Artikel
The challenge of musculoskeletal tissue engineering – from cell cultures to large animal models
Die Herausforderung des Tissue Engineering von Skelettmuskelgewebe – von Zellkulturen zu großen Tiermodellen
Suche in Medline nach
Autoren
Veröffentlicht: | 18. August 2015 |
---|
Gliederung
Abstract
Engineering functional skeletal muscle tissue still remains a major challenge. So far clinically relevant sizes of functional skeletal muscle tissue could not be engineered yet. One of the obstacles to overcome is the development of a suitable scaffold for muscle tissue engineering in vivo, another is the lack of differentiation in expanded adult muscle precursor cells. Materials and different architectures of scaffolds which are used for engineering functional skeletal muscle are presented here as well as approaches to the differentiation challenge. Finally the translation from cell culture over small to large animal models for engineering axially vascularized musculoskeletal tissues will be described.
Zusammenfassung
Das Tissue Engineering von Skelettmuskelgewebe stellt noch immer eine große Herausforderung dar. Eine Generierung von funktionellem Neo-Muskelgewebe in klinisch relevanter Größe konnte bis heute noch nicht gezeigt werden. Die wichtigsten Hindernisse, die es zu überwinden gilt, sind die Entwicklung eines auch für die in vivo Anwendung geeigneten Scaffolds sowie der Verlust an Differenzierungsfähigkeit expandierter adulter Muskelvorläuferzellen. Material und Aufbau bereits für das Muskel Tissue Engineering verwendeter Scaffolds sowie eigene und durch andere Gruppen erforschte Ansätze zur Überwindung des Differenzierungsproblems sollen in dieser Übersicht dargestellt werden. Schließlich soll anhand des Stands der derzeitigen Forschung sowie unter Einbeziehung eigener Arbeiten der Weg von der Zellkultur über entsprechende Kleintier- bis hin zu Großtiermodellen aufgezeigt werden, welcher in Zukunft zu der angestrebten Anwendbarkeit beim Menschen führen könnte.
Introduction
Generation of functional skeletal muscle neo-tissue remains a major challenge in tissue engineering. So far functional skeletal muscle tissue in a clinically relevant size could not be created yet. However, there are multiple options for clinical applications so that the skeletal muscle tissue engineering is still a developing field [1]. In the past, tissue engineering of skeletal muscle first evolved in the 80s and early 90s when Vandenburgh et al. engineered contracting muscle cells in vitro for the first time [2]. Thereafter, the myooids – muscle tissue engineered in vitro with a maximum thickness of 1 mm [3] – introduced by Strohman et al. [4], raised hopes that clinically relevant sizes of functional skeletal muscle may be engineered in the near future [1]. However, one of the remaining obstacles is to develop a suitable scaffold for muscle tissue engineering in vitro and in vivo [5]. The materials and different architectures which have been used for functional skeletal muscle in the past and presence will be discussed in this review. Furthermore, the specific demands for skeletal muscle tissue engineering concerning the cell differentiation challenge as well as the task of axial vascularization for future clinical application will be highlighted.
Clinical application for tissue engineering of skeletal muscle
Tissue engineering of skeletal muscle has been in focus of different studies for the treatment for inherent muscular diseases, in particular Duchenne’s muscular dystrophy (DMD) [1]. Though promising in the beginning, the implantation of muscle precursor cells or engineered skeletal muscle tissue did not meet the expectations. Other applications, such as gene therapy with antisense oligonucleotides (AONs) e.g. could also be promising [6], [7].
In plastic and reconstructive surgery, 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 as well as loss of volume at the donor site, known as donor site morbidity. Moreover, the transferred tissue has to be supplied with blood by a microsurgically created anastomosis between the transplant’s (“pedicle”) vessels and the recipient vessels. This comes along with the risk 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 site morbidity [1], [5].
Under specific circumstances the transplanted muscle tissue itself is not only used for covering exposed vulnerable structures, but also as a functional substitute for paralysed, missing or denervated muscles, thus called “free functional muscle transfer”. The most common clinical situation for such demand is facial nerve palsy. The transfer of innervated free flaps, e.g. the gracilis muscle of the thigh as a free transplant or the temporal muscle as a pedicled transfer, is routinely used in clinics to reanimate the facial mimic [8]. Though the results are encouraging, this technique often does not lead to satisfactory results [9]. Thus, in such special situations engineered muscle tissue could be designed to more closely resemble the original muscle in size and shape. Ideally the engineered muscle neo-tissue should also encompass a motor nerve with an adequate calibre to create a microsurgical anastomosis with a motoric nerve at the recipient site [1], [5].
