Journal of Investigative Medicine:
Symposium and Meeting Reports
Using a Type 1 Collagen-Based System to Understand Cell-Scaffold Interactions and to Deliver Chimeric Collagen-Binding Growth Factors for Vascular Tissue Engineering
Pang, Yonggang MD, PhD*†‡; Greisler, Howard P. MD*†§
From the *Department of Surgery, Loyola University Medical Center, Maywood; †Edward Hines Jr VA Hospital, Hines; ‡Department of Biomedical Engineering, Illinois Institute of Technology, Chicago; and §Department of Cell Biology, Neurobiology and Anatomy, Loyola University Medical Center, Maywood, IL.
Received May 4, 2010.
Accepted for publication June 22, 2010.
Reprints: Howard P. Greisler, MD, Department of Surgery, Loyola University Medical Center, 2160 S First Ave, Maywood, IL 60153. E-mail: firstname.lastname@example.org.
Supported by grants from the NIH R01-HL41272 (HPG), Department of Veteran's Affairs (HPG), and Chicago Association for Research and Education in Science (HPG). This symposium was supported in part by a grant from the National Center for Research Resources (R13 RR023236).
Vascular tissue engineering should provide more biocompatible and functional conduits than synthetic vascular grafts. Understanding cell-scaffold interactions and developing an efficient delivery system for growth factors and other biomolecules to control the signaling between the cells and the scaffold are fundamental issues in a wide range of tissue engineering research fields. Type 1 collagen is a natural scaffold extensively used in vascular tissue engineering and is a widely used vehicle in biomolecule delivery. In this article, we will discuss type 1 collagen as a vascular tissue engineering scaffold, describe strategies for elucidating the interaction between cells and type 1 collagen scaffolds using various imaging techniques, and summarize our work on the development of a chimeric collagen-binding growth factor-based local delivery system.
In the United States, cardiovascular disease has been the leading cause of death for many years; nearly 2300 Americans died of cardiovascular disease each day, and coronary heart disease caused approximately 1 of every 6 deaths in the United States in 2006.1 Bypasses are widely used to treat vascular diseases, and in 2006, 448,000 inpatient bypass procedures were performed in the United States.1 However, there are still a large number of patients who do not have suitable vessels for the surgery2 and thus require synthetic vessels as alternatives.3,4 Because of the drawbacks of the synthetic vessels, efforts have been made in vascular tissue engineering to develop biocompatible functional conduits.5,6 Cells, scaffolds, and biomolecular signals are the 3 principle components of tissue engineering.7 An ideal scaffold should mimic the biochemical and biomechanical properties of the extra cellular matrix of the tissue to be engineered and should support vascular cell viability and optimal phenotypic characterization. An ideal tissue engineering system also requires better control of the biomolecular signals between cells and scaffold. An efficient way to regulate and target the signal is to deliver desired growth factors locally to the cells. In this article, we will focus on type 1 collagen, one of the most widely used natural scaffolds. We will cover the application of type 1 collagen as a vascular tissue engineering scaffold, describe strategies for elucidating the interaction between cells and type 1 collagen-based 3-dimensional (3-D) scaffolds, and discuss a local growth factor delivery strategy based on a chimeric collagen-binding growth factor using type 1 collagen as the delivery vehicle.
APPLICATION OF TYPE 1 COLLAGEN AS SCAFFOLD FOR VASCULAR TISSUE ENGINEERING
Both synthetic and natural materials have been used as scaffolds for vascular tissue engineering. Although the chemical properties of synthetic materials are controllable and reproducible, biocompatibility, biomechanical stability, immunoreactivity and infectability, and inflammation remain as unmet challenges. Natural scaffolds are commonly large molecule components of the extracellular matrix (ECM) in vivo or have similar macromolecular properties to natural ECM and include collagen,8,9 fibrin,10 hyaluronic acid,11 alginate,12 and chitosan.13 Collagens constitute 25% of the total protein mass of most mammals and have minimal cross-species immunological reactivity.14 Most hydrogel-based scaffolds do not naturally adhere cells nor promote cell function. Collagen is a rare exception because it is rich of integrin-binding domains that enhance cell attachment and growth. At least 19 different types of collagen have been confirmed. The collagen molecule is a 3-stranded structure consisting of 3 polypeptide chains twined around one another, and each chain has an individual twist in opposite directions.14 The strands are held together primarily by hydrogen bonds and also by covalent bonds providing supplementary binding. The molecular weight of the basic collagen molecule is approximately 300 kd, and the length and the width of a rod-shaped collagen fiber are 300 and 1.5 nm, respectively.14 Among all the types of collagen, type 1 collagen is the most favorable for vascular tissue engineering because it is a major ECM protein within the wall of a blood vessel, and smooth muscle cells (SMCs) and fibroblasts naturally reside in collagen-rich matrices. It is also relatively easily shaped into tubular configurations.15,16 Therefore, it has been extensively used in vascular tissue engineering from the first tissue-engineered blood vessel17 until today.
