Advanced Drug Delivery Reviews 33 (1998) 15–30
L
Culture of organized cell communities
Lisa E. Freed*, Gordana Vunjak-Novakovic
Division of Health Sciences and Technology, Massachusetts Institute of Technology, 45 Carleton Street, Cambridge, MA 02139, USA
Received 8 September 1997; received in revised form 28 October 1997; accepted 30 December 1997
Abstract
Cells cultured in vitro will tend to retain their differentiated phenotype under conditions that resemble their natural in vivo
environment, for example, when cultured on polymer scaffolds in tissue culture bioreactors. In this chapter, we define
organized cell communities as three-dimensional in vitro grown cell–polymer constructs that display important structural and
functional features of the natural tissue. We review representative studies in which the research goal was to culture organized
cell communities resembling cartilage, bone, skeletal muscle or cardiac-like tissue. These constructs can potentially serve as
tissue equivalents for in vivo transplantation or as a model system for the in vitro testing of cell and tissue-level responses to
molecular, mechanical or genetic manipulations.  1998 Elsevier Science B.V. All rights reserved.
Keywords: Scaffold; Bioreactor; Tissue culture; Tissue engineering; Cartilage; Bone; Skeletal muscle; Cardiac muscle
Contents
1. Introduction ............................................................................................................................................................................
2. Representative cultures of organized cell communities ...............................................................................................................
2.1. Cartilage .........................................................................................................................................................................
2.2. Bone ...............................................................................................................................................................................
2.3. Striated muscles ...............................................................................................................................................................
2.3.1. Skeletal muscle ......................................................................................................................................................
2.3.2. Cardiac muscle ......................................................................................................................................................
3. Effects of environmental factors on tissue formation in vivo and in vitro .....................................................................................
3.1. Scaffold-related factors.....................................................................................................................................................
3.1.1. Effects of cell shape and density..............................................................................................................................
3.1.2. A representative scaffold ........................................................................................................................................
3.2. Bioreactor-related factors..................................................................................................................................................
3.2.1. Mass transfer .........................................................................................................................................................
3.2.2. Mechanical factors .................................................................................................................................................
3.2.3. A representative bioreactor .....................................................................................................................................
4. Conclusion and future research directions .................................................................................................................................
4.1. The cells .........................................................................................................................................................................
4.2. The scaffold ....................................................................................................................................................................
4.3. The in vitro culture environment .......................................................................................................................................
Acknowledgments .......................................................................................................................................................................
References ..................................................................................................................................................................................
*Corresponding author. Tel.: 1 617-253-3858; Fax: 1 617-258-8827; E-mail: [email protected]
0169-409X / 98 / $ – see front matter  1998 Elsevier Science B.V. All rights reserved.
PII: S0169-409X( 98 )00017-9
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L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
1. Introduction
The loss of a tissue or its function due to congenital defects, disease or trauma is one of the most
difficult, frequent and costly problems in human
medicine [52]. Current treatment modalities include
autografts (e.g. skin, blood vessels), allografts (e.g.
heart, kidney transplants) and artificial prostheses
(e.g. joints, heart valves). However, each of the
above methods has its limitations, which include
shortage of donor tissue, immune rejection and
pathogen transfer, and limited service life, respectively. The advent of tissue engineering has been
motivated by the challenge to produce tissue substitutes that can restore the structural features and
physiological functions of natural tissues in vivo
[55].
There have been two basic approaches to tissue
engineering: (a) in vitro cultivation of cell–polymer
constructs for in vivo implantation and (b) direct in
vivo implantation of isolated cells and / or biomaterials. We will discuss only the former category here,
i.e. tissue constructs obtained by growing isolated
cells on polymer scaffolds using various in vitro
tissue culture bioreactors. We will define an organized cell community as an in vitro grown construct
that displays important structural and functional
characteristics of a natural tissue.
In addition to serving as tissue equivalents for
clinical transplantation, organized cell communities
can be used for in vitro studies of normal or
pathological tissue function. In particular, constructs
based on either normal or genetically engineered
cells can be used to study cell differentiation
and tissue formation under well controlled conditions, assess tissue responses to biochemical and
mechanical stimuli and identify the functions
of specific genes or gene products that can be
either over-expressed or knocked-out. Organized cell communities thus represent a model
system for the direct testing of cell and tissue-level
responses to molecular, mechanical or genetic
manipulations.
In the following review, we will discuss representative examples of organized cell communities as
well as the effects of environmental factors on in
vivo and in vitro tissue development.
2. Representative cultures of organized cell
communities
Representative studies in which the research goal
was to create in vitro a three-dimensional (3D)
equivalent of cartilage, bone, skeletal muscle or
cardiac muscle are listed in Table 1. These four
tissue types were selected because the feasibility of
culturing organized communities of their component
cells has recently been demonstrated by independent
research groups, and because their successful engineering is expected to have a major clinical impact.
Several key parameters varied greatly from study to
study: (1) cell source (e.g. articular cartilage, bone
marrow, skeletal or cardiac muscle, or cell line) and
age of donor (embryo, neonate, adult), (2) scaffold
material (agarose, collagen, synthetic polymer) and
form (gel, fibrous mesh, porous sponge), (3) cultivation vessel (static or mixed Petri dishes, static or
well-mixed spinner flask, perfused cartridge, rotating
vessels; Fig. 1), (4) in vitro cultivation time (one
week to seven months) and (5) methods used to
assess the resulting tissue construct (e.g. histological,
biochemical, molecular, functional).
