Analysis of Connective Tissue Progenitor Cell Behavior on Polydimethylsiloxane Smooth and Channel Micro-Textures

Abstract

Growth of human connective tissue progenitor cells (CTPs) was characterized on smooth and microtextured polydimethylsiloxane (PDMS) surfaces. Human bone marrow derived cells were cultured for nine days under conditions promoting osteoblastic differentiation on Smooth PDMS and PDMS Channel microtextures (11 μm high, 45 μm wide channels, and separated by 5 μm wide ridges). Glass tissue culture dish surfaces were used as controls. Cell numbers per colony, cell density within colonies, alignment of cells, area of colonies, and colony shapes were determined as a function of substrate surface topography. An alkaline phosphatase stain was used as a marker for osteoblastic phenotype. CTPs attached, proliferated, and differentiated on all surfaces with cell process lengths of up to 80 μm. Cells on the Smooth PDMS and control surfaces spread and proliferated as colonies in proximity to other cells and migrated in random directions creating colonies that covered significantly larger areas (0.96 and 1.05 mm², respectively) than colonies formed on PDMS Channel textures (0.64 mm²). In contrast, cells on PDMS Channel textures spread, proliferated, aligned along the channel axis, and created colonies that were more dense, and with lengths of longest colony axes that were significantly longer (3252 μm) than those on the Smooth PDMS (1265 μm) and control surfaces (1319 μm). Cells on PDMS Channel textures were aligned at an angle of 14.44° relative to the channel axis, and the resulting colonies exhibited a significantly higher aspect ratio (13.72) compared to Smooth PDMS (1.57) and control surfaces (1.51).

Introduction

The outcome of normal bone fracture healing or bone grafting procedures is highly dependent upon a proper osteoactive environment provided by the local fracture hematoