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Liquid Crystalline Collagen: A Self-Assembled Morphology for the Orientation of Mammalian Cells John E. Kirkwood and Gerald G. Fuller* Department of Chemical Engineering, Stanford UniVersity, Stanford, California 94305 ReceiVed NoVember 11, 2008. ReVised Manuscript ReceiVed January 4, 2009 We report the creation of collagen films having a cholesteric banding structure with an orientation that can be systematically controlled. The action of hydrodynamic flow and rapid desiccation was used to influence the orientation of collagen fibrils, producing a film with a twisted plywood architecture. Adult human fibroblasts cultured on these substrates orient in the direction of the flow deposition, and filopodia are extended onto individual bands. Atomic force microscopy reveals the assembly of 30 nm collagen fibrils into the uniform cholesteric collagen films with a periodic surface relief. The generation of collagen with a reticular, “basket-weave” morphology when using lower concentrations is also discussed.
Introduction Biopolymers play a central role in the drive to create materials that mimic natural production to provide innovative personal and medical care products. There is a need for organized, threedimensional structures for tissue replacement and biomemetic material assembly. Creating these complex material formats requires control over cellular function and mobility. Collagen is the most prevalent protein in the human body and is found in connective tissues and load-bearing structures such as bone and teeth, making it a premier candidate for this purpose. Collagen has found wide use in cosmetic surgery, as a constituent in artificial skin and bone implants and is a common ingredient in personal care products. There is a wide selection of materials with partial or full collagen content on the market for cell culture and tissue engineering applications, but few of them consist of oriented collagen structures necessary for directional control of cell mobility. The collagen molecule is a rod about 300 nm in length and 1.5 nm in diameter formed from three polypeptide strands wound in a left-handed triple helix. This relatively stiff, filament-shaped molecule can self-assemble into strong fibrous bundles with the high tensile strength derived from its triple helical structure. This molecule is distinctive in the regular pattern of amino acids that are arranged to form each strand and is characterized by a typical sequence of Gly-X-Pro or Gly-X-Hyp, where X can be any of the amino acids commonly found in proteins. Collagen has the remarkable ability of organizing into a hierarchy of fibrils with diameters up to several hundred nanometers and complex suprafibrillar, liquid-crystal-like architectures.1 Molecular solutions of this rodlike protein exhibit the behavior of a lyotropic liquid crystal with a spontaneous phase transition from an isotropic to crystalline state as the concentration is increased. The chirality of the collagen molecule creates a rotation in the director of the nematic state, forming a twisted nematic, cholesteric state.2 The composites in nature that appear geometrically similar to liquid crystalline geometries are known as “biological analogues” or twisted plywood architectures.2,3 In this case, the collagen fibrils * To whom correspondence should be addressed. Phone: (650) 723-9243. E-mail:
[email protected]. (1) Hulmes, D. J. S. J. Struct. Biol. 2002, 137, 2–10. (2) Neville, A. C. Biology of Fibrous Composites: Cambridge University Press: Cambridge, U.K., 1993. (3) Neville, A. C. Tissue Cell 1988, 20, 133–143.