2.1. Cartilage
Cartilage is an avascular, aneural tissue that
consists of relatively few cells, collagen ( | 60% of
the dry weight, dw), proteoglycan (PG, | 25% dw),
and other proteins and glycoproteins ( | 15% dw)
[11]. A network of fibrillar collagen (predominately
type II) provides form and a framework that immobilizes PGs and swells with water [11]. New
approaches to cartilage repair have recently been
reviewed [12,58] and a cell-based procedure for the
repair of human knee injuries [8] was approved by
the Food and Drug Administration (FDA) in August
1997. Tissue-engineered cartilage would need to be
several mm thick to repair damaged human knee
joints [11] and at least 4 mm thick for some nasal
reconstructive surgery applications [38].
Cartilaginous constructs have been cultivated in
vitro using isolated cells [e.g. chondrocytes and bone
marrow stromal cells (BMSC)], biomaterials (e.g.
agarose, synthetic polymers) and bioreactor vessels
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
17
Table 1
Selected in vitro tissue engineering studies
Cell
(source)
Scaffold
(initial dimensions)
Cultivation
conditions
Cultivation
time
Results
References
I. Cartilage
1
Chondrocyte
(bovine articular)
2
Chondrocyte
(bovine articular)
3
Chondrocyte
(bovine articular)
4
Chondrocyte
(bovine articular)
5
Chondrocyte
(rabbit articular)
6
Chondrocyte
(bovine articular)
7
Chondrocyte
(bovine articular)
8
Bone marrow stromal cell
BMSC, chick embryo
Agarose gel
(16 mm diameter 3 1 mm thick)
Fibrous PGA and porous PLLA
(10 3 5–10 3 2–3 mm)
Fibrous PGA
(10 mm diameter 3 5 mm thick)
Fibrous PGA
(10 mm diameter 3 5 mm thick)
Fibrous PGA
(10 mm diameter 3 2 mm thick)
Fibrous PGA and porous collagen
(no dimensions reported)
Fibrous PGA
(5 mm diameter 3 2 mm thick)
Fibrous PGA
(5 mm diameter 3 2 mm thick)
Static dish
10 weeks
Mechanically functional
Buschmann et al. [15]
Static dish
6 weeks
PGA better than PLLA
Freed et al. [24]
Mixed dish
8 weeks
Freed et al. [27]
Spinner flask
8 weeks
Perfused cartridge
4 weeks
Perfused cartridge
5 weeks
3.5 mm thick mixed
dish better than static
4.8 mm thick mixed
flask better than static
More tissue at edge
than at center
Cartridge better than static dish
Rotating vessel
7 months
Mixed dish
II. Bone
1
MC3T3
(osteogenic line)
2
MBA-15
(osteogenic line)
3
BMSC
(mouse)
4
BMSC
(rat)
5
BMSC
(chick embryo)
Fibrous collagen
(200 mm 2 surface)
Fibrous polyester
2000 mm 2 surface
Porous collagen
(16 mm 2 surface)
Porous PLGA
(7 mm diameter 3 1.9 mm thick)
Fibrous PGA
(5 mm diameter 3 2 mm thick)
III. Muscle
A. Skeletal
1
Skeletal myoblast
(chick embryo)
2
C2C12 with rhGH gene
(myogenic line)
B. Cardiac
1
Cardiomyocyte6b-gal
(chick embryo)
2
3
Heart cells
neonatal rat
Heart cells
neonatal rat
Vunjak-Novakovic et al. [86]
Dunkelman et al. [19]
Grande et al. [36]
4 weeks
4–8 mm thick
mechanically functional
Cartilaginous
Martin et al. [54]
Static dish
8 weeks
Osteogenesis
Casser-Bette et al. [16]
Mixed dish
4 weeks
Osteogenesis
Benayahu et al. [4]
Static dish
4 weeks
Osteogenesis
Schoeters et al. [68]
Static dish
8 weeks
Ishaug et al. [45]
Mixed dish
4 weeks
0.24 mm thick
osteoid layer
0.3–0.5 mm thick
mineralized layer
Collagen–silastic membrane
(25 3 10 mm)
Collagen/matrigel
silastic membrane
(30 3 5 mm)
Static
under
Static
under
4 weeks
30–35 3 1–2 mm
response to stretch
18 3 2.5 mm
rhGH secretion
Vandenburgh et al. [80]
Collagen gel
(15 3 13 3 5 mm)
Static dish
under tension
Fibrous PGA
(10 mm diameter 3 2 mm thick)
Fibrous PGA
(5 mm diameter 3 2 mm thick)
Rotating vessel
3 weeks
Spinner flask
1 week
dish
tension
dish
tension
(Fig. 1) as follows. Bovine articular chondrocytes
cultured in agarose gels in static Petri dishes produced a mechanically functional extracellular matrix
3 weeks
11 days
15 3 0.18 mm
contractile
b-gal activity; response
to stretch and drugs
Contractile
cardiomyocytes
Contractile
cardiomyocytes
Freed et al. [33]
Martin et al. [54]
Vandenburgh et al. [81]
Eschenhagen et al. [20]
Freed and Vunjak-Novakovic
[32]
Bursac et al. [14]
(ECM) containing glycosaminoglycan (GAG) [15].
