Article pubs.acs.org/Biomac
Microfabrication of Custom Collagen Structures Capable of Guiding Cell Morphology and Alignment Eun-A Kwak, Suji Ahn, and Justyn Jaworski* Department of Chemical Engineering and Institute of Nanoscience and Technology, Hanyang University, 222 Wangsimni-ro, Seongdong-gu, Seoul 133-791, South Korea S Supporting Information *
ABSTRACT: The patterning of biological components into structural analogues of native tissues to simulate an environment for directing cell growth is one important strategy in biomaterials fabrication. It is widely accepted that chemical, mechanical, and topological cues from the extracellular matrix (ECM) provide important signals for guiding cells to exhibit characteristic polarity, orientation, and morphology. To fully understand the delicate relationship between cell behavior and ECM features, biomaterials fabrication requires improved techniques for tailoring nano/microstructured patterns from relevant biological building blocks rather than using nonbiological materials. Here we reveal a unique approach for the nano/microfabrication of custom patterned biomaterials using collagen as the sole building material. With this new fabrication technique, we further revealed that custom collagen patterns could direct the orientation and morphology of fibroblast growth as a function of vertex density and pattern spacing. Our findings suggest that this technique may be readily adopted for the free form fabrication of custom cell scaffolds purely from natural biological molecules including collagen, among other relevant ECM components.
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INTRODUCTION The use of biological materials for microfabrication have traditionally been explored through microcontact printing,1,2 self-assembly,3 photolithographic patterning,4 and microfluidic patterning.5 These methods have demonstrated significant merits in terms of reproducibility and large area 2D patterning; however, methods for prototyping custom 2D or even 3D structures from biological materials remain highly desired. Achievements including hierarchical patterning have shown that microfluidic delivery may be used to generate layer-by-layer construction of cells and cell-embedded collagen gels.6 Recently, we have also demonstrated an effective technique for generating tailored 3D microstructures using filamentous virus as the lone building material.7 In the following work, we demonstrate that this approach can be extended to other biomaterials, namely collagen, for the direct writing of custom collagen microstructures as a scaffolds for directing cellular patterning and morphology. Moreover, we present several findings made with respect to the effects of vertex density (i.e., the number of intersecting collagen fibers per unit area) in layer by layer fabricated collagen patterns as well as the effects of pattern spacing on controlling cell spreading and alignment. Collagen has frequently been employed in the engineering of surfaces for cellular attachment.8−11 Depending on its structure, collagen provides important optical and mechanical properties12−14 as well as playing key roles in guiding cell migration and proliferation.15 Type I collagen provides one of the most important extracellular networks for the support of cell-laden tissues in nature,16 as it contains several sites for cell attachment via surface integrins. As with most networks, the topology or © 2015 American Chemical Society
arrangement of collagen is important. The topologies and alignment of collagen fiber networks depend on conditions including applied loads, cross-linking, and cell-induced reorganization.17 Recent works have shown that tuning the surface topology could induce effects on cellular polarization, orientation, and migration.18−20 In a specific example, researchers have implemented grooved silicone topographies coated with proteins involved in cell-binding to reveal alignment of cells along parallel microgrooves21 and also remodeling of underlying extracellular matrix (ECM).22 Correspondingly, microfabrication of custom cellular microenvironments with topological cues as well as biochemical cues is an important goal for orienting cells of different tissue types.23 Recent biological printing methods have proved capable of creating large patterns;24,25 however, the implementation of biomolecular building blocks alone in the direct writing of nano/microstructures has, with few exceptions,26 been largely unexplored. In this work, we utilize pure collagen as a standalone material and also as a composite material for the formation of custom patterned microstructures. Notwithstanding, there exist microfluidic techniques that have achieved patterning of cells with feature sizes of 0.1−3 mm by flow of hydrogel precursors albeit along with cell suspensions.25 Similarly, cell scaffolds could be generated by dispensing hydrogel precursors as mixtures with fibrinogen27 and RGDReceived: March 4, 2015 Revised: May 5, 2015 Published: May 8, 2015 1761