are aligned in a way geometrically similar to the arrangement of molecules in a cholesteric liquid crystal but have lost their viscous character.4 This lyotropic phase transition offers the possibility of creating liquid crystalline materials with collagen. Indeed, it is possible to produce nematic, precholesteric, and cholesteric phases at concentrations above approximately 20 mg/ mL.5-9 Liquid crystalline patterns that can be created with collagen have been hypothesized as the source of many similar patterns that are observed in tissue,5,10,1 suggesting that these tissues are also created with a liquid crystalline assembly process in vivo. Martin et al.11 have hypothesized that the suprafibrillar tissue architecture is determined by the ability of soluble precursor molecules to form liquid crystalline arrays, prior to fibril assembly. In another study, they have also demonstrated the ability to stabilize the chiral fluid phase of collagen by increasing the pH through ammonia vapor gelation. The gel that is produced in this way maintains the helical symmetry of the fluid counterpart and is structurally similar to collagen matrixes built by various cell types in vivo.8,12 Understanding the organization and cellular deposition of these complex biological structures is an important step in creating materials to replace or repair them. The ability to direct cellular growth through physical, contact guidance13-15 or chemical means16-19 is desirable in many (4) Cowin, S. C. Meccanica 1999, 34, 77–102. (5) Giraud-Guille, M. J. Biomech. 2003, 36, 1571–1579. (6) Giraud-Guille, M. M. J. Mol. Biol. 1992, 224, 861–873. (7) Giraud-Guille, M. M.; Besseau, L.; Chopin, C.; Durand, P.; Herbage, D. Biomaterials 2000, 21, 899–906. (8) Belamie, E.; Mosser, G.; Gobeaux, F.; Giraud-Guille, M. M. J. Phys.: Condens. Matter 2006, 18, 115–129. (9) Mosser, G.; Anglo, A.; Healry, C.; Bouligand, Y.; Giraud-Guille, M. M. Matrix Biol. 2006, 25, 3–13. (10) Cowin, S. J. Non-Newtonian Fluid Mech. 2004, 119, 155–162. (11) Martin, R.; Farjanel, J.; Eichenberger, B.; Colige, A.; Kessler, E.; Hulmes, D. J. S.; Giraud-Guille, M. M. J. Mol. Biol. 2000, 301, 11–17. (12) Giraud-Guille, M. M.; Belamie, E.; Mosser, G.; Helary, C.; Gobeaux, F.; Vigier, S. C. R. Chim. 2008, 11, 245–252. (13) Oakley, C.; Brunette, D. M. J. Cell Sci. 1993, 106, 343–354. (14) Sniadecki, N. J.; Desai, R. A.; Ruiz, S. A.; Chen, C. S. Ann. Biomed. Eng. 2006, 34, 59–74. (15) Vernon, R. B.; Gooden, M. D.; Lara, S. L.; Wight, T. N. Biomaterials 2005, 26, 3131–3140. (16) Beningo, K. A.; Dembo, M.; Wang, Y. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 52. (17) von Philipsborn, A. C.; Lang, S.; Bernard, A.; Loeschinger, J.; David, C.; Lehnert, D.; Bastmeyer, M.; Bonhoeffer, F. Nat. Protoc. 2006, 1, 1322–1328. (18) Jiang, X.; Buzewicx, D. A.; Wong, A. P.; Piel, M.; Whitesides, G. M. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 975–978.
10.1021/la803736x CCC: $40.75 2009 American Chemical Society Published on Web 02/10/2009
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situations from wound repair to biomedical device design. In nature, collagen is a guide for cell growth in the extracellular matrix.20,21 Cells are observed to grow parallel to collagen fibers that form the framework for aligned tissues. Methods to create anisotropic films of collagen have received a great deal of attention because of the ability of aligned collagen to orient the growth of cells. A review of the literature reveals that anisotropic films and gels of collagen fibrils have been produced using several methods such as dip-pen nanolithography,22 reverse dialysis,23 high-strength magnetic field orientation,24-26 surface-modified magnetic beads,27 and electrospinning of a fibrous mat onto a spinning disk.28 The use of nonfibrillar collagen has been given a lot of attention by Giraud-Guille and co-workers,6,9,29,30 who have studied the formation of liquid crystalline textures by collagen at an air interface. The method employed by this group involves the continuous injection of collagen solutions into small microchambers, concentrating the collagen as the solvent evaporates, which subsequently produces liquid crystalline textures at the interface with air. The response of mammalian cells to these liquid crystalline collagen films has not been reported, and to further connect observations on the natural ordering of collagen, a few pertinent questions must be answered. Can the cholesteric liquid-crystal-like states of collagen induce the contact guidance of cells? Can a methodology be developed to control the orientation of the liquid crystalline textures? In this paper, we describe the generation of highly uniform, cholesteric liquid-crystal-like collagen films that are capable of inducing contact guidance in adult human fibroblasts. The fibroblasts are guided along individual cholesteric bands, parallel to the direction of flow. The technique is capable of creating thin films on different substrates such as glass and rubber.