An alternative approach is to use a synthetic polymer
instead of a naturally occuring material (e.g. agarose)
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Fig. 1. Isolated cells, e.g. from cartilage, bone, muscle or bone marrow are seeded onto polymer scaffolds and cultured in vitro, e.g. in Petri
dishes, spinner flasks, perfused cartridges or rotating vessels. The resulting engineered constructs can be used either as implants or for in
vitro research.
to allow greater flexibility and reproducibility in
scaffold design and production [78]. Cartilaginous
cell–polymer constructs based on bovine articular
chondrocytes and fibrous meshes of polyglycolic
acid (PGA) or porous sponges of poly-L-lactic acid
(PLLA) were grown in static Petri dishes [24]. Both
scaffolds had high surface areas for cell attachment
and high void volumes (97% for PGA and 91% for
PLLA) for the deposition of GAG and type II
collagen. Higher rates of cell growth and GAG
deposition were observed when chondrocytes were
grown on PGA meshes rather than on PLLA
sponges; these findings were attributed to differences
in polymer geometry (fibrous mesh vs. porous
sponge, pore size distribution) and degradation rate
(higher for PGA than PLLA). PGA scaffolds were
subsequently optimized and characterized in detail,
and a procedure was established for their reproducible processing on a commercial scale [26]. Cartilaginous tissue constructs were also formed by
expanding embryonic avian BMSCs in monolayers
in the presence of fibroblast growth factor (FGF-2)
and culturing the first passage cells on PGA scaffolds
in mixed Petri dishes [54].
Chondrocyte–PGA constructs grown in orbitally
mixed Petri dishes were thicker (up to 3.5 mm) and
more cartilaginous than those grown statically
[26,27]. The use of magnetically stirred spinner
flasks further increased the efficiency of cell seeding
to essentially 100% [87] and yielded 4.8 mm-thick
constructs [86]. The use of perfused cartridges
resulted in cartilaginous chondrocyte–PGA constructs with more cells and ECM peripherally than
centrally [19]. Another study done in perfused
cartridges showed that the choice of scaffold affected
ECM synthesis rates such that PGA and collagen
scaffolds respectively enhanced GAG and protein
synthesis rates, however, final construct dimensions
were not reported [36]. The use of rotating bioreactors in longterm (6 weeks to 7 months) cultivation of chondrocytes on PGA scaffolds resulted in
constructs that were histologically cartilaginous
throughout their entire cross-sections (5–8 mm
thick), with the exception of fibrous outer capsules
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
(0.15–0.45 mm thick) and resembled cartilage tissue
biochemically and biomechanically [33,34].
The above studies indicate that the cell source,
biomaterial scaffold and bioreactor vessel all play
important roles during the culture of organized
communities of chondrogenic cells (see Section 3).
2.2. Bone
Bone is a vascularized tissue that contains relatively few cells, an organic matrix consisting of type
I collagen and other proteins ( | 30% dw) and
inorganic mineral consisting mainly of carbonaterich hydroxyapatite ( | 70% dw) [56]. New approaches to bone repair were recently reviewed [94]
and feasibility studies have been carried out using
human BMSCs in nude mice [41]. In contrast to
cartilage, bone is expected to require relatively high
rates of mass transfer during in vitro cultivation, e.g.
by vascularization, either prior to or following in
vivo implantation.
Osteogenesis has been studied in vitro using a
variety of cells (osteoblasts, BMSCs), scaffolds
(collagen, synthetic polymers) and biochemical factors (b-glycerophosphate, dexamethasone), as follows. In vitro production of bone-specific markers
was demonstrated for MC3T3 cells grown on fibrous
collagen in static dishes [16], BMSCs grown on
porous collagen (Colla-Tec  ) in static dishes [68],
and MBA-15 cells grown on fibrous polyester
(Fibro-Cel  ) in orbitally mixed dishes [4]. In the
above three studies, cell densities decreased with
distance from the construct surface, but the overall
thicknesses of the osteoid tissue were not reported.
Bone-like tissues with mineralized surfaces (0.24
mm thick) and relatively acellular interiors were
produced by culturing BMSCs on 7 mm diameter 3
1.9 mm thick, 90% porous sponges of 75:25 poly
DL-lactic–co-glycolic acid [45] in static dishes. In
another study, bone-like tissues with mineralized
surfaces (0.3–0.5 mm thick) and interiors that resembled hypertrophic cartilage were produced by culturing BMSCs on 5 mm diameter 3 2 mm thick, 97%
porous PGA meshes in mixed Petri dishes [54].
The above studies imply that the mass transfer of
biochemical factors plays an important role during
19
the culture of organized communities of osteogenic
cells.
