Fisher John P. e.a. (ed.) Tissue Engineering

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27-4 Tissue Engineering

(b)

10 msec

100 msec

1 sec

Ion channel

current

Cell

action potential

Tissue

anisotropic propagation

Heart ECG

Heart

Cardiac bundles and sheets

5 cm

Anisotropic cell network

100 mm

Intercalated disk

2 mm

Gap junction plaque

50 nm

5 nm

Cx-4 3

(a)

Gap junction

channel

FIGURE 27.1 Levels of anatomical and electrophysiological organization in cardiac muscle. (a) Intercalated disk is

specialized end-to-end connection between cardiac cells. Gap junction plaque is shown in cross-section and en face.

Cx-43 is gap junction protein connexin-43. Note that structural complexity in heart spans many orders of magnitude

from nanometer-size scale in single channels to centimeter-size scale in the heart. (b) Time constants of electro-

physiological function in cardiac muscle range from nanoseconds for a single channel gating to seconds for heart

beats. L and T denote longitudinal and transverse direction, respectively. Pulse sign denotes site of stimulus.

in the heart [59]. In addition, presence of noncontractile scar in heart milieu can cause locally increased

stress gradients, which through mechano-electric feedback may yield in stretch-induced arrhythmias [60].

27.3 Current State of Cardiac Tissue Engineering

Over the last several years different strategies have been developed to design engineered cardiac tissues

that could be used for pharmacological, genetic, and functional studies in vitro and possible implantation

in vivo, as outlined in Table 27.1. These studies have shown that structure and function of cardiac tissue

constructs depend on the animal species used for the cell dissociation [61–63], composition of seeded

cells [14,64], initial cell seeding density [62,63,65,66], scaffold characteristics [40,67–70], composition

of culture medium [69], type of bioreactor [63,65,69], and applied physical forces [61,71]. Most of these

results are based on the evaluation of general histology, and assessment of cellular properties including cell

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Cardiac Tissue Engineering 27-5

TABLE 27.1 Cardiac Tissue Engineering In Vitro and In Vivo

In vitro

Cell source Scaffold Bioreactor Assessment References

Neonatal rat

ventricle

Microcarrier

beads

HARV Immunohistology, ultrastructure,

pharmacology

[77,78]

Embryonic chick

ventricle

Planar collagen gel

with

supplements

Static petri dish

attachment to velcro

Immunohistology,

pharmacology, ultrastructure,

gene manipulation, mechanical

contractile force

[79]

Neonatal rat

ventricle

planar collagen gel

with

supplements

Static petri dish

attachment to velcro

Immunohistology,

pharmacology, ultrastructure,

contractile force

[62]

Neonatal rat

ventricle

Collagen gel ring

with

supplements

Static petri dish and

cyclic stretch

Immunohistology,

pharmacology, ultrastructure,

contractile force

[72]

Rat smooth

muscle, skin

ﬁbroblasts, fetal

ventricle, human

atria and

ventricle

Rectangular

gelatin mesh

Static petri dish Histology, cell proliferation [86]

Young human

ventricle

Rectangular

gelatin mesh

Cyclic stretch in dish Histology, proliferation,

mechanical

[71]

Neonatal rat

ventricle

Rectangular

collagen scaffold

(“tissue ﬂeece”)

Static petri dish RT-PCR, pharmacology,

ultrastructure, mechanical

[73,87]

Neonatal rat

ventricle

Fibrin glue and

thick collagen gel

around aorta

Unperfused and

perfused through

aorta

FDG-PET, Immunohistology [88]

Neonatal rat

ventricle

Cross-linked

collagen mesh

HARV Immunohistology, ultrastructure [89]

Fetal and neonatal

rat ventricle

Electrospun

tubular collagen

scaffold

HARV Immunohistology, ultrastructure

mechanical stress–strain curves

[74,90]

Neonatal rat

ventricle

No scaffold Static petri dish Immunohistology, ultrastructure,

electrical connectivity,

mechanical, subcutaneous

implantation

[75,92]

Embryonic chick

and neonatal rat

ventricle

Fibrous PGA disk Static petri dish,

spinner ﬂask, HARV

Viability, metabolic activity,

immunohistology,

ultrastructure

[63]

Neonatal rat

ventricle

Fibrous PGA disk Spinner ﬂask Viability, metabolic activity,

immunohistology,

ultrastructure, tissue scale

electrophysiology

[64]