Skeletal muscle tissue
The typical architecture of mature skeletal muscle is based on 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 fibers is a pre-requisite to enable a longitudinal force generation which constitutes the exact function of the muscle [1]. Furthermore, skeletal muscle tissue is one of the most vulnerable tissues regarding its tolerance of hypoxia. Hence 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 paragraphs [10]. While stability is one of the main points of any extracellular matrix (ECM) in general, the ECM of skeletal muscle tissue in particular should exhibit good elasticity to support and conduct the contraction of the muscle fibers. Thus, a potential scaffold for skeletal muscle tissue should balance stability and elasticity and enable the parallel alignment of muscle precursor cells [10]. Besides the necessary mechanical impact on contractility itself, it was also shown that myogenic differentiation is significantly influenced by the stiffness of the scaffold [11].
In vivo, injured muscle tissue is regenerated by satellite cells [12]. These cells were first described by Mauro et al. and named as satellite cells because of their localization beneath the basal lamina of muscle fibers [13]. Today, the “satellite cell” is identified by its expression of the transcription factor Pairedbox 7 (Pax7) [1], [14]. 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 [15]. 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 [16], [17]. 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 [1]. Therefore, satellite cells have been used mainly as cell source for skeletal muscle tissue engineering in the past [18], [19]. Only a minor sub-population of approximately 10 percent 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 [1], [20]. The regenerative potential of the Pax+/MyoD- sub-population is astonishing, since whole muscle bundles can be regenerated in vivo [21]. Though, isolated satellite cells show a significant loss of their proliferative potential when cultured in vitro [22]. 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 [23]. Thus, the generation of a suitable number of muscle-precursor cells for skeletal muscle tissue in vitro remains a challenge. Gilbert and his group demonstrated that the satellite cell niche can be mimicked in vitro by cultivating isolated satellite cells on laminin cross-linked PEG hydrogels with an elasticity of 12 kPa which exactly equals the elasticity of the basal lamina in skeletal muscle tissue [24]. Therefore, these muscle precursor cells isolated from mature muscle tissue seem to be the most suitable cell source for skeletal muscle tissue engineering. Though, the isolation of an adequate number of precursor-cells and the preservation of the proliferation rate in vitro are still an obstacle that has to be overcome before a clinical application could be realized [25].
The use of mesenchymal stem cells (MSC) has often been proposed because of their higher proliferation rate in vitro [26]. Adipose-derived mesenchymal stem cells (ADSCs) in particular promise to offer an easy accessible and highly potential cell source for application in a clinical setting [27]. However, myogenic differentiation of mesenchymal stem cells is challenging in vitro as well as in vivo. Brazelton et al. reported a poor incorporation rate of 5–10% of transplanted MSCs in skeletal muscle tissue in vivo [28]. 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 [29]. The secretion of anti-inflammatory, anti-apoptotic and angiogenic factors by transplanted MSCs constitute these paracrine effects which support the local regeneration of injured skeletal muscle tissue [30]. The pro-angiogenic effect is also seen when the secretome of MSCs is added in vivo [31]. However, mesenchymal stem cells also fuse with co-cultured with myoblasts in vitro [25] which underlines their versatile contribution to muscle regeneration. As an interesting feature, mesenchymal stem cells can be transplanted allogenically due to their low immunogenicity [32], [33]. Recently, induced pluripotent stem cells (iPSCs) were introduced into tissue engineering, displaying an even higher proliferation rate but also a seriously augmented risk of dedifferentiation and tumorigenicity in vivo [5]. Therefore, this cell source has only rarely been studied for tissue engineering applications in vivo, yet [1].
Scaffolds in skeletal muscle tissue engineering
A plethora of materials have been applied and characterized for different tissue engineering purposes, but only few have met the specific demands for skeletal muscle tissue engineering. In the first place, biocompatibility and the absence of tumorigenicity are crucial issues for application in vivo [5]. 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 [34], [35]. Natural components of the native (ECM) of skeletal muscle, e.g. collagen I, elastin or Laminin are promising and clinically applicable candidates for tissue engineering of skeletal muscle in vivo [36]. Their use in vivo is non-hazardous and bovine collagen I shows a very low immunogenicity in xenogenous models in vivo [37]. 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 [38]. Since vascularization, neurotization and myogenic differentiation of implanted myoblasts into mature muscle fibers takes several months, these materials have to be modified in order to make their way into any clinical setting. Concerning stability, biodegradable synthetic materials are a cost-effective and easy to handle alternative. Materials like poly-l-lactic acid (PLLA) or poly-e-caprolactone (PCL) are stable over approximately one year in vivo [39], [40]. Though both materials are biocompatible, the acidic degradation products of PLLA can lead to cell toxic effects [41]. Furthermore, the disadvantages of PCL are mostly its high hydrophobicity and low elasticity [36]. Furthermore, PCL can be coated or blended with materials like collagen [42], [43] or gelatine [44] to enhance cell-attachment. The aim of designing scaffolds for skeletal muscle tissue engineering is to create a three-dimensional architecture mimicking the natural ECM with its biologic as well as mechanic properties as closely as possible. Hence, potential scaffolds for tissue engineering of skeletal muscle should reflect the parallel alignment of native myotubes and myofibrils as well as the elasticity of its native surrounding ECM [1]. Following the cell guidance theory, as first postulated by Curtis and Wilkinson [45], the myogenic differentiation and parallel alignment of myogenic cells can thus be enhanced [46]. Hence matrices like fibrin or other hydrogels with random orientation may not render best results. A possibility to gain parallel alignment within a scaffold is the unidirectional freeze-drying of hydrogels. Thus, ice crystals form in a spatially orientated pattern leading to orientated pores afterwards [1]. This method has successfully been used for materials like collagen and silk fibroin [47] and the pore-size can be controlled by the freezing temperature [48]. As a disadvantage, the alignment of the pores remains spatial and the scaffold itself shows a random architecture, though.
