| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Pfizer Lecture in Human Physiology |
1 Division of Molecular Physiology, University of Dundee, MSI/WTB Dow Street, Dundee DD1 5EH, UK
Abstract
Tissue engineering is a discipline of great promise. In some areas, such as the cornea, tissues engineered in the laboratory are already in clinical use. In other areas, where the tissue architecture is more complex, there are a number of obstacles to manoeuvre before clinically relevant tissues can be produced. However, even in areas where clinically relevant tissues are decades away, the tissues being produced at the moment provide powerful new models to aid the understanding of complex physiological processes. This article provides a personal view of the role of tissue engineering in advancing our understanding of physiology, with specific attention being paid to musculoskeletal tissues.
(Received 19 June 2005;
accepted after revision 3 August 2005; first published online 9 August 2005)
Corresponding author K. Baar: Division of Molecular Physiology, University of Dundee, MSI/WTB Dow Street, Dundee DD1 5EH, UK. Email: k.baar{at}dundee.ac.uk
The goal of tissue engineering is to use scaffolding materials and multipotent stem cells to produce tissue replacements (Vunjak-Novakovic et al. 2004). While great progress has been made towards this goal, a number of obstacles remain, including: (1) vascularization of the engineered tissues; (2) maturation of the tissues, since most are developmentally arrested; (3) transdifferentiation of the donor stem cells to simultaneously produce all of the cells required to form a complete tissue; and (4) creation of effective interfaces between biological and artificial materials.
While these limitations currently prevent large-scale growth of tissues for use in regenerative medicine, the tissues that are currently being made are potentially powerful tools for studying tissue physiology in a controlled environment.
Need for multiple models to enable understanding of tissue physiology
Our understanding of human physiology has benefited greatly from a reductionist approach. Use of model organisms, such as non-human primates, pigs, rats, mice and flies, has given physiologists a degree of control that would never be possible in human experiments. Further reduction, studying isolated cells in culture, has given even more control by allowing us to look at the response of an individual cell type to a physiological intervention. As with all models, none of these is a true representation of what occurs in vivo within the intact human body, but they provide a great deal of initial information about the basic mechanisms underlying human physiology.
The goal of any model is to have distinct experimental advantages while remaining as close as possible to the physiological reality. This is especially important in musculoskeletal tissues. Tissues such as muscle, tendon, ligament and bone are distinct from many other tissues in the body in that their function is largely mechanical. Bone provides the rigid scaffold that gives our bodies shape and provides the levers that allow movement. Tendons and ligaments are mechanical interfaces between tissues, functioning to absorb and return force. Muscles, whether smooth, cardiac or skeletal, produce the forces required to pass foodstuffs, circulate the blood and allow us to breathe and move. Surrogate measures of function have been created for these tissues so that in vitro experiments can be linked to the physiological function. For bone, mitrogen activated protein (MAP) kinase activation, prostaglandin production or nitric oxide synthesis symbolize the adaptation to loading (Smalt et al. 1997). In tendon and ligament, collagen synthesis is correlated to functional changes (Banes et al. 1999). In skeletal muscle, protein-to-DNA ratio is used to note hypertrophy or atrophy, and this in turn is correlated to force (Rommel et al. 2001). In cardiac muscle, the expression of neonatal genes and elevation of protein synthesis are markers for growth (Boluyt et al. 1997). While these markers have their roles, what we as physiologists want to know is whether the bone is stronger, whether the tendon is stiffer or whether the muscle produces more force. To answer these questions, traditional cell culture models are insufficient.
One new model that will allow physiologists to make these determinations in vitro is three-dimensional (3D) tissue engineering. Three-dimensional tissue engineering involves isolating or transdifferentiating the desired cell type and then either providing a 3D scaffold (Radisic et al. 2004) or promoting the reorganization of a monolayer of cells into a self-organized 3D engineered tissue (Baar et al. 2005). Following the formation of the 3D construct, a variety of physiological parameters can be determined. For muscle, force production, time to peak tension, half-relaxation time and excitability (chronaxie and rheobase) can be measured by electrically stimulating the constructs while attached to a force transducer (Dennis & Kosnik, 2000; Dennis et al. 2001; Kosnik et al. 2001; Dennis & Kosnik, 2002) For tendon, tangent stiffness, ultimate tensile strain and stored energy can be measured during a stress–strain test (Garvin et al. 2003; Calve et al. 2004). For bone, tests for compression stiffness, three- or four-point bending and tensile strength are possible (Thomson et al. 1995). Using these tests it is possible to determine the effect of altered genetics or chemical or mechanical stimuli on the physiological function of the tissue. The other distinct advantage of 3D engineered tissues is that the cells can be allowed to proliferate for a short period without losing the unique mechanical properties of the cell (Hecker et al. 2005). This means that tissue from a single biopsy can be expanded to produce 30–60 genetically identical 3D engineered tissues (Huang et al. 2005). This is especially important in human experiments, where the starting material is extremely valuble. To illustrate the utility of these tools, the early results of studies from our laboratory as well as others will be discussed below.