Neonatal rat

ventricle

Fibrous PGA disk Perfusion cartridge Viability, metabolic activity,

immunohistology,

ultrastructure

[94,95]

Neonatal rat

ventricle C2C12

myoblasts

collagen sponge

disk with

matrigel

Orbitally mixed dish,

Perfusion cartridge

Viability, metabolic activity,

immunohistology,

pharmacology, excitation

threshold, capture rates

[65,96]

Neonatal rat

ventricle

Surface

hydrolyzed,

laminin-coated

PGA disk

Spinner ﬂask,

3D gyrator, HARV

Immunohistology,

immunoblotting, ultrastructure,

viability, metabolic activity,

tissue electrophysiology

[69]

Continued

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27-6 Tissue Engineering

TABLE 27.1 Continued

In vitro

Cell source Scaffold Bioreactor Assessment References

Neonatal rat

ventricle

Surface hydrolyzed,

laminin-coated

PGA disk

HARV Immunohistology,

immunoblotting,

pharmacology, cell

electrophysiology

[76]

Neonatal rat

ventricle

Fibronectin coated

PLGA disk

HARV Histology, ultrastructure,

optical mapping of

action potentials

[121,122]

Fetal rat ventricle Alginate disk Static petri dish Viability, metabolic

activity, histology

[66]

In vivo implantation in heart

Postoperative

Cell source Scaffold Site of implantation assessment References

Fetal rat ventricle Gelatin mesh cube Infarct in cryoinjured

left rat ventricle

Histology, ultrastructure,

ventricular pressure in

Langendorf preparation

[97]

Rat aortic smooth

muscle

Gelatin mesh, PTFE

patch, PGA, and

PCLA sponge

Defect in right

outﬂow ventricular

tractinrat

Immunostaining, cell

proliferation,

morphometry

[67,98]

Rat aortic smooth

muscle

PCLA sponge Postinfarct aneurysm

in rat left ventricular

Immunohistology,

echocardiography,

ventricular pressure

[99]

Fetal rat ventricle Alginate disk Rat coronary

occlusion site

Immunohistology,

echocardiography

[100]

Neonatal rat

ventricle

Collagen gel ring with

supplements

Perimeter of healthy

rat ventricle

Immunohistology,

ultrastructure,

pharmacology,

echocardiography

[85]

HARV — high-aspect-ratio-vessel, HFDG-PET — Fluor-Deoxy-Glucose-Positron-Emission-Tomography, PLGA —

poly(lactic-co-glycolic) acid, PTFE — polytetraﬂuoroethylene, PCLA — ε-caprolactone-co-l-lactide reinforced with knitted

poly-l-lactide fabric.

number, viability, metabolic activity, expression of cardiac-speciﬁc proteins, and ultrastructural features.

Few groups also focused on measurements of contractile force at tissue scale [62,72–75], while only

one group has studied in detail microscopic and macroscopic electrical properties of tissue constructs

[64,69,76]. The following paragraphs will give an overview of existing in vitro and in vivo efforts in the

emerging ﬁeld of cardiac tissue engineering.

27.3.1 Cardiogenesis In Vitro

Akins et al. [77,78] have shown that neonatal rat ventricular myocytes can form multilayered inter-

connected structures when cultivated on ﬁbronectin-coated polystyrene beads or collagen ﬁbers inside

high-aspect-ratio-vessel (HARV) bioreactors. After 6 days in culture, cardiac cells formed small, several

layers thick clusters in the regions between the beads, exhibited presence of sarcomeres and gap junctions,

and rhythmically contracted at rates that were slower in the presence of propranolol. The nonmyocytes

were distributed throughout the tissue clusters, with most of the endothelial cells lining on the interface

between the cluster and culture medium.