The most promising method to create strictly parallel aligned structures could be the electrospinning technique [49]. 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 [43], [50]. Thus, a plethora of biomaterials can be spun to nano- or microfibers like hyaluronic acid, collagen I, elastin as well as synthetic polymers [1], [51]. In order to enhance the cell-attachment in vitro and in vivo post-spinning modifications like coating [52] or plasma treatment [53] have been developed. However, these methods are limited to few fiber-layers. For three-dimensional nanofibrous scaffolds synthetic polymers can be blended with biopolymers like collagen, hyaluronic acid or elastin. 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 [1]. Zhang et al. demonstrated that the core-shell spinning technique using PCL as core and collagen I as shell could be superior to a post-spinning coating of PCL-fibers with collagen [54]. However, control of the “pore-size” or better “interspaces between the fibers” remains a major challenge and a significant disadvantage of the electrospinning method in general. The more parallel the fibers are aligned the denser they are packed after spinning. As a consequence the absence of adequate interspaces interferes with cell migration and – more vital for in vivo application – also ingrowth of vessels into the scaffolds [43], [55]. 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 [56]. Taken together, PCL-collagen blend nanofibers are – also based on our own experimental results [36], [43] – promising candidates as scaffolds for muscle tissue engineering, pending the development of sufficient interspaces by sacrificial fibers of other means.
Skeletal muscle tissue engineering in vivo
In order to generate functional neo-tissue in clinically relevant size for later clinical application, at least 1 mm in thickness of such neo-muscle has to be achieved. However, thes is usually the limitation for tissue engineering in vitro, based on diffusion characteristics which are generally believed to be limited to a distance of 500 µm. Hence the generation larger neo-tissue constructs needs a sufficient vascularization of the implanted scaffold and thus the engineered tissue in vivo [57]. Usually the scaffolds are implanted first and the cells are added in a second operation after complete vascularization of the matrix or small scaffolds are pre-seeded prior to implantation. With the latter approach apoptosis of implanted cells due to a lack of initial nutrient is likely to occurred, while with the first approach, one may observe less cell death due to prior vascularization [58]. The duration of the pre-vascularization period, i.e. the time until vessels haven sufficiently grown into the center of the scaffold depends on the material and architecture of the scaffold in general and the pore size in particular [1]. Hence 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 [59], [60]. Regarding muscle precursor cells, the pore size of the scaffold should ideally range between 50 µm and 200 µm [61]. These pre-requisites 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 parallel electrospun scaffolds in particular, the size and interconnectivity of the interspaces remains a challenge despite multiple methods described to increase the interspaces as discussed above [1], [43].
In a rat AV-loop in vivo study we could characterize the vascularization of randomly versus parallel spun PCL-collagen blend nanofiber scaffolds in vivo [43]. In this study, an AV-loop was microsurgically created as first described by Erol and Spira in 1980 in the groin of rats and implanted into the matrix inside an isolation chamber [62]. Though technically 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 [43]. Therefore, the AV-loop model is a technique applicable for a potential clinical use in tissue engineering in general. Though the total number of vessels inside the scaffolds was higher within the randomly spun nanofiber scaffolds, the parallel spun group showed a more 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 even after 8 weeks in vivo [43]. Interestingly, the migration of cells through smaller pore sizes has been observed by Zhang et al. with fibroblasts cultivated on randomly spun PCL/collagen scaffolds before [54]. As an explanation, the dynamic structure of nanofiber scaffolds was postulated: due to the flexible and fibrous structure, cells can push the fibers aside and thus migrate through the scaffold. Herein, parallel spun scaffolds are even more flexible since the fibers are not attached to each other. This could possibly explain the more consistent vascularization of the scaffolds [1].