Engineering adult skeletal muscle: role of electrical stimulation
Chronic low-frequency electrical stimulation (CLFS) was originally used to test whether motor neurone-specific impulse patterns could affect the contractile speed of skeletal muscles (Salmons & Vrbova, 1967). A tonic stimulus that mimicked the impulse pattern of a slow motor neurone was found to slow the speed of both contraction and relaxation in fast-twitch muscles. The importance of electrical activity on the phenotype of skeletal muscle was reinforced when Salmons and Sréter showed that the slow-to-fast muscle transition following cross-reinnervation of the soleus with the peroneal nerve could be reversed by CLFS (Salmons & Sreter, 1976). This experiment showed conclusively that the role of the nerve in determining muscle phenotype was largely dependent on electrical, not chemical input. Since that time, the CLFS-induced fast-to-slow transition has been shown to be the result of a change in the isoforms of the contractile proteins (myosin heavy chain, Brown et al. 1983), regulatory proteins (myosin light chain, Sreter et al. 1973; tropomyosin, Roy et al. 1979; and troponin, Hartner & Pette, 1990) and calcium-sequestering proteins (parvalbumin, Green et al. 1984; and sarco/endoplasmic reticulum Ca-ATPase (SERCA), Ohlendieck et al. 1991). The shift in protein isoforms, as well as the concomitant mitochondrial biogenesis (Takahashi & Hood, 1993), results in a fatigue-resistant muscle with an improved duty cycle.
While these in vivo experiments have given us great insight into the phenotype resulting from electrical stimulation, they are limited as to determining the molecular mechanisms underlying this change. The most promising molecular candidate underlying the fast-to-slow shift in muscle phenotype is the calcium-dependent phosphatase, calcineurin. Overexpression of calcineurin induces a fast-to-slow fibre transformation (Chin et al. 1998), and electrical stimulation is thought to activate calcineurin (Dunn et al. 2001; Kubis et al. 2002). However, in vivo it is difficult to determine whether the effects of electrical stimulation require calcineurin activation or whether electrical stimulation and calcineurin activate distinct molecular pathways that have a similar effect on muscle phenotype. To address this question directly, we have recently undertaken experiments using 3D engineered muscles treated with 2 weeks of CLFS, cyclosporin A (CsA, to block calcineurin), or both interventions together, in order to determine the effects on force production and contraction and relaxation time. To perform these experiments we formed 3D engineered muscles from primary rat soleus muscle cells and a fibrin gel scaffold (Huang et al. 2005). These constructs were either electrically stimulated with 5–20 Hz pulses every 4 s, treated with 500 ng ml–1 CsA, or given both interventions. Cyclosporin A treatment decreased the time-to-peak tension (TPT) without significantly affecting half-relaxation time (1/2RT, Fig. 1, Y. C. Huang, R. G. Dennis & K. Baar, unpublished observations). Slow electrical stimulation did not affect either the TPT or the 1/2RT of the constructs. Interestingly, both CLFS and CsA increased force production in the constructs. When the two stimuli were added together the TPT remained faster and the force produced by the constructs was 3.4-fold higher than in controls (Fig. 1). These results suggest three important things about the mechanism of CLFS/calcineurin induced changes in muscle phenotype. First, cyclosporin A treatment increases the rate of skeletal muscle contraction, probably through modulation of myosin heavy chain expression. Second, cyclosporin A treatment alone has no effect on the rate of skeletal muscle relaxation, suggesting that a permissive level of another factor is required for the shift in calcium sequestration. Lastly, inhibition of calcineurin and electrical stimulation has independent and additive effects on muscle force production, suggesting that the two stimuli use distinct anabolic mechanisms. These observations are consistent with observations by Zádor and colleagues that neither innervation nor calcineurin directly controls the expression of SERCA2a (Zador & Wuytack, 2003; Zador et al. 2005).