Materials and Methods Rat tail collagen, type I (BD Biosciences) was purchased at stock concentrations of 3.6 and 10 mg/mL in 0.02 N acetic acid (pH ≈ 3.5). The 10 mg/mL solution was then dialyzed against polyethylene glycol (Fluka) for 20 min at 4 °C until the solution reached a final concentration of approximately 20 mg/mL. The substrate used was silica glass (Fisher) cleaned by sonication (30 min) in a 1.5% Deconex 12-PA (Technotrade) cleaning solution at 60 °C, rinsed with copious amounts of deionized water (Millipore Direct-Q 5), and stored in a clean environment. The silica glass was alternately cleaned by plasma treatment using a plasma cleaner (Gala Instrumente, prep 5) on 50% power for 5 min. Collagen films were created using a custom deposition system consisting of a three-axis robotic arm (I&J Fisnar 500LN) and a syringe pump (Harvard Apparatus, milliliter OEM pump). The design of the apparatus allowed the writing of the collagen solution onto a variety of substrates by programming the robotic arm to follow (19) Thakkar, R. G.; Ho, F.; Ngan, F. H.; Liepmann, D.; Li, S. Biochem. Biophys. Res. Commun. 2003, 307, 883–890. (20) Mutsaers, S. E.; Bishop, J. E.; McGrouther, G.; Laurent, G. J. Int. J. Biochem. Cell Biol. 1997, 29, 5–17. (21) Leitinger, B.; Hohenester, E. Matrix Biol. 2007, 26, 146–155. (22) Wilson, D. L.; Martin, R.; Hong, S.; Cronin-Golomb, M.; Mirkin, C. A.; Kaplan, D. L. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 13660–13664. (23) Knight, D. P.; Nash, L.; Hu, X. W.; Haffegee, H.; Ho, M.-W. J. Biomed. Mater. Res. 1998, 41, 185–191. (24) Torbet, J.; Malbouyres, M.; Builles, N.; Justin, V.; Roulet, M.; Damour, O.; Oldberg, A.; Ruggiero, F.; Hulmes, D. J. S. Biomaterials 2007, 28, 4268– 4276. (25) Dickenson, R. B.; Guido, S.; Tranquillo, R. T. Ann. Biomed. Eng. 1994, 22, 342–356. (26) Rosner, B. I.; Siegel, R. A.; Grosberg, A.; Tranquillo, R. T. Ann. Biomed. Eng. 2003, 31, 1383–1401. (27) Guo, C.; Kaufman, L. J. Biomaterials 2007, 28, 1105–1114. (28) Zhong, S.; Teo, W. E.; Zhu, X.; Beuerman, R. W.; Ramakrishna, S.; Yung, L. Y. L. J. Biomed. Mater. Res., A 2006, 79A, 456–463. (29) Besseau, L.; Giraud-Guille, M. M. J. Mol. Biol. 1995, 251, 197–202. (30) Giraud-Guille, M. M. Mol. Cryst. Liq. Cryst. 1987, 153, 15–30.
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Figure 1. Schematic of robotic deposition onto a glass slide. The syringe tip is curved parallel to the glass surface. Fluid exits the orifice with velocity in the direction opposite that of the syringe movement.
prescribed paths on the surface. The arm of the robot supports a disposable syringe needle connected to an external syringe pump, which dispenses the solutions. The syringe tip is curved so that the exiting fluid is ejected parallel to the target surface, Figure 1. The opposing directions of fluid and robotic movement create an extensional flow component in the fluid exiting the syringe. The extensional flow due to the relative motion of the syringe tip and the substrate, coupled with the pressure-driven flow out of the syringe, provides significant velocity gradients to orient the collagen molecules during deposition. The draw ratio, the ratio of the substrate velocity to the fluid velocity out of the syringe nozzle, can be calculated to compare the relative effect of altering the flow rate or robot velocity. The deposition speed of the robot, Vr, was adjusted between 20 to 100 mm/s. The syringe needle orifice size could also be adjusted by using different needle sizes, 18-27 gauge (inner diameter of 0.84-0.19 mm). The flow rate of collagen used was in the range of 0.05-0.5 mL/min. The 22 gauge needle was used with the highest frequency and produced the most consistent films for the viscosity of the highly concentrated collagen solutions. A 22 gauge needle at a flow rate of 0.3 mL/min and robot speed of 100 mm/s produces a draw ratio of 9.7. Prior to use, an aliquot of the collagen solution was sonicated at 4 °C to reduce the number of collagen aggregates, and this procedure has been shown not to destroy the triple helical nature of the molecule.31,32 Sonication was carried out according to the procedure used by Giraud-Guille33 with two 10 min pulses of sonication and a 10 min rest between the pulses. The collagen solution was deposited under controlled flow conditions onto the glass substrates and allowed to dry under ambient conditions. Acidic conditions are maintained in the solution throughout the entire procedure to prevent the formation of fibrils and fibers. As desiccation of the solution occurs, the concentration of the solution increases while the area of contact remains the