Group of Eschenhagen has done some of the most comprehensive work in the ﬁeld, using mixtures of

embryonic chick [79] or neonatal rat cardiac cells [62,72] and gels made of collagen type I supplemented

with matrigel, chick embryo extract, and horse serum. Their initial work was based on Vandenburgh’s

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Cardiac Tissue Engineering 27-7

approach for engineering of skeletal muscle [80], where cell–gel mixture was cast in the thin planar

geometry between two parallel Velcro-coated glass tubes. Firm attachment to Velcro-imposed static stress

on free edges of the gel resulting in thin biconcave tissue construct (8×15×0.18 mm

) with loose, aligned

cardiac cell network formed mostly along the construct edges [79]. The alignment and density of this net-

work was improved by use of the chronic cyclic stretch during cultivation [61]. In their current approach

[72], cardiac constructs termed engineered heart tissues (EHTs) are made by embedding neonatal rat

ventricular cells in circularly molded collagen gels, which are subsequently cultivated in static conditions

for 7 days and subjected to chronic cyclic stretch (10%, 2 Hz) for additional 7 days. Resulting submillimeter

thick rings of tissue contain aligned cardiomyocytes organized in loose but uniform tissue-like network

with frequently forming 20 to 50 µm thick cardiac ﬁbers [72]. Myocytes in this network spontaneously

contract at steady rates of ∼2 Hz, and exhibit differentiated cardiac-speciﬁc ultrastructure including par-

allel sarcomeres, T-tubules, SR vesicles, formed dyads, and basement membrane [72]. The initial seeding

of unpuriﬁed cell mixture (no differential preplating) result in the presence of microphages and abundant

ﬁbroblasts, scattered throughout the EHT, as well as endothelial and smooth muscle cells, packed more

densely in the outer compared to inner region. When electrically and pharmacologically stimulated, EHTs

exhibit cardiac-speciﬁc mechanical properties including Frank–Starling behavior, a positive inotropic

response to extracellular calcium and isoprenaline, and negative inotropic effect to carbachol. Although

recorded twitch amplitudes of 1 to 2 mN/mm

are an order of magnitude lower than those found in native

cardiac tissues [81], the twitch to resting tension ratio is larger than 1, similar to native muscle. The use of

rat cells, horse serum, chick embryo extract, matrigel, and unpuriﬁed cell seeding mixture are all found

to increase the maximum developed force and mechanical integrity of EHTs, while increase in collagen

content seems to decrease twitch tension [14,70]. Up to now, EHTs have been used for studying the effect

of genetic and pharmacological manipulations on cardiac contractile function [62,82–84], and were also

implanted in vivo (see work by Zimmermann et al. [85]).

Group of Li [86] seeded biodegradable gelatin meshes with different cell types including stomach

smooth muscle cells, skin ﬁbroblasts and fetal ventricular myocytes from rat, and adult atrial and

ventricular myocytes from humans. Rat cells and human atrial, but not ventricular, cells proliferated

over 3 to 4 weeks in culture. All cells migrated in a 300 to 500 µm thick outside layer of gelatin scaffold,

which slowly degraded with the highest degradation rate found in the presence of ﬁbroblasts. In separate

in vitro study [71], the same group showed that 2 weeks of cyclic mechanical stretch improved cell prolif-

eration, distribution, and mechanical strength of tissue constructs made using gelatin scaffolds and heart

cells isolated from children who underwent repair of Tetralogy of Fallot.

Koﬁdis et al. [73,87], used 20 × 15 × 2mm

commercially available collagen-based scaffolds (“tissue

ﬂeece”) that were inoculated with neonatal rat cardiac cells and cultured in petri dishes. The randomly

distributed cells formed sparse synchronously contractile networks, and exhibited cardiac speciﬁc mech-

anical responses to stretch, extracellular calcium, and epinephrine. In an attempt to increase the thickness

of the engineered cardiac tissue, the same group recently embedded a rat aorta in the 8.5 mm thick mixture

of collagen gel and cardiac cells, and used pulsatile ﬂow through the aorta for 2 weeks as a vehicle for

nutrition and oxygen delivery [88]. The aorta remained patent throughout the culture and viability was

increased compared to unperfused controls.

van Luyn et al. [89] have also used neonatal rat cells and commercially available cross-linked collagen I

bovine matrices, and cultured them in HARV bioreactors for up to 3 weeks. Spatially scattered cells

exhibited immature sarcomeres, gap junctions, and stained for troponin-T.

In recent studies, Evans et al. [90] and Yost et al. [74] cultured embryonic and neonatal rat cardiac

cells on ﬁbronectin coated aligned tubular scaffolds (15 mm long, 4 mm inner, 5 mm outer diameter)

made from extruded collagen I ﬁbers [91]. After 3 to 6 weeks in HARV bioreactors, cardiac cells aligned,

contracted spontaneously, formed few interconnected cell layers (with total thickness of ∼20 µm) on

the inside and outside lumen of the tube, and exhibited registered sarcomeres and randomly distributed

gap junctions. Tubular collagen scaffolds exhibited viscoelastic properties qualitatively resembling those

of native papillary muscle [74] only when seeded with cardiac cells, as inferred from the shape of the

stress–strain hysteresis loops.