|
Mechanical loading is one of the most potent stimulators of musculoskeletal growth and adaptation (Benjamin & Hillen, 2003). A number of in vitro models have been developed to test the effect of stretch on a variety of cell types in two dimensions. Most of these techniques have stretched a monolayer of cells using flexible-bottomed culture dishes that were placed under static load (Vandenburgh & Kaufman, 1979) or dynamically loaded using either a screw-driven piston (Vandenburgh et al. 1995) or a vacuum-based loading device (Banes et al. 1985). These experiments have provided valuable information; however, as discussed above, the functional changes that result are much less well understood. More recently, stretch has been used on 3D engineered musculoskeletal tissues and the functional consequences of the stretch described.
Eschenhagen and his group (Eschenhagen et al. 1997) have used mechanical loading to improve the function of their 3D engineered heart tissues (EHT). These tissues are created by a gelation process in which neonatal rat hearts are dissociated, the cardiomyocytes are mixed with a solution containing collagen I and matrigel, and then pipetted into molds of various sizes and shapes (Eschenhagen et al. 1997; Fink et al. 2000; Zimmermann et al. 2000, 2002a,b). The original description of the effect of stretch on these tissues used rectangular molds containing a piece of silicone tubing covered with Velcro® at each end to promote adhesion with the collagen gel. After 4 days in culture, the constructs were placed in a bioreactor, a device that controls the chemical, mechanical and electrical environment of the constructs, and a unidirectional stretch of between 1 and 20% was applied at a frequency of 2 Hz. The mechanical stretch induced a 40% increase in cardiomyocyte cross-sectional area and a two- to fourfold increase in force production (Fig. 2). This finding is extremely interesting since it suggests that the cardiomyocyte growth observed was compensatory and not decompensatory hypertrophy. The initial phase of heart muscle hypertrophy, in response to increased load, results in an increase force production to maintain wall stress (Frey & Olson, 2003). Prolonged strain results in a further increase in the size of the cardiomyocytes without a concomitant increase in force production. This switch, from compensatory to decompensatory growth, is a hallmark of disease progression (Czubryt & Olson, 2004). Since the best determinant of this shift is decreased coupling of cell size and force production it has been difficult to model in vitro. However, stretch of 3D engineered cardiac tissue provides a culture model that may allow identification of the molecular mechanism underlying the shift towards decompensatory growth and heart failure.
|
Engineered skeletal muscle with improved fatigue resistance (overexpression studies)
One of the strengths of using engineered tissues lies in the ability to transfect the composite cells and rapidly determine the functional importance of overexpressing or knocking down specific proteins within the cells. We have used this technique in our engineered muscles to determine the effects of a coordinator of mitochondrial biogenesis, PPAR
coactivator (PGC-1
), on muscle function. Myoblasts were transfected with the PGC-1 cDNA, driven by the myosin light chain 2 slow promoter to achieve muscle-specific expression of the cDNA. Constructs were then engineered from the transfected cells and fatigue resistance was determined using a 5 min contraction protocol consisting of a single 1 s tetanus every 5 s for 5 min. Overexpression of PGC-1 resulted in a 2.75-fold increase in force at the end of the 5 min contraction period (Fig. 3). This finding mimics quite closely what is seen in mice overexpressing PGC-1 in muscle (Lin et al. 2002). Muscles from these mice showed an 
2.7-fold increase in their fatigue index compared to littermate controls. The close approximation of the effect of transfection in engineered muscle with the physiological effects of overexpression in transgenic animals provides validation for using engineered muscle as a functional model of in vivo physiology. Furthermore, when knocking out a gene of interest produces embryonic lethality in mice, knockdown experiments in engineered tissues provide a possible alternative to tissue-specific knockouts for determining the functional effects of a specific protein.
|
Promoting increased size of engineered tissues (perfusion/vascularization).