same. This rapid volume reduction occurs in less than 15 min. The entire apparatus was in a laminar flow hood to maintain sterility for cell culture. After drying, the samples were examined between crossed polarizers on an optical microscope (Nikon TE300). Human fibroblasts (ATCC CRL-2091) were cultured on substrates in DMEM medium. The medium was supplemented with 0.1% FBS, 0.01% 100× penicillin/streptavidin, 0.01% 100× glutamine, 0.01% MEM nonessential amino acids, and 0.01% 100× sodium pyruvate (DMEM medium and supplements purchased from Invitrogen). Cells were plated at a density of approximately 10 000 cells/mL and grown for 12-48 h at 37 °C with 5% CO2. The cells were fixed in a solution of 10% formaldehyde (Sigma-Aldrich) in 1× phosphate-buffered saline (PBS; Gibco) for 10 min. Images were captured with a 10× phase contrast objective using a Nikon TE300 microscope. Atomic force microscopy was performed on the collagen films using a Veeco MultiMode atomic force microscope in tapping mode using Nanosensors tips (PPP-BSI) with a nominal force constant of 0.1 N/m and resonance frequency of 28 kHz or Tap300Al tips (Budget Sensors Tap300Al) with a nominal force constant of 40 mN/m and (31) Nishihara, T.; Doty, P. Proc. Natl. Acad. Sci. U.S.A. 1958, 44, 411–417. (32) Hodge, A. J.; Schmitt, F. O. Proc. Natl. Acad. Sci. U.S.A. 1958, 44, 418–424. (33) Besseau, L.; Coulomb, B.; Lebreton-Decoster, C.; Giraud-Guille, M. M. J. Biomech. 2002, 23, 27–36.
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Figure 2. Schematic representation of the collagen films on a glass slide (left). Cholesteric banding morphology of the generated collagen film viewed under crossed polarizers at 40× magnification (right).
resonance frequency of 300 kHz. Scanning was performed at a frequency of 0.5-2 Hz. For fluorescent imaging of the cellular actin fibers, the cells were stained with Alex Fluor 488 phalloidin (Invitrogen). The staining protocol is as follows. The growth medium is aspirated, and the plates are washed one time in PBS and fixed with 3% formaldehyde in PBS for 10 min. The cells are permeabilized with a solution of 1% Triton (Sigma-Aldrich) in PBS for 10 min and washed twice with a solution of 0.1% Triton in PBS. Blocking buffer, composed of 5% horse serum (Gibco) and 0.1% Triton in 1× PBS, is then added to the plates for 20 min. The fluorescent dye is dispersed in blocking buffer and set on the plates for 1 h. The plates are washed three times in a 0.1% Triton in PBS solution, mounted with Vectashield (Vector Laboratories) and a glass coverslip, and stored in the dark at -20 °C until use. Fluorescent dyes are excited with a Mercury lamp on a Nikon Microphot and Pentamax cooled CCD camera and images recorded with Metamorph software.
Results and Discussion The synergetic effects of hydrodynamic forces and rapid desiccation of a drawn solution of concentrated, nonfibrillar collagen produces a film with the banding of a cholesteric liquid crystal. These cholesteric bands are aligned with the long axis parallel to the flow direction. In Figure 2, we show one such film in a polarizing light microscope. In this particular example, a solution with a concentration of 20 g/mL was deposited using a draw ratio of approximately 3. Only a portion of the line of collagen drawn onto the glass substrate is shown here, and the well-oriented, parallel banding structure persisted along the entire length of the deposition path (50 mm). This pattern of banding structure is in distinct contrast to the common phenomenon of shear banding that arises following the cessation of shear flow of many polymeric liquid crystals. In this latter case, the bands are formed perpendicular to the flow direction, whereas the present results produced bands parallel to the direction of deposition.34,35 A birefringent image of the collagen stripe is shown in Figure 3, where two sections of collagen in the cholesteric liquid crystal phase (C) are separated by a region in the center of the stripe where collagen is isotropically oriented (I) outlined by the dotted lines. Immediately following deposition, the contact line (solid lines) of the collagen solution with the glass is pinned. As desiccation of the solution progresses, we observe the contact line to move toward the center of the stripe and a thin film of collagen is deposited in the cholesteric liquid crystalline state. The contact line of the collagen solution and glass provides a uniform constraining layer for the deposition of the collagen.2 (34) Picken, S. J.; Moldenaers, P.; Berghmans, S.; Mewis, J. Macromolecules 1992, 25, 4759–4767. (35) Olmsted, P. D. Rheol. Acta 2008, 47, 283–300.
