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Tissue Engineering - John P. Fisher

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26-8

Tissue Engineering

26.5 Bioreactors for Mechanical Conditioning

In vivo, the pulsatile nature of blood flow imposes radial pressure upon the vessel wall, which subjects SMCs within the medial layer to cyclic strain. Thus, a great deal of research has examined SMC behavior in response to cyclic stretch and found such stimuli important in the fabrication of vascular tissue, particularly with respect to ECM synthesis and tissue organization. For example, SMCs seeded on purified elastin membranes and exposed to 2 days of cyclic stretching (10% beyond the resting length) have been shown to incorporate hydroxyproline into protein three to five times more rapidly than stationary controls, indicating increased collagen synthesis in response to strain [Leung et al., 1976]. Cyclic strain also increased the synthesis of collagen types I and III and chondroitin-6-sulfate without stimulating DNA synthesis. Another study also detected enhanced matrix production in collagen constructs seeded with rat aortic SMCs and subjected to 7% cyclic strain [Kim et al., 1999]. Over 20 weeks of culture under cyclic strain, SMCs upregulated expression of elastin and collagen type I. Elastin content from these SMCs increased 49% over unstretched controls. Furthermore, organization of the tissue was observed, as evidenced by perpendicular alignment of SMCs to the direction of the applied strain.

Similar results have been obtained in three-dimensional constructs. Tubular collagen constructs seeded with SMCs were cultured over thin-walled silicone sleeves and subsequently exposed to regulated intraluminal pressures to stretch the vessel in a repeatable fashion for up to 8 days. The 10% cyclic distension in diameter caused the scaffolds to contract, SMCs and bundles of collagen fibers to align circumferentially around the vessel, and improvement of the scaffold’s mechanical properties [Seliktar et al., 2000]. Moreover, this model system was employed to investigate the remodeling capacity of these constructs via the activity of matrix metalloproteinases (MMPs) known to cleave solubilized type I collagen fragments [Seliktar et al., 2001]. Constructs mechanically conditioned for 4 days contained five times higher amounts of MMP-2 compared to static controls and increased MMP-2 activity. The increases in MMP-2 levels correlated favorably with improvements in mechanical strength and material modulus as a result of cyclic strain. When a nonspecific inhibitor of MMP-2 was added to the culture media, MMP-2 levels decreased and mechanical properties were reduced, negating the benefits of mechanical conditioning. These studies indicate that strain-mediated remodeling of collagen scaffolds is essential for improved construct of mechanical properties.

Because of the profound effects of cyclic strain on SMC orientation, ECM production, and tissue organization, preculture of vascular graft constructs in a pulsatile flow bioreactor system may help recreate the natural structure of native vessels and allow one to better achieve the mechanical properties required of the construct. A schematic of a typical pulsatile flow bioreactor system is shown in Figure 26.3. The mechanical stimuli from pulsatile flow could generate the cyclic strain necessary to alter ECM production, thereby creating a histologically organized, functional construct with satisfactory mechanical characteristics for implantation. To develop a blood vessel substitute, Niklason et al. [1999], cultured PGA constructs in a pulsatile blow bioreactor generating 165 beats per minute (bpm) and 5% radial strain. The pulse frequency of this system was chosen to mimic a fetal heart rate, believed to possibly provide optimal conditions for new tissue formation. However, most mechanical conditioning investigations mentioned above conducted strain studies at 60 bpm, more representative of an adult heart rate, with promising outcomes. Therefore, the optimal bioreactor culture conditions for the development of a TEVG remain to be elucidated. Nevertheless, such a system shows promise for the production of a blood vessel substitute with the necessary mechanical and biochemical components.

26.6 Conclusions

In the past couple of decades, a great deal of progress on TEVGs has been made. Still, many challenges remain and are currently being addressed, particularly with regard to the prevention of thrombosis and the improvement of graft mechanical properties. In order to develop a patent TEVG that grossly resembles native tissue, required culture times in most studies exceed 8 weeks. Even with further advances in

Tissue Engineered Vascular Grafts

26-9

 

Compliance

TEVG

 

chamber

 

Pulsatile

Check valves

Graft

 

pump

 

chamber

 

 

5% CO2

Reservoir

FIGURE 26.3 Diagram of a typical pulsatile flow bioreactor for culture of TEVGs.

the field, TEVGs will likely not be used in emergency situations because of the time necessary to allow for cell expansion, ECM production and organization, and attainment of desired mechanical strength. Furthermore, TEVGs will probably require the use of autologous tissue to prevent an immunogenic response, unless advances in immune acceptance render allogenic and xenogenic tissue use feasible. TEVGs have not yet been subjected to clinical trials, which will determine the efficacy of such grafts in the long term. Finally, off the shelf availability and cost will become the biggest hurdles in the development of a feasible TEVG product.