The primary limitation to the development of large well-organized 3D engineered tissues is the delivery of nutrients and oxygen to the cells at the core of the construct. In all cases for non-perfused engineered tissues, there is a depth limit from the surface of the construct beyond which the cells will fail to survive. We term this the ‘viable radius’. When the radius of the tissue construct exceeds the viable radius for that tissue, necrosis is readily visible in the core. The viable radius of tissue constructs varies with the construction and the metabolic rate of the cells. In self-organizing cardiac tissues the viable radius appears to be in the range of 40–50 µm (Baar et al. 2005), whereas in skeletal muscle and tendon constructs it is 150–200 µm (Dennis & Kosnik, 2000; Calve et al. 2004). The gelation technique has overcome this limitation to some degree by promoting the formation of strands of cells within the scaffolding material (Zimmermann et al. 2002b). However, this has the undesirable side-effect of decreasing the function of the construct (i.e. the specific force produced). The Vunjak-Novakovic (Carrier et al. 2002a,b) and Vandenburgh laboratories (Chromiak et al. 1998) have reported temporary solutions. Both groups have successfully grown bigger constructs using a low shear stress mass perfusion apparatus (Carrier et al. 2002a,b). By mass perfusing a dense scaffolding material containing neonatal cardiac muscle cells with a flow rate from 0.6 to 3 ml min–1, Carrier et al. (2002a) were able to increase the number of cells deep within the matrix. They report that a flow rate of 0.6 ml min–1 produces a uniform cell density up to 1 mm deep.
While experiments using perfusion can slightly increase some metabolic parameters and construct size, new methods will be required to provide the level of circulating nutrients and oxygen required for the formation of larger tissues. The most frequent method of clinical neovascularization uses an extrinsic blood supply (Cassell et al. 2002). Here, an avascular structure is implanted into an animal or human near a rich vascular bed, such as a subcutaneous region, in the mesentery, or in the omentum. Vascularization of the implanted tissue occurs as a result of the inflammatory wound healing process in response to the surgery. Implantation of 3D muscle tissues is known to result in a high degree of vascularization (Okano & Matsuda, 1998; Li et al. 1999; van Wachem et al. 1999; Leor et al. 2000; Sakai et al. 2001; Eschenhagen et al. 2002; Zimmermann et al. 2002a); however, it is unclear whether the new vascular tissue allows the constructs to grow larger upon the return to cell culture or aids in the development of a more mature phenotype.
Another possible technique that can be completed in vitro involves co-culturing the engineered tissues with embryoid bodies (Wartenberg et al. 2001). Embryoid bodies are grown from pluripotent embryonic stem cells that aggregate to form 200–400 µm balls of cells. These cells can differentiate into cells of any of the three germ layers. When embryoid bodies were cultured in contact with tumour spheroids where central necrosis was apparent, within 5 days a branched network of capillary-like structures sprouted (Wartenberg et al. 2001). This neovascularization resulted in a decrease in central necrosis and a 557% increase in the size of the tumour spheroid.
Engineering physiologically relevant tissue interfaces. The most extreme difference between engineered tissues and in vivo tissues is at the interfaces (Fig. 4). In vivo, the transition from muscle to bone is seamless. The regional variability of tendon allows it to transfer mechanical force and power from the compliant muscle to the stiff bone with high fidelity (E. M. Arruda, S. Calve, R. G. Dennis, K. Mundy & K. Baar, unpublished observations). This natural transition prevents impedance mismatching and therefore decreases the likelihood of injury at the transition. While this aspect of physiology is essential to protecting muscle and allowing the transfer of force and power, it is very poorly understood. In fact, while 13 genes are known to be involved in determining muscle attachment in Drosophila melanogaster, no genes are known to play this role in mammals. Tendons initially develop independently from muscle in the limb bud mesoderm (Edom-Vovard & Duprez, 2004). The tendon cells only contact the somatically derived muscle cells later in development. Once linked, the presence of gradients of fibroblast growth factors, such as FGF-4 and FGF-8 (Edom-Vovard et al. 2001, 2002), and growth and differentiation factors (Wolfman et al. 1997; Aspenberg & Forslund, 1999; Lou et al. 2001; Rickert et al. 2001) are believed to give cues that are important for the formation of the tissue interface. This type of tissue development is well suited for tissue engineering. Gradients of growth factors can be immobilized on the semisolid scaffolds that form the muscle anchors (DeLong et al. 2005). Preloading the scaffolds with fibroblasts and then placing them in a muscle cell culture would recapitulate much of the developmental biology involved in tendon formation. However, whether this technique is sufficient to produce a graded non-elastic tissue like tendon has yet to be determined.