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modified alginate.28 For direct patterning of collagen, electrohydrodynamic writing has been successful using large nozzles (100−200 μm diameter), similar to those employed in microfluidic printing techniques, to prevent blockages. While the minimum feature sizes of the pattern were inherently larger than the size of a cell, the researchers nonetheless showed successful cell attachment to the collagen surfaces.29 Certainly, these systems have proven value in providing a means for facilitating growth of viable cells within a tailored scaffold; however, the morphology, polarization, and orientation of the patterned cells remain largely uncontrollable. Prior works on contact guidance of fibroblasts have shown that cell orientation can be correlated with the alignment of the underlying collagen fibers.30 A number of techniques for orienting collagen fibers across large surface areas have been explored, including the use of stretch-induced orientation,31 the application of high strength magnetic fields during fibrillogenesis,32 and more recently electrospinning.33 Microfluidic channels have also shown linear collagen patterns (10−400 μm wide) will result in cultured endothelial cells orienting in the direction of the fiber alignment in channel,34 with the most dramatic alignment occurring at the smallest channel width of 10 μm in the direction of the flow. Given the increased predominance of shear forces at smaller channel diameters, it may be clear that the orienting shear forces enhanced fiber alignment, which may be seen in related processing techniques such as extrusion.35 It is important to note the term collagen fibers is used here in the general sense indicating the bundled collagen structures extruded by our technique having 0.8−2 μm diameter. For us to realize direct writing of collagen, we utilized a motorized stage and capillary tip capable of providing controlled movement in three orthogonal axes. Using a microcapillary tip filled with collagen, we could initiate microstructure formation by contacting a substrate to form a meniscus of collagen which rapidly solidified to form the foundation for subsequent extrusion of collagen from the capillary upon movement of the tip/stage system. Free form extrusion was capable with this system in order to allow custom patterns of collagen to be produced within several minutes. A postfabrication heat processing step require several days was found to be necessary to stabilize the structures so that they may be implemented under prolonged cell culture conditions. By controlling the direction and spacing of the patterned collagen structures, we find that cultured cells (i.e., murine fibroblasts) exhibited guidance in their alignment allowing orientation of their growth upon contact with the microstructures. In addition, their morphology was dramatically influenced with respect to polarization depending on the microstructured topology, most strikingly by vertex density with respect to the extent of intersecting (or layer by layer generated) collagen fiber structures. Because this approach may be used to generate customizable patterns capable of directing cell shape and orientation, it is expected to significantly benefit research in cellular biomechanics/dynamics and may soon provide a means for generating well-defined tissue engineering scaffolds. In what follows, we provide details of our unique fabrication strategy implementing collagen as a biological building block for custom microscale patterning of biomaterials, and we anticipate future adaptation of this system with other biomolecules of interest.
Article
EXPERIMENTAL PROCEDURES
Preparation of Collagen and Collagen−Polydiacetylene Suspensions. Type 1 elongated collagen fibrils from bovine achilles tendon (powder, C9879 Sigma-Aldrich) were dissolved in 0.1 M acetic acid (pH 2.8) to a final concentration of 1.6, 3.3, 6.5, 13, and 26 mg/ mL by heating at 90 °C for 1−2 h followed by tip-sonication for 10 min. Pure collagen suspensions were used directly for fabrication or were premixed into a composite with 20% (v/v) of 1 mM polydiacetylene (PDA) vesicles for the purpose of fluorescent visualization. Preparation of polydiacetylene vesicles has been documented extensively in prior literature,36 but a brief overview of the protocol used is as follows. Approximately 5.6 mg of 10,12pentacosadiynoic acid (PCDA) (powder, GFS Chemicals, Ohio, USA) was dissolved in 200 uL of dimethyl sulfoxide (DMSO) and added to 15 mL HEPES (10 mM, pH 7.4). The solution was then heated to 80 °C for 15 min followed by tip-sonication for 15 min. After sonication, the liquid was filtered with a 0.8 μm syringe filter while warm and stored immediately at 4 °C overnight to allow for self-assembly of the vesicles. UV polymerization of the self-assembled vesicles using a hand-held UV lamp (1 mW/cm2 for 30 min) afforded a blue colored solution of nonfluorescent PDA which was mixed 20% (v/v) with the collagen solution to provide the collagen−PDA composite for fabrication. It is important to note that the PDA vesicles will become their red, fluorescent form after heat treatment,37 which is conducted after fabrication. Direct Writing with Collagen and Collagen−PDA. Borosilicate capillaries with the following dimensions (10 cm, 1 mmOD, 0.5 mmID) were purchased from Sutter Instrument Co. and were further processed to have an inner diameter typically of 0.8−2 μm (in addition 5 μm and 8 μm tips were created as well by the same technique) using a micropipette puller Model P-97 also from Sutter Instrument Company. Glass slides possessing aldehyde or amine coatings were purchased from LumiNano Co. (Korea) as substrates on which to fabricate the patterned collagen structures and were cut into thirds prior to use. To fabricate the biomaterial patterns, collagen or collagen-PDA solutions were loaded into the microcapillaries which were then placed into the holder