2.3. Striated muscles
Striated muscles are highly vascularized tissues
composed of fibers with longitudinally positioned
sarcomeres (the contractile units) that contain actin,
myosin and tropomyosin. In skeletal muscle, the
fibers are derived from end-to-end fusion of embryonic myoblasts [56], whereas in cardiac muscle,
they consist of cells joined end-to-end by specialized
junctions (intercalated discs) [57]. Cell- and tissuebased approaches to the repair of damaged or
defective skeletal muscle [46,64,81] and cardiac
muscle [48,74,75] are currently under investigation.
Skeletal and cardiac muscle receive | 25% of the
total cardiac output (CO) at rest and | 75% during
strenuous exercise, as compared to cartilage and
bone, which, respectively, receive 2 and 10% of the
CO at rest and during exercise [82]. Engineered
muscles are thus expected to require substantially
higher mass transfer rates than cartilage or bone.
2.3.1. Skeletal muscle
‘Kinetically engineered’ skeletal muscle was produced by culturing embryonic avian skeletal muscle
myoblasts on a collagen-coated elastic substratum,
either quiescently or under tension [80]. The amplitude, direction and time history of the applied
forces were designed to mimic those during in vivo
embryogenesis, i.e. static tension due to skeletal
growth and dynamic tension due to muscular contraction. Cultured cells were first subjected to unidirectional stretch, then constant tension and, finally,
to unidirectional stretch with superimposed dynamic
stretch / relaxation. This resulted in the formation of a
monolayer of multinucleated myotubes that were
orientated in the direction of stretch and which
subsequently detached from the substratum and
formed a rod-like ‘organoid’. Organoids grown in
the presence of mechanical forces were 1–2 mm in
diameter 3 30–35 mm long and contained parallel
networks of unbranched fibers, a well-defined epimysium and developing tendons [80]. The engineered muscle contracted and increased its axial
tension by 91% in response to potassium. In contrast,
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organoids grown quiescently were only 0.5–1.0 mm
long, consisted of disorganized, branching fibers and
could not perform directed work.
A related study reported the formation of tissueengineered muscle using C2C12 cells that were
genetically modified to secrete recombinant human
growth hormone (rhGH) to obtain organoids that
could potentially be used for gene therapy [81]. The
cells were mixed with collagen and Matrigel  , cast
in semicircular molds and cultivated under tension.
This resulted in the formation of rod-like organoids
that measured 2.5 mm diameter 3 18 mm long and
consisted of a 0.18-mm-thick surface layer of longitudinally oriented fibers and an inner phase with a
low density of unoriented fibers. Organoids secreted
pharmacological levels of rhGH. Loss of tension
during in vitro cultivation led to significant fiber
atrophy and loss of cellular proteins.
The above studies indicate that mechanical factors
play an important role during the culture of organized communities of skeletal myoblasts.
2.3.2. Cardiac muscle
Cardiac-like tissue constructs were recently engineered by cultivating embryonic avian cardiomyocytes in a collagen gel under tension [20].
Construct contractile function (force, frequency) was
measured by connecting one end of the construct to
an electrode (for electrophysiological stimulation)
and the other to an isometric force transducer (for
recording). Final constructs measured 15 mm 3 (6–
10) mm 3 0.18 mm and contained sarcomeres and
intercalated discs. There was a layer of longitudinally oriented cells at the construct edges and a
relatively acellular, disorganized inner tissue phase.
More than 80% of the cells displayed myocytespecific markers (actin, sarcomeric tropomyosin).
Constructs containing cells transfected with Lac-Z
expressed b-galactosidase activity. Constructs contracted spontaneously at approximately 72 beats per
min and generated tension in response to specific
mechanical, electrical and chemical stimuli.
In another set of studies, cardiac-like tissue constructs that contracted spontaneously and synchronously at 30–130 beats per min were formed by
cultivating neonatal rat heart cells on PGA meshes in
rotating vessels [32]. Final constructs measured 6
mm diameter 3 1.3 mm thick and consisted of an
approximately 0.10 mm thick, compact surface zone
with multiple layers of elongated cells and a porous
interior consisting of interconnected cells and polymer fibers. More recently, heart cell-PGA constructs
grown for 1 week in mixed flasks were shown to
have a 150 mm thick outer region containing differentiated, electromechanically coupled cardiomyocytes around a central, mostly acellular region [14].
The above studies imply that mechanical factors
(e.g. tension, hydrodynamic shear) may play a role
during the culture of organized communities of
cardiomyocytes.
3. Effects of environmental factors on tissue
formation in vivo and in vitro
The structures and functions of cells, tissues and
organs within the human body are in part determined
by environmental factors. It is likely that the same
factors that affect in vivo tissue development,
maintenance and remodeling are important during
the in vitro culture of organized cell communities.
The selected examples described above (Section 2)
suggest that at least two sets of factors can influence
cell growth and differentiation in vitro: (1) scaffoldrelated factors, including spatial cell arrangement
(i.e. 2D vs. 3D cultivation), cell morphology following attachment (e.g. flat vs. spherical), and cell
density and distribution (e.g. high or low, uniform or
nonuniform) and (2) bioreactor-related factors, including the mass transfer of chemical species to and
from the cells, and physicochemical signals (e.g.
mechanical, biochemical). The specific factors
needed during in vitro cultivation are likely to
depend on the tissue type and developmental stage.
