평활근의 조직공학

김병수

한양대학교 응용화학공학부

Tissue Engineering of Smooth Muscle

Byung-Soo Kim

Division of Applied Chemical Engineering, Hanyang University

INTRODUCTION

Smooth muscle (SM) is an important functional element of a number of cardiovascular (e.g., blood vessels), gastrointestinal (e.g, esophagus, stomach, and intestines), and urinary tissues (e.g., bladder and ureters). The functions of the SM element in these tissues include providing appropriate mechanical properties to withstand fluid pressures (e.g., blood pressure and urinary pressure) and contracting to regulate blood pressures or transfer fluids (e.g., digestive substance and urine). In this study, functional SM tissues with appropriate mechanical properties and contractility were fabricated from cultured smooth muscle cells (SMCs) and biodegradable polymers using a tissue engineering technique.

MATERIALS AND METHODS

Scaffolds

Polyglycolic acid (PGA) fiber-based scaffolds (fibers approximately 12mm in diameter) assembled into non-woven arrays were purchased from Albany International Inc. (Taunton, MA, USA). The porosity of the PGA matrix was approximately 97%. To stabilize scaffolds, a solution of PLLA (Boehringer Ingelheim, Winchester, VA, USA) dissolved in chloroform (5% w/v) was placed in an atomizer (Devilbus Corp., USA) and sprayed over both sides of PGA matrix from a distance of 3 inches using a nitrogen stream (18psi) [1]. The solvent was subsequently allowed to evaporate, and scaffolds were lyophilized to remove residual solvent. Type I collagen sponges were purchased from Integra Life Science Inc. (Plainsboro, NJ, USA). These sponges were fabricated by lyophilizing bovine type I collagen solution and cross-liking with glutaradehyde.

SMC isolation

SMCs were isolated from rat aortas using an adaptation of a previously published technique [2]. In brief, the descending aortas of 300-350g adult male Lewis rats after the removal of endothelium, adventitia, fat, and connective tissue, were cut into multiple small pieces and incubated for 90 minutes at 37□C in a sterile spinner flask (100 ml, Bellco Glass Inc., Vineland, NJ, USA) containing an enzymatic dissociation buffer. This buffer contains 0.125 mg/ml elastase (Sigma Chemical Co., St. Louis, MO), 1.0 mg/ml collagenase (CLS type I, 204 units/mg, Worthington Biochemical Corp., Freehold, NJ), 0.25 mg/ml soybean trypsin inhibitor (type 1-S, Sigma), and 2.0 mg/ml crystallized bovine serum albumin (BSA, Gibco/Life Technologies, Gaithersburg, MD). The resultant tissue suspension was filtered through a 100 □m Nitex filter (Tetko, Inc., Briarcliff Manor, NY) and centrifuged at 200 g for 5 minutes. The pellet was resuspended in growth medium (Medium 199 (Gibco) supplemented with 20% fetal bovine serum (FBS)(Gibco), 2 mM L-glutamine (Gibco), 100 units/ml penicillin (Gibco), and 0.1 mg/ml streptomycin (Gibco). SMCs were maintained in growth media containing 20% FBS until the first passage, while all subsequent cultures were grown in the presence of 10% FBS. SMCs between passage 3 and 5 were used in this study.

Engineering SM tissues under cyclic strain condition

A cell suspension (3.5x107 cells/ml, 3 ml per scaffold) was injected onto polymer scaffolds (50 x 35 x 0.25 mm) in petri dishes. The dishes containing the seeded polymer scaffolds were agitated at 100 rpm with an orbital shaker for 24 hours in a humidified 5% CO2 atmosphere. This dynamic seeding method enhanced the cell seeding efficiency dramatically compared to static seeding methods [3]. The seeded scaffolds were subsequently washed with PBS (Gibco), transferred to new petri dishes, and maintained in a humidified 5% CO2 atmosphere for 2 days with the medium changed every day. SMCs were subjected to cyclic strain in vitro using custom-made strain units. The seeded scaffolds were clamped in the tissue culture chamber and subjected to cyclic strain by movement of the crank back and forth as the eccentric disk connected to the crank rotates. The strain units are placed in a humidified incubator with 5% CO2 at 370C. The seeded scaffolds were subjected to cyclic strain at a frequency of 1 Hz (1 cycle per second) and an amplitude of 7% of initial length, which are similar to those of SM tissues in vivo, for up to 20 weeks. As a control, seeded scaffolds were fixed at only one end of the clamps and moved back and forth at the same frequency and amplitude as the mechanical strain conditions.

Histological analyses

For histological analysis, tissue samples were fixed in 10% (v/v) buffered formalin, dehydrated with a series of ethanol, paraffin embedded, sectioned (5□m thick), and stained with Verhoeff's staining. For transmission electron microscopic analysis, tissue samples were fixed in 2% glutaraldehyde for 2 hours at 40C, postfixed in 1% osmium tetroxide for 30 minute at 40C, dehydrated with acetone, and embedded in polybed 812 resin (Analytical Standards, Kungsbacka, Sweden). Thin sections (70nm thick) were cut by a Reichert Ultracut ultramicrotome and stained with lead citrate and uranyl acetate. The specimens were examined in a Jeol 100CX electron microscope (Jeol, Tokyo, Japan).