As a further prerequisite for later clinical application, the newly generated skeletal muscle tissue should be innervated by a motor nerve. In order to enable axial vascularization and neurotization at the same time in vivo, we developed and characterized a new small animal AV-loop model, i.e. the EPI-loop model. In this microsurgical rat model, the AV-loop is based on a different vascular axis (the epigastric vessels) as well as motor branch of the obturator nerve. After myoblasts and MSCs coimplantation in a prevascularized isolation chamber, we could observe areas of myogenic differentiation with alpha-sarcomeric actin and MHC expression in the constructs. Quantitative PCR analysis showed an expression of myogenic markers in all specimens [63]. Thus, neurotization and addition of bFGF and dexamethasone allow myogenic differentiation of MSC in an axially vascularized in vivo model for the first time. These findings are a new step towards clinical applicability of skeletal muscle tissue engineering and display its potential for regenerative medicine [63].
Future perspectives of skeletal muscle tissue engineering
In the future skeletal muscle tissue engineering could be further augmented by means of MSC- and/or ADSC cocultivation with muscle precursor cells (myoblasts). Furthermore the differentiation process could be supported by means of smart matrices, i.e. functionalized nanofiber scaffolds releasing myogenic transcription factors or specific components of the ECM/basal lamina of native skeletal muscle tissue. Possibly the architecture of the fibers themselves might also play a pivotal role in the future for myogenic differentiation.
For in vivo application in general and later clinical application in particular, one has to address the problems outlined above in adequate large animal models first, before possible translation into humans. Herefore the development of a large animal (sheep) model for axial vascularization during the last years could be an important step towards later clinical application: after first development of the sheep AV-loop model itself [64], it was further characterized [65] and applied for tissue engineering of vascularized bone [66], [67], including the autologous MSC implantation [68] as well as incorporation of BMP-2 [69], [70] and an extrinsic vascularization pattern [71]. In conclusion results obtained from cell cultures and small animal models could once not only be translated to this microsurgical large animal vascularization model in the field of bone Tissue Engineering, but also for the even more challenging task of skeletal muscle tissue engineering.
References
- 1.
- Klumpp D, Horch RE, Beier JP. Skeletal muscle tissue engineering. In: Boccaccini AR, Ma RX, editors. Tissue engineering using ceramics and polymers. 2nd ed. Amsterdam et al.: Woodhead Publ., Elsevier; 2014. p. 524-40. DOI: 10.1533/9780857097163.3.524
- 2.
- Vandenburgh HH, Karlisch P. Longitudinal growth of skeletal myotubes in vitro in a new horizontal mechanical cell stimulator. In Vitro Cell Dev Biol. 1989 Jul;25(7):607-16. DOI: 10.1007/BF02623630
- 3.
- Dennis RG, Kosnik PE 2nd, Gilbert ME, Faulkner JA. Excitability and contractility of skeletal muscle engineered from primary cultures and cell lines. Am J Physiol, Cell Physiol. 2001 Feb;280(2):C288-95.
- 4.
- Strohman RC, Bayne E, Spector D, Obinata T, Micou-Eastwood J, Maniotis A. Myogenesis and histogenesis of skeletal muscle on flexible membranes in vitro. In Vitro Cell Dev Biol. 1990 Feb;26(2):201-8. DOI: 10.1007/BF02624113
- 5.
- Klumpp D, Horch RE, Kneser U, Beier JP. Engineering skeletal muscle tissue--new perspectives in vitro and in vivo. J Cell Mol Med. 2010 Nov;14(11):2622-9. DOI: 10.1111/j.1582-4934.2010.01183.x
- 6.
- Williams JH, Schray RC, Sirsi SR, Lutz GJ. Nanopolymers improve delivery of exon skipping oligonucleotides and concomitant dystrophin expression in skeletal muscle of mdx mice. BMC Biotechnol. 2008;8:35. DOI: 10.1186/1472-6750-8-35
- 7.
- Nelson SF, Crosbie RH, Miceli MC, Spencer MJ. Emerging genetic therapies to treat Duchenne muscular dystrophy. Curr Opin Neurol. 2009 Oct;22(5):532-8. DOI: 10.1097/WCO.0b013e32832fd487
- 8.
- Terzis JK, Konofaos P. Nerve transfers in facial palsy. Facial Plast Surg. 2008 May;24(2):177-93. DOI: 10.1055/s-2008-1075833
- 9.
- Terzis JK, Noah ME. Analysis of 100 cases of free-muscle transplantation for facial paralysis. Plast Reconstr Surg. 1997 Jun;99(7):1905-21. DOI: 10.1097/00006534-199706000-00016
- 10.
- Klumpp D, Horch RE, Bitto F, Boos AM, Kneser U, Beier JP. Tissue Engineering von Skelettmuskelgewebe – Stand und Perspektiven [Skeletal muscle tissue engineering – current concepts and future perspectives]. Handchir Mikrochir Plast Chir. 2010 Dec;42(6):354-9. DOI: 10.1055/s-0030-1261888
- 11.