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Figure 3. Picture of a single collagen stripe. The stripe has two separate morphologies of collagen present, cholesteric liquid crystal domains (C) and isotropic domains (I). The cholesteric domains persist from the contact line with the glass (solid line) to the center of the stripe (dotted line), and band formation is parallel to the deposition path. The isotropic domain in the center of the stripe (dotted lines) is a result of a lower concentration phase of collagen.
Figure 4. Cholesteric banding of a collagen film observed through crossed polarizers at 100× magnification. The director rotates as one moves across the banding structure.
This boundary condition was found necessary for the production of uniform liquid crystalline phases in numerical simulation of chiral liquid crystals.36,37 In contrast to the birefringence of the cholesteric phase, the isotropic region of collagen in the center of the stripe does not appear birefringent under the light microscope. The presence of an isotropic region in the center of the stripe suggests that the molecular orientation imparted by the flow has relaxed before cholesteric banding could develop. The boundary condition along with the deposition technique provides a methodology for the formation of a monodomain liquid crystalline structure. The banding structure indicates the presence of cholesteric twisting of the director prescribing local, molecular orientation in the collagen materials. As one moves across the bands, the director rotates with the cholesteric pitch of the material, the distance over which the molecule undergoes a full rotation, and light transmission through crossed polarizers alternates between bright and dark stripes, Figure 4. The helical half-pitch of this structure is ∼3.5 µm. Recently, Mosser et al.9 were able to produce small regions of cholesteric banding by continuous injection and desiccation of highly concentrated collagen into a glass microchamber over long time scales (3 weeks to 3 months). However, the result was a heterogeneous collection of liquid crystalline organizations from spherulites to banded structures of varying pitches. In the process described here, the solutions are subjected to a sequence of events that allow for successful manipulation of (36) De Luca, G.; Rey, A. D. Mater. Res. Soc. Symp. Proc. 2003, 735, 7.4.17.4.6. (37) De Luca, G.; Rey, A. Phys. ReV. E 2004, 69, 011706.
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Figure 5. Height mode atomic force microscopy images of collagen cholesteric banding structures at decreasing length scales. The direction of deposition was from the bottom to the top of the picture on cleaned silica glass. Key: (A) 60 µm × 60 µm image, 350 nm height scale, (B) 20 µm × 20 µm image, 350 nm height scale, scan of selected area in image (A), (C) 10 µm × 10 µm, 250 nm height scale (the director rotates from the upper left to the upper right across the image with arrows shown as guides), (D) 3 µm × 3 µm image, 75 nm height scale.
the orientation of the bands. The initial 20 mg/mL solutions, although quite concentrated, did not produce evidence of cholesteric banding. Birefringence prior to desiccation was not observed in fluid solutions with concentrations as high as 50 mg/mL. At sufficiently high draw ratios, the initial flow deposition causes unidirectional orientation of the rodlike, collagen chains. This would most likely occur even if the material was initially in a cholesteric state because of the ability of velocity gradients within a flow to overcome cholesteric rotations of the director.38,39 The deposited film undergoes two rate processes that ultimately lead to the observed banding structure. In the absence of hydrodynamic forces, orientation of the chains in the deposited films will relax. This will occur concurrently with desiccation of the film, which will concentrate the collagen and induce an isotropic-to-cholesteric phase transition. If the time scale for desiccation is shorter than the orientational relaxation time scale, highly oriented, cholesteric bands will result. Measuring the topology of the cholesteric banding structure with the aid of an atomic force microscope (Figure 5) reveals the striking self-assembly of the collagen molecules into a higher (38) Rey, A. D. Phys. ReV. E 1996, 53, 4198–4201. (39) Porter, R. S.; Barrall, E. M.; Johnson, J. F. J. Chem. Phys. 1966, 45, 1452–1456.
