue is also seen selectively when stromal-cell derived factor 1α (SDF-1α) is overexpressed (Haider et al., 2008) or released by the implanted matrix (Grefte et al., 2007). SDF-1α is also a downstream factor which is activated by IGF-1. Furthermore, a less well-known but also potential factor is akirin-1 which activates quiescent satellite cells. Thus, muscle regeneration is accelerated but also the myogenic differentiation of implanted satellite cells in vivo increases. The pro-myogenic effect of akirin-1 is the known result of IGF-2 activation (Marshall et al., 2008). Thus, different aspects of muscle regeneration and differentiation can be addressed by multiple factors. However, in a clinical setting the application of growth factors in humans should be discussed critically. Though a definite risk of neoplasma has not been scientifically proven in vivo yet, the potential tumorigenicity of the different factors has to be analyzed by future research.
Searching for a safe alternative for clinical application, the newly described microRNAs (miR) could offer an interesting option. These non-coding RNAs show a length of 20–22 nucleotides and certain miRNAs are muscle specific, i.e. miR-1, miR-133 and miR-206 (Callis et al., 2008). Injection of all three miRNAs into regenerating muscle tissue in vivo results in accelerated regeneration with less formation of fibrous tissue (Nakasa et al., 2010). Herein, microRNAs concurrently regulate proliferation and differentiation in regenerating muscle tissue. Whereas miR-133 increases the proliferation of muscle precursor cells, miR-1 and miR-206 promote myogenic differentiation (Chen et al., 2006; Kim et al., 2006). Since both miRNAs only enhance myogenic differentiation and even decrease the proliferation rate of muscle precursor cells, these miRNAs could be a safe alternative in a clinical setting. Though, the long-term effects and potential tumorigenicity have to be analyzed in further studies.
Apart from the factors discussed before, Wilson and Harris showed in 1993 that myogenic differentiation is also induced by electrical stimulation. Electrical stimulation has therefore been studied with different scaffolds in vitro (Stern-Straeter et al., 2005) and in vivo (Fujita et al., 2007) in general. Especially in case of electrospun scaffolds, the spinning of electrical conductive fibers is possible. Therefore, the synthetic polymers polypyrrole as well as poly(aniline) (PANi) are conductive and biodegradable (Gomez and Schmidt, 2007; Borriello et al., 2011). Though cell attachment on both materials is poor due to their hydrophobicity, the co-spinning of electrical conductive fibers into an established nanofiber scaffold could multiply the advantages of a nanofibrous scaffold for the engineering of skeletal muscle tissue, in particular (Li et al., 2006). However, the spinning of such a composite scaffold is technically challenging. In a study of Ghasemi-Mobarakeh et al. (2009), the authors showed enhanced neurite outgrowth when neural cells were cultivated on PANi/PCL/gelatine blend fibers and stimulated electrically. Electrically conductive nanofiber scaffolds also enhance myogenic differentiation which has been shown by Jun and coworkers in vitro (2009). They found a significant up-regulation of myogenin, an early marker of myogenesis, as a result of electrical stimulation.
17.6 Electrospun scaffolds in vivo/arteriovenous (AV)-loop models in the rat
The initial aim of tissue engineering was to generate functional tissues in clinically relevant sizes. Therefore, the generated tissue must clearly exceed the threshold of 1 mm thickness which is the limit for tissue engineering in vitro. Since nutrient supply via sole diffusion is possible up to a distance of 500 μm, the generation of clinically relevant sizes inevitably asks for vascularization of the implanted scaffold and thus the engineered tissue in vivo (Kannan et al., 2005). Therefore, the scaffold is usually implanted first and the cells are added in a second operation after complete vascularization of the matrix. Thus, apoptosis of implanted cells due to a lack of nutrient supply can be prevented (Arkudas et al., 2007). Herein, the pre-vascularization time until vessels are grown into the whole scaffold depends on the material and architecture of the scaffold in generally and the pore size especially. Therefore, a porosity of approximately 90% of a scaffold with high interconnectivity and an adequate pore size enable the migration of precursor and endothelial cells so that vascularization as well as tissue formation is possible not only at the periphery of a matrix but also at the center (Freed et al., 1994; Ishaug-Riley et al., 1998). Regarding muscle precursor cells, the pore size of the scaffold should ideally range between 50 μm and 200 μm (Lee et al., 2008). These prerequisites are well presented in hydrogels or sponges with high porosity where cells can freely migrate through the matrix by degrading the hydrogel. However, in scaffolds with parallel alignment in general and scaffolds electrospun in parallel in particular, the size and interconnectivity of the interspaces remains a challenge, though multiple methods exist to enlarge the interspaces as discussed above.
