Fisher John P. e.a. (ed.) Tissue Engineering
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15-10 Tissue Engineering
Electrical current has been shown to stimulate bone regeneration and enhance its healing [72–75].
Bioreactors have been designed to electrically stimulate cells seeded on conductive surfaces [75]. After
culturing calvarial osteoblasts under an electric field with 10 µA and 10 Hz for 6 h/day, the cell number
was higher by 46% and the amount of calcium deposited was raised by 307% after 21 days, compared to
nonstimulated cells. Upregulation of mRNA expression for collagen type I was also observed.
As demonstrated by several studies, the usage of flow perfusion greatly enhances the formation of bone
matrix. By doing this, mechanotransduction mechanisms related to the differentiation of osteoblastic cells
are potentially activated. Great challenges are still encountered however. What are the actual mechanisms
that transduce the external shear forces and influence the cellular behavior? Currently, only estimates of the
shear rates at the interior of three-dimensional scaffolds have been provided, and detailed mathematical
modeling needs to be conducted. This will allow the determination of the actual values of the shear rate
inside the constructs and their spatial distribution.
Using larger scaffolds can represent another problem; is it possible to achieve a completely homogeneous
distribution of matrix in larger constructs? What would be the necessary culturing conditions to achieve
this? How long must the cells be cultured to produce an osteoinductive enough matrix? And how long
and under what conditions must the cells be precultured prior to the application of mechanical forces to
avoid their detachment?
15.4.3 Cartilage
Cartilage is a tissue with limited capabilities of regeneration when damaged. Between the two general types
of cartilage, articular cartilage is continuously exposed to mechanical forces and needs to dissipate loads
under physiological conditions. On the other hand, in other parts of the body the elastic properties of
the cartilage are more important. Cartilaginous tissue has been engineered using spinner flasks, rotating-
wall bioreactors, perfusion systems, and compression bioreactors that better mimic the physiological
environment of the cartilage tissue. The most widely used cell types for the regeneration of cartilage
are mesenchymal stem cells or primary chondrocytes. Regarding the scaffold, different materials have
been used; synthetic and natural hydrogels are among the most popular choices, including collagen, poly
α-hydroxy esters, and their copolymers. Fibrinogen-based and glycosaminoglycan (GAG)-based matrices
are also being employed [76,77]. Some of the markers used to evaluate the formation of cartilaginous
matrix are cellularity, production of collagen type II, and production of GAGs [8]. It has to be pointed
out that the synthesis of native cartilage dramatically varies in different locations of the body.
Vunjak-Novakovic et al. [10] compared the performance of different bioreactors (static, spinner flask,
and rotating wall) in the formation of cartilaginous matrix. Highly porous PGA scaffolds were seeded
with chondrocytes and cultured statically, in a spinner flask, and a rotating-wall vessel for 6 weeks. At
the end of the culture period, GAG and collagen type II levels were five times greater than the values
obtained in early culture. The spinner flask induced a larger production of GAGs and collagen type II;
however, an even greater difference was observed in the rotating bioreactor compared to the static culture.
Histomorphometric studies revealed the formation of tissue in the periphery of the constructs cultured
statically, while those in the spinner flask had an outer fibrous capsule in spite of the increment in
mass transport. Scaffolds cultured in the rotating-wall bioreactor, on the other hand, showed a better
distribution of the matrix, but a gradient in concentration of GAGs was observed in all the constructs.
Gooch et al. [78] studied the effect of shear stress on the formation of cartilage matrix in a spinner flask
and compared it with a static culture. Chondrocytes were dynamically seeded for 3 days on fibrous PGA
matrices with a fiber diameter of 13 µm using a spinner flask bioreactor. Cultivation was carried out for
42 days at different mixing intensities. Production of GAGs was greater in the spinner flask, and increased
along with the mixing intensity. Collagen production showed a similar behavior although no significant
difference was observed among the different mixing intensities.
The effect of shear stress has also been conducted in a rotating-wall vessel [11]. Variation of the rotation
speed of the mobile wall when culturing chondrocytes in poly(lactic acid) (PLA) foams for 28 days did
not cause a change in cell density among scaffolds cultured at different rotation rates although they were
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Tissue Engineering Bioreactors 15-11
larger than those found in static conditions. However, the deposition of matrix GAGs decreased at higher
shear rates, whereas the collagen production showed a contrary behavior. The production of collagen
increased along with the shear rate and time of cultivation, showing a more dramatic dependence on the
rotation speed.
It is important to point out that the shear stress induced in a rotating-wall bioreactor occurs only at
the surface of the scaffold; therefore, perfused cartridges have been used to produce stress stimulation at
the interior of the construct. Pazzano et al. [79] cultured calf chondrocytes seeded on PGA matrices for
4 weeks under static and flow perfusion conditions. Not only did the perfusion system increase the amount
of GAGs by 180% compared with the static conditions, but it also created an organized, homogeneous
matrix. After the 4 weeks of culture, chondrocytes were aligned in the direction of flow, creating a structure
similar to that of some regions of native articular cartilage [80].
Native articular cartilage withstands up to 20 MPa due to compression in vivo; therefore, bioreactors
using this kind of stimulus have been implemented obtaining promising results [6,81–84]. Mizuno et al.
[85] used a culture system that consisted of a perfusion column and a pressurizer. Medium is peristaltically
pumped through the perfusion column were the scaffolds are kept. Downstream from the column there is a
back-pressure regulator. Cyclic pressure was controlled by using a needle valve and a computer-controlled
spring. Chondrocytes were statically seeded in porous collagen matrices for 3 days. Posteriorly, the sponges
were introduced in the perfusion chamber, and flow was started at the rate of 0.33 ml/min. The culture
conditions were pure perfusion, cyclic compression (0.015 Hz) 2.8 MPa, and constant compression at
2.8 MPa of hydrostatic fluid pressure. Production of GAGs and collagen type II were enhanced by the
application of pressure load even though there were no significant differences in cellularity. Hydrostatic
pressure produced about 3.1 times more sulfated GAGs than the controls, whereas the cyclic pressure
produced 2.7 times more sulfated GAGs than the controls. In addition, a native-like matrix was observed,
containing lacunae that entrapped the chondrocytes and a uniform spatial distribution [85]. Carver et al.
[86] had found similar results when comparing the influences of cyclic pressure, flow perfusion, and
culture in spinner flasks, in the formation of cartilaginous matrix. They found that the combination of
flow perfusion and intermittent pressurization seemed to accelerate the matrix formation. In agreement
with these findings, Hung et al. [84] reported the production of cartilage-like tissue after culturing
chondrocytes seeded on agarose gels for 8 weeks under physiologic deformational loading.
The variability in the synthesis and the mechanical properties of cartilage in different locations of the
body and zones of the same location introduces a great challenge in the regeneration of cartilage tissue. Is
the mechanical environment the controlling factor in the formation of zone and location-specific ECM?
Or in different locations of the body, are chondrocytes preprogrammed with different genetic information
to generate a specific type of extracellular matrix? How do chondrocytes in the growth plate cartilage
recognize a differentiation pattern that leads to bone formation? The elucidation of these questions will
provide valuable information to cartilage tissue engineers.
15.4.4 Anterior Cruciate Ligament and Tendons
Ligaments and tendons operate in the presence of various types and levels of mechanical strains that
are expected to influence the behavior of cells residing in these tissues. Mesenchymal stem cells (MSCs)
have been shown to preferentially differentiate into ligament fibroblasts in the presence of mechanical
forces when cultured in bioreactors that generated mechanical strains that resemble the native ligament
stretching conditions [87].
The most widely used material as scaffolding for the regeneration of anterior cruciate ligament (ACL)
and tendon is collagen; other materials such as polymeric fibrous matrices have been utilized [88].
However, the mechanical properties of collagen bundles are considerably inferior to those of the native
tissue. This problem can be partially overcome by preferentially orienting the collagen fibers of the matrix
through the seeding of cells and the application of mechanical strain [89].
The principle of a bioreactor that can mimic the biomechanical conditions of ligament and tendons is
shown in Figure 15.5. The construct is attached to a moving platform that produces mechanical strain
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Construct
Moving platform
FIGURE 15.5 Principle of cyclic strain bioreactor for ligament and tendon engineering.
through stretching. Langelier et al. [89] designed a cyclic traction device that can produce these mechanical
strains on cell-seeded constructs. This machine consists of an actuating unit that controls the cyclic motion
frequency (0.01 to 1 cycle/min) and amplitude (0 to 2 cm) for collagenous matrices with a length between
2 and 10 cm. A transmission unit connects the actuating unit to the testing unit, and its purpose is to
transmit motion to the testing unit through a rotating shaft. The testing compartment unit, shown in
Figure 15.4, is where the matrix is located. The two walls at both extremes are made out of stainless
steel, and the material chosen for the other surfaces was acrylic to allow visualization of the construct.