- Boontheekul T, Hill EE, Kong HJ, Mooney DJ. Regulating myoblast phenotype through controlled gel stiffness and degradation. Tissue Eng. 2007 Jul;13(7):1431-42. DOI: 10.1089/ten.2006.0356
- 12.
- Snow MH. Myogenic cell formation in regenerating rat skeletal muscle injured by mincing. II. An autoradiographic study. Anat Rec. 1977 Jun;188(2):201-17. DOI: 10.1002/ar.1091880206
- 13.
- Mauro A. Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol. 1961 Feb;9:493-5. DOI: 10.1083/jcb.9.2.493
- 14.
- Seale P, Sabourin LA, Girgis-Gabardo A, Mansouri A, Gruss P, Rudnicki MA. Pax7 is required for the specification of myogenic satellite cells. Cell. 2000;102(6):777-86. DOI: 10.1016/S0092-8674(00)00066-0
- 15.
- Weintraub H, Davis R, Tapscott S, Thayer M, Krause M, Benezra R, Blackwell TK, Turner D, Rupp R, Hollenberg S. The myoD gene family: nodal point during specification of the muscle cell lineage. Science. 1991 Feb;251(4995):761-6. DOI: 10.1126/science.1846704
- 16.
- Beier JP, Kneser U, Stern-Sträter J, Stark GB, Bach AD. Y chromosome detection of three-dimensional tissue-engineered skeletal muscle constructs in a syngeneic rat animal model. Cell Transplant. 2004;13(1):45-53. DOI: 10.3727/000000004772664888
- 17.
- Beier JP, Stern-Straeter J, Foerster VT, Kneser U, Stark GB, Bach AD. Tissue engineering of injectable muscle: three-dimensional myoblast-fibrin injection in the syngeneic rat animal model. Plast Reconstr Surg. 2006 Oct;118(5):1113-21; discussion 1122-4. DOI: 10.1097/01.prs.0000221007.97115.1d
- 18.
- Stern-Straeter J, Bach AD, Stangenberg L, Foerster VT, Horch RE, Stark GB, Beier JP. Impact of electrical stimulation on three-dimensional myoblast cultures – a real-time RT-PCR study. J Cell Mol Med. 2005 Oct-Dec;9(4):883-92. DOI: 10.1111/j.1582-4934.2005.tb00386.x
- 19.
- Otto A, Collins-Hooper H, Patel K. The origin, molecular regulation and therapeutic potential of myogenic stem cell populations. J Anat. 2009 Nov;215(5):477-97. DOI: 10.1111/j.1469-7580.2009.01138.x
- 20.
- Kuang S, Kuroda K, Le Grand F, Rudnicki MA. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell. 2007 Jun;129(5):999-1010. DOI: 10.1016/j.cell.2007.03.044
- 21.
- Le Grand F, Rudnicki MA. Skeletal muscle satellite cells and adult myogenesis. Curr Opin Cell Biol. 2007 Dec;19(6):628-33. DOI: 10.1016/j.ceb.2007.09.012
- 22.
- Yaffe D. Retention of differentiation potentialities during prolonged cultivation of myogenic cells. Proc Natl Acad Sci USA. 1968 Oct;61(2):477-83. DOI: 10.1073/pnas.61.2.477
- 23.
- Boonen KJ, Post MJ. The muscle stem cell niche: regulation of satellite cells during regeneration. Tissue Eng Part B Rev. 2008 Dec;14(4):419-31. DOI: 10.1089/ten.teb.2008.0045
- 24.
- Gilbert PM, Havenstrite KL, Magnusson KE, Sacco A, Leonardi NA, Kraft P, Nguyen NK, Thrun S, Lutolf MP, Blau HM. Substrate elasticity regulates skeletal muscle stem cell self-renewal in culture. Science. 2010 Aug;329(5995):1078-81. DOI: 10.1126/science.1191035
- 25.
- Beier JP, Bitto FF, Lange C, Klumpp D, Arkudas A, Bleiziffer O, Boos AM, Horch RE, Kneser U. Myogenic differentiation of mesenchymal stem cells co-cultured with primary myoblasts. Cell Biol Int. 2011 Apr;35(4):397-406. DOI: 10.1042/CBI20100417
- 26.
- Deans TL, Elisseeff JH. Stem cells in musculoskeletal engineered tissue. Curr Opin Biotechnol. 2009 Oct;20(5):537-44. DOI: 10.1016/j.copbio.2009.10.005
- 27.
- Zhu Y, Liu T, Song K, Fan X, Ma X, Cui Z. Adipose-derived stem cell: a better stem cell than BMSC. Cell Biochem Funct. 2008 Aug;26(6):664-75. DOI: 10.1002/cbf.1488
- 28.