order structure. The bands are composed of organized groups of small fibrils approximately 30 nm in diameter. These 30 nm fibrils can be precursors to higher order fibril and fiber structures. However, the use of a low-pH solution arrests the fiber assembly in this study. Within Figure 5A,B, individual bands display a braided texture along their length. These textures are oscillations in height that occur on length scales on the order of the individual band widths. At this time, the origin of these features is not known. The fibril arrangement within the banding structure is highly organized (Figure 5C), and inspection of the images reveals that they are consistent with that of a cholesteric liquid crystal with the director orientation rotating ∼45° between individual bands. The rotation across the banding is equivalent to previous reports of liquid crystalline collagen (see Figure 17 in ref 6, Figure 1d in ref 8, and Figure 1 in ref 9). In Figure 5D, an AFM image of a single band is shown; the fibrils appear tightly packed with sporadic defects visible on the surface as dark ellipses. The individual fibrils lack the 67 nm D-periodicity found on fibers formed under neutral pH.40-42 This is a consequence of the acidic conditions of the deposition procedure. The formation of collagen fibrils without the banding has been shown in ref 43 to be a result of the electrolytes present in solution. More specifically, potassium
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Figure 6. Sample depth analysis measurement of cholesteric banding shown using an AFM height mode image (right) and a surface profile (left). Red arrows signify the center points of each band, and the height profile displays a 100 nm vertical height difference between these two points.
ions are needed in solution for the banding structure to form although the mechanism for this dependence is unclear. The height profile of the collagen banding area was analyzed using height mode images from the atomic force microscope. The height profile across a large region of the film is not flat as one might expect. Instead, the profile has periodic undulations in height with peaks and troughs registering with the centers of individual bands. Height undulations between individual bands were found to be on the order of 150 nm in depth, an order of magnitude smaller than the 3500 nm half-pitch of the cholesteric band. A sample height calculation is given in Figure 6, where a cross section of adjacent cholesteric bands is shown. The depth of the lower band with respect to its neighbors is calculated by selecting the center point of each band. Each band is constructed of a specific orientation of collagen fibrils with the neighboring band composed of fibrils with an orientation rotated by 45°. For the bands with the lowest height, the collagen molecules are oriented orthogonal to the long axis of the banding structure. These height differences may be created by modulations in the surface energy at the solution-air interface and influenced by the anchoring and chirality of the molecule. This was discussed by Meister et al.44 for a small cyclic siloxane oligomer which was much smaller than the protein in this work. The cholesteric organization of the oligomer displayed a half-pitch of 145 nm and periodic surface relief of 1 nm. This periodic relief was also seen by Maeda45,46 working with evaporating droplets of collagen (mixture of type I and type III). The drops produced concentric rings of alternating collagen molecular orientation visible under a polarizing optical microscope. The width of each ring ranged from 3 to 120 µm depending on the initial concentration (5-19 mg/mL). The ring width also varied with the distance from the center of the drop, with the width of each ring increasing as the distance decreased.45 This variation and nonuniformity of the ring width across the drop are distinctly different from the uniform liquid crystalline structures reported previously (see Figure 5f in ref 9 and Figure 9 in ref 30). We observe uniform structures to form upon desiccation with a regular periodicity described by the pitch of the cholesteric band. (40) Cisneros, D. A.; Hung, C.; Franz, C. M.; Muller, D. J. J. Struct. Biol. 2006, 154, 232–245. (41) Bozec, L.; Horton, M. Biophys. J. 2005, 88, 4223. (42) Gutsmann, T.; Fantner, G. E.; Venturoni, M.; Ekani-Nkodo, A.; Thompson, J. B.; Kindt, J. H.; Morse, D. E.; Fygenson, D. K.; Hansma, P. K. Biophys. J. 2003, 84, 2593–2598. (43) Jiang, F.; Horber, H.; Howard, J.; Muller, D. J. J. Struct. Biol. 2004, 148, 268–278. (44) Meister, R.; Dumoulin, H.; Halle, M. A.; Pieranski, P. J. Phys. II 1996, 6, 827–844. (45) Maeda, H. Langmuir 1999, 15, 8505–8513. (46) Maeda, H. Langmuir 2000, 16, 9977–9982.
