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2007, 111, 14602-14606 Published on Web 09/20/2007
Capillary Differentiation of Endothelial Cells on Microgrooved Surfaces Hua Wang,† Shengfu Chen,† Buddy D. Ratner,*,† E. Helene Sage,*,‡ and Shaoyi Jiang*,† Department of Chemical Engineering, UniVersity of Washington, Seattle, Washington 98195 and Hope Heart Program, Benaroya Research Institute at Virginia Mason, Seattle, Washington 98101 ReceiVed: July 22, 2007; In Final Form: September 3, 2007
We investigated the capillary differentiation of endothelial cells cultured on microgrooved polydimethylsiloxane (PDMS) substrates coated with fibronectin. Bovine aortic endothelial (BAE) cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) on PDMS substrates produced photolithographically. Phase contrast images of BAE cells were taken to assess the overall morphology of cells. Confocal microscopy was used to examine the existence of lumen inside cord-like and tube-like structures developed at different stages of cell culture. Herein we show for the first time that microgrooved PDMS substrates of appropriate dimensions coated with fibronectin are capable of inducing capillary differentiation of endothelial cells in the absence of angiogenic growth factors.
Introduction Angiogenesis, the formation of new blood vessels, plays a critical role in many physiological and pathological processes, such as embryogenesis, wound healing, and tumor growth.1,2 In tissue engineering, angiogenesis is required to ensure adequate oxygen and nutrients to and effective removal of waste products from the new tissue.3 Until now, successful tissue engineering applications have been restricted to relatively thin (i.e., less than 100-200 µm) tissues or avascular structures, such as skin, where local diffusion can provide sufficient mass transport. For thicker, metabolically demanding organs, such as heart, brain, and liver, angiogenesis remains a critical obstacle to tissue regeneration. Better understanding and control of angiogenesis is an important step in tissue engineering. Since the early observations of angiogenesis in vitro by Folkman and Haudenschild4 25 years ago, a number of in vitro angiogenesis assays have been established. It has been demonstrated that angiogenesis can be initiated through the action of many soluble factors such as fibroblast growth factor (FGF), vascular endothelial growth factor (VEGF), and tumor necrosis factor.5 In addition to these angiogenic factors, in vitro capillary formation is also shown to depend on the local matrix environment. For example, endothelial cells seeded onto plastic culture dishes coated with adhesive proteins formed cord-like structures slowly (2-3 weeks) on top of a confluent monolayer of cells.6 A network of tube-like structures was observed within 1-3 days when endothelial cells were cultured on top of a gel made of extracellular matrix (ECM) components, such as collagen and fibrin.7,8 * To whom correspondence should be addressed. Shaoyi Jiang e-mail:
[email protected]. Phone: (206) 543-4548. Fax: (206) 685-3451. Buddy D. Ratner e-mail:
[email protected]. Phone: (206) 685-1005. Fax: (206) 616-9763. Helene Sage e-mail: hsage@ benaroyaresearch.org. Phone: (206) 903-2026. Fax: (206) 903-2144. † University of Washington. ‡ Benaroya Research Institute at Virginia Mason.
10.1021/jp075746z CCC: $37.00
The mechanism by which a matrix exerts its effect on the formation of capillary-like structures has been under active investigation. Different ECM components can stimulate distinct biochemical signaling events through specific binding interactions between matrix proteins and cell surface receptors. For example, Madri and Williams9 showed that endothelial cells proliferated when seeded on type-I or -III collagen, whereas they differentiated on type-IV or -V collagen. Chalupowicz et al.10 reported that maximal capillary tube formation was observed with fibrin II (lacking both fibrinopeptides A and B), minimal tube formation with fibrin I (lacking only fibrinopeptide A), and complete absence of tube formation with fibrin 325 (lacking the NH2- terminal β15-42 sequence, in addition to fibrinopeptides A and B). Other studies suggested that biomechanical tension between endothelial cells and matrices might regulate the development of endothelial cells to capillaries by switching between cell proliferation and differentiation. For example, Vailhe et al.8 demonstrated that malleable fibrin gels induced more capillary differentiation of endothelial cells than rigid fibrin gels. Gamble et al.11 showed that anti-integrin antibodies that inhibit cell-matrix interactions converted endothelial cells from a proliferative phenotype toward a differentiation phenotype, leading to enhanced capillary tube formation. Mechanochemical switching between proliferation and differentiation was also achieved by altering the coating densities of fibronectin.12 Besides these biochemical and biomechanical studies, recent work with micropatterned flat substrates coated with adhesive islands of extracellular matrix demonstrated that endothelial cells could be geometrically switched between growth, apoptosis, and differentiation depending on cell shape (i.e., degree of cell extension).13 A recent study also showed the formation of capillary tube-like structures by micropatterned endothelial cells on 20-µm lines of bare chitosan.14 In living systems, basement membranes possess a complex, three-dimensional topography known to affect cell behavior.15 © 2007 American Chemical Society
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J. Phys. Chem. C, Vol. 111, No. 40, 2007 14603
Figure 1. Schematic diagram of the photolithographic process.