ELUCIDATION OF CELL-TYPE 1 COLLAGEN SCAFFOLD INTERACTIONS
In 1986, Weinberg and Bell17 established a model for vascular tissue engineering that used SMC-populated collagen to construct the media layer of blood vessel. It was a milestone in tissue engineering history. However, they failed to show the requisite mechanical strength. In 1993, L'Heureux et al18 modified this model using cells from different sources but encountered the same problem. Recently, scientists began to understand that the biomechanical strength of a hydrogel-based tissue-engineered blood vessel is dependent largely upon hydrogel compaction by SMCs or fibroblasts within the scaffolds. Various methods have been used to increase the compaction by SMCs and thereby increase the mechanical strength of the engineered vessel, including in vitro mechanical preconditioning or biochemical stimulation of SMCs. Smooth muscle cells that contribute most of the contractility of an artery and the 3-D ECM, where SMCs reside, is composed primarily of type 1 collagen. It is therefore critical to understand the mechanisms by which cells interact with the matrix to facilitate the engineering of a better tissue-engineered blood vessel.
Collagen remodeling has been extensively studied with some traditional techniques, performed on a fixed sample with an electron microscope or at a macroscopic level by measuring the changes of length, width, and height of the collagen-cell constructions.19,20 Both methods have provided limited information. Cells remodel scaffolds by continuously rearranging the microstructure of the scaffold while cells remodel the surrounding matrix either in vivo or in vitro. Therefore, it is important to understand their interaction dynamically. Moreover, in growth factor-incorporated systems, cells will alter the release of growth factors dynamically. Growth factors are in the nanometer range, and although they are commonly clustered, the aggregated growth factor microspheres are still at the micrometer level. Therefore, it is also important to study these dynamics with high resolution. We have developed a new microscopic approach to better understand their temporal and spatial interaction dynamics in 6 dimensions: 3 spatial dimensions (x, y, and z), a time dimension, a spectral dimension (multichannel of fluorophores), and a multiposition dimension (mosaic imaging).9 First live time-lapse phase imaging enables a long-term dynamic study of collagen remodeling, and the locomotion characteristics of the cells were studied with computer-assisted cell tracking. Fluorescent and reflection confocal microscopic techniques were used to acquire images and for quantification of microscale changes of both SMCs and collagen fibers during remodeling. Although the time-lapse z-stack images were acquired, visualizing and presenting the 3-D structure dynamically are still challenging. In our laboratory, we developed 2 image processing techniques to accomplish real 3-D visualization. First, 3-D volume-rendered images were generated at each time point, and a 3-D video was generated using the 3-D image frames (Fig. 1). This 3-D presentation is the first report in the literature as far as we know. Another image processing technique that we developed for dynamic visualization is the multiwindow dynamic view that enables viewing the multiple focal planes at each time point.