In the context of a material coating, the surface profile of the collagen film resembles a series of grooves and ridges with a periodicity of the helical half-pitch of the cholesteric band (see Figure 10, which is published as Supporting Information). This self-assembled topology extends the length of the cholesteric structure, producing long-range ordering. This topology is reminiscent of substrates produced using lithographic techniques, where grooves and ridges are physically etched onto silicon or rubber substrates.14,47,48 The architecture of the collagen film in the cholesteric state is uniform and organized. However, a different structure of collagen is produced either when the initial concentration is too low for the isotropic-cholesteric phase transition to occur or if the flow-imparted molecular orientation relaxes before complete desiccation. We found that depositing collagen at a concentration of 10 mg/mL was inadequate for the production of cholesteric banding. The films do not appear birefringent when observed under crossed polarizers. Analyzing the film with the atomic force microscope reveals a reticular organization of the collagen fibrils, Figure 7. This basket weave of collagen fibrils is highly packed, with aggregated bundles separated by pits and crevices. The reticular formation provides a unique two-dimensional surface for mammalian cell growth. The surface has a very low porosity due to the highly interconnected bundles of small fibrils. The deposition of cells to cholesteric collagen films is shown in Figure 8 with adult human fibroblasts cultured for 24 h. The bodies of the fibroblasts stretch out and become spindle-like with their long axes in the direction of the banding. The anisotropic growth of the fibroblasts demonstrates the ability of the cholesteric banding architecture to induce the contact guidance of the fibroblast cells. While the idea of contact guidance was first proposed in 1934,49 to our knowledge there have not been reports on the guidance of fibroblasts by liquid crystalline banding structures. The liquid crystalline collagen structure has two physical features that are cues for contact guidance, the height difference between bands and the orientations of the individual collagen fibrils. To investigate the contact guidance cues directing cellular motility, the fibroblast cells were fluorescently stained with phalloidin, illuminating the actin fibers in the cell body (a fluorescent image of the stress fiber alignment in a polarized fibroblast is given in Figure 11 of the Supporting Information). When the fibroblast cell first attaches to the surface, it explores its surroundings by sending out extensions in every direction. (47) Loesberg, W. A.; Riet, J.; te van Delft, F. C. M. J. M.; Schon, P.; Figdor, C. G.; Speller, S.; van Loon, J. J. W. A.; Walboomers, X. F.; Jansen, J. A. Biomaterials 2007, 28, 3944–3951. (48) Teixeira, A. I.; Abrams, G. A.; Bertics, P. J.; Murphy, C. J.; Nealey, P. F. J. Cell Sci. 2003, 116, 1881–1892. (49) Weiss, P. J. J. Exp. Zool. 1934, 68, 393–448.
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Figure 7. AFM height mode image of reticular formation of collagen fibrils (10 µm × 10 µm image, 250 nm height scale). The collagen film was formed with a lower initial concentration (10 mg/mL) and is constructed of a basket weave of fibers without a specific orientation.
Figure 8. Adult human fibroblasts after 24 h in culture on a cholesteric collagen film (phase contrast microscopy, 10× magnification). The cells are polarized parallel to the banding structure of collagen.
These filopodial extensions are used to sense the topographical and chemical features surrounding the cell.50-52 Once the filopodia of the cell find a suitable feature, they establish focal adhesion contacts (groupings of integrin molecules) and deposit extracellular matrix material. The fibroblast then forms lamellipodia, which move the cell to the site of the feature. It has also been shown that a single filopodial contact can direct the motion of the entire cell,53 suggesting that only a single contact on the banding structure is needed to draw the cell into alignment. We observe the fibroblasts grown on the cholesteric banding structure to not only orient parallel to the bands but also extend filopodia onto individual bands while the bulk of the cell body may or may not be contained on a single band. The size of each fibroblast is greater than the width of each band, and individual cells are observed to follow the guidance of multiple bands. The (50) Dalby, M. J.; Riehle, M. O.; Johnstone, H.; Affrossman, S.; Curtis, A. S. G. Cell Biol. Int. 2004, 28, 229–236. (51) Iijima, M.; Devreotes, P. Cell 2002, 109, 599–610. (52) Walboomers, X. F.; Ginsel, L. A.; Jansen, J. A. J. Biomed. Mater. Res. 2000, 51, 529–534. (53) O’Connor, T. P.; Duerr, J. S.; Bentley, D. J. Neurosci. 1990, 10, 3935– 3946.