- Brazelton TR, Nystrom M, Blau HM. Significant differences among skeletal muscles in the incorporation of bone marrow-derived cells. Dev Biol. 2003;262(1):64-74. DOI: 10.1016/S0012-1606(03)00357-9
- 29.
- Satija NK, Singh VK, Verma YK, Gupta P, Sharma S, Afrin F, Sharma M, Sharma P, Tripathi RP, Gurudutta GU. Mesenchymal stem cell-based therapy: a new paradigm in regenerative medicine. J Cell Mol Med. 2009 Nov-Dec;13(11-12):4385-402. DOI: 10.1111/j.1582-4934.2009.00857.x
- 30.
- Meirelles Lda S, Nardi NB. Methodology, biology and clinical applications of mesenchymal stem cells. Front Biosci (Landmark Ed). 2009 Jan 1;14:4281-98.
- 31.
- Estrada R, Li N, Sarojini H, An J, Lee MJ, Wang E. Secretome from mesenchymal stem cells induces angiogenesis via Cyr61. J Cell Physiol. 2009 Jun;219(3):563-71. DOI: 10.1002/jcp.21701
- 32.
- García-Castro J, Trigueros C, Madrenas J, Pérez-Simón JA, Rodriguez R, Menendez P. Mesenchymal stem cells and their use as cell replacement therapy and disease modelling tool. J Cell Mol Med. 2008 Dec;12(6B):2552-65. DOI: 10.1111/j.1582-4934.2008.00516.x
- 33.
- Rossignol J, Boyer C, Thinard R, Remy S, Dugast AS, Dubayle D, Dey ND, Boeffard F, Delecrin J, Heymann D, Vanhove B, Anegon I, Naveilhan P, Dunbar GL, Lescaudron L. Mesenchymal stem cells induce a weak immune response in the rat striatum after allo or xenotransplantation. J Cell Mol Med. 2009 Aug;13(8B):2547-58. DOI: 10.1111/j.1582-4934.2009.00657.x
- 34.
- Madden L, Juhas M, Kraus WE, Truskey GA, Bursac N. Bioengineered human myobundles mimic clinical responses of skeletal muscle to drugs. Elife. 2015;4:e04885. DOI: 10.7554/eLife.04885
- 35.
- Vukicevic S, Kleinman HK, Luyten FP, Roberts AB, Roche NS, Reddi AH. Identification of multiple active growth factors in basement membrane Matrigel suggests caution in interpretation of cellular activity related to extracellular matrix components. Exp Cell Res. 1992;202(1):1-8. DOI: 10.1016/0014-4827(92)90397-Q
- 36.
- Beier JP, Klumpp D, Rudisile M, Dersch R, Wendorff JH, Bleiziffer O, Arkudas A, Polykandriotis E, Horch RE, Kneser U. Collagen matrices from sponge to nano: new perspectives for tissue engineering of skeletal muscle. BMC Biotechnol. 2009;9:34. DOI: 10.1186/1472-6750-9-34
- 37.
- Peng YY, Glattauer V, Ramshaw JA, Werkmeister JA. Evaluation of the immunogenicity and cell compatibility of avian collagen for biomedical applications. J Biomed Mater Res A. 2010 Jun;93(4):1235-44. DOI: 10.1002/jbm.a.32616
- 38.
- Arkudas A, Pryymachuk G, Hoereth T, Beier JP, Polykandriotis E, Bleiziffer O, Horch RE, Kneser U. Dose-finding study of fibrin gel-immobilized vascular endothelial growth factor 165 and basic fibroblast growth factor in the arteriovenous loop rat model. Tissue Eng Part A. 2009 Sep;15(9):2501-11. DOI: 10.1089/ten.tea.2008.0477
- 39.
- Gunatillake PA, Adhikari R. Biodegradable synthetic polymers for tissue engineering. Eur Cell Mater. 2003 May 20;5:1-16.
- 40.
- Bölgen N, Menceloğlu YZ, Acatay K, Vargel I, Pişkin E. In vitro and in vivo degradation of non-woven materials made of poly(epsilon-caprolactone) nanofibers prepared by electrospinning under different conditions. J Biomater Sci Polym Ed. 2005;16(12):1537-55. DOI: 10.1163/156856205774576655
- 41.
- Ignatius AA, Claes LE. In vitro biocompatibility of bioresorbable polymers: poly(L, DL-lactide) and poly(L-lactide-co-glycolide). Biomaterials. 1996;17(8):831-9. DOI: 10.1016/0142-9612(96)81421-9
- 42.
- Schnell E, Klinkhammer K, Balzer S, Brook G, Klee D, Dalton P, Mey J. Guidance of glial cell migration and axonal growth on electrospun nanofibers of poly-epsilon-caprolactone and a collagen/poly-epsilon-caprolactone blend. Biomaterials. 2007 Jul;28(19):3012-25. DOI: 10.1016/j.biomaterials.2007.03.009
- 43.