3.1. Scaffold-related factors
Structural templates appear to modulate and
coordinate tissue formation. For example, complete
restoration of the original tissue structure and function occurs in vivo if a toxin destroys the gastrointestinal epithelial cells but spares their basement
membrane, while a scar forms if the injury destroys
both the cells and their substratum [43,85]. Most
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
normal cells are anchorage-dependent and their
growth is affected by their interaction with a substratum [21]. In general, substrates that promote cell
proliferation tend to suppress differentiation and vice
versa [44].
3.1.1. Effects of cell shape and density
Several in vitro studies have associated chondrocyte differentiation with round cell shape and / or
high cell seeding density. Benya and Shaffer [5]
found that chondrocytes that had dedifferentiated by
serial passage in monolayers redifferentiated (i.e.
reacquired a spherical shape, ceased dividing and
resumed the synthesis of PG and type II collagen)
when transferred into 3D agarose gel cultures. Some
investigators correlated increased PG synthesis with
round chondrocyte shape [34,40,59], while others
associated type II collagen synthesis with cell–cell
and cell–matrix interactions rather than cell shape
[83,84]. Chondrocyte dedifferentiation can also be
slowed or prevented by cultivation in suspension
[2,10], in aggregates [3], in alginate [7,40,66], in the
presence of ECM macromolecules [47] and on
polymer scaffolds [26,78].
It has been suggested that the early stages of
chondrocyte differentiation might be regulated by
cell-to-cell contacts in an environment that is capable
of activating the differentiation program, e.g. the
reduction of intercellular spaces during cell condensation in the precartilaginous region of the limb
[62,76]. Watt [90,91] attributed stabilized chondrocyte phenotype at high cell densities to cell-tocell contacts and / or soluble mediators and suggested
that a cell cultured in vitro will tend to retain its
differentiated phenotype under conditions that resemble its natural in vivo environment. This hypothesis
is consistent with the observed temporal and spatial
patterns of chondrogenesis in chondrocyte–PGA
constructs [34]. Possible underlying mechanisms
include changes in gene transcription and / or translation caused by cell shape changes resulting from
cell–cell and / or cell–ECM interactions [9,90,95].
Abbot and Holzer [1] hypothesized that: (a) the
effective cross-talk between cells is not dependent
upon cell density alone, but also on the presence of
homotypic differentiated cells in the immediate cell
environment, (b) the absence of physical interaction
between cells promotes DNA synthesis in order to
21
achieve the cell density needed for the cells to
communicate and express their differentiated phenotype and (c) a cell committed to the synthesis of
specialized somatic molecules will not concurrently
engage in DNA synthesis and cell multiplication.
The term ‘community effect’ was later coined by
Gurdon [39], who suggested that the ability of a cell
to respond to phenotypic induction is enhanced by,
or even dependent on, other neighboring cells differentiating in the same way at the same time.
Cultivation in 2D monolayers causes dedifferentiation in several cell types (e.g. chondrocytes, osteoblasts, cardiomyocytes) [4,5,20,84] and does not permit the complete differentiation of other cells (e.g.
skeletal myoblasts) [79]. The use of 3D scaffolds for
culturing organized cell communities provides a way
of overcoming some of these problems. Ideally, a
scaffold should meet all of the following criteria: (1)
reproducible processing into complex, 3D shapes,
(2) highly porous structure that permits a spatially
uniform cell distribution during cell seeding and
minimizes diffusional constraints during in vitro
cultivation, (3) potential to tailor its chemical, structural and mechanical properties and (4) controlled
biodegradation for long-term biocompatibility (in the
case when the organized cell community is meant to
be implanted in vivo). The scaffold surface (e.g.
chemistry, wettability) affects cell spreading and
proliferation, while scaffold structure (e.g. dimensions, porosity, pore size) may affect the spatial cell
arrangement and the transmission of biochemical and
mechanical signals.
3.1.2. A representative scaffold
One example of a scaffold that was specifically
designed for tissue engineering is shown in Fig. 2
[26]. PGA was the material selected since it met all
of the above criteria, and was FDA approved for
clinical use as an absorbable surgical suture [23].
PGA fibers were extruded and processed to form a
highly porous (96–97% void volume), mechanically
stabilized mesh of 13 mm diameter fibers formed as
5–10 mm diameter 3 2–5 mm thick discs (Fig. 2b).
Chondrocytes (bovine, rabbit, human; articular, costal), bone marrow stromal cells (embryonic chick)
and heart cells (neonatal rat) readily attached to the
PGA fibers, and retained their spherical morphology
and contractile function (Fig. 2a). Scaffold mass
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[25,31,32,86]. Flow and mixing within bioreactors
are expected to affect tissue formation in at least two
ways: (1) enhanced mass transfer (e.g. of gases,
nutrients, metabolites, growth factors) and (2) mechanical stimulation (e.g. by hydrodynamic pressure
or shear).
Fig. 2. (a) Photomicrographs of single PGA fibers with attached
chondrocytes and heart cells; (b) Scanning electron micrograph of
a fibrous polyglycolic acid (PGA) scaffold.
decreased to about 50% of the initial level over four
weeks of in vitro cultivation [26]. The time constants
for scaffold degradation and cartilaginous ECM
deposition were of the same order of magnitude, a
situation associated with enhanced tissue regeneration according to the hypothesis of isomorphous
tissue replacement [93].