The top of the compartment is a removable acrylic plate screwed to the walls and is removed when the
scaffold needs to be manipulated. Generally, the collagen gel is formed over two bone anchors for adequate
mechanical attachment to the traction machine [89]. The construct can be subjected to cyclic stretching at
a frequency of up to 1 Hz and amplitude from 0 to 30 mm for any extended period of time. The system is
controlled via special software, and the experimental conditions of stretching and amplitude can be easily
changed [90].
MSCs seeded on collagen gels were cultured under translational and rotational strain concurrently.
After 21 days of culture, a ligament-like morphology was observed in the constructs, with elongated, well-
organized collagen types I and III in the longitudinal direction. Unlike the constructs cultured statically,
those under mechanical strain presented an ECM that contained tenascin-C, which is a marker of ligament
ECM [87]. A similar behavior was observed by Awad et al. [91] when MSC-seeded collagen scaffolds were
cultured under a continuously strained environment; formation of organized collagen bundles was also
observed.
Cyclic stretching has been also implemented in tendon tissue engineering and has demonstrated a
significant effect on the alignment of collagen bundles [92]. Avian tendon internal fibroblasts were incor-
porated into type-I collagen matrices at a rate of 2×10
5
cells per scaffold. Mechanical loading was applied
1 h per day with 1% elongation and 1 Hz for up to 11 days. Loaded constructs presented aligned cells that
spread throughout the matrix, with elongated nuclei and cytoplasmic extensions. Moreover, the modulus
of elasticity of the mechanically loaded constructs was 2.9 times greater than those of the nonstimulated
controls [92].
One of the challenges in tendon and ligament tissue engineering is the selection of cell source for their
repair. Although MSCs have been shown to have the ability to differentiate into tendon and ligament cells
in the presence of mechanical forces, the use of specific growth factor combinations that can initiate the
differentiation toward these phenotypes in regular cell cultures would minimize the time of tissue regen-
eration in bioreactors. In addition to that, the specific cellular mechanisms that transduce the mechanical
signals to these cells and control their behavior are not yet clearly understood.
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15.4.5 Other Tissues
Apart from the previously mentioned applications where bioreactors have been widely utilized, bioreactors
have been used for the culture of cell types involved in the regeneration of other tissues, including
hepatocytes and neuronal cells. The high metabolic requirements of hepatic cells make the use of perfusion
systems for their culture ideal. Powers et al. [93] cultured hepatocytes in the presence of continuous flow
perfusion and the cells remained viable for up to 2 weeks. The use of electromagnetic fields in neuronal
tissue regeneration has also provided elegant ways for the alignment of neuronal cells in the guided
regrowth of axons [94].
15.5 Design Considerations
When designing a bioreactor, the first factor that must be taken into account is the kind of stimulation to
be emulated. Therefore, the apparatus must be able to cause similar effects to those found physiologically
and at matching conditions, in addition to ensuring proper transportation of nutrients. A temperature of
37
◦
C and a humid atmosphere with 5% CO
2
are required [95]. Other important factors are:
1. Easy handling and assembling, lowering risks of contamination
2. Sterilizability
3. Fit into an incubator and operate at the given conditions
4. Allow change of culture medium aseptically
5. Facilitate monitoring of tissue formation
6. Automatically operated in order to ensure reproducibility of operation conditions
7. Ability to produce several constructs at the same time
8. Ease of scaling up
15.6 Challenges in Bioreactor Technologies
Implementation of bioreactorsin the process of tissue engineering has been driven by the need to mimic the
physiological conditions in which cells operate at different locations of the human body. Most bioreactors
in reality are biomimetic environments that induce morphological arrangements similar to those found
in native tissues. To a certain degree, the new technologies have been successful in achieving the initial
goals: creation of native tissue-like matrices; nevertheless, many obstacles are yet to be overcome in order
to produce efficient and practical constructs that can be used for the regeneration of damaged or lost
organs.
The transport of nutrients and oxygen still remains an issue in the processing of many tissues. Some cells
have a high metabolic demand; such is the case of hepatic tissue, thus requiring an efficient transportation
of these components to the place of growth. As discussed in this chapter, part of this problem has been
overcome by the utilization of perfusion systems. However, scaling up the size of the construct may pose
new challenges. When increasing the dimensions of the scaffolding, other interrogatives arise. Would it
still be possible to obtain a homogeneous distribution of cells and ECM throughout the scaffold? What is
the flow rate necessary to guarantee the delivery of oxygen and nutrients to the interior of the construct?