of a guided extrusion system. The extrusion systems was comprised of a set of movable DC driven platforms (STM-1−50) for moving the stage (X,Z directions) and capillary tip (Y,Z directions). The system was setup on a vibration isolation table and the motors were driven using a pair of controllers from Namil Optical Components. For a typical fabrication process, the patterning commenced by first lowering the capillary tip at slow speeds (0.1−0.2 mm/min) until it was near enough to form a liquid bridge meniscus with the underlying substrate (i.e., amine coated glass). After liquid bridge formation, the capillary tip was then raised several micrometers at the same pulling rate of (0.1−0.2 mm/min). The lateral extrusion of the collagen fibers was then conducted at a slow pulling rate of 0.2 mm/min for a distance of approximately 50 μm, and the tip was then lowered to again contact the substrate. At this point, the writing process was either continued in the same manner by initially pulling up, then laterally, and then recontacting the surface, or the writing process was chosen to cease by initiating a fast tip speed of 10 mm/min which was sufficient to sever the liquid bridge. Each of the patterns used in this work were generated by this same technique to allow controlled fabrication of the collagen into different patterns. After completion of a pattern, the glass slide as transferred to a heater for incubation at 90 °C for various time points ranging from 1 to 96 h as described in the results. Cell Culture and Growth on Patterned Substrates. Prior to harvesting and placement on collagen patterns, NIH 3T3 murine fibroblast cells were cultured in 1x DMEM (Gibco) containing 4.5 g/L D-glucose, L-glutamine, 110 mg/L sodium pyruvate, 10%(v/v) newborn calf serum, and 1%(v/v) penicillin/streptomycin (Gibco). Cells were maintained in T-25 culture flasks in a 37 °C incubator with 5% CO2 and were passaged every 2−3 days upon reaching 80−90% confluence. For culturing of fibroblasts onto the patterned surfaces, 1 mL of trypsin was first added to a T-25 flask of 70−80% confluent fibroblasts, and the culture was incubated briefly at 37 °C before 1762
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Figure 1. (A) Overview of extrusion guided microfabrication technique for generating collagen patterned substrates. Specifically, a capillary (having 0.8−2 μm tip diameter) is filled with collagen suspension and lowered to form a liquid bridge contact with a glass substrate. Writing commences by raising the tip followed by controlled lateral movement in any direction. Continuous writing of the structure may be finalized by providing rapid movement of the capillary. While a range of custom collagen microstructures can be drawn by this technique, we restricted our analysis of the cell directing capabilities of these patterns to only (B) linear parallel fibers and (C) layer-by-layer generated crosshatch structures in order to assess the importance of pattern spacing and vertex density. The collagen patterns are observable from phase contrast, and the use of a fluorescent tracer can also be implemented to improve visualization.
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resuspension with 5 mL of warm DMEM media containing serum and counting by hemocytometer. Cells were harvested at room temperature by centrifugation at 220 rcf for 5 min followed by removal of the media and resuspension in approximately 3 mL of new serumcontaining DMEM media. Next, the patterned glass slide was place in a 35 mm Petri dish, and 100 μL of cell suspension was added on top of the glass slide. After remaining untouched for 20 min at room temperature, an addition 2 mL of serum-containing DMEM media was added to the Petri dish which was then placed in the 37 °C incubator with 5% CO2 where it remained until further analysis or changing of media. Calcein Staining, Microscopy, and Image Analysis. After incubating the fibroblasts cultured on collagen and collagen-PDA patterned surfaces, the media was removed and 1% Calcein AM (v/v) with 1× phosphate buffered saline was added to the Petri dish followed by incubation at 37 °C for 10 min. Imaging of the fluorescently stained fibroblasts was performed using an Olympus Ix71 microscope with 10×, 20×, and 40× objectives. Images were obtained with phase contrast or use of a green excitation filter set to observe the presence of cells. A Texas Red excitation filter set (or MCherry filter set if specified as such) was also employed to visualize the presence of collagen-PDA patterned substrates due to the fluorescence of heated, red phase PDA vesicles embedded in the patterns. The images were further processed by ImageJ (NIH, USA) for overlaying the images in Figure 6c,d and for analyzing the orientation distribution of cells cultured on unpatterned and patterned substrates (Figure 5e,f) as well as providing a color survey of the orientation (Figure 5b,d) by using the OrientationJ Distribution Plug-in (Version 19.11.2012 as written by Daniel Sage of the Biomedical Imaging Group at Ecole Polytechnique Fédérale de Lausanne, Switzerland). OrientationJ was run with the following parameter set for obtaining the color orientation survey for Figure 5b,d: hue=orientation; saturation=coherency; brightness=original-image. Analysis of the projected cell surface area was measured using imageJ region of interest manager and converting the measured units to a known control image to obtain the area as micrometers squared.