3.2. Bioreactor-related factors
Bioreactors appear to permit the in vitro culture of
larger, better organized cell communities than can be
achieved using standard tissue culture techniques
3.2.1. Mass transfer
The transport of chemical species lies at the heart
of physiology and, to a large extent, determines
tissue structure [37,53]. Cells communicate with
each other via a combination of diffusion and
convective flow, which are in turn driven by hydrodynamic, concentration and osmotic gradients. In
vivo, mass transfer to chondrocytes within cartilage
is thought to involve diffusion in conjunction with
convective transport by a ‘physiological pump’, i.e.
the fluid flow that accompanies tissue loading and
unloading [61]. Mass transfer has been shown to
limit the size and determine the composition of in
vitro-grown chondrocyte–PGA constructs in which
cell proliferation increases the mass transfer requirements whereas the accumulation of tissue components decreases the porosity. In particular, the diffusional permeability of constructs decreased dramatically (e.g. to 3% of initial levels over four weeks)
and in proportion to the amount of deposited ECM
[13,28].
In mixed cultures, fluid motion generates dynamic
shear at the construct surface and can be expected to
simultaneously increase mass transfer and hydrodynamically stimulate the cells (see Section 3.2.2).
Constructs grown in orbitally mixed Petri dishes
were larger and contained more ECM than constructs
grown in static dishes [27,86]. When large chondrocyte–PGA constructs were first grown in spinner
flasks and then transferred to static flasks, a necrotic
center developed, whereas when similar constructs
were first grown statically and then transferred to
spinner flasks, ECM deposition increased markedly
(Vunjak-Novakovic, unpublished data). Cultivation
conditions present in mixed and static flasks were
found to be aerobic and anaerobic, respectively, as
assessed by lactate-to-glucose ratios and ammonia
production rates [60]. The addition of insulin-like
growth factor I (IGF-I) to the culture medium
increased the size and improved the composition of
chondrocyte–PGA constructs and the effects of the
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
growth factor were further enhanced when constructs
were grown in rotating vessels instead of static and
mixed flasks [35]. Hydrodynamic factors and IGF-I
thus acted synergistically, which might be explained
by bioreactor-enhanced transport of the growth factor
to the construct surface.
In vivo, most cells are no more than 100 mm from
the nearest capillary [82] that serves to supply
oxygen and nutrients, remove waste products and
transport biochemical signals. Capillaries provide an
effective means of mass transfer because their small
diameter, of approximately 6–8 mm, ensures that the
residence time of the blood is greater than or equal to
the radial diffusion time of the chemical species
within the tissue [92]. Vascularization of in vitro
cultured organized cell communities has not yet been
achieved, to the best of our knowledge. Tissues that
are normally vascularized in vivo (e.g. bone, muscle)
are thus even more likely to be mass transfer limited
during in vitro cultivation than is the normally
avascular cartilage. As described above, the respective thicknesses of engineered cartilaginous, bonelike and cardiac-like tissues grown in vitro to date
were 5.0, 0.5 and 0.18 mm (Table 1). It is likely that
these thicknesses are determined by metabolic rates
of the component cells in conjunction with the
permeability of the construct.
3.2.2. Mechanical factors
The form of any tissue may be thought to represent a diagram of underlying forces transmitted
across the ECM to the individual cells [77]. In vivo,
mechanical forces arise from diverse sources, including dynamic compression of cartilage, interstitial
fluid flow in bone, and tension in muscles. Cartilage
in load-bearing regions of a joint contains the highest
amount of GAG, an essential component for compressive stiffness [71]. Cartilage explants responded
to physiological levels of dynamic compression (i.e.
amplitudes of 1–5% and frequencies of 0.01–1 Hz)
with increased rates of GAG and protein synthesis
[63,67]. Likewise, bone actively remodels in response to loading such that its structure and mechanical properties are governed by the distribution
of stress [18], and muscles grow and hypertrophy in
response to passive stretch during development and
to active tension later in life [49]. The mechanism by
which bone responds to mechanical loading has been
23
proposed to involve the following steps, which might
be extended to other tissues: (1) mechanocoupling,
e.g. mechanical forces induce cell deformation, (2)
mechanotransduction, e.g. cytoskeletal alterations
affect gene expression, (3) signal transduction, e.g.
gene expression affects protein synthesis and (4)
overall response, e.g. the cell number and amount of
ECM increases and the tissue grows [18].
In vitro, intermittent forces due to the motion of
the medium in roller bottles stimulated chondrocytes
to form cartilaginous nodules [50,51], cyclic hydrostatic pressure (5 MPa, 0.25 Hz) enhanced GAG
synthesis in cartilage explants [63], and fluid shear
enhanced PG size and synthesis rate in chondrocyte
monolayers [72]. Flow-induced shears (up to 1 dyn /
cm 2 ) applied to osteoblasts cultured on macroporous
collagen beads using a fluidized bed reactor increased mineralization as compared to static cultures
[42]. It was thus suggested that hydrodynamic forces
can stimulate cells via pressure fluctuations, stretching of the cell membrane and / or shear stress [6].