Furthermore, biomaterial surfaces are rarely flat at the molecular level.16 Besides biochemical, biomechanical, and 2D geometric properties, the 3D topographic features of substrates also influence fundamental cell-matrix interactions. For example, adhesion, migration, and ECM production have been shown to be generally greater on rougher surfaces for a wide variety of cells.17,18 Investigations of grooved substrates also revealed that cells aligned to the long axis of the grooves and became more oriented with increasing depth and decreasing width of the grooves.19-21 Most of the substrates currently under investigation are with micrometer-scaled features, to fully understand the behavior of cell adhering to basement membranes, substrates with smaller, denser, nanometer-scale features need to be fabricated because the topographic features of the basement membranes have been shown to possess features in the nanometer size range. Although previous studies have provided valuable insight into the interactions of cells with substrates of different topographies, little is known about the topographic effects of synthetic microstructured surfaces on the capillary differentiation of endothelial cells. In the present study, we examined the capillary differentiation of endothelial cells via topographic control by culturing bovine aortic endothelial (BAE) cells on microstructured poly(dimethy siloxane) (PDMS) substrates coated with fibronectin, which assists initial cell adhesion. Microgrooves with depths of 30 µm, and widths of 30-200 µm were produced by photolithography. Herein, we report that microgrooved surfaces can induce the capillary differentiation of endothelial cells, depending on the widths of the grooves. Materials and Methods Substrate Preparation. Patterned PDMS substrates were cast from photolithographically microfabricated silicon masters,10 as shown in Figure 1. The silicon masters possess parallel surface grooves with widths ranging from 30 to 200 µm and depths of 30 µm. PDMS was prepared by mixing prepolymer and catalyst (Sylgard 184 kit, Dow Corning, Midland, MC) in a 10:1 w/w ratio. The mixture was cast onto the patterned silicon master and placed under vacuum to eliminate bubbles generated during mixing. PDMS was then cured by baking for 1 h at 55 °C. After
Figure 2. Phase contrast images of BAE cells cultured in DMEM supplemented with 10% FBS for 1 day on microgrooved PDMS substrates coated with fibronectin. The depths of the grooves are 30 µm. The widths are (a) 200 µm, (b) 100 µm, and (c) varied from 200, 150, 100, and 50 to 30 µm. The ridges of the grooves are marked as “R”. Original magnification is 10× (bars ) 50 µm).
cooling to room temperature, the PDMS gel was peeled from the silicon master and punched into 1/4-inch circular slides. Before cell culture, the micropatterned PDMS slides were UV ozone cleaned and coated with 200 µg/mL fibronectin (Sigma Aldrich, St Louis, MO) at 37 °C for 2 h. Endothelial Cell Culture. BAE cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), passaged once a week and discarded after 15 passages as described previously.22 Cell culture medium and reagents were obtained from Gibco (Carlsbad, CA). Micropatterned PDMS substrates coated with fibronectin were transferred into a 24-well culture plate. Then, 2 mL of BAE cell suspension was plated in DMEM with 10% FBS at 100 000 cells/mL. Microscopy. Phase contrast images of living cells were recorded using an inverted microscope (Nikon TE200) with a ×10 objective at the University of Washington Engineered Biomaterials (UWEB) Microscopy and Image Analysis Center (Seattle, WA). Confocal microscopy was carried out with a confocal imaging system (LSM 510 Zeiss). Capillary-like structures were fixed with 4% paraformaldehyde for 5 min, stained with 0.2% phloxine (Sigma, St. Louis, MO) for 2 min, and washed with PBS.