RESULTS AND DISCUSSION While exciting methods have recently been develop for the larger scale extrusion of whole cells embedded in hydrogel precursors as discussed above, these approaches offer little or no control over nano/microstructured patterning of the biomaterial network.25,27−29,38 To address and understand the importance of surface/network topology on cell growth and function, nano/microscale fabrication techniques are required. Dip-pen techniques have shown it possible to create submicron surface features of thiolated collagen by transfer from an atomic force microscopy tip onto a solid support and in doing so provided a new level of control for direct writing of biomaterial.26,39 In contrast, we demonstrate here the continuous fabrication of collagen with micron to submicron scale feature sizes by extrusion from a microcapillary. As depicted schematically in Figure 1, this approach utilizes the pure biomaterial (i.e., collagen without any precursors or cells) for fabrication by pulled extrusion from the microcapillary after adhesion to the substrate. Control over the pulling direction of the liquid bridge can provide custom direct writing of the nano/microstructured collagen. The 3D extrusion device relies on three orthogonal motor controlled axes to provide precise positioning of a glass microcapillary filled with a biological material. Here we find that this technique could be adapted to the use of collagen as the biological material; thus, we explored this process for patterning of collagen structures and further examined their potential for use under cell culture conditions. The success of the microfabrication of collagen depends on the ability of a nanoscale liquid bridge to be formed upon contact between a surface and the microcapillary filled with collagen solution. From the liquid bridge point of contact, a solid foundation of collagen is produced, and upon initiating movement of the microcapillary tip in a desired direction, we 1763
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Figure 2. Phase contrast images of parallel patterned collagen fibers before and after washing with 10 times with phosphate buffered saline and after 37 °C incubation in cell culture media. Each column represents the different durations of postfabrication dehydrothermal treatment (90 °C) that were used to stabilize the collagen patterns prior to washing. While 6 h of heating was insufficient in stabilizing the patterns, we find that heating for 48 h provided full retention of the collagen (scale bar: 100 μm).
may control the extrusion of collagen into a custom geometry. In order to carry out effective direct writing of collagen microstructures, the collagen must be continuously extruded as can be controlled by using a suitable speed for moving the capillary which can vary depending on the given tip diameter being used (Supporting Information Figure 1). For instance, when implementing a 2 μm tip, collagen structures can be patterned at writing speeds from 0.01 to 0.2 mm/min for a 13 mg/mL collagen suspension. By contrast, for 5 μm and 8 μm diameter tips, a minimum writing speed of 0.5 mm/min is found to be necessary to fabricate patterns of collagen at the same concentration. When increasing the tip diameter to 5 μm and 8 μm, we found that it was possible to extrude collagen patterns with larger feature sizes corresponding to the approximate size of the capillary tip diameter in use (Supporting Figure 2). Once the preferred structure has been fabricated, the writing process can be terminated by a rapid movement of the tip (>10 mm/min) in order to cut the drawn fiber from the prior structure, and a subsequent structure may be produced at this point. This approach offers the benefit of flexibility through direct writing of the biological materials and does not require any filler material. In contrast to conventional microfabrication approaches, this technique requires no sacrificial materials/layers, masks, complex equipment, or even a preconceived design strategy. As such, prototyping of structural features can be carried out whether for