- Klumpp D, Rudisile M, Kühnle RI, Hess A, Bitto FF, Arkudas A, Bleiziffer O, Boos AM, Kneser U, Horch RE, Beier JP. Three-dimensional vascularization of electrospun PCL/collagen-blend nanofibrous scaffolds in vivo. J Biomed Mater Res A. 2012 Sep;100(9):2302-11. DOI: 10.1002/jbm.a.34172
- 44.
- Kim MS, Jun I, Shin YM, Jang W, Kim SI, Shin H. The development of genipin-crosslinked poly(caprolactone) (PCL)/gelatin nanofibers for tissue engineering applications. Macromol Biosci. 2010 Jan;10(1):91-100. DOI: 10.1002/mabi.200900168
- 45.
- Curtis A, Wilkinson C. Topographical control of cells. Biomaterials. 1997;18(24):1573-83. DOI: 10.1016/S0142-9612(97)00144-0
- 46.
- Gingras J, Rioux RM, Cuvelier D, Geisse NA, Lichtman JW, Whitesides GM, Mahadevan L, Sanes JR. Controlling the orientation and synaptic differentiation of myotubes with micropatterned substrates. Biophys J. 2009 Nov;97(10):2771-9. DOI: 10.1016/j.bpj.2009.08.038
- 47.
- Madaghiele M, Sannino A, Yannas IV, Spector M. Collagen-based matrices with axially oriented pores. J Biomed Mater Res A. 2008 Jun;85(3):757-67. DOI: 10.1002/jbm.a.31517
- 48.
- Schoof H, Apel J, Heschel I, Rau G. Control of pore structure and size in freeze-dried collagen sponges. J Biomed Mater Res. 2001;58(4):352-7. DOI: 10.1002/jbm.1028
- 49.
- Ayres C, Bowlin GL, Henderson SC, Taylor L, Shultz J, Alexander J, Telemeco TA, Simpson DG. Modulation of anisotropy in electrospun tissue-engineering scaffolds: Analysis of fiber alignment by the fast Fourier transform. Biomaterials. 2006 Nov;27(32):5524-34. DOI: 10.1016/j.biomaterials.2006.06.014
- 50.
- Boudriot U, Dersch R, Greiner A, Wendorff JH. Electrospinning approaches toward scaffold engineering – a brief overview. Artif Organs. 2006 Oct;30(10):785-92. DOI: 10.1111/j.1525-1594.2006.00301.x
- 51.
- Sell SA, McClure MJ, Garg K, Wolfe PS, Bowlin GL. Electrospinning of collagen/biopolymers for regenerative medicine and cardiovascular tissue engineering. Adv Drug Deliv Rev. 2009 Oct;61(12):1007-19. DOI: 10.1016/j.addr.2009.07.012
- 52.
- Riboldi SA, Sampaolesi M, Neuenschwander P, Cossu G, Mantero S. Electrospun degradable polyesterurethane membranes: potential scaffolds for skeletal muscle tissue engineering. Biomaterials. 2005 Aug;26(22):4606-15. DOI: 10.1016/j.biomaterials.2004.11.035
- 53.
- Martins A, Duarte AR, Faria S, Marques AP, Reis RL, Neves NM. Osteogenic induction of hBMSCs by electrospun scaffolds with dexamethasone release functionality. Biomaterials. 2010 Aug;31(22):5875-85. DOI: 10.1016/j.biomaterials.2010.04.010
- 54.
- Zhang YZ, Venugopal J, Huang ZM, Lim CT, Ramakrishna S. Characterization of the surface biocompatibility of the electrospun PCL-collagen nanofibers using fibroblasts. Biomacromolecules. 2005 Sep-Oct;6(5):2583-9. DOI: 10.1021/bm050314k
- 55.
- Telemeco TA, Ayres C, Bowlin GL, Wnek GE, Boland ED, Cohen N, Baumgarten CM, Mathews J, Simpson DG. Regulation of cellular infiltration into tissue engineering scaffolds composed of submicron diameter fibrils produced by electrospinning. Acta Biomater. 2005 Jul;1(4):377-85. DOI: 10.1016/j.actbio.2005.04.006
- 56.
- Baker BM, Gee AO, Metter RB, Nathan AS, Marklein RA, Burdick JA, Mauck RL. The potential to improve cell infiltration in composite fiber-aligned electrospun scaffolds by the selective removal of sacrificial fibers. Biomaterials. 2008 May;29(15):2348-58. DOI: 10.1016/j.biomaterials.2008.01.032
- 57.
- Kannan RY, Salacinski HJ, Sales K, Butler P, Seifalian AM. The roles of tissue engineering and vascularisation in the development of micro-vascular networks: a review. Biomaterials. 2005 May;26(14):1857-75. DOI: 10.1016/j.biomaterials.2004.07.006
- 58.