We have used several culture systems (Fig. 1) to
study the combined effects of hydrodynamic forces
and increased mass transfer on the formation of
engineered cartilage [31]. In the spinner flask system,
constructs fixed to needles were subjected to turbulent mixing. In particular, a magnetic stir bar generated unidirectional fluid motion and spatially
nonuniform distributions of fluid velocity, pressure
and shear. Fluid motion was characterized by isotropic turbulence [86] at an average intensity level
below that previously reported to cause cell damage
[17]. Compared to constructs grown in orbitally
mixed Petri dishes, constructs grown in spinner
flasks were larger and contained higher amounts of
tissue components, especially collagen [86]. However, constructs grown in mixed dishes and in
spinner flasks formed fibrous outer capsules that
were approximately 300 mm thick after 6 to 8 weeks
of cultivation, and might be attributed to the effects
of mechanical forces. In particular, cells exposed to
external forces tend to flatten and activate stress–
protection mechanisms in order to remain firmly
attached to their substrate [22] and increase their
stiffnesses by cytoskeletal rearrangements [89].
The use of rotating vessels yielded constructs with
a continuously cartilaginous, mechanically functional
ECM containing GAG and type II collagen [33,34].
24
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
In this bioreactor, constructs were exposed to a
laminar flow field by adjusting the vessel rotation
speed such that the constructs were maintained in a
state of continual free-fall during cultivation [30].
Flow-visualization studies showed that fluid mixing
in rotating vessels was generated by gravitational
construct settling and the associated oscillations and
tumbling [31].
3.2.3. A representative bioreactor
Some advantages of using the rotating bioreactor
for the culture of organized cell communities are
shown in Fig. 3 and Table 2. In particular, engineered cartilage (i.e. constructs made using bovine
articular chondrocytes and 5 mm diameter 3 2 mm
thick PGA scaffolds) and natural cartilage (equally
sized bovine articular cartilage explants) were cultivated either statically or in rotating vessels for six
weeks. Constructs grown in rotating vessels consisted of a cartilaginous tissue matrix with only a few
cell layers at the surface, while constructs grown
statically contained more ECM peripherally than
centrally (Fig. 3). Compared to constructs grown
statically, constructs grown in rotating vessels were
larger, contained more cells, GAG and total collagen
(Table 2), and had better biomechanical properties
(i.e. higher stiffness during radially confined compression, lower dynamic permeability; VunjakNovakovic, unpublished data). Similar advantages of
rotating bioreactors over static cultures were observed the same in studies of cartilage explants and
chondrocyte–PGA constructs. These results imply
that the structure and function of both natural and
engineered cartilage can be modulated by flow and
mixing during in vitro cultivation. In particular, an
appropriately designed bioreactor can both enhance
mass transfer and provide hydrodynamic stimulation
at the tissue–fluid interface, e.g. by fluctuations in
fluid velocity and stresses.
4. Conclusion and future research directions
Over the last ten years, much progress has been
made in the in vitro culture of organized cell
communities for potential use as implants and / or
studies of normal and pathological tissue function.
This progress is due both to an improved understanding of in vivo tissue growth and the use of 3D
scaffolds in conjunction with bioreactors to regenerate tissue equivalents from isolated cells.
We considered organized cell communities to be
in vitro-grown 3D constructs that displayed important structural and functional characteristics of natural tissues. However, specific requirements for an
organized cell community, i.e. to what extent must it
resemble a natural tissue, remain to be defined. In
general, engineered tissues for clinical use should: (a)
resemble natural tissues structurally and functionally,
(b) be available in a variety of sizes and shapes, (c)
continue to develop following in vivo implantation
and (d) completely integrate with the surrounding
host tissue. Functional requirements will depend on
the particular tissue, e.g. load-bearing for cartilage
and bone, and contractility for skeletal and cardiac
muscle. Some of the basic and practical problems
that need to be addressed for the optimization of in
vitro-cultured organized cell communities are summarized below.
4.1. The cells
A cell cultured in vitro will tend to retain its
differentiated phenotype under conditions that resemble its natural in vivo microenvironment [90]. In
particular, the combination of high cell density and
an appropriate substratum are needed to induce
cooperative cell–cell and cell–matrix interactions
[39,90]. In the case of cartilage tissue engineering,
possible cell sources include mature chondrocytes
[8,70] and BMSCs [54,65]. The potential use of
BMSCs for autologous cartilage implants has several
advantages: (a) a marrow aspirate is easier to obtain
than a cartilage biopsy, (b) BMSCs are quickly
amplified in monolayers and dedifferentiation is not
an issue, while chondrocytes proliferate slowly and
tend to dedifferentiate, (c) the mitotic potential of
BMSCs remains high while that of chondrocytes
decreases with the age of the donor and (d) in
contrast to chondrocytes, BMSCs can potentially be
used to engineer cartilage–bone composites for the
repair of defects extending from the articular surface
into the underlying bone [54,88].
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
Fig. 3. Photomicrographs of chondrocyte–PGA constructs cultured for six weeks in either static flasks or rotating vessels.