the purpose of proof-of-concept development or optimization of surface features. Based on our intuition gained from working with this direct writing system, we first assessed a range of collagen concentrations that seemed plausible for effective fabrication. It was clear that the concentration of the collagen suspension was important in dictating the quality of the resulting fiber and therefore the practical range for this fabrication strategy. Due to the high viscosity of the collagen suspension, we were unable to fill the microcapillary with collagen having concentrations of 26 mg/mL or higher. Reducing the concentration to 13 mg/mL of collagen facilitated effective microcapillary loading and also provide sustainable fabrication of continuously written collagen patterns when using the 2 μm tip at 0.2 mm/min (Supporting Figure 3c). By contrast, 6.5 mg/mL of collagen did not allow
continuous direct writing at 0.2 mm/min (Supporting Figure 3b), and at lower concentrations, only an unstable foundation of collagen could be formed resulting in immediate breaking of the structure upon capillary movement or changing of the writing direction (Supporting Figure 3a). At slower writing speeds of 0.01 mm/min and 0.05 mm/min, however, the 6.5 mg/mL collagen suspension could successfully be used for generating patterns from the 2 μm tip. The stability of writing using a 2 μm tip under different collagen concentrations with respect to pulling speed is summarized in Supporting Figure 3d. Based on these results, we utilized a collagen concentration of 13 mg/mL and writing speed of 0.2 mm/min with 0.8−2 μm diameter tips to generate all of the patterned substrates in the remainder of this work unless otherwise noted. Following from this, the next variable to consider was the choice of substrate, wherein we surveyed aldehyde coated, amine coated, and uncoated glass slides for their ability to adhere with the patterned collagen fibers. As it was our intention to implement the collagen structures under extended cell culturing conditions, we examined the written collagen lines on the various substrates with respect to their retention and stability after washing/incubation in an aqueous buffer environment. Initially, we found that the patterns were predominantly unstable, hence we implemented an additional heat treatment step of 1 h to promote adhesion of the collagen to the substrate. The resulting structures were found to have improved stability particularly for the case of the amine treated glass (Supporting Figure 4). Amine-treated glass was from thereon used as the primary substrate. Since 1 h of heating was found to provide some improvement in substrate adhesion and retention in pattern geometry, we pursued more prolonged heat treatments, which have been referred to in literature as dehydrothermal treatments. Dehydrothermal treatments are often used to remove water (typically at least 90 °C for 24 h) and subsequently form intermolecular cross-links in collagen bundles.40,41 Observation of the samples by scanning electron microscopy (SEM; Supporting Figure 5) before and after dehydrothermal treatment showed no effect on the diameter or morphology of the collagen fibers; however, this does not eliminate the possible occurrence of molecular scale rearrange1764
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Figure 3. Phase contrast images of a collagen patterned substrate (A) before addition of fibroblasts, (C) 2 days after culturing of fibroblasts, and (D) 5 days after culturing of fibroblasts reveal that cells contacting the patterned area have preferential alignment in the orientation of the long fiber axis. (F) Calcein staining provides better visualization of the bipolar morphology for cells located along the parallel fibers, while at the vertex of two fibers more of a multipolar morphology appears to be exhibited. The location of the patterned collagen fibers (B) before and (E) after cell culturing can be observed using a red excitation filter, as red fluorescent PDA was incorporated with the collagen suspension to provide a fluorescent tracer.