- Arkudas A, Beier JP, Heidner K, Tjiawi J, Polykandriotis E, Srour S, Sturzl M, Horch RE, Kneser U. Axial prevascularization of porous matrices using an arteriovenous loop promotes survival and differentiation of transplanted autologous osteoblasts. Tissue Eng. 2007 Jul;13(7):1549-60. DOI: 10.1089/ten.2006.0387
- 59.
- Freed LE, Marquis JC, Langer R, Vunjak-Novakovic G. Kinetics of chondrocyte growth in cell-polymer implants. Biotechnol Bioeng. 1994 Mar;43(7):597-604. DOI: 10.1002/bit.260430709
- 60.
- Ishaug-Riley SL, Crane-Kruger GM, Yaszemski MJ, Mikos AG. Three-dimensional culture of rat calvarial osteoblasts in porous biodegradable polymers. Biomaterials. 1998;19(15):1405-12. DOI: 10.1016/S0142-9612(98)00021-0
- 61.
- Lee M, Wu BM, Dunn JC. Effect of scaffold architecture and pore size on smooth muscle cell growth. J Biomed Mater Res A. 2008 Dec;87(4):1010-6. DOI: 10.1002/jbm.a.31816
- 62.
- Erol OO, Spira M. New capillary bed formation with a surgically constructed arteriovenous fistula. Surg Forum. 1979;30:530-1.
- 63.
- Bitto FF, Klumpp D, Lange C, Boos AM, Arkudas A, Bleiziffer O, Horch RE, Kneser U, Beier JP. Myogenic differentiation of mesenchymal stem cells in a newly developed neurotised AV-loop model. Biomed Res Int. 2013;2013:935046. DOI: 10.1155/2013/935046
- 64.
- Beier JP, Horch RE, Arkudas A, Polykandriotis E, Bleiziffer O, Adamek E, Hess A, Kneser U. De novo generation of axially vascularized tissue in a large animal model. Microsurgery. 2009;29(1):42-51. DOI: 10.1002/micr.20564
- 65.
- Weigand A, Boos AM, Ringwald J, Mieth M, Kneser U, Arkudas A, Bleiziffer O, Klumpp D, Horch RE, Beier JP. New aspects on efficient anticoagulation and antiplatelet strategies in sheep. BMC Vet Res. 2013;9:192. DOI: 10.1186/1746-6148-9-192
- 66.
- Beier JP, Hess A, Loew J, Heinrich J, Boos AM, Arkudas A, Polykandriotis E, Bleiziffer O, Horch RE, Kneser U. De novo generation of an axially vascularized processed bovine cancellous-bone substitute in the sheep arteriovenous-loop model. Eur Surg Res. 2011;46(3):148-55. DOI: 10.1159/000324408
- 67.
- Beier JP, Horch RE, Hess A, Arkudas A, Heinrich J, Loew J, Gulle H, Polykandriotis E, Bleiziffer O, Kneser U. Axial vascularization of a large volume calcium phosphte ceramic bone substitute in the sheep AV loop model. J Tissue Eng Regen Med. 2010 Mar;4(3):216-23. DOI: 10.1002/term.229
- 68.
- Boos AM, Loew JS, Deschler G, Arkudas A, Bleiziffer O, Gulle H, Dragu A, Kneser U, Horch RE, Beier JP. Directly auto-transplanted mesenchymal stem cells induce bone formation in a ceramic bone substitute in an ectopic sheep model. J Cell Mol Med. 2011 Jun;15(6):1364-78. DOI: 10.1111/j.1582-4934.2010.01131.x
- 69.
- Boos AM, Loew JS, Weigand A, Deschler G, Klumpp D, Arkudas A, Bleiziffer O, Gulle H, Kneser U, Horch RE, Beier JP. Engineering axially vascularized bone in the sheep arteriovenous-loop model. J Tissue Eng Regen Med. 2013 Aug;7(8):654-64. DOI: 10.1002/term.1457
- 70.
- Boos AM, Weigand A, Deschler G, Gerber T, Arkudas A, Kneser U, Horch RE, Beier JP. Autologous serum improves bone formation in a primary stable silica-embedded nanohydroxyapatite bone substitute in combination with mesenchymal stem cells and rhBMP-2 in the sheep model. Int J Nanomedicine. 2014;9:5317-39. DOI: 10.2147/IJN.S66867
- 71.
- Weigand A, Beier JP, Hess A, Gerber T, Arkudas A, Horch RE, Boos AM. Acceleration of vascularized bone tissue-engineered constructs in a large animal model combining intrinsic and extrinsic vascularization. Tissue Eng Part A. 2015 May;21(9-10):1680-94. DOI: 10.1089/ten.TEA.2014.0568