|
A number of physiological issues must be addressed in order to ensure the future development of 3D engineered musculoskeletal tissues. (1) The developmental state of the cells within the 3D constructs needs to be determined. The force of contraction, time to peak tension and half-relaxation time of muscle are determined by the isoform of myosin, calcium-handling machinery and regulatory proteins that are expressed. There are no reports in the literature on 3D skeletal muscles, and very little information in the literature on cardiac tissues, that focuses on the expression pattern of these proteins in engineered tissue. (2) Mechanisms for shifting the phenotype of the 3D engineered tissues to that of the mature tissue need to be developed. This would include: in muscle, the expression of adult contractile proteins (mature myosin heavy chain (MHC)), the proper regulatory proteins (in heart, cardiac-specific tropomyosin and troponins) and mature calcium-handling machinery (in heart, SERCA2a and phospholamban), and the desired functional architecture, such as pennation angle and fibre length; in tendon and ligament, the type of collagen and proteoglycans expressed and the regional mechanical variability of the tissue. (3) Novel techniques need to be developed to make it possible to engineer suitable tissue-to-tissue interfaces, such as the myotendonous junction. This is essential in order to provide mechanical impedance matching and to create the complex structures that are involved in the transduction of mechanical signals.
If we can answer these difficult questions, not only will we have developed functionally important engineered tissues, but we will also have greatly advanced our understanding of physiology.
Footnotes
The Pfizer Lecture in Human Physiology was given at the Physiological Society Meeting at Kings College London 18th December 2004.
References
Aspenberg P & Forslund C (1999). Enhanced tendon healing with GDF 5 and 6. Acta Orthop Scand 70, 51–54.[Medline]
Baar
K, Birla
R, Boluyt
MO, Borschel
GH, Arruda
EM
&
Dennis
RG (2005). Self-organization of rat cardiac cells into contractile 3-D cardiac tissue. Faseb J
19, 275–277.
Banes AJ, Gilbert J, Taylor D & Monbureau O (1985). A new vacuum-operated stress-providing instrument that applies static or variable duration cyclic tension or compression to cells in vitro. J Cell Sci 75, 35–42.[Abstract]
Banes AJ, Weinhold P, Yang X, Tsuzaki M, Bynum D, Bottlang M & Brown T (1999). Gap junctions regulate responses of tendon cells ex vivo to mechanical loading. Clin OrthopRelat Res 367, S356–S370.
Benjamin M & Hillen B (2003). Mechanical influences on cells, tissues and organs – ‘Mechanical Morphogenesis’. Eur J Morph 41, 3–7.
Boluyt
MO, Zheng
JS, Younes
A, Long
X, O'Neill
L, Silverman
H, Lakatta
EG
&
Crow
MT (1997). Rapamycin inhibits 1-adrenergic receptor-stimulated cardiac myocyte hypertrophy but not activation of hypertrophy-associated genes. Evidence for involvement of p70, S6 kinase. Circ Res
81, 176–186.
Brown
WE, Salmons
S
&
Whalen
RG (1983). The sequential replacement of myosin subunit isoforms during muscle type transformation induced by long term electrical stimulation. J Biol Chem
258, 14686–14692.
Calve S, Dennis RG, Kosnik PE II, Baar K, Grosh K & Arruda EM (2004). Engineering of functional tendon. Tissue Eng 10, 755–761.[CrossRef][Medline]
Carrier RL, Rupnick M, Langer R, Schoen FJ, Freed LE & Vunjak-Novakovic G (2002a). Effects of oxygen on engineered cardiac muscle. Biotechnol Bioeng 78, 617–625.[CrossRef][Medline]
Carrier RL, Rupnick M, Langer R, Schoen FJ, Freed LE & Vunjak-Novakovic G (2002b). Perfusion improves tissue architecture of engineered cardiac muscle. Tissue Eng 8, 175–188.[CrossRef][Medline]
Cassell OC, Hofer SO, Morrison WA & Knight KR (2002). Vascularisation of tissue-engineered grafts: the regulation of angiogenesis in reconstructive surgery and in disease states. Br J Plast Surg 55, 603–610.[CrossRef][Medline]
Chin
ER, Olson
EN, Richardson
JA, Yang
Q, Humphries
C, Shelton
JM, Wu
H, Zhu
W, Bassel-Duby
R
&
Williams
RS (1998). A calcineurin-dependent transcriptional pathway controls skeletal muscle fiber type. Genes Dev
12, 2499–2509.