fluorescence visualization of our patterned structures after fabrication (Figure 1c and Figure 3b). To gain insight as to how the collagen patterns would behave in the presence of cells, we evaluated various geometries, lengths, and spacing of patterned collagen as substrates for the culture of mouse fibroblasts (NIH 3T3). Because fibroblasts are known to remodel extracellular matrix proteins such as collagen, we expected depletion of the original collagen patterns; however, we found that this was not the case with the patterned collagen remaining even after several days of culture (Figure 3). Several interesting observations were made evident as the duration of culture time increased, particularly that as the fibroblasts migrate along the glass surface they tend to stop and adhere at the patterned collagen locations which may be expected given the presence of cell binding domains within the collagen fiber. Furthermore, once the cells were attached to an individual collagen fiber they appeared to preferentially orient themselves tangentially relative to the direction of the patterned fiber. From Figure 3d, we can see the contrast in morphology for the fibroblasts bound to the glass in relation to those bound to the collagen fibers. Calcein staining (Figure 3f) provides a clearer view of the cells transition from a multipolar to extended bipolar morphology upon interacting with the linear collagen fiber; however, near the vertices where two perpendicular fibers have met, this transition is not as pronounced. The presence of the fibers was invisible when using the green excitation filter, so to observe the collagen− PDA composite pattern, a red excitation filter was used (Figure 3e). The broad emission from the Calcein stain can also be seen in observations with our red filter set which may make it difficult to observe the presence of the patterned fibers. Therefore, we have provided additional images (Supporting Figures 6 and 7) that show the fluorescence of collagen/PDA patterned lines before culture and during culture of fibroblasts. Further testing the possibility of the fibers directing the alignment of cell growth, we examined fibroblasts cultured on
ment due to heating. Here we examined the patterned collagen after various durations of heating at 90 °C in order to identify a postfabrication treatment suitable for retaining the shape and stability of the patterned collagen fibers. We found upon only 6 h of thermal treatment that the presence of collagen patterns underwent significant changes in morphology and extensive pattern loss after washing. By contrast, 48 and 96 h of thermal treatment provided ample retention in the shape of the patterned collagen even after washing and an additional 4 days of incubation under cell culturing conditions (Figure 2). Because almost no collagen loss was observed after extensive heat treatment, the incorporation of a 48 h postfabrication heating step was necessarily implemented for each of the collagen patterned substrates to be used in cell culture. It may be seen in some instances that the patterns appear to undergo a change in shape near the points used for initiating and ending the fiber fabrication process; however, this is a factor of the semimanual fabrication process, as we have not yet automated this technique (specifically, this results from a superfluous gap between the fiber and glass substrate at these locations). Regions in direct contact with the glass in contrast will exhibit no change in their patterned shape if given a sufficiently long dehydrothermal treatment. Visual examination of the patterned collagen proved demanding during the optimization process due to the small feature sizes of the collagen fibers. Accordingly, in the subsequent experiments we implemented a comixture of collagen and fluorescent particles, specifically polydiacetylene (PDA) vesicles, in order to provide better visualization of the resulting collagen patterns. The polymerized PDA were chosen as the cocomponent, since we previously found it to be noncytotoxic and to produce a bright red fluorescence after exposure to stimuli such as heating.42 Because the pretreatment process elicited the formation of the red fluorescent phase of the PDA vesicles, incorporating PDA as an additive to the collagen suspension offered an effective means for direct 1765
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Figure 4. (A) Parallel collagen-PDA patterns were fabricated in order to assess the effect of various line spacing (10, 30, 60, and 120 μm) on cell growth. (B) Prior to heat treatment the PDA component was nonfluorescent; however, (C) after heat treatment the fluorescent response of the PDA tracer could be readily observed. After 2 days of culturing of fibroblasts on the patterned surface followed by Calcein staining (D), phase contrast and fluorescence images with (E) red excitation or (F) green excitation filters show the alignment of cells in the patterned area to be congruent with the patterned structures for each of the line spacing. Bridging of patterned lines was observed for the close line spacing of 10 μm, while cells can be found slightly between the parallel lines (not fully interacting with the collagen) at the wide line spacing of 120 μm.
Figure 5. After 2 days of culture, fluorescence images of Calcein-stained fibroblasts from (A) unpatterned and (C) collagen−PDA patterned surfaces (equivalent sample to Figure 4f) were obtained and further surveyed in regards to their cell orientation (B and D, respectively), using the OrientationJ plug-in of ImageJ. Using the same software, the distribution of cell orientation was estimated and compared for the (E) unpatterned and patterned surfaces. Additionally, (F) regions having different pattern spacing were analyzed separately to provide their respective distributions of cell orientation, which indicate preferred alignment of cells with the parallel patterned collagen lines in the case of each spacing distance.
parallel collagen fiber patterns having a range of interfiber spacing. As seen in Figure 4a, the layout consisted of micron diameter fibers having line spacing of 10, 30, 60, and 120 μm. After heat treatment of the collagen−PDA composite pattern, the PDA vesicles exhibited a red fluorescence which was used to observe their presence during cell culturing (Figure 4b,c). After 2 days of cell culture (Figure 4d−f), the parallel patterned
collagen fibers once again appeared to direct alignment of cell growth orientation, and moreover exhibited an effect on the cell density where in the unpatterned area was less sparse. Initially, we suspected contact inhibition to play a role;43 nonetheless, this disparity in cell density has not yet been determined as we have not yet assessed the rates of cell proliferation or migration with respect to line spacing. By analyzing the distribution of 1766
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Figure 6. (A) Phase contrast and fluorescence images (using red or green excitation filters) of fibroblasts after 2 days of culture on a crosshatch structure of collagen−PDA. The patterns were generated using layer by layer microfabrication (as outlined in Figure 1c) with varying pattern spacing in order to control the surface density of vertices (intersecting collagen fibers). (B) Image analysis reveals that the closer pattern spacing (in which the number of vertices per unit area is increasing) causes an increase in cell spreading, represented by the projected cell area. The original red fluorescent image arising from the patterned structure at 120 μm spacing is provided as an overlay with phase contrast images after (C) 1 day and (D) 2 days of cell culture to better illustrate that the wider pattern spacing promotes cells to align with the patterned collagen (scale bar: 50 μm).