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27-8 Tissue Engineering

Very elegant studies by Shimizu et al. [75,92,93] have demonstrated that cardiac cells can form 3D

multilayer tissue-like structures without the use of any type of scaffold. Isotropic monolayers of puri-

ﬁed cardiac cells were cultured to conﬂuence on the surfaces made of temperature responsive polymer

poly(N-isopropylacrylamide). This polymer is slightly hydrophobic and cell adhesiveat 37

◦

C and becomes

hydrophilic and cell repellent when cooled below 32

◦

C. After 4 days, up to four cardiac sheets were detached

(together with secreted extracellular matrix) from polymer surface by cooling to 20

◦

C and overlaid using

pipette or polyethylene mesh. Overlaid sheets exhibited uniform gap junction distribution, connected

electrically, and formed compact multilayered spontaneously contractile cardiac constructs with area

of1cm

and thickness of up to 50 µm. After subcutaneous implantation, cardiac constructs survived

up to 12 weeks, spontaneously contracted, appeared vascularized and at 3 weeks exhibited twitch tension

of 1.2 mN [75].

Group of Freed and Vunjak-Novakovic has utilized various approaches to engineering of cardiac tissue

based on the use of biodegradable polymer scaffolds and different tissue culture bioreactors. Initial studies

of Bursac et al. [64], and Carrier et al. [63] have demonstrated that cardiac cells formed tissue-like con-

structs when seeded on 5 mm diameter×2 mm thick ﬁbrous poly(glycolic) acid (PGA) disks inside spinner

ﬂask bioreactors. Cells in the outer 50 to 70 µm thick region were randomly oriented and connected in

the relatively dense multilayer network. These cells expressed cardiac speciﬁc proteins (α-sarcomeric actin

and troponin-T), end ultrastructural features characteristic of cardiac myocytes (parallel sarcomeres, all

types of specialized junctions, dense mitochondria, glycogen granules). The cells in the interior of these

tissue constructs were sparsely distributed and often necrotic. Spontaneous macroscopic contractions

were observed at days 2 to 4 of culture and generally ceased thereafter, with occasional activity at rates of

less then 1 Hz on culture day 7. Action potential propagation and electrical excitability were studied using

linear array of eight metal microelectrodes with 500 µm spatial resolution (Figure 27.2a). Constructs were

electrically excitable and exhibited isotropic, macroscopically continuous electrical propagation with velo-

cities as high as 60% of those found in native ventricles [64]. Use of puriﬁed cell mixture (after differential

preplating) for seeding resulted in superior electrophysiological properties including higher velocity of

propagation, increased maximum rates of tissue capture and lower excitation threshold compared with

use of unpuriﬁed (no preplating) cell mixture. In addition, use of neonatal vs. chick cardiac cells, dynamic

seeding and cultivation in bioreactors vs. static petri dishes, and increase in the number of seeded cells up

to 8×10

cells per scaffold have all improved cell packing density, metabolic activity, and electrophysiolo-

gical properties of constructs [63,64]. In further studies, Carrier et al. [94] looked in the use of perfusion

through tissue construct as means to improve the cellularity and tissue architecture, and studied effect of

oxygen deprivation on engineered cardiac muscle [95]. In the most recent studies from the same group,

Radisic et al. [65,96] used cell–matrigel mixture to densely inoculate cardiac cells inside collagen sponge

scaffolds, and employed similar perfusion bioreactor for construct cultivation. The viability, metabolic

activity, and cellular density through ∼1 mm thick region were higher than in constructs cultured in

orbital shakers and those from studies by Carrier et al. In a different study, Papadaki et al. [69] have

shown signiﬁcant improvements in structure and function of cardiac constructs when PGA scaffolds were

hydrophilized and coated with laminin, percentage of serum in culture medium reduced after 2 days

of cultivation, constructs seeded with concentrated cell suspension in rotating gyrators, and cultivation

performed in HARV bioreactors. Compared to previous studies, tissue constructs exhibited better cell

viability yielding thicker (120 to 160 µm) cardiac-like outer region, and higher cellular expression of dif-

ferentiation marker proteins including creatine kinase-MM (involved in metabolism), sarcomeric myosin

heavy chain (involved in contractile function), and gap junction protein Connexin-43. Tissue scale elec-

trical properties approached those found in native muscle with conduction velocity at basic stimulation

rate of 1 Hz and maximum capture rate reaching 90 and 70% of those found in donor neonatal ventricles,

respectively (Figure 27.2a). Macroscopic electrical propagation was effectively isotropic due to random

cell orientation and uniform gap junction distribution. Further electrophysiological and pharmacological

studies at microscopic scale by Bursac et al. [76] demonstrated that action potentials in cardiac cells from