It is well known that cell contractility, which is mediated by the cytoskeletal system, plays a major role in matrix remodeling, cell-matrix interaction, and growth factor release. However, little has been reported on direct simultaneous viewing of cell cytoskeleton component changes and related scaffold changes at the microscale level. In our laboratory, we used our confocal microscopic techniques to directly visualize the α-smooth muscle actin fibers and collagen fibers. The directional relationship between actin fibers and collagen fibers was analyzed by fast Fourier transform, and the results provided quantitative information showing that actin fibers play an important role in collagen scaffold remodeling. Reflection confocal microscopy provides nonlabel and noninvasive means to visualize collagen fiber reorganization at high resolution. However, one limitation is that under the high magnification objective and within one visual field, very few cells, often only one, can be viewed. Data based on few cells are too limited, and arbitrary judgments may occur. To balance the resolution and field-of-view requirements, mosaic imaging is a good choice. Some 2-D mosaic techniques have been successfully used in studies of ocean-floor21 and of x-ray22 images. Applying mosaic techniques to 3-D collagen matrix and cells is challenging because of the complex structure. Moreover, most of the previous mosaic imaging studies were based on motor-controlled stages at low magnification. Image stitching using a motor-controlled stage may generate mismatched images because of the limitation of the accuracy of the stage and the hydrogel sample that may independently move as the stage moves. These errors are tolerable in low-magnification images but not in high-magnification images. The solution for this 3-D hydrogel mosaic imaging is to use software image registration to stitch the images. In other words, the software finds the overlay parts of 2 adjacent images and corrects image position including tilt. The speed and the accuracy were also improved by generating a position file from the cell channels, which have simpler structure compared with the collagen channel. The position information in the file was then used to adjust the images in the collagen channel. Stack mosaic imaging was simply achieved by using the same position file in the other focal planes. Thus, we created a simple and cost-effective way to generate multichannel, multifocal plane mosaic imaging without using an expensive motor-controlled stage while simultaneously eliminating the mechanical errors.
Quantification of the degree of scaffold remodeling is another step we took to study the cell-scaffold interaction. A traditional way of quantifying the spatial changes of collagen scaffolds is measuring the dimension changes of the collagen gel globally, an approach which limited the study to the macroscale level. Low accuracy and limited information generated from macroscale studies have led to an incomplete understanding of the remodeling process. In our laboratory, we perform the quantification at the microscale level by analyzing the digitized confocal images at the micrometer level, enabling a better understanding of the mechanism of remodeling. We quantify not only the compaction in the remodeling process but also, by fast Fourier transform, the alignment, which cannot be accomplished in more traditional techniques. After the transform, total orientation of both cells and collagen fibers is clearly shown, and the alignment index is generated from the spectral plot, which provides quantitative information.
APPLICATION OF A GROWTH FACTOR DELIVERY STRATEGY USING TYPE 1 COLLAGEN AS THE DELIVERY VEHICLE
Natural polymers and their derivatives in the form of gels and sponges have been used extensively as delivery vehicles. Collagen in particular is a readily available ECM component that allows cell infiltration and remodeling, making it highly suitable for biomolecule delivery. To the best of our knowledge, the first description of using collagen as a delivery vehicle was reported in 1973.23 Since then, various collagen-based delivery systems have been used (Table 1).
Most of the collagen-based delivery system just traps the biomolecules inside the collagen fibril networks, and release largely depends on microdimension of the collagen fibers and growth factors. A major drawback to this delivery strategy is that the time course is not well controlled. There is typically a burst release within the first few hours, and total release time is comparatively short. This is partly due to the large pore size of the scaffold material compared with the relatively small size of most growth factors. For example, the pore size of type 1 collagen scaffolds commonly varies from 66.8 μm (at 1.67-mg/mL concentration) to 37.5 μm (at 2.5-mg/mL concentration),34 whereas the size of most growth factors are at the nanometer level or lower. Decreasing the pore size of collagen by increasing the gel concentration may extend the retention of the growth factor inside the collagen hydrogel. However, this reduces the nutrient transport and limits cell growth and viability. To overcome these problems, we constructed a chimeric collagen-binding fusion growth factor, R136K-CBD (Fig. 2). It consists of R136K, which is a relatively thrombin-resistant mutant derivative of FGF-1, and a collagen-binding domain (CBD).35 We demonstrated that R136K-CBD has a significantly higher binding affinity to a 3-D collagen scaffold than FGF-1 and R136K. Furthermore, R136K-CBD showed still greater binding affinity compared with FGF-1 and R136K in a cell-populated collagen gel. We then fluorescently labeled R136K-CBD, and by confocal microscopic imaging and analysis, we found that R136K-CBD shows uniform distribution throughout collagen gel, which suggests that incorporated R136K-CBD would likely have an essentially uniform biologic effect on SMCs. Growth factor internalization was directly visualized by our microscopic techniques. Both R136K-CBD and the SMCs were fluorescently labeled (different labels) and incubated within the collagen hydrogel. The SMCs were then isolated from the collagen, and the internalized R136-CBD was found to be localized to cytoplasm and the perinuclear region. To better understand the delivery mechanism, the internalization was also studied in 3-D in an on-site visualization mode without isolating SMCs from the collagen hydrogel. R136K-CBD binding to collagen fibers was visualized by reflection and fluorescent confocal microscopy. By image volume rendering of SMCs and collagen fibers, their spatial relationships were clearly visualized. More importantly, R136K-CBD elicited a greater mitogenic effect on SMCs in the 3-D collagen scaffold. Up to 7 days, R136K-CBD-stimulated groups showed continuous increasing cell proliferation, whereas the proliferation rate dropped in R136K- and FGF-1-stimulated groups. At day 7, R136K-CBD demonstrated a 2.0- and 2.1-fold greater mitogenic effect than R136K and FGF-1, respectively.