25
26
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
Table 2
Cultivation of cartilage constructs and explants in static flasks or rotating vessels a
Group cultivation vessel
Constructs
Not applicable
Explants
Spinner flask
Static flask
Rotating vessel
Static flask
Rotating vessel
Mixing mechanism
Mass transfer rate
Fluid shear at tissue surfaces
Convection
High turbulent
Diffusion
Low
None
Convection
High
laminar
dynamic
Diffusion
Low
None
Convection
High
laminar
dynamic
Cultivation time:
3 days
6 weeks
6 weeks
0 (fresh cartilage)
6 weeks
6 weeks
Dry weight (mg)
Wet weight (mg)
Cells (mg)
Glycosaminoglycan (mg)
Total collagen (mg)
5.960.9
78.2613.8
0.760.33
0.660.1
0.460.1
14.660.7
197.169.2
0.9760.01
5.360.6
2.860.2
24.762.1
237.1620.7
1.4460.20
11.261.4
8.260.7
8.060.6
54.167.6
0.4860.10
3.360.7
3.961.4
19.462.7
166.6617.2
0.8960.16
4.260.3
8.260.8
34.860.2
225.3632.0
0.7260.09
15.360.5
16.764.9
a
Data represent the average6standard deviation of six independent measurements.
4.2. The scaffold
The in vitro growth of large organized cell communities requires optimization of scaffold seeding
methods with respect to: (a) yield, to maximize cell
utilization, (b) kinetic rate, to minimize the time in
suspension for anchorage-dependent and shear-sensitive cells and (c) uniformity of cell distribution, to
permit spatially uniform tissue regeneration. These
requirements can be met using a highly porous
scaffold with large pores in conjunction with a
bioreactor that provides convective transport of the
suspended cells to the scaffold interior. In addition,
controlled biodegradation of the scaffold is desired to
match the rate of tissue growth and to provide
long-term biocompatibility in vivo. The scaffold
shown in Fig. 2 was specifically designed for cartilage tissue engineering; it is likely that further
modifications of the surface (e.g. the addition of
specific functional groups into the polymer backbone), 3D structure (e.g. size and orientation of
polymer fibers) and mechanical properties (e.g. elasticity, stiffness) will be required for other tissue
types. For example, scaffolds containing specific
cellular recognition molecules are being developed
for liver and nerve tissue engineering [69].
4.3. The in vitro culture environment
In cultures of organized cell communities, the
entire surface of a growing tissue should be exposed
to well-mixed medium in order to minimize diffusional constraints. Mixing both maintains a uniform
concentration of chemical species (e.g. gases, nutrients) in the bulk phase and increases the mass
transfer rate at the construct surface. These mixing
requirements can be met using various bioreactors
(e.g. spinner flasks, rotating vessels), which have
been shown to improve the structure, composition
and function of engineered cartilage [29,31,34]. The
biochemical and mechanical factors required during
in vitro cultivation are expected to depend on the
tissue type and its developmental stage. For example,
the addition of specific factors (e.g. growth factors,
hormones, metabolites) during cell expansion and
tissue cultivation induced BMSCs to form either
cartilaginous or bone-like tissue [54].
In general, a structural and functional tissue
equivalent can be regenerated in vitro only if the
bioreactor provides a balanced tissue culture environment with the appropriate cues for maintenance of
specific cell functions. A perfused bioreactor system
is expected to have several advantages over batchoperated bioreactors, including maintaining the concentrations of chemical species at desired levels.
Such steady-state conditions more closely approximate in vivo cell and tissue homeostasis than the
step-changes in medium composition that occur
during the operation of fed-batch bioreactors. The
use of automated control systems, which might
include biosensors to trigger appropriate increases in
the supply rates of medium and gas components,
L.E. Freed, G. Vunjak-Novakovic / Advanced Drug Delivery Reviews 33 (1998) 15 – 30
may further stabilize the bioreactor microenvironment. Other ongoing work involves the correlation of
hydrodynamic parameters obtained using numerical
fluid structure analysis [73] with cell- and tissuelevel responses in engineered cartilage. Such analysis
is needed to test the hypothesis that hydrodynamic
fluctuations in pressure and velocity enhance the
growth of engineered cartilage in a manner similar to
that previously reported during dynamic mechanical
loading of natural cartilage explants [63,67].
In summary, organized cell communities can
potentially serve either as tissue equivalents for
clinical transplantation or as an in vitro model
system for direct testing of cell- and tissue-level
responses to molecular, mechanical or genetic manipulations. We described organized cell communities of cartilage, bone, skeletal muscle and
cardiac muscle that were grown in vitro using
isolated cells, polymer scaffolds and bioreactors. At
this time, our ability to quantitatively assess the
functional behavior of engineered tissues and our
understanding of the mechanisms underlying in vitro
tissue formation are at an early stage and are based
largely on empirical data, intuitive approaches and
qualitative descriptions [37]. However, the increasing
use of mathematical models to describe the individual components of tissue engineering systems
(e.g. cell–ECM interactions, mechanical stimulation
of cell function, mass transfer of chemical species,
tissue biomechanics) is expected to help in the
optimization of cultivation parameters, the development of structure–function correlations and in the
design of further hypothesis-driven experiments.
[3]
[4]
[5]
[6]
[7]
[8]
[9]
[10]
[11]
[12]
[13]
[14]
Acknowledgments
This work was supported by the National
Aeronautics and Space Administration (Grant No.
NAG9-836).
[15]
[16]
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