Cell source is a factor that has not been studied in detail. It is not clear, for the engineering of some
tissues, whether cells extracted from different locations could have different responses under identical
stimulation. In the case of bone the question arises, as to whether cells extracted from an area that is not
load bearing respond in similar fashion to those harvested from a load-bearing site like the femur? Similar
questions are unanswered for cartilage and ligaments.
Bone, cartilage, vascular grafts, and ligaments, among others, require potent mechanical properties
that have not been matched by the engineered constructs to this day. Establishing effective mechanical
properties depends not only on the biomaterial used but also in the bioreactor design and time of culture.
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Mechanical stimulations have proved to be the path to generate mechanically efficient implants, but not all
biomaterials can be exposed to mechanical strains without generating some degree of structural damage.
Standard criteria for the evaluation of the quality of the final construct are necessary from a mechanical,
chemical, biological, and morphological point of view. Large variations on the conditions of operation
and culture in different bioreactor designs to engineer a given tissue prevent the comparison of different
studies and thereby the drawing of definite conclusions regarding the optimization of these conditions.
In addition, histological evaluation of the regenerated tissue (or in vitro generated graft) should always be
complemented by appropriate mechanical tests.
Time also represents a significant concern when engineering tissue grafts. Given the fact that the
engineered constructs will ultimately be transplanted into diseased patients, the time of production
becomes a critical factor. The ultimate goal is to produce, in a short period of time, a construct that can
induce the formation of new tissue in vivo. It is obvious that the definition of a “short period of time”
depends upon the tissue being engineered and the urgency for a replacement. Some tissues like bone or
cartilage could offer some flexibility in the time of production depending on the type of injury, but in
life-threatening situations, such as failing heart valves, it is important to rapidly produce a construct.
Consequently, tissue engineering strategies should always attempt to minimize the time of production.
The answer to shortening the time of generation of engineered tissues may lie in the combination of
mechanical and chemical signals. Both principles have been studied separately; certain growth factors
such as bone morphogenetic proteins, vascular endothelial growth factor, and insulin like growth factors,
among others, have been shown to greatly help the formation of tissues in vitro and in vivo. Therefore, it is
expected that the combination of growth factors and mechanical stimulation in a bioreactor could generate
mature matrices at a faster pace. It is important as well to determine what the intracellular mechanisms
related to the signaling pathways are. A better understanding of the biological mechanisms involved in the
mechanosensing processes will give a new perspective in the design of new experiments and improvement
of those already existent. Moreover, new directions could be taken based on these mechanisms.
Acknowledgment
We would like to acknowledge the Seed Grant Program of the Bioengineering Center at the University of
Oklahoma.
References
[1] Bonassar, J. and Vacanti, C. Tissue engineering: the first decade and beyond. J. Cell Biochem. Suppl.
30/31, 297, 1998.
[2] Boden, S.D. Bioactive factors for bone tissue engineering. Clin. Orthop., 367S, S84, 1999.
[3] Lieberman, J.R., Daluiski, A., and Einhorn, T.A. The role of growth factors in the repair of bone.
J. Bone Joint Surg., 84-A, 1032, 2002.
[4] Wang, E.A. et al. Recombinant human bone morphogenetic protein induces bone formation. Proc.
Natl Acad. Sci. USA, 87, 2220, 1990.
[5] Trivedi, N. et al. Improved vascularization of planar diffusion devices following continuous infusion
of vascular endothelial growth factor (VEGF). Cell Transplant, 8, 175, 1999.
[6] Stoltz, J.F. et al. Influence of mechanical forces on cells and tissues. Biotechnology, 37, 3, 2000.
[7] Sikavitsas, V.I., Bancroft, G.N., and Mikos, A.G. Formation of three-dimensional cell/polymer
constructs for bone tissue engineering in a spinner flask bioreactor and a rotating wall Bessel
bioreactor. J. Biomed. Mater. Res., 62, 136, 2002.
[8] Darling, E.M. and Athanasiou, K.A. Articular cartilage bioreactors and bioprocesses. Tissue Eng.,9,
9, 2003.
[9] Goldstein, A.S. et al. Effect of convection on osteoblastic cell growth and function in biodegradable
polymer foam scaffolds. Biomaterials, 22, 1279, 2001.
mikos: “9026_c015” — 2007/4/9 — 15:51 — page 15 — #15
Tissue Engineering Bioreactors 15-15
[10] Vunjak-Novakovic, G., Obradovic, B., and Free, L.E. Bioreactor studies of native and tissue
engineered cartilage. Biorheology, 39, 259, 2002.