Figure 9. (A) Fluorescence image of three adult human fibroblasts on a collagen film with cholesteric banding (40× magnification). The colors of the image are inverted to enhance contrast; the cells appear dark. Cholesteric banding of collagen is visible as dark striations across the image from left to right. Growth extensions of the fibroblasts follow the cholesteric banding. (B) Fluorescent image superimposed with background under low light.
fluorescently labeled fibroblasts (Figure 9; see also Figure 12, which is given as Supporting Information) are superimposed with the cholesteric banding in the background. The pictures were produced by simultaneously fluorescently illuminating cells stained with phalloidin and illuminating the background with white light. The excited fluorophores appear white, while a very low intensity white light illuminates the cholesteric banding in the background. The banding appears in the picture as an artifact to the low-intensity light used, enhancing image contrast. The inversion of fluorescence in Figure 9A creates the illusion of dark cells on the background. Inspection of these cells provides clear evidence that the filopodia extend preferentially onto the bands that appear darker on the light microscope, suggesting that these bands provide stronger guidance cues over the neighboring bands. Comparing the fluorescent images of Figure 9 with the atomic force micrographs of the cholesteric collagen structure given
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earlier (Figure 5), we were able to determine that the darker bands are composed of collagen oriented orthogonal to the direction of cell growth. The width of the darker bands on the light microscope is approximately 1.5 µm, registering with the size of the bands of lower height from the AFM height mode images. From the topographical analysis, we also found that these “darker” cholesteric bands are approximately 150 nm lower in height than neighboring bands. This depression creates a channel in the surface capable of guiding cell growth. The composition of collagen in the channel is orthogonal to the direction of filopodial extension, signifying that the topographical depression is guiding the cell growth and not the presence of the oriented collagen fibrils. The 150 nm channel depth should be sufficient to induce contact guidance as it is larger than the lower topographical limit of 35 nm found by Loesberg et al.47 but smaller than the 1000 nm limit found in ref 15. Moreover, previous work in the literature has shown that the filopodia of the cell can respond to topographical features as small as 10 nm in size.50,54,55
Conclusions Highly concentrated collagen solubilized in acidic acid was drawn onto glass substrates and quickly desiccated. The deposition process induces a combination of shear and extensional flow to orient the monomers of collagen in solution. During the desiccation process the molecular solution assembles into cholesteric liquid crystalline structures, leaving a thin collagen film. A defect-free cholesteric banding structure was found to persist along the length of the contact line of the deposited solution, parallel to the flow deposition. The surface relief of the cholesteric banding was found to create periodic undulations of ∼150 nm with a helical half-pitch of ∼3.5 µm. These topographical features are able to induce the contact guidance of adult human fibroblasts. Actin fibers of fibroblasts were fluorescently stained, and filopodial extensions were found to track individual cholesteric bands and align the actin fibers within the cell parallel to the (54) Dalby, M. J.; Childs, S.; Riehle, M. O.; Johnstone, H. J. H.; Affrossman, S.; Curtis, A. S. G. Biomaterials 2003, 24, 927–935. (55) Berry, C. C.; Dalby, M. J.; McCloy, D.; Affrossman, S. Biomaterials 2005, 26, 24.
Kirkwood and Fuller
banding. The cells follow the collagen around regions where the bands have a radius of curvature, suggesting an ability to not only guide cells in a planar manner but guide them through a wide variety of written designs. In this deposition process, the individual fibrils are formed under acidic conditions and lack the 67 nm D-periodicity characteristic of collagen fibers formed in physiological buffers. Poole et al.56 have reported that this periodicity is a key component of collagen-induced contact guidance and necessary for directed cellular growth. This result suggests that the orientation capability of the cholesteric banding structure could be enhanced if the fibrils possessed this periodicity possibly by adjusting the initial pH of the collagen solution. The self-assembled collagen scaffolding offers a unique mechanism for contact guidance and biomaterial tissue production. The generated collagen scaffolding can be deposited on a variety of substrates and could be used as a coating on materials ranging from metals to rubber. It could also provide an experimental pathway to the understanding of the assembly process of cholesteric phases in nature. We also observe a strong interplay between concentration and relaxation during the deposition procedure. A reticular morphology is observed when the concentration is below the nematiccholesteric transition point. This reticular morphology is reminiscent of the collagen organization in the human dermis. One potential application for this film is the creation of a substrate to promote extracellular matrix deposition that models the original tissue. Acknowledgment. We thank Michael Paukshuto for his helpful discussions at the start of the project and Jaykumar Rajadas for assisting with the mammalian cell culture. Partial funding for the project was provided by the Center on Polymer Interfaces and Macromolecular Assemblies (CPIMA) at Stanford University. Supporting Information Available: Three figures providing additional images from the AFM and cell growth experiments. This material is available free of charge via the Internet at http://pubs.acs.org. LA803736X (56) Poole, K.; Khairy, K.; Friedrichs, J.; Franz, C.; Cisneros, D. A.; Howard, J.; Mueller, D. J. Mol. Biol. 2005, 349, 380–386.