Recently, the authors have analyzed the vascularization of randomly and PCL–collagen blend nanofiber scaffolds spun in parallel in vivo (Klumpp et al., 2012). In this study, an arteriovenous (AV)-loop was microsurgically created as first desribed by Erol and Spira in 1979 in the groin of rats and implanted into the matrix inside an isolation chamber. Though techniqually challenging and time-consuming, the advantage of this model lies in the strict axial vascularization of the whole matrix. Thus, vessels sprout from the loop vessels inside the chamber only and the engineered tissue inside the chamber can be transplanted with a microsurgical anastomosis of the pedicle to the site where the engineered tissue is needed. Therefore, the AV-loop model is a vital technique for a potential clinical application of tissue engineering in general.
Herein, the authors have recently analyzed the axial vascularization of randomly and scaffolds electrospun in parallel in vivo (Klumpp et al., 2012). In this study, PCL–collagen blend scaffolds were implanted into the AV-loop model in rats and the vascularization after 4 and 8 weeks was analyzed three-dimensionally with micro-CT scans. The electrospun nanofiber scaffolds both showed a long pre-vascularization time of over 4 weeks. Though the total number of vessels inside the scaffolds was higher in the randomly spun nanofiber scaffolds, the group spun in parallel showed a constant vascularization of the center whereas the vessels in the randomly spun group sprouted in the periphery of the scaffolds without an adequate vascularization of the center after 8 weeks in vivo. The more consistant vascularization of parallel nanofiber scafolds is contrary to the small interspaces of between the fibers. Interestingly, the migration of cells through smaller pore sizes has been observed by Zhang et al. (2005) with fibroblasts cultivated on randomly spun PCL–collagen scaffolds before. As an explanation, the dynamic structure of nanofiber scaffolds was postulated: owing to the flexible and fibrous structure, cells can push the fibers aside and thus migrate through the scaffold. Herein, scaffolds spun in parallel are even more flexible since the fibers are not attached to each other. This could possibly explain the more consistant vascularization of the scaffolds.
17.7 Conclusions and future trends
Though the engineering of skeletal muscle tissue is still challenging due to the complex structure and mechanical demands of the contracting tissue, its application in vivo is interesting and clinically relevant. The directional contraction of skeletal muscle asks for a strict parallel alginment of the matrix. Thus, electrospun nanofiber scaffolds can mimick the natural ECM of skeletal muscle tissue very closely and the mechanical properties of electrospun scaffolds can be adjusted to the demands of the tissue. A disadvantage of the electrospinning technique is the poor control of the resulting pore size which is crucial for ingrowing vessels. Therefore, the long period of pre-vascularization in nanofiber scaffolds complicates their application in vivo. Thus, enlarging the pore sizes of electrospun scaffolds as well as accelerating the vascularization through the addition of proangiogenic factors will be vital points for future research. The further development of electrospun scaffolds into drug delivery systems will be an important and interesting field of research in the future. Thus, the challenge of myogenic differentiation of muscle precursor cells in vivo may be facilitated by pro-myogenic factors like IGF-1 or miRNAs.