[11] Saini, S. and Wick, T.M. Concentric cylinder bioreactor for production of tissue engineered cartilage:
effect of cell density and hydrodynamic loading on construct development. Biotechnol. Prog., 19,
510, 2003.
[12] Sutherland F.W. et al. Advances in the mechanisms of cell delivery to cardiovascular scaffolds:
comparison of two rotating cell culture systems. ASAIO J., 48, 346, 2002.
[13] Vunjak-Novakovic, G. et al. Microgravity studies of cells and tissues. Ann. N Y Acad. Sci., 974, 504,
2002.
[14] Begley, C.M. and Kleis, S.J. The fluid dynamic and shear environment in the NASA/JSC rotating
wall perfused-vessel bioreactor. Biotechnol. Bioeng., 70, 32, 2000.
[15] Williams, K.A. et al. Computational fluid dynamics modeling of steady-state momentum and mass
transport in a bioreactor for cartilage tissue engineering. Biotechnol. Prog., 18, 951, 2002.
[16] Mizuno, S., Allemann, F., and Glowacki, J. Effects of medium perfusion on matrix production by
bovine chondrocytes in three-dimensional collagen sponges. J. Biomed. Mater. Res., 56, 368, 2001.
[17] Navarro, F.A. et al. Perfusion of medium improves growth of human oral neomucosal tissue
constructs. Wound Repair Regen., 9, 507, 2001.
[18] Bancroft, G.N., Sikavitsas, V.I., and Mikos, A.G. Design of a flow perfusion bioreactor system for
bone–tissue engineering applications. Tissue Eng., 9, 549, 2003.
[19] Bancroft, G.N. et al. Fluid flow increases mineralized matrix deposition in 3D perfusion culture
of marrow stromal osteoblasts in a dose-dependent manner. Proc. Natl Acad. Sci. USA, 99, 12600,
2002.
[20] Cartmell, S.H. et al. Effects of medium perfusion rate on cell seeded three-dimensional bone
constructs in vitro. Tissue Eng., 9, 1197, 2003.
[21] Martin, I., Wendt, D., and Herberer, M. The role of bioreactors in tissue engineering. Trends
Biotechnol., 22, 80–86, 2004.
[22] Li, Y. Effects of filtration seeding on cell density, spatial distribution, and proliferation in nonwoven
fibrous matrices. Biotechnol. Prog., 17, 935, 2001.
[23] Holy, C.E., Shoichet, M.S., and Davies, J.E. Engineering three-dimensional bone tissue in vitro using
biodegradable scaffolds: investigating initial cell-seeding density and culture period. J. Biomed.
Mater. Res., 51, 376, 2000.
[24] Vunjak-Novakovic, G. et al. Dynamic cell seeding of polymer scaffolds for cartilage tissue
engineering. Biotechnol. Prog., 14, 193, 1998.
[25] Sucosky, P. et al. Fluid mechanics of a spinner-flask bioreactor. Biotechnol. Bioeng., 85, 34, 2004.
[26] Burg, K.J.L. et al. Comparative study of seeding methods for three-dimensional polymeric scaffolds.
J. Biomed. Mater. Res., 51, 642, 2000.
[27] Wendt, D. et al. Oscilating perfusion of cell suspensions through three-dimensional scaffolds
enhances cell seeding efficiency and uniformity. Biotechnol. Bioeng., 84, 205, 2003.
[28] Sodian, R. et al. Tissue-engineering bioreactors: a new combined cell-seeding and perfusion system
for vascular tissue engineering. Tissue Eng., 8, 863, 2002.
[29] Carrier, R.L. et al. Cardiac tissue engineering: cell seeding, cultivation parameters, and tissue
construct characterization. Biotechnol. Bioeng., 64, 580, 1999.
[30] Tranquillo, R. The tissue-engineered small-diameter artery. Ann.NYAcad. Sci., 961, 251, 2002.
[31] L’Heureux, N. et al. A completely biological tissue-engineered human blood vessel. FASEB J., 12,
47, 1998.
[32] Guyton, A. and Hall, J. Textbook of Medical Physiology, 10th ed., Saunders Company, 2000, p. 145.
[33] Braddock, M. et al. Fluid shear stress modulation of gene expression in endothelial cells. New
Physiol. Sci., 13, 241, 1998.
[34] Nerem, R. and Sliktar, D. Vascular tissue engineering. Annu. Rev. Biomed. Eng., 3, 225, 2001.