Chromiak JA, Shansky J, Perrone C & Vandenburgh HH (1998). Bioreactor perfusion system for the long-term maintenance of tissue-engineered skeletal muscle organoids. In Vitro Cell Dev Biol Anim 34, 694–703.[Medline]
Czubryt
MP
&
Olson
EN (2004). Balancing contractility and energy production: the role of myocyte enhancer factor 2 (MEF2) in cardiac hypertrophy. Recent Prog Horm Res
59, 105–124.
DeLong SA, Moon JJ & West JL (2005). Covalently immobilized gradients of bFGF on hydrogel scaffolds for directed cell migration. Biomaterials 26, 3227–3234.[CrossRef][Medline]
Dennis RG & Kosnik PE (2000). Excitability and isometric contractile properties of mammalian skeletal muscle constructs engineered in vitro. In Vitro Cell Dev Biol Anim 36, 327–335.[Medline]
Dennis RG & Kosnik PE (2002). Mesenchymal cell culture: instrumentation and methods for evaluating engineered muscle. In Methods in Tissue Engineering, ed. Atala A & Lanza R, pp. 307–316.
Dennis RG, Kosnik PE II, Gilbert ME & Faulkner JA (2001). Excitability and contractility of skeletal muscle engineered from primary cultures and cell lines. Am J Physiol 280, C288–C295.
Dunn
SE, Simard
AR, Bassel-Duby
R, Williams
RS
&
Michel
RN (2001). Nerve activity-dependent modulation of calcineurin signaling in adult fast and slow skeletal muscle fibers. J Biol Chem
276, 45243–45254.
Edom-Vovard F, Bonnin M & Duprez D (2001). Fgf8 transcripts are located in tendons during embryonic chick limb development. Mech Dev 108, 203–206.[CrossRef][Medline]
Edom-Vovard F & Duprez D (2004). Signals regulating tendon formation during chick embryonic development. Dev Dyn 229, 449–457.[CrossRef][Medline]
Edom-Vovard F, Schuler B, Bonnin MA, Teillet MA & Duprez D (2002). Fgf4 positively regulates scleraxis and tenascin expression in chick limb tendons. Dev Biol 247, 351–366.[CrossRef][Medline]
Eschenhagen T, Didie M, Munzel F, Schubert P, Schneiderbanger K & Zimmermann WH (2002). 3D engineered heart tissue for replacement therapy. Basic Res Cardiol 97 (Suppl. 1), I146–I152.
Eschenhagen T, Fink C, Remmers U, Scholz H, Wattchow J, Weil J, Zimmermann W, Dohmen HH, Schafer H, Bishopric N, Wakatsuki T & Elson EL (1997). Three-dimensional reconstitution of embryonic cardiomyocytes in a collagen matrix: a new heart muscle model system. Faseb J 11, 683–694.[Abstract]
Fink
C, Ergun
S, Kralisch
D, Remmers
U, Weil
J
&
Eschenhagen
T (2000). Chronic stretch of engineered heart tissue induces hypertrophy and functional improvement. Faseb J
14, 669–679.
Frey N & Olson EN (2003). Cardiac hypertrophy: the good, the bad, and the ugly. Annu Rev Physiol 65, 45–79.[CrossRef][Medline]
Garvin J, Qi J, Maloney M & Banes AJ (2003). Novel system for engineering bioartificial tendons and application of mechanical load. Tissue Eng 9, 967–979.[CrossRef][Medline]
Green HJ, Klug GA, Reichmann H, Seedorf U, Wiehrer W & Pette D (1984). Exercise-induced fibre type transitions with regard to myosin, parvalbumin, and sarcoplasmic reticulum in muscles of the rat. Pflugers Arch 400, 432–438.[CrossRef][Medline]
Hartner KT & Pette D (1990). Fast and slow isoforms of troponin I and troponin C. Distribution in normal rabbit muscles and effects of chronic stimulation. Eur J Biochem 188, 261–267.[Medline]
Hecker L, Baar K, Dennis RG & Bitar KN (2005). Development of a 3-dimensional physiological model of the internal anal sphincter bioengineered in-vitro from isolated smooth muscle cells. Am J Physiol Gastrointest Liver Physiol. 289, 188–196.