Our current microscopic studies on dynamic interactions between cells and collagen are on samples without mechanical stimulation. Because mechanical stimulation has been shown to increase the mechanical strength of collagen-cell constructs, we are developing techniques to dynamically image the sample while under mechanical stimulation. For CBD-based delivery systems, more complex delivery strategies are being developed to accomplish both short- and long-term delivery of multiple peptides, including the encapsulation of the CBD growth factor in biodegradable microspheres, allowing its binding to the collagen after its release from the microspheres and eventually internalization by the cells.
1. Lloyd-Jones D, Adams RJ, Brown TM, et al. Heart disease and stroke statistics-2010 update: a report from the American Heart Association. Circulation. 2010;121(7):e46-e215.
2. Campbell JH, Efendy JL, Campbell GR. Novel vascular graft grown within recipient's own peritoneal cavity. Circ Res. 1999;85(12):1173-1178.
3. Bader A, Steinhoff G, Strobl K, et al. Engineering of human vascular aortic tissue based on a xenogeneic starter matrix. Transplantation. 2000;70(1):7-14.
4. Xue L, Greisler HP. Biomaterials in the development and future of vascular grafts. J Vasc Surg. 2003;37(2):472-480.
5. Stegemann JP, Kaszuba SN, Rowe SL. Review: advances in vascular tissue engineering using protein-based biomaterials. Tissue Eng. 2007;13(11):2601-2613.
6. Thomas AC, Campbell GR, Campbell JH. Advances in vascular tissue engineering. Cardiovasc Pathol. 2003;12(5):271-276.
7. Griffith LG, Naughton G. Tissue engineering-current challenges and expanding opportunities. Science. 2002;295(5557):1009-1014.
8. Pang Y, Wang X, Ucuzian AA, et al. Local delivery of a collagen-binding FGF-1 chimera to smooth muscle cells in collagen scaffolds for vascular tissue engineering. Biomaterials. 2010;31(5):878-885.
9. Pang Y, Ucuzian AA, Matsumura A, et al. The temporal and spatial dynamics of microscale collagen scaffold remodeling by smooth muscle cells. Biomaterials. 2009;30(11):2023-2031.
10. Xue L, Greisler HP. Angiogenic effect of fibroblast growth factor-1 and vascular endothelial growth factor and their synergism in a novel in vitro quantitative fibrin-based 3-dimensional angiogenesis system. Surgery. 2002;132(2):259-267.
11. Tan H, Ramirez CM, Miljkovic N, et al. Thermosensitive injectable hyaluronic acid hydrogel for adipose tissue engineering. Biomaterials. 2009;30(36):6844-6853.
12. Stevens MM, Qanadilo HF, Langer R, et al. A rapid-curing alginate gel system: utility in periosteum-derived cartilage tissue engineering. Biomaterials. 2004;25(5):887-894.
13. Jiang T, Kumbar SG, Nair LS, et al. Biologically active chitosan systems for tissue engineering and regenerative medicine. Curr Top Med Chem. 2008;8(4):354-364.
14. Lee CH, Singla A, Lee Y. Biomedical applications of collagen. Int J Pharm. 2001;221(1-2):1-22.
15. Stegemann JP, Nerem RM. Altered response of vascular smooth muscle cells to exogenous biochemical stimulation in two- and three-dimensional culture. Exp Cell Res. 2003;283(2):146-155.
16. Miller EJ, Rhodes RK. Preparation and characterization of the different types of collagen. Methods Enzymol. 1982;82(pt A):33-64.