[35] Thompson, C.A. et al. A novel pulsatile, laminar flow bioreactor for the development of tissue-
engineered vascular structures. Tissue Eng., 8, 1083, 2002.
mikos: “9026_c015” — 2007/4/9 — 15:51 — page 16 — #16
15-16 Tissue Engineering
[36] Sodian, R. et al. New pulsatile bioreactor for fabrication of tissue-engineered patches. J. Biomed.
Mater. Res. (Appl. Biomater.), 58, 401, 2001.
[37] Dumont, K. et al. Design of new pulsatile bioreactor for tissue engineered aortic heart valve
formation. Artif. Organs, 26, 710, 2002.
[38] Hoerstrup, S.P. et al. New pulsatile bioreactor for in vitro formation of tissue engineered heart
valves. Tissue Eng., 6, 75, 2000.
[39] Weinberg, C.B. and Bell, E. A blood vessel model constructed from collagen and cultured vascular
cells. Science, 23, 397, 1986.
[40] Niklason, L.E. et al. Functional arteries grown in vitro. Science, 284, 489, 1999.
[41] Lian, X. and Howard, P.G. Blood vessels, in Principles of Tissue Engineering, Lanza, R., Langer, R.,
and Vacanti, J. (Eds.), 2nd ed., Academic Press, New York, 2000, chap. 32.
[42] Niklason L.E. and Langer, R.S. Advances in tissue engineering of blood vessels and other tissues.
Transp. Immunol., 5, 303, 1997.
[43] Hoerstrup, S.P. et al. Tissue engineering of small caliber vascular grafts. Eur. J. Cardiothorac. Surg.,
20, 164, 2001.
[44] Selitkar, D. et al. Dynamic mechanical conditioning of collagen–gel blood vessel constructs induces
remodeling in vitro. Ann. Biomed. Eng., 28, 351, 2000.
[45] Peng, X. et al. In vitro system to study realistic pulsatile flow and stretch signaling in cultured
vascular cells. Am. J. Physiol. Cell Physiol., 279, C797, 2000.
[46] Barron, V. et al. Bioreactors for cardiovascular cell and tissue growth: a review. Ann. Biomed. Eng.,
31, 1017, 2003.
[47] Mann, B.K. and West, J.L. Tissue engineering in the cardiovascular system: progress toward a tissue
engineered heart. Anat. Rec., 263, 367, 2001.
[48] Schenke-Layland, K. Complete dynamic repopulation of decellularized heart valves by application
of defined physical signals — an in vitro study. Cardiovasc. Res., 60, 497, 2003.
[49] Mol, A. The relevance of large strains in functional tissue engineering of heart valves. Thorac.
Cardiovasc. Surg., 51, 78, 2003.
[50] Goldstain, A.S. et al. Effect of osteoblastic culture conditions on the structure of poly(dl-Lactic-co-
Glycolic acid) foam scaffolds. Tissue Eng., 5, 421, 1999.
[51] Botchwey, E.A. et al. Bone tissue engineering in a rotating bioreactor using a microcarrier matrix
system. J. Biomed. Mater. Res., 55, 242, 2001.
[52] Bancroft, G.N. and Mikos, A.G. Bone tissue engineering by cell transplantation, in Polymer Based
Systems on Tissue Engineering, Replacement and Regeneration, Reis, R.L. and Cohn, D. (Eds.), Kluwer
Academic Publishers, Boston, 2002, p. 251.
[53] Nam, Y.S., Yoon, J.J., and Park, T.G. A novel fabrication method of macroporous biodegradable
polymer scaffolds using gas foaming salt as a porogen additive. J. Biomed. Mater. Res. (Appl.
Biomater.), 53, 1, 2000.
[54] Hollinger, J.O. and Battistone, G.C. Biodegradable bone repair materials. Synthetic polymers and
ceramics. Clin. Orthop. Relat. Res., 207, 290, 1986.
[55] Nefussi, J.R. et al. Sequential expression of bone matrix proteins during rat calvaria osteo-
blast differentiation and bone nobule formation in vitro. J. Histochem. Cytochem., 45, 493,
1997.
[56] Ishaug, S.L. et al. Bone formation by three-dimensional stromal osteoblast culture in biodegradable
polymer scaffolds. J. Biomed. Mater. Res., 36, 17, 1997.
[57] Shea, L. et al. Engineered bone development from pre-osteoblast cell line on three-dimensional
scaffold. Tissue Eng., 6, 605, 2000.
[58] Pavlin D. et al. Mechanical loading stimulates differentiation of periodontal osteoblasts in a mouse
osteoinduction model: effect on type I collagen and alkaline phosphatase genes. Calcif. Tissue Int.,
67, 163, 2002.