Mechanical test

The tensile mechanical properties of the SM tissues engineered with the collagen matrices in vivo for 10 weeks (n=3) were tested using a mechanical tester (model 5542, Instron corp., Canton, MA, USA). Longitudinal matrix strips (40 mm in length, 8 mm in width, and 0.1mm in thickness) were used for the test. A 500-Newton-maximum load cell was used, and the cross-head speed was 0.5 inch/min. The Youngs modulus was obtained from the slope of the initial linear section of the stress-strain curve.

Tissue contractility test

The engineered SM tissues were placed in Tyrodes solution (125.0mM NaCl, 23.8mM NaHCO3, 26.8mM KCl, 0.5mM MgCl2, 0.4mM NaH2PO4, 0.2mM CaCl2, 0.6mM glucose) in an organ bath (Randnoti Glass Technology, Monrovia, CA, USA) with a working volume of 50mL. The pH of the Tyrodes solution was maintained at 7.4 during all experiments by constant bubbling with 95% O2 and 5% CO2. The organ bath was double-chambered to maintain the temperature constant at 37oC with an external heating circuit. Changes in isometric tension were recorded with a recorder (model 1325, Bio-rad Laboratories, Hercules, CA, USA). The specimens were suspended from an electrode (Randnoti Glass Technology) in the organ bath and attached to a 2g force transducer (Randnoti Glass Technology). A passive force of 1g was applied. The specimens were allowed to equilibrate for 60 min prior electrical field or drug stimulation. Electrical field stimulation was applied with the following parameters; 80V, 5-msec duration, 32, 64, and 128Hz frequency, and 3 min intervals between 20 sec stimulations. Carbachol (Sigma) was used at a concentration of 10-4 M for drug stimulation. To minimize artifacts related to tissue exhaustion or drug interference, each specimen was subjected to only one series of electrical field stimulations followed by administration of the drug

RESULTS

The culture of cell-seeded polymer scaffolds resulted in the formation of newly-formed SM tissues. SMCs proliferated on the polymer scaffolds and produced extracellular matrices such as collagen and elastin, while the polymer scaffolds degraded during the culture, resilting in the formation of natural SM tissues with predefined size and shape.

Application of cyclic mechanical stimuli resulted in a great enhancement in mechanical properties of the engineered SM tissues. After being subjected to cyclic strain for 20 weeks, engineered SM tissues had 12±3-fold and 34±9-fold increases in ultimate tensile strength and Young's modulus, respectively, compared to control tissues [4].

Cyclic mechanical stimuli significantly altered SMC phenotype on polymer scaffolds. SMCs in tissues engineered under cyclic strain exhibited the properties of contractile phenotype, which is the phenotype of SMCs in native SM tissues, as analyzed by transmission electron microscopy, while cells in control tissues exhibited the properties of the synthetic, non-differentiated phenotype. Bundles of myofilaments coalescing in dense bodies, a contractile apparatus, were evident in SMCs under cyclic strain, whereas the cytoplasm of cells in control tissues was dominated by significant amounts of rough endoplasmic reticulum, a synthetic organelle [5].

Organ bath experiments were performed to determine if the engineered SM tissues exhibit contractility, which is one of SM-specific functions. The preliminary results indicated that the SM tissues engineered under cyclic strain condition contracted in response to both electric and biochemical stimulation, whereas the SM tissues engineered under the control condition (no strain) did not contracted in response to both types of stimulation.

CONCLUSIONS

This study demonstrated that functional SM tissues with appropriate mechanical properties and contractility can be engineered from cultured  SMCs and biodegradable polymers. This study may aid in defining condition required to engineer functional SM tissues and lead to clinically relevant engineered SM-containing tissues such as blood vessels, intestines, and bladder.

REFERENCES

1.	Kim BS, Mooney DJ (1998). Engineering smooth muscle tissue with a predefined structure. Journal of Biomedical Materials Research 41: 322-332.

2.	Rothman A, Kulik TJ, Taubman MB, Berk BC, Smith CWJ, Nadal-Ginard B (1992) Development and characterization of a cloned rat pulmonary arterial smooth muscle cell line that maintains differentiated properties through multiple subcultures. Circulation, 86: 1977-1986.

3.	Kim BS, Putnam AJ, Kulik TJ, Mooney DJ (1998). Optimizing seeding and culture methods to engineer smooth muscle tissue on biodegradable polymer matrices. Biotechnology & Bioengineering 57: 46-54.

4.	Kim BS, Nikolovski J, Bonadio J, Mooney DJ (1999). Cyclic mechanical strain regulates the development of engineered smooth muscle tissue. Nature Biotechnology 17: 979-983.

5.	Kim BS, Mooney DJ (2000). Scaffolds for engineering smooth muscle tissues under cyclic mechanical strain conditions. Journal of Biomechanical Engineering 122: 210-215.