7-day constructs were comparable to those in 2-day old donor ventricles with respect to depolarization

upstroke, amplitude, and resting potential. The major difference was prolonged action potential duration

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Cardiac Tissue Engineering 27-9

(b)

Tissue

construct

Neonatal

ventricle

Cardiac action potential

20 mV

100 ms

1mm

Tissue sample

Tyrode’s

T =30°C

Cell scale recording

Tissue scale recording

–

1mm

Tissue sample

pH = 7.32

T =37°C

1 2

3 4 5 6

(a)

Electrical impulse propagation

030 300

Tissue construct

Neonatal ventricle

Time (msec)

FIGURE 27.2 Tissue and cell scale electrophysiological recordings in 7-day old cardiac tissue constructs and 2-day

old neonatal ventricles. (a) Custom-built linear array of two stimulating and eight recording microelectrodes (only six

are shown) was used for assessment of macroscopic impulse propagation. S and R in two tracings denote stimulus

artifact and responses, respectively. Note relatively smooth biphasic shapes of recorded extracellular waveforms in

constructs and ventricles. The response amplitude in constructs is an order of magnitude lower than in ventricles due

to smaller number of cardiac cell layers present. Propagation velocities in constructs and ventricles are comparable

(i.e., times for propagation from electrode 1 to 6 are similar). (b) Glass capillary microelectrodes are used for action

potential recordings from single cells in the tissue sample. Note fast upstroke and similar resting potential, but longer

action potential in constructs than in ventricles. S and R denote stimulus artifact and response, respectively.

(Figure 27.2b) and absence of early repolarization notch due to downregulation of transient outward

potassium current. In addition, cardiac cells cultured in 3D tissue constructs maintained more differen-

tiated phenotype (higher expression of marker proteins) and more in-vivo like action potential features

compared to those cultured in 2D monolayers under similar cultivation conditions.

27.3.2 In Vivo Implantation for Cardiac Repair

By April 2004, in vivo implantation of engineered cardiac tissue in infracted heart was attempted by only

three groups.

Li et al. [97] cultured fetal rat myocytes on biodegradable gelatin meshes (15 ×15 ×5mm

)for7days

and implanted cardiac constructs over the scar area in cryoinjured syngeneic rat hearts. Cells populated

sparse interstices of gelatin meshes (see in vitro work by Li’s group) and continued to proliferate and

spontaneously contract in vitro for at least 26 days. Epicardially implanted grafts survived for 5 weeks,

exhibited increased cellularity, slight degradation, and moderate degree of vascularization. Left ventricu-

lar developed pressure, showed no improvement over the control animals. In other studies [67,98], the

same group evaluated use of various scaffold materials seeded with aortic smooth muscle cells (used to

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27-10 Tissue Engineering

presumably increase elasticity of the patch) for repair of defect in the right ventricular outﬂow tract in

syngeneic rats. Eight weeks postimplantation constructs made of ε-caprolactone-co-l-lactide reinforced

with knitted poly-l-lactide fabric (PCLA) outperformed those made of gelatin, PGA, and polytetraﬂuoro-

ethylene (PTFE) with respect to cellularity, elastin content, and preserved thickness. In the next study

[99], grafts made of PCLA and smooth muscle cells were used to repair left ventricular aneurysm in the

rat hearts after transmural infraction. Cell-seeded grafts reduced abnormal chamber distensibility and

improved ventricular function compared with implanted cell-free grafts, as assessed by echocardiography

and constant pressure measurements in Langendorff preparation.