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Figure 3. Phase contrast images of BAE cells cultured in DMEM supplemented with 10% FBS for 5 days on microgrooved PDMS substrates coated with fibronectin. The depth of the grooves is 30 µm. The widths are (a) 200 µm, (b) 100 µm, and (c) varied from 200, 150, 100, and 50 to 30 µm. The ridges of the grooves are marked as “R”. Original magnifications are 10× for a-c and 20× for d-f, which are three zoom-in images of c (bars ) 50 µm).
Figure 4. Confocal micrographs of the cord-like structures on top of cell monolayers. Phloxine staining was used to localize the whole cell body. Figures A-H are optical sections located at 4.93, 6.05, 7.17, 8.29, 9.41, 10.53, 11.65, and 12.77 µm from the top edge of the cells, respectively. Original magnification is 40×.
Results and Discussion Within 1 day of culture, BAE cells cultured on microgrooved substrates exhibited a typical polygonal morphology and formed confluent monolayers (Figure 2). There were no obvious differences between the cells located on the ridges and in the
grooves. After 4-6 days of culture, a cell morphology, which was termed by Iruela-Arispe et al. as “sprouting”,23 was observed (Figure 3). The sprouting cells organized into cordlike structures on top of a confluent monolayer of cells, and the cord-like structures were better aligned to the long axis of
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Figure 5. Phase contrast images of BAE cells cultured in DMEM supplemented with 10% FBS for 2 weeks on microgrooved PDMS substrates coated with fibronectin. The depth of the grooves is 30 µm. The widths are varied from 200, 150, 100, and 50 to 30 µm. The ridges of the grooves are marked as “R”. Original magnifications are 10× for a and b and 20× for c and d (bars ) 50 µm).
Figure 6. Confocal micrographs of the tube-like structures shown in the marked region of Figure 5b, demonstrating the existence of lumens. Phloxine staining was used to localize the whole cell body. Figures A-H are optical sections located at 4.61, 5.53, 6.46, 7.38, 8.30, 9.22, 10.15, and 11.07 µm from the top edge of the cells, respectively. Figure 1 is the cross section of the tubes. Original magnification is 40×.
the grooves as the widths of the grooves decreased (Figure 3df). However, hollow structures were not observed inside these cord-like structures, as shown by confocal microscopy (Figure 4). The mechanism of the “sprouting” phenomenon remains to be elucidated. The fact that these cord-like structures were observed at the bottom of the grooves rather than on the neighboring ridges could be attributed to limited mass transportation in the grooves and the potent effect of the grooves
for cell alignment. Previous studies revealed that cell alignment along the long axis of the grooves was often associated with the reorganization of actin and other cytoskeletal elements in an orientation parallel to the grooves.15 Changes in the shape of endothelial cells could further alter intracellular metabolic pathways that might be required for the formation of cord-like structures.12 Interestingly, previous studies showed that the degree of cell orientation increased with increasing groove depth
14606 J. Phys. Chem. C, Vol. 111, No. 40, 2007 and decreasing groove width. In our study, we also found more and better-aligned cord-like structures formed with decreasing groove width. In cell cultures plated for a longer period of time (e.g., 2-3 weeks), another pattern of cells was observed. Most (>95%) of the endothelial cells detached from the substrate, while tubelike structures were formed with the ends of the tubes attached to those cells that remained adherent to the ridges of the grooves (Figure 5). The tubes organized into a network, sometimes forming “bridges” across the grooves, possibly related to cell traction force balance. Results from confocal microscopy (Figure 6) demonstrated the existence of lumens inside these tube-like structures because labeling by phloxine was not observed in the inner part of the tubes. The network of tubes could be easily detached from the substrate by washing with PBS, indicating that most cells lost their contacts with the substrate. We speculate that cell elevation and retraction play important roles in the formation