Although many obstacles still exist in the effort to develop a small-diameter TEVG, the potential benefits of such an achievement are exciting. In the near future, a nonthrombogenic TEVG with sufficient mechanical strength may be developed for clinical trials. Such a graft will have the minimum characteristics of biological tissue necessary to remain patent over a time period comparable to current vein graft therapies. As science and technology advance, TEVGs may evolve into complex blood vessel substitutes. TEVGs may become living grafts, capable of growing, remodeling, and responding to mechanical and biochemical stimuli in the surrounding environment. These blood vessel substitutes will closely resemble native vessels in almost every way, including structure, composition, mechanical properties, and function. They will possess vasoactive properties, able to dilate and constrict in response to stimuli. Close mimicry of native blood vessels may ultimately aid in the engineering of other tissues dependent upon vasculature to sustain function. With further understanding of the factors involved in cardiovascular development and function combined with the foundation of knowledge already in place, the development of TEVGs should one day lead to improved quality of life for those with vascular diseases and other life threatening conditions.

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27

Cardiac Tissue

Engineering: Matching

Native Architecture

and Function to

Develop Safe and

Efficient Therapy

 

27.1

Introduction. . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

27-1

 

 

Cellular Cardiomyoplasty Tissue Cardiomyoplasty

27-3

 

27.2

Cardiac Architecture and Function . . . . . . . . . . . . . . . . . . . . .

 

27.3

Current State of Cardiac Tissue Engineering . . . . . . . . . .

27-4

 

 

Cardiogenesis In Vitro In Vivo Implantation for Cardiac

 

 

 

Repair

 

 

 

27.4

Design Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

27-10

 

 

Cell Source and Immunology Cellular Composition

 

 

 

Tissue Architecture

Tissue Thickness Electrical

 

 

 

Function and Safety

Spontaneous Activity

27-15

 

27.5

Future Work . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

 

27.6

Conclusions . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

27-16

Nenad Bursac

Acknowledgment. . . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

27-16

Duke University

References . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

27-17

27.1 Introduction

After an acute myocardial infarction, lost cardiomyocytes are replaced by a noncontractile fibrous tissue. Although it is suggested that heart has a small regenerative potential via cell proliferation [1], or stem cell recruitment [2], the rate of renewal is insufficient to compensate for myocyte loss. As a result, altered workload of a surviving myocardium may ultimately lead to deterioration in contractile function and congestive heart failure (CHF). Besides traditional pharmacological therapies (diuretics, β-blockers, angiotensine, and aldosterone inhibitors) [3] or heart transplant [4], investigators are evaluating innovative approaches for treatment of CHF including mechanical assist devices [5], dynamic cardiomyoplasty [6],

27-1

27-2

Tissue Engineering

transmyocardial laser revascularization [7], and artificial heart [8]. Nevertheless, in end stage disease, heart transplant remains the only option with good long-term results [4]. However, inadequate availability of donor organs ( 10% of current needs [9]) requires new strategies for treatment of increasing number of heart failure patients.

One promising approach is augmentation of the number of functional myocytes in the diseased heart using methodologies for cardiomyocyte cell cycle activation [10], adult stem cell mobilization [11], or cellular transplantation [12]. Cellular transplantation at the site of injury in the heart can be accomplished either by injecting isolated cells (“cellular cardiomyoplasty”) [13], or by implanting a cardiac tissue patch engineered in vitro (“tissue cardiomyoplasty”) [14].

27.1.1 Cellular Cardiomyoplasty

More than 10 years ago, pioneering studies in the laboratory of Lauren Field have shown feasibility of cell transplantation in the heart [15,16]. Since then, different investigators have used cardiac [15] or skeletal myoblast cell lines [16], fetal [17,18], neonatal [19], and adult [20] cardiac myocytes, autologous [21,22] and syngeneic [23] skeletal myoblasts, smooth muscle cells [24], endothelial cells [25], native [26] or genetically altered fibroblasts [27], embryonic [28], bone marrow [29], mesenchymal [30], or heart derived [31] stem cells, as potential donor cells. As a result, treated hearts have shown improvement in diastolic function, almost independent of transplanted cell type [26]. The improvement in the systolic function (generation of active force) on the other hand required use of cells with the myogenic (contractile) potential [26,32]. The possible therapeutic benefit of cellular cardiomyoplasty stems from structural remodeling of scar region [33], enhancement of myocardial revascularization [12], and direct structural and functional integration of donor cells with the host myocardium [18]. Presently, clinical studies in the United States, Europe, and Asia are under way to investigate feasibility and safety in using autologous bone marrow derived stem cells and skeletal myoblasts in treatment of postinfarction left ventricular disfunction [13,34,35]. Initial results are promising, but reveal risk for ventricular arrhythmias [34], limiting in some studies the pool of patients to only those that already have internal defibrillators. Since no systematic studies have been performed to assess the electrical performance of the heart postcardiomyoplasty, and little data exists on electrical interaction between donor and host cells in vivo or in vitro [36], the causes of the postoperative electrical instability are not known. Plausible explanations include inflammatory response and subsequent fibrosis at implantation site [35], possible electrical coupling in conjunction with different electrophysiology between implanted cells and cardiomyocytes [18,34], and possible stimulation of sympathetic nerve sprouting and overexpression of neurotransmitters after the cell transplantation [37].