Leor et al. [100] cultured fetal rat myocytes on porous biodegradable alginate disks (6 mm diameter,

1 mm thick) inside 96-well plates for 4 days, and implanted cardiac tissue constructs over infracted region

in rat hearts 7 days after permanent occlusion of left main coronary artery. Nine weeks postimplant-

ation, cardiac constructs survived, while alginate scaffold substantially degraded. Cardiac grafts were

neovascularized, contained inﬁltrated macrophages and lymphocytes due to use of allogenic cells and no

immunosuppression, and exhibited small number of sparsely distributed cardiac cells presumably due

to low initial seeding density. Echocardiography revealed attenuated left ventricular dilatation and main-

tained contractile function, although it was not clear if implanted cell-free scaffolds would have produced

similar results. Further in vitro study from the same group [66] focused on methods to increase cell density

in alginate scaffolds by applying moderate centrifugal forces during seeding.

Zimmermann et al. [85] implanted 12-day old ring-shaped EHTs (see in vitro work from Eschehagen’s

group) around the circumference of healthy syngeneic rat hearts. Two weeks after implantation, EHTs

were vascularized, innervated, expressed differentiatedcardiac phenotype, and did not alter left ventricular

function compared to preoperative state, as assessed by echocardiography. Spontaneous contractions were

preserved in vivo, but no intercellular coupling of EHTs and host tissue could be demonstrated. Despite the

syngeneic approach, EHTs were completely degraded in the absence of immunosuppression, presumably

due to presence of allogenic components in reconstitution mixture (e.g., matrigel, horse serum, chick

embryo extract).

In all of the described in vivo attempts no electrophysiological studies of engineered patch were done

pre- or postimplantation.

27.4 Design Considerations

Ultimate success of cardiac tissue repair with an implanted cardiac patch depends on thorough under-

standing of the key parameters of tissue design in vitro, and careful deﬁnition of the desired tissue

engineering outcomes.

27.4.1 Cell Source and Immunology

One of the crucial aspects for successful engineering of cardiac tissue is a choice of implanted cells.

Experiences from cellular cardiomyoplasty show that for the improvement of heart systolic function

implanted cells need to be (or be capable of becoming) contractile [26]. Although fetal and neonatal

cardiac cells are clearly shown to functionally incorporate in the myocardium [17–19], they are not cells of

choice due to limited proliferation potential, immunological, and ethical issues. For this reason, their use

will probably stay limited to in vitro model systems and proof-of-concept in vivo studies. Possible “ideal”

cell source may be human embryonic or adult stem cells. Although human embryonic stem cells are shown

to differentiate into cardiac myocytes [101,102], their immunogenic and tumorogenic nature, and low

efﬁciency and speciﬁcity of differentiation (<1% of cells differentiate into mixture of atrial, ventricular,

and nodal cells), as well as ethical issues, mayﬁnally preclude their clinical use. Some hope lies in the nuclear

transfer technology (“therapeutic cloning”) [103], and genetic knock-out of major histocompatibility

complexes [104], which may offer strategies for preventing immune rejection. Autologous adult stem cells

from skeletal muscle, peripheral blood, or bone marrow appear as better choice for cell transplantation

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Cardiac Tissue Engineering 27-11

than embryonic stem cells. For example, autologous skeletal myoblasts are easy to proliferate in vitro

and implant in vivo and cause no immune response, which currently makes them one of the cell types

used in clinical trials [33]. Unfortunately, they do not express gap junction proteins and are still not

shown to functionally couple with host cardiac tissue when implanted [105,106], although this may be

resolved with stable transgene expression of connexin molecules [107]. Mesenchymal or hematopoietic

stem cells derived from bone marrow may represent an ideal cell source [29,108]. However, the efﬁciency

of their transdifferentiation into cardiac myocytes remains controversial in light of recent ﬁndings that

question techniques used to quantify the number of transdifferentated cells in heart [109,110]. Still

their beneﬁcial effect may come from induced neovascularization in implantation sites. Current research

efforts are focused on increase in percentage of embryonic or adult stem cells that commit to cardiac

phenotype by use of different growth factors or media compositions, introduction of early cardiac genes

in undifferentiated stem cells in vitro, or coculture of stem cells with differentiated cardiac myocytes.

Another promising alternative is in vitro genetic reprogramming of differentiated somatic cells (for review

see Reference 111).