[59] Sikavitsas, V.I., Temenoff, J.S., and Mikos, A.G. Biomaterials and bone mechanotransduction.
Biomaterials, 22, 2581, 2001.
mikos: “9026_c015” — 2007/4/9 — 15:51 — page 17 — #17
Tissue Engineering Bioreactors 15-17
[60] Reich, K.M. and Frangos, J.A. Effect of flow on prostaglandin E2 inositol trisphosphate levels in
osteoblasts. Am. J. Physiol., 261, C429, 1991.
[61] Klein-Nulend, J. et al. Nitric oxide response to shear stress by human bone cell cultures is endothelial
nitric oxide synthase dependent. Biochem. Biophys. Res. Commun., 250, 108, 1998.
[62] Owan, I. et al. Mechanotransduction in bone: osteoblasts are more responsive to fluid forces than
mechanical strain. Am. J. Physiol., 273, C810, 1997.
[63] Knothe-Tate, M.L., Knothe, U., and Niederer, P. Experimental elucidation of mechanical load-
induced fluid flow and its potential role in bone metabolism and functional adaptation. Am. J. Med.
Sci., 316, 189, 1998.
[64] Burger, E.H. and Klein-Nulend, J. Mechanotransduction in bone: role of the lacuno-canalicular
network. FASEB J., 13S, S101, 1997.
[65] Cowin, S.C. Bone poroelasticity. J. Biomech., 32, 217, 1999.
[66] Van den Dolder, J. et al. Flow perfusion culture of marrow stromal osteoblasts in titanium fiber
mesh. J. Biomed. Mater. Res., 64A, 235, 2003.
[67] Meinel, L. et al. Bone tissue engineering using human mesenchymal stem cells: effects of scaffold
material and medium flow. Ann. Biomed. Eng., 32, 112, 2004.
[68] Sikavitsas, V.I. et al. Mineralized matrix deposition by marrow stromal osteoblasts in 3D perfu-
sion culture increases with increasing fluid shear forces. Proc. Natl Acad. Sci. USA, 100, 14683,
2003.
[69] Neidlinger-Wilke, C., Wilke, H.J., and Claes, L. Cyclic stretching of human osteoblasts affects
proliferation and metabolism: a new experimental method and its application. J. Orthop. Res., 12,
70, 1994.
[70] Winter, L.C. et al. Intermittent versus continuous stretching effects on osteoblast-like cells in vitro.
J. Biomed. Mater. Res., 67A, 1269, 2003.
[71] Shimko, D. et al. A device for long term, in vitro loading of three-dimensional natural and engineered
tissues. Ann. Biomed. Eng., 31, 1347, 2003.
[72] Yonernori, K. Early effects of electrical stimulation on osteogenesis. Bone, 19, 173, 1996.
[73] Brighton, C.T. et al. Signal transduction in electrically stimulated bone cells. J. Bone Joint Surg. Am.,
83-A, 1514, 2003.
[74] Bassett, C.A., Pawluk, R.J., and Pilla, A.A. Augmentation of bone repair by inductively coupled
electromagnetic fields. Science, 184, 575–577, 1984.
[75] Supronowicz, P.R. et al. Novel current-conducting composite substrates for exposing osteoblasts to
alternating current stimulation. J. Biomed. Mater. Res., 59, 499, 2002.
[76] Temenoff, J.S. and Mikos, A.G. Review: tissue engineering for regeneration of articular cartilage.
Biomaterials, 21, 431, 2000.
[77] Hendrickson, D.A. et al. Chondrocyte-fibrin matrix transplants for resurfacing extensive articular
cartilage defects. J. Orthop. Res., 12, 485, 1994.
[78] Gooch, K.J. et al. Effects of mixing intensity on tissue-engineered cartilage. Biotechnol. Bioeng., 72,
402, 2001.
[79] Pazzano, D. et al. Comparison of chondrogenesis in static and perfused bioreactor culture.
Biotechnol. Prog., 16, 893, 2000.
[80] Kuettner, K.E. Biochemistry of articular cartilage in health and disease. Clin. Biochem., 25, 155,
1992.
[81] Shieh, A.C. and Athanasiou, K.A. Principles of cell mechanics for cartilage tissue engineering. Ann.
Biomed. Eng., 31, 1, 2003.
[82] Buschmann, M.D. et al. Stimulation of aggrecan synthesis in cartilage explants by cyclic loading is
localized to regions of high interstitial fluid flow. Arch. Biochem. Biophys., 366, 1, 1999.