of the tube-like structures. After weeks of culture, endothelial cells, especially those at the bottom of the grooves, detached themselves from the poorly adhesive substrates. They retracted and migrated up along the sidewalls of the grooves, while maintaining their lateral contacts with neighboring cells. As a result of multicellular retraction, portions of the cells became elevated, aligned, and stretched, leading to the formation of the tube-like structures. Conclusions The capillary differentiation of endothelial cells induced by microgrooved PDMS substrates coated with fibronectin in the absence of angiogenic growth factors is demonstrated here. We show for the first time that besides biochemical, biomechanical, and 2D geometric properties, the 3D topographic features also influence fundamental cell-matrix interactions such that capillary differentiation of endothelial cells can be induced by topographic cues alone. The differentiation depends on the dimension of the grooves. Cord-like structures on top of a confluent monolayer of cells and a network of tube-like structures were identified at different stages of cell culture, with lumens inside the tube-like structures verified by confocal microscopy. Further work is needed to elucidate the mechanism of the capillary differentiation of endothelial cells observed. However, there are four factors that are implicated in this mechanism: mass transport limitations at the bottom of the grooves; the potent effect of the grooved structures for cell
Letters alignment; elevation of the cells at the bottom of the grooves; and multicellular retraction. Our observations suggest that micronmeter-scale featured devices could be designed to direct the angiogenic integration of medical implants and tissue engineering constructs. Acknowledgment. This work is supported by NSF EEC9529161 through the University of Washington Engineered Biomaterials (UWEB)-Engineering Research Center and NSF CAREER Award (CTS-0092699). We thank the UWEB Optical Microscopy and Image Analysis Shared Resource for help with optical microscopy and the Center for Nanotechnology at the University of Washington for help with photolithography and confocal microscopy. References and Notes (1) Arnold, F.; West, D. Pharmacol. Ther. 1991, 52, 407-422. (2) Folkman, J.; Shing, Y. Angiogenesis. J. Biol. Chem. 1992, 267, 10931-10934. (3) Jain, R.; Au, P.; Tam, J.; Duda, D.; Fukumura, D. Nat. Biotechnol. 2005, 23, 821-823. (4) Folkman, J.; Haudenschild, C. Nature 1980, 288, 551-556. (5) Folkman, J.; Klagsbrun, M. Science 1987, 235, 442-447. (6) Feder, J.; Marasa, J.; Olander, J. J. Cell. Physiol. 1983, 116, 1-6. (7) Vernon, R.; Lara, S.; Drake, C.; Iruela-Arispe, M.; Angello, J.; Little, C.; Wight, T.; Sage, E. In Vitro Cell DeV. Biol. 1995, 31, 120-131. (8) Vailhe, B.; Ronot, X.; Tracqui, P.; Usson, Y.; Tranqui, L. In Vitro Cell. DeV. Biol.: Anim. 1997, 33, 763-773. (9) Madri, J.; Williams, S. J. Cell Biol. 1983, 98, 153-165. (10) Chalupowicz, D.; Chowdhury, Z.; Bach, T.; Barsigian, C.; Martinez, J. J. Cell Biol. 1995, 130, 207-215. (11) Gamble, J.; Matthias, L.; Meyer, G.; Kaur, P.; Russ, G.; Faull, R.; Berndt, M.; Vadas, M. J. Cell Biol. 1993, 121, 931-943. (12) Ingber, D.; Folkman, J. J. Cell Biol. 1989, 109, 317-330. (13) Dike, L.; Chen, C.; Mrksich, M.; Tien, J.; Whitesides, G.; Ingber, D. In Vitro Cell DeV. Biol. 1999, 35, 441-448. (14) Carlos, C.; Wang, Y.; Ho, C. J. Am. Chem. Soc. 2005, 127, 15981599. (15) Flemming, R.; Murphy, C.; Abrams, G.; Goodman, S.; Nealey, P. Biomaterials 1999, 20, 573-588. (16) Curtis, A.; Wilkinson, C. Biomaterials 1997, 18, 1573-1583. (17) Lampin, M.; Warocquier-Clerout, R.; Legris, C.; Degrange, M.; Sigot-Luizard, M. J. Biomed. Mater. Res. 1997, 36, 99-108. (18) Groessner-Screiber, B.; Tuan, R. J. Cell Sci. 1992, 101, 209-217. (19) Dunn, G.; Brown, A. J. Cell Sci. 1986, 83, 313-340. (20) Wojciak-Stothard, B.; Madeja, Z.; Korohoda, W.; Curtis, A.; Wilkinson, C. Cell Biol. Int. 1995, 19, 485-490. (21) Meyle, J.; Gultig, K.; Brich, M.; Hammerle, H.; Nisch, W. J. Mater. Sci. Mater. Med. 1994, 5, 463-466. (22) Liu, L.; Chen, S.; Giachelli, C.; Ratner, B.; Jiang, S. J. Biomed. Mater. Res. 2005, 74A, 23-31. (23) Iruela-Arispe, M.; Hasselaar, P.; Sage, E. Lab. InVest. 1991, 64, 174-186.