27.4.2 Cellular Composition

The heart is composed of different types of cells. Engineering of functional tissue patch requires selection

of the appropriate composition of seeded cell mixture. For example, if an implant were to be localized

in the ventricle, cardiac myocytes that comprise a main fraction of seeded cells should be of ventricular

origin or with ventricular characteristics. The percentage of nonmyocytes in the seeding mixture is a

parameter that can be varied. Eschenhagen’s group found that it was necessary to use higher percentage

of nonmyocytes (no preplating after cell isolation) to improve mechanical integrity and twitch tension of

EHTs [14]. In contrast, scaffold-free constructs by Shimizu et al. [75] developed similar twitch tensions

despite the fact that they were made of puriﬁed cardiac cell mixture. This suggests that the percentage of

“needed” nonmyocytes may actually depend on the presence and type of scaffold. It is possible that in

EHTs, higher number of ﬁbroblasts contributed initial contraction of collagen gel [112], which in turn

increased the proximity and intercellular connectivity between myocytes, yielding increased contractile

force. In addition, Bursac et al. [64] have shown that use of unpuriﬁed cardiac cell mixture decreased

propagation velocity and compromised electrical properties in cardiac constructs. Therefore, for a given

type of scaffold, the percentage of seeded nonmyocytes should be selected to optimize for both mechanical

and electrical function of cardiac constructs.

27.4.3 Tissue Architecture

The anisotropic architecture and dense cell packing of native cardiac tissue impose important design

rules in engineering of functional cardiac patch. For instance, velocity of electrical propagation and

mechanical stiffness in healthy adult human ventricles are on average 2 to 3 times larger in longitudinal

(ﬁber) than in transverse (cross-ﬁber) direction [113,114]. This can vary widely with location in the

heart, age of individual, and heart disease. Therefore, only a cardiac patch with dense 3D network of

elongated and aligned cardiac myocytes that mimic architecture of native tissue can generate desirable

spatio-temporal distribution of electrical and mechanical activity, and result in efﬁcient and safe therapy.

Aligned growth of cardiac cells can be induced by static or dynamic stretch [61,115,116], presence of free

tissue boundaries [117], or by cell guidance with oriented surface topography [49,118,119] and anisotropic

distribution of chemical cues for cell attachment [49,120]. Eschenhagen’s group applied cyclic stretch and

ring geometry creating a sparse network of oriented cardiac cells in the form of a thin (submillimeter

diameter) cardiac cable. Repair of larger injured area in the heart will, however, require engineering of an

anisotropic slab of 3D cardiac-like tissue with controllable shape, size, and geometry.

Bursac et al. [49] have recently shown that anisotropic monolayers of cardiac cells (that mimic longit-

udinal sections of native cardiac tissue) can be designed with highly controllable architecture using surface

microabrasion or micropatterning of extracellular matrix proteins (e.g., ﬁbronectin) (Figure 27.3b). These

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27-12 Tissue Engineering

(b)

Isochrones

2 mm

10 m

Sarcomeric a-actin and DAPI

Fibronectin

Excitation

light source

sample

Photodetectors

Recorded signals

Fiber optic

bundle

Data acquisition

Experimental chamber

(cross-section)

(a)

(c)

Isochrones

2 mm

Scanning electron micrograph

50 m

200 m

Light micrograph

17 mm

FIGURE 27.3 Architecture and impulse propagation in 2D and 3D anisotropic cultures of neonatal rat ventricular

cells. (a) Optical mapping setup. Tissue samples stained with voltage sensitive dye were positioned over 61 hexagonally

arranged ﬁbers and transilluminated with excitation light. Two seconds of optically recorded action potentials from

a cardiac cell monolayer are shown on the right. Gray circles denote active ﬁbers. Square denotes a recording site in

the bundle and corresponding voltage trace (for details see Reference 49). (b) Anisotropic monolayer of cardiac cells.

Cells were cultured on micropatterned lines of ﬁbronectin. On the left, culture is deliberately scratched to expose

patterned lines and cell-deposited ﬁbronectin. Aligned cells exhibit prominent sarcomeres and elongated nuclei

(middle). Elliptical isochrones demonstrate anisotropic propagation (right). ∗ denotes recording sites. Pulse symbol

denotes site of stimulus. Thedegree of anisotropy can be systematically varied by controlling the amount of intercellular

clefts and cell co-alignment [49]. (c) Anisotropic tissue construct. Oriented ﬁbrous architecture in PLGA scaffolds

(left) was accomplished by leaching sucrose from a polymer-coated template made of aligned sucrose ﬁbers [121].