ogy of the 30 μm cross-hatch pattern as compared to that of the aligned, bipolar morphology observed in the case of 10 μm parallel spaced fibers exhibiting an even higher ligand density. Similarly, when the vertex density is decreased, we found the cell morphology to become less multipolar and eventually form the extended bipolar morphology once again at 120 μm spacing. In contrast to the lack of directional preference observed for narrow cross-hatch patterns (higher vertex densities), at the lowest vertex density of the 120 μm crosshatch pattern, which is far larger than the cell itself, fibroblast growth once again appeared to be directed preferentially in alignment with the orientation of the collagen fibers. While these results provide a very good case for how control of the vertex density and line spacing can help to influence the cell morphology, we insist that this does not eliminate the possibility of other factors (perhaps more significant biochemical cues) being used in conjunction with this approach for dictating the shape and behavior of patterned cell growth. In looking into foundational studies of topological effects on fibroblast growth, it becomes apparent that their morphology represent adaptation to their local environment.44 When isolating the effects of surface topology as determined from parallel grooved surfaces, fibroblasts are known to exhibit preferential alignment in the direction of the parallel surface features.45 This is supported by fibroblasts exhibiting contact guidance in which anisotropic topographic features induce alignment as cells extend and retract preferentially along the direction of surface features, and this effect is further amplified when cultures are grown on patterned substrates supplemented with serum which contains proteins that adsorb to the surface before the cells adhere. 46 Such biochemical cues can significantly improve cell adhesion, as increases in cell binding domains present on a surface are known to enhance cell spreading.47 Similarly, works examining parallel surface topologies with fibronectin coatings give evidence that filopodia formation and cell alignment increase with density and depth of patterned grooves resulting in elongated cell morphologies.48 Based on this information, we may venture that the bipolar
oriented cells with image processing tools, we could confirm the preferential fibroblast alignment in the direction of the collagen fiber axis as compared to unpatterned surfaces (Figure 5e). With respect to fiber line spacing, we found that at the largest spacing of 120 μm between patterned collagen lines, there was a slight decrease in the proportion of fibroblasts oriented in the direction of the fibers (Figure 5f). This lower distribution of parallel cell alignment for the 120 μm fiber spacing condition may be attributed in part by the existence of several cells between the parallel fibers that had little to no contact with the patterned collagen. Interestingly, at 10 μm we see a slightly wider distribution for the cell orientation, as the close proximity of parallel fibers more easily allowed cells to associate with more than one neighboring fiber. For each of the patterned line spacing, we found significant cell extension into a bipolar morphology compared to the unpatterned substrate, although, as alluded to previously, the appearance of a vertex (or intersection) in the collagen pattern could potentially mitigate this effect. To examine the effect of vertices on cell morphology and orientation, we fabricated layer by layer collagen patterns (Figure 1c and Figure 6) in which the fiber spacing could be used to control the effective density of vertices on the substrate surface. Specifically, we used direct writing of 30, 60, and 120 μm spaced collagen-PDA fibers orthogonally written in a crosshatch pattern to achieve different surface densities of vertices (number of intersecting collagen fibers per unit area). The culture of fibroblasts on the collagen patterned surface revealed an interesting effect of vertex density on cell morphology in terms of cell spreading and polarization. The high vertex density patterns produced from 30 μm pattern spacing revealed significantly large projected cell area compared to the 60 and 120 μm pattern spacing condition. We could question whether this is a direct effect of increasing the relative amount of cell binding domains which exist on the underlying collagen substrate or if this is attributed to the vertex density. To support that this observation may be an effect of pattern morphology rather than purely ligand density driven, we point out the demarcation between the spread, multipolar morphol1767
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to exploit our findings of the influence vertex density and pattern spacing has on controlling cell spreading, polarity, and alignment for not only fibroblasts but examining their influence in other cell types. In addition, we expect this unique proof-ofconcept in biomaterials fabrication to be of use for investigators researching topics including the role of mechanical signaling in controlling cell shape and motility, the control of cell spacing, and the remodeling of custom designed scaffolds. This technique may hold broad interest in providing customization of the topology and alignment of collagen in a local nano/ microenvironment. Because this approach offers the freedom to design a stable scaffold purely out of biological materials, we anticipate the adaptation of this system to generating other biomaterial patterns of interest and look forward to the extension of the layer-by-layer technique to examining the possibility of generating 3D scaffolds in the near future.