[83] Guilak, F. et al. The effects of matrix compression on PG metabolism in articular cartilage explants.
Osteoarthr. Cartil., 2, 91, 1994.
[84] Hung, C. et al. A paradigm for functional tissue engineering of articular cartilage via applied
physiologic deformational loading. Ann. Biomed. Eng., 32, 35, 2004.
mikos: “9026_c015” — 2007/4/9 — 15:51 — page 18 — #18
15-18 Tissue Engineering
[85] Mizuno, S. et al. Hydrostatic fluid pressure enhances matrix synthesis and accumulation by bovine
chondrocytes in three-dimensional culture. J. Cell Physiol., 39, 319, 2002.
[86] Carver, S.E. and Heath, C.A. Influence of intermittent pressure, fluid flow, and mixing on the
regenerative properties of articular chondrocytes. Biotechnol. Bioeng., 65, 274, 1999.
[87] Altman, G.H. et al. Cell differentiation by mechanical stress. FASEB J., 16, 270, 2002.
[88] Goh, J.C. et al. Tissue-engineering approach to the repair and regeneration of tendons and ligaments.
Tissue Eng., 9, S31, 2003.
[89] Langelier, E. et al. Cyclic traction machine for long-term culture of fibroblast-populated collagen
gels. Ann. Biomed. Eng., 27, 67, 1999.
[90] Goulet, F. et al. Tendons and ligaments, in Principles of Tissue Engineering, Lanza, R., Langer, R.,
and Vacanti, J. (Eds.), 2nd ed., Academic Press, New York, 2000, chap. 50.
[91] Awad, H.A. et al. In vitro characterization of mesenchymal stem cell-seeded collagen scaffolds for
tendon repair: effects of initial seeding density on contraction kinetics. J. Biomed. Mater. Res., 51,
233, 2000.
[92] Garvin et al. Novel system for engineering bioartificial tendons and application of mechanical load.
Tissue Eng., 9, 130–132, 2003.
[93] Powers, M.J. et al. A microfabricated array bioreactor for perfused 3D liver cultures. Biotechnol.
Bioeng., 78, 257, 2002.
[94] Eguchi, Y., Ogiue-Ikeda, M., and Ueno, S. Control of orientation of rat Schwann cells using an 8-T
static magnetic field. Neurosci. Lett., 351, 130, 2003.
[95] Miller, W.M. Bioreactor design considerations for cell therapies and tissue engineering. In WTEC
Workshop on Tissue Engineering Research in the United States, Proceedings, MD: International
Technology Research Institute, Loyola College, Baltimore, 2000, p. 5.
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16
Animal Models for
Evaluation of
Tissue-Engineered
Orthopedic Implants
Lichun Lu
Esmaiel Jabbari
Michael J. Moore
Michael J. Yaszemski
Mayo Clinic College of Medicine
16.1 Introduction.............................................. 16-1
16.2 Animal Model Selection ................................. 16-2
16.3 Commonly Used Animals ............................... 16-2
16.4 Specific Animal Models ................................. 16-3
Biocompatibility • Biodegradation • Osteogenesis •
Chondrogenesis
16.5 Experimental Studies .................................... 16-5
Experimental Design • Evaluation Methods
References ....................................................... 16-6
16.1 Introduction
Animal models are an indispensable tool in tissue engineering research as they provide important inform-
ation that may lead to eventual development of clinically useful treatment of diseases. Current tissue
engineering strategies often involve three main components: cells, scaffolds, and bioactive factors for the
repair or regeneration of a specific tissue type. Research in animal models thus bridges the gap between
in vitro studies (such as scaffold degradation, cell–scaffold interactions, and scaffold toxicity) and human
clinical trials. Animal models have been and will continue to be used to develop an understanding of each of
the primary components separately, in combination, and ultimately in pathologic orthopedic conditions.
As an example, the appropriate sequence of studies for the development of a new biodegradable bone
regeneration material might be (1) biomaterial synthesis and structural characterization; (2) scaffold
fabrication and measurements of mechanical and degradation properties in vitro; (3) biocompatibility
and bone formation assessed by cell culture in vitro; (4) biocompatibility and degradation of the material
in vivo, typically by subcutaneous implantation in a lower-level animal model such as a rat; (5) if no
significant toxic effect is observed, the material is then evaluated for its intended application such as an
appropriate bone defect model in a higher-level animal such as a rabbit; (6) if the material functions well
16-1