Huang
YC, Dennis
RG, Larkin
L
&
Baar
K (2005). Rapid formation of functional muscle in vitro using fibrin gels. J Appl Physiol
98, 706–713.
Kosnik PE, Faulkner JA & Dennis RG (2001). Functional development of engineered skeletal muscle from adult and neonatal rats. Tissue Eng 7, 573–584.[CrossRef][Medline]
Kubis
HP, Scheibe
RJ, Meissner
JD, Hornung
G
&
Gros
G (2002). Fast-to-slow transformation and nuclear import/export kinetics of the transcription factor NFATc1 during electrostimulation of rabbit muscle cells in culture. J Physiol
541, 835–847.
Leor J, Aboulafia-Etzion S, Dar A, Shapiro L, Barbash IM, Battler A, Granot Y & Cohen S (2000). Bioengineered cardiac grafts: a new approach to repair the infarcted myocardium? Circulation 102, 56–61.
Li RK, Jia ZQ, Weisel RD, Mickle DA, Choi A & Yau TM (1999). Survival and function of bioengineered cardiac grafts. Circulation 100, 63–69.
Lin
J, Wu
H, Tarr
PT, Zhang
CY, Wu
Z, Boss
O, Michael
LF, Puigserver
P, Isotani
E, Olson
EN, Lowell
BB, Bassel-Duby
R
&
Spiegelman
BM (2002). Transcriptional co-activator PGC-1 drives the formation of slow-twitch muscle fibres. Nature
418, 797–801.[CrossRef][Medline]
Lou J, Tu Y, Burns M, Silva MJ & Manske P (2001). BMP-12 gene transfer augmentation of lacerated tendon repair. J Orthop Res 19, 1199–1202.[CrossRef][Medline]
Ohlendieck K, Briggs FN, Lee KF, Wechsler AW & Campbell KP (1991). Analysis of excitation-contractioncoupling components in chronically stimulated canine skeletal muscle. Eur J Biochem 202, 739–747.[Medline]
Okano T & Matsuda T (1998). Muscular tissue engineering: capillary-incorporated hybrid muscular tissues in vivo tissue culture. Cell Transplantation 7, 435–442.[CrossRef][Medline]
Radisic
M, Park
H, Shing
H, Consi
T, Schoen
FJ, Langer
R, Freed
LE
&
Vunjak-Novakovic
G (2004). Functional assembly of engineered myocardium by electrical stimulation of cardiac myocytes cultured on scaffolds. Proc Natl Acad Sci U S A
101, 18129–18134.
Rickert M, Jung M, Adiyaman M, Richter W & Simank HG (2001). A growth and differentiation factor-5 (GDF-5)coated suture stimulates tendon healing in an Achilles tendon model in rats. Growth Factors 19, 115–126.[Medline]
Rommel C, Bodine SC, Clarke BA, Rossman R, Nunez L, Stitt TN, Yancopoulos GD & Glass DJ (2001). Mediation of IGF-1-induced skeletal myotube hypertrophy by PI(3)K/Akt/mTOR and PI(3)K/Akt/GSK3 pathways. Nature Cell Biology 3, 1009–1013.[CrossRef][Medline]
Roy RK, Mabuchi K, Sarkar S, Mis C & Sreter FA (1979). Changes in tropomyosin subunit pattern in chronic electrically stimulated rabbit fast muscles. Biochem Biophys Res Commun 89, 181–187.[Medline]
Sakai
T, Li
RK, Weisel
RD, Mickle
DA, Kim
ET, Jia
ZQ
&
Yau
TM (2001). The fate of a tissue-engineered cardiac graft in the right ventricular outflow tract of the rat. J Thorac Cardiovasc Surg
121, 932–942.
Salmons S & Sreter FA (1976). Significance of impulse activity in the transformation of skeletal muscle type. Nature 263, 30–34.[CrossRef][Medline]
Salmons S & Vrbova G (1967). Changes in the speed of mammalian fast muscle following longterm stimulation. J Physiol 192, 39P–40P.