extended morphology of fibroblasts exhibited in the presence of parallel collagen fibers could be a result of contact guidance supported through cell binding domains present on the collagen fiber. Similarly, we may expect this to be the case for cross-hatch patterned collagen fibers when the pattern spacing is well above the size of the fibroblast. However, at a narrow cross-hatch pattern spacing (increased vertex density) comparable to the size of the cell, it may be that the local environment appears less anisotropic (i.e., exhibiting less contact guidance) as compared to the case of parallel or larger spaced cross-hatch patterns. Also, as we have pointed out the narrower cross-hatch pattern inherently provides increased sites for cell binding. Nonetheless, more closely spaced parallel fibers, providing a higher surface density of collagen, still exhibited the bipolar morphology rather than the spread (multipolar) morphology of the dense cross-hatch pattern, which implies that not only the surface ligand density but also the surface anisotropy is a key factor. Taking this into consideration, it is also important to note that a vast amount of prior works on the effect of surface topology has suggested that cell morphology largely depends on cell type in addition to feature dimension.49 Given the discussions of the complexity of this research area, we hope that our new technique for generating custom surface topologies from pure collagen structures may provide a means for gaining further insight into determining the topological constraints for regulating cell shape and perhaps more complex processes including migration within different physical environments. In our resulting approach, we find this technique can yield a multilayer patterned surface in less than an hour; however, as we have shown the collagen structures require an additional several days for the dehydrothermal treatment step in order to be sufficiently stable for use in cell culture. We anticipate this approach will open the way for patterning of collagen and other biological materials, but it is important not to overlook that purely biological components have already been implemented in several nonpatterning fabrication processes including electrospinning, electro-blowing, and wet-spinning among others to create fiber-derived meshes and membranes.50,51 Whether the use of purely biopolymeric components is of benefit to future cell scaffold applications depends on the biomaterial being employed, as controlling the mechanical properties, degradation rates, and biocompatibility of some natural biopolymer may be less reliable compared to well-characterized synthetic polymers. Nonetheless, the innate functionality of natural biopolymers and the possibility of incorporating new functionality via genetic and chemical modification could prove advantageous. For now, the ability to generate custom submicron/micron sized patterns in real-time from a pure biological material such as collagen is seen as a valuable step forward.
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ASSOCIATED CONTENT
S Supporting Information *
Additional images of direct writing with various concentrations of collagen and on different glass surface chemistries as well as a SEM comparison of fiber morphology before and after dehydrothermal treatment as well as SEM of fibers produced from different diameter capillary tips are provided. The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.5b00295.
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AUTHOR INFORMATION
Corresponding Author
*Tel.: +82-(0)2-2220-2339; Fax: +82-(0)2-2220-1935; E-mail:
[email protected]. Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was supported by the Basic Science Research Program through the National Research Foundation (NRF) funded by the Ministry of Science, ICT & Future Planning (2013R1A1A1076117), and also by the Priority Research Centers Program through the NRF funded by the Ministry of Education (2012R1A6A1029029).
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ABBREVIATIONS PDA, polydiacetylene; PCDA, 10,12-pentacosadiynoic acid
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CONCLUSION In summary, we have demonstrated that our direct writing technique may be used for the microfabrication of custom collagen patterns capable of sustaining cell growth and more significantly providing control over cellular orientation and morphology. Because the collagen microarchitectures proved stable under extended culture conditions, we expect this approach to be of broad interest to areas in which prototyping of a biomaterial design is desired, such as tissue engineering, biological assay development, and even traditional cell biology studies regarding cell-matrix signaling. Looking forward, we aim
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