Cardiac cells were aligned in numerous regions along the direction of PLGA ﬁbers (middle). Macroscopic propagation

was anisotropic (albeit moderately), as assessed by optical mapping of transmembrane voltage (right) [122].

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Cardiac Tissue Engineering 27-13

methodologies enabled systematic control over the degree of anisotropy (longitudinal-to-transverse velo-

city anisotropy ratios from 1 to 5.6), ﬁber direction, and amount of longitudinal intercellular clefts in the

2D cardiac monolayers. Optically recorded action potentials with voltage sensitive dye RH237 (at 61 sites

over2cm

area) were used to map propagation of electrical activity and degree of functional anisotropy

(Figure 27.3a). It would be ideal if techniques for 3D cardiac tissue culture could be developedwith a similar

level of architectural control as those used for 2D cell culture. In most recent studies Bursac et al. [121,122]

extruded and baked sucrose to form 3D aligned ﬁbrous templates in an attempt to induce anisotropic

architecture in poly(lactic-co-glycolic) acid (PLGA) scaffold disks (Figure 27.3c). After 2 weeks of culture

in HARV bioreactors, 12 mm diameter, 0.7 mm thick cardiac constructs exhibited regions with aligned

cells, and moderate degrees of functional anisotropy (i.e., longitudinal-to-transverse velocity ratio of

up to 2), as assessed by optical mapping of electrical propagation (Figure 27.3c). Using similar techniques

to align elastomeric polymers [123] instead of PLGA in conjunction with chronic mechanical stimula-

tion may yield 3D cardiac patches with physiological degrees of structural and functional anisotropy. An

alternative approach is the use of electrospun scaffolds [74,91,124] providing that obtained degrees of

porosity can support deeper cell penetration during seeding than achievable by current methodologies.

27.4.4 Tissue Thickness

Another important parameter in the design of a functional cardiac patch is thickness of the tissue con-

struct. Compared to other cells in the body, cardiac myocytes require high oxygen and nutrient supply

due to continuous contractile activity. High metabolic demand in myocardium is sustained by dense

vascularization with average arterial intercapillary distances that range from 15 to 50 µm depending on

the size of the heart and basal heart rate [125,126]. Absence of vasculature in tissue constructs is a limiting

factor for engineering thicker (>200 to 300 µm) cardiac patches with physiological cell packing densities.

Nonetheless, high cell density is necessary for establishment of proper intercellular communication, which

in turn enables efﬁcient generation of mechanical force and fast, electrically safe impulse propagation.

Mixing and perfusion of culture medium, and/or cyclic mechanical stimulation are some of the methods

that can alleviate diffusional limits of oxygen and nutrient supply, but only to a certain extent. Neverthe-

less, thin but dense engineered cardiac tissue is still a good in vitro approximation of a viable portion of

explanted ventricular slice after several hours of superfusion [127], and thus could be used for different

tissue-scale functional studies in vitro. These studies would be more versatile and technically easier than

studies in monolayer cultures (e.g., they would enable variety of force measurements, and yield substan-

tially higher signal-to-noise ratio during extracellular and optical recordings of electrical propagation).

Moreover, implantation of even 10 to 20 well-coupled anisotropic cardiomyocyte layers over the infracted

area may still have a signiﬁcant therapeutic value regarding the facts that after epicardial infarction only

several disarrayed cell layers may survive above the scar, and that increased thickness of survived layers is

directly correlated with decreased incidence of arrhythmias [128]. This is one of the reasons why approach

by Shimizu et al. [75] with scaffold-free cardiac multilayers deserves close attention. On the other hand,

engineering of thicker cardiac slices will depend upon the development of externally perfusable, patent

microcapillary-like networks inside the tissue constructs. Recent work by Vacanti’s group [129] represents

an interesting approach to this problem.

27.4.5 Electrical Function and Safety

One of the most important criteria for successful design of cardiac patch is the issue of electrical and

mechanical safety. It is important to understand that haphazardly adding donor cells or transplanting

a poorly designed tissue patch into an already compromised heart may only increase the likelihood for

aneurisms, tissue rupture, or arrhythmias. In general, the injection of donor cells or implantation of a tissue

patch introduces structural and functional heterogeneity in the cardiac milieu that depends on (1) electro-

mechanical characteristics of donor cells, (2) the density, coupling, and geometrical arrangement of

donor cells within the implant, (3) the degree of interaction between the donor and host cells, and