17. Weinberg CB, Bell E. A blood vessel model constructed from collagen and cultured vascular cells. Science. 1986;231(4736):397-400.
18. L'Heureux N, Germain L, Labbe R, et al. In vitro construction of a human blood vessel from cultured vascular cells: a morphologic study. J Vasc Surg. 1993;17(3):499-509.
19. Li S, Moon JJ, Miao H, et al. Signal transduction in matrix contraction and the migration of vascular smooth muscle cells in three-dimensional matrix. J Vasc Res. 2003;40(4):378-388.
20. Franco C, Ho B, Mulholland D, et al. Doxycycline alters vascular smooth muscle cell adhesion, migration, and reorganization of fibrillar collagen matrices. Am J Pathol. 2006;168(5):1697-1709.
21. Marks RL, Rock SM, Lee MJ. Real-time video mosaicking of the ocean floor. IEEE J Ocean Eng. 1995;20(3):229-241.
22. Loo BW Jr, Meyer-Ilse W, Rothman SS. Automatic image acquisition, calibration and montage assembly for biological x-ray microscopy. J Microsc. 2000;197(pt 2):185-201.
23. Rubin AL, Stenzel KH, Miyata T, et al. Collagen as a vehicle for drug delivery. Preliminary report. J Clin Pharmacol. 1973;13(8):309-312.
24. Friedl P, Maaser K, Klein CE, et al. Migration of highly aggressive MV3 melanoma cells in 3-dimensional collagen lattices results in local matrix reorganization and shedding of alpha2 and beta1 integrins and CD44. Cancer Res. 1997;57(10):2061-2070.
25. Brewster LP, Washington C, Brey EM, et al. Construction and characterization of a thrombin-resistant designer FGF-based collagen binding domain angiogen. Biomaterials. 2008;29(3):327-336.
26. Kay JS, Litin BS, Jones MA, et al. Delivery of antifibroblast agents as adjuncts to filtration surgery-part II: delivery of 5-fluorouracil and bleomycin in a collagen implant: pilot study in the rabbit. Ophthalmic Surg. 1986;17:796-801.
27. Phinney RB, Schwartz SD, Lee DA, et al. Collagen-shield delivery of gentamicin and vancomycin. Arch Ophthalmol. 1988;106:1599-1604.
28. Moursi AM, Winnard PL, Fryer D, et al. Delivery of transforming growth factor-beta2-perturbing antibody in a collagen vehicle inhibits cranial suture fusion in calvarial organ culture. Cleft Palate Craniofac J. 2003;40:225-232.
29. Letic-Gavrilovic A, Piattelli A, Abe K. Nerve growth factor beta(NGF beta) delivery via a collagen/hydroxyapatite (Col/HAp) composite and its effects on new bone ingrowth. J Mater Sci Mater Med
30. Sun JS, Lin FH, Wang YJ, et al. Collagen-hydroxyapatite/tricalcium phosphate microspheres as a delivery system for recombinant human transforming growth factor-beta 1. Artif Organs
31. Hu Y, Zhang C, Zhang S, et al. Development of a porous poly(l-lactic acid)/hydroxyapatite/collagen scaffold as a BMP delivery system and its use in healing canine segmental bone defect. J Biomed Mater Res A
32. Govender S, Csimma C, Genant HK, et al. Recombinant human bone morphogenetic protein-2 for treatment of open tibial fractures: a prospective, controlled, randomized study of four hundred and fifty patients. J Bone Joint Surg Am. 2002;84-A:2123-2134.
33. Takezawa T, Takeuchi T, Nitani A, et al. Collagen vitrigel membrane useful for paracrine assays in vitro and drug delivery systems in vivo. J Biotechnol. 2007;131:76-83.
34. Friedl P, Maaser K, Klein CE, et al. Migration of highly aggressive MV3 melanoma cells in 3-dimensional collagen lattices results in local matrix reorganization and shedding of alpha2 and beta1 integrins and CD44. Cancer Res. 1997;57:2061-2070.
35. Brewster LP, Washington C, Brey EM, et al. Construction and characterization of a thrombin-resistant designer FGF-based collagen binding domain angiogen. Biomaterials. 2008;29:327-336.
type 1 collagen; growth factor; confocal microscopy; local delivery; scaffold
© 2010 American Federation for Medical Research
Highlight selected keywords in the article text.