27.1.2 Tissue Cardiomyoplasty

Some of the major hurdles in restoring heart function by cellular cardiomyoplasty include limited survival of injected cells in the region of scar tissue, and no architectural repair of the infarcted area. An alternative approach is tissue cardiomyoplasty, which involves in vitro cultivation of compact three-dimensional (3D) cardiac tissue patch, and subsequent implantation over or instead of the infracted scar tissue. Although surgically more challenging compared to cellular cardiomyoplasty, this methodology has a potential to improve efficiency and localization of tissue repair in larger size cardiac injury such as infarction of major coronary vessels, or congenital heart defects [38]. Ideally, based on the location, shape, and size of injury and architecture of surrounding tissue (assessed by ultrasound, MRI, or other noninvasive technique [39]), functional cardiac patch with needed geometry and 3D structure is engineered in vitro starting from selected cell type, natural or synthetic scaffold, and appropriate culturing vessel (bioreactor). Inside the bioreactor, cells attach to biocompatible (and possibly degradable) scaffold, interconnect, and assemble in three dimensions to reconstitute an in vivo-like cardiac tissue equivalent (construct). The combination of biochemical and physical stimuli during culture is designed to best mimic physiological state of tissue, and to support cell differentiation or transdifferentiation and desired 3D tissue architecture. At a proper

Cardiac Tissue Engineering

27-3

time point, tissue construct is removed from the bioreactor and surgically implanted in the site of injury in order to restore or improve electrical and mechanical function of diseased heart.

In reality, however, the successful reconstitution of cardiac-like tissue patch in vitro starting from single cells is an extremely challenging problem due to limited proliferation potential and high metabolic demand of cardiac cells, as well as complex anisotropic architecture and electro-mechanical function of native cardiac tissue. While recent reviews on cardiac tissue engineering [14,40] have focused on scaffold biomaterials, bioreactors, and cultivation conditions, this chapter will provide emphasis on the electrophysiological considerations and role of tissue architecture in the development of functional cardiac patch. It is this author’s view, that these factors will play an important role in the design of efficient and safe therapies, despite the fact that they are frequently neglected in current in vitro and in vivo studies.

27.2 Cardiac Architecture and Function

The crucial architectural and functional feature of cardiac muscle tissue (Figure 27.1) is anisotropy, that is, anatomically and biophysically, properties of cardiac muscle vary in different directions [41]. Microscopic structural anisotropy in cardiac tissue results from the spatial alignment of elongated cardiac myocytes (Figure 27.1a), and the preferential location of intercellular junctions (e.g., fascia adherence, gap junctions, desmosomes) in end-to-end vs. side-to-side cell connections [42]. Macroscopic anisotropy is a result of the presence of aligned cardiac muscle fibers and sheets that transmurally rotate inside the heart wall (180rotation from endoto epicardium) [43]. The unique anisotropic architecture of cardiac tissue enables an orderly sequence of electrical and mechanical activity, and efficient pumping of blood from the heart. Beside architecture, electrical membrane properties of cardiac myocytes also vary substantially depending on the location in the heart with distinct differences between atria and ventricles, endoand epicardial regions, left and right heart, base and apex, etc. [44–46]. Moreover, the heart contains a large variety of nonmyocytes (e.g., fibroblasts, endothelial cells, smooth muscle cells, neural cells, leukocytes) with specific roles in the cardiac function that are still not fully elucidated.

Structural anisotropy and intercellular continuity of the excitable cardiac substrate have profound effect on electrical and mechanical functioning of the heart. For example, anatomical anisotropy in heart tissue causes a larger intracellular resistance per unit length in the transverse (across fiber) than longitudinal (along fiber) direction, resulting in smaller velocity but larger maximum slope of action potential upstroke and safer electrical propagation in the transverse direction [47,48]. As a consequence, electrical stimulation of a small region in the heart tissue results in development of elliptical rather than circular propagating wavefront [49] (Figure 27.1b). Directly related to anisotropy is evidence that cardiac impulse conduction at the microscopic level is discontinuous at sites of gap junctions, and even stochastic due to small local variations in ion channel function, gap junction distribution, and adjoining tissue architecture [50,51]. In contrast to these small physiological variations, larger variations of electrical properties (e.g., action potential duration, intercellular coupling, electrical load) at the cellular and tissue level may result in increased susceptibility to propagation slowing and conduction block [46,51]. Slow propagation velocity and unidirectional block are some of the main prerequisites for the initiation of reentrant cardiac arrhythmias [51,52].

The degree of anatomical and functional anisotropy depends on location in the heart and age of the individual, with main determinants being cell size and geometry, type, amount, and distribution of cell junctions in membrane, and macroscopic tissue architecture [53–55]. Electrical anisotropy changes in certain cardiac pathologies such as ischemia, infarction, and heart failure [56,57]. This change is a consequence of the altered gap junction distribution and expression, as well as formation of longitudinal collagenous septa between the cardiac fibers which result in discontinuous transverse propagation (“nonuniform anisotropy”) and increased susceptibility to reentrant arrhythmias [58]. In particular, in canine hearts, “border zone” between infracted and normal tissue exhibits disarray of cardiac cells and gross change in anisotropy, resulting in conduction slowing, block, and reentrant “figure of eight” circuits