Smalt R, Mitchell FT, Howard RL & Chambers TJ (1997). Mechanotransduction in bone cells: induction of nitric oxide and prostaglandin synthesis by fluid shear stress, but not by mechanical strain. Adv Exp Med Biol 433, 311–314.[Medline]
Sreter FA, Gergely J, Salmons S & Romanul F (1973). Synthesis by fast muscle of myosin light chains characteristic of slow muscle in response to long-term stimulation. Nat New Biol 241, 17–19.[Medline]
Takahashi
M
&
Hood
DA (1993). Chronic stimulation-induced changes in mitochondria and performance in rat skeletal muscle. J Appl Physiol
74, 934–941.
Thomson RC, Yaszemski MJ, Powers JM & Mikos AG (1995). Fabrication of biodegradable polymer scaffolds to engineer trabecular bone. J Biomater Sci Polym Ed 7, 23–38.[Medline]
Vandenburgh
H
&
Kaufman
S (1979). In vitro model for stretch-induced hypertrophy of skeletal muscle. Science
203, 265–268.
Vandenburgh
HH, Shansky
J, Solerssi
R
&
Chromiak
J (1995). Mechanical stimulation of skeletal muscle increases prostaglandin F2 production, cyclooxygenase activity, and cell growth by a pertussis toxin sensitive mechanism. J Cell Physiol
163, 285–294.[CrossRef][Medline]
van Wachem PB, Brouwer LA & van Luyn MJ (1999). Absence of muscle regeneration after implantation of a collagen matrix seeded with myoblasts. Biomaterials 20, 419–426.[CrossRef][Medline]
Vunjak-Novakovic G, Altman G, Horan R & Kaplan DL (2004). Tissue engineering of ligaments. Annu Rev Biomed Eng 6, 131–156.[CrossRef][Medline]
Wartenberg
M, Donmez
F, Ling
FC, Acker
H, Hescheler
J
&
Sauer
H (2001). Tumor-induced angiogenesis studied in confrontation cultures of multicellular tumor spheroids and embryoid bodies grown from pluripotent embryonic stem cells. Faseb J
15, 995–1005.
Wolfman NM, Hattersley G, Cox K, Celeste AJ, Nelson R, Yamaji N, Dube JL, DiBlasio-Smith E, Nove J, Song JJ, Wozney JM & Rosen V (1997). Ectopic induction of tendon and ligament in rats by growth and differentiation factors 5, 6, and 7, members of the TGF-ß gene family. J Clin Invest 100, 321–330.[Medline]
Zador E, Fenyvesi R & Wuytack F (2005). Expression of SERCA2a is not regulated by calcineurin or upon mechanical unloading in skeletal muscle regeneration. FEBS Lett 579, 749–752.[CrossRef][Medline]
Zador E & Wuytack F (2003). Expression of SERCA2a is independent of innervation in regenerating soleus muscle. Am J Physiol 285, C853–C861.
Zimmermann WH, Didie M, Wasmeier GH, Nixdorff U, Hess A, Melnychenko I, Boy O, Neuhuber WL, Weyand M & Eschenhagen T (2002a). Cardiac grafting of engineered heart tissue in syngenic rats. Circulation 106, I151–I157.
Zimmermann WH, Fink C, Kralisch D, Remmers U, Weil J & Eschenhagen T (2000). Three-dimensional engineered heart tissue from neonatal rat cardiac myocytes. Biotechnol Bioeng 68, 106–114.[CrossRef][Medline]
Zimmermann
WH, Schneiderbanger
K, Schubert
P, Didie
M, Munzel
F, Heubach
JF, Kostin
S, Neuhuber
WL
&
Eschenhagen
T (2002b). Tissue engineering of a differentiated cardiac muscle construct. Circ Res
90, 223–230.
Acknowledgements
I would like to thank R. G. Dennis and Y. C. Huang for outstanding support and assistance. This work was supported by a grant from the defense advanced research projects agency (Navy) contract no. N66001 [GenBank] –02-C-8034.
This article has been cited by other articles:
|
|
 |
|
  C. K. Hager-Ross, C. S. Klein, and C. K. Thomas Twitch and Tetanic Properties of Human Thenar Motor Units Paralyzed by Chronic Spinal Cord Injury J Neurophysiol, July 1, 2006; 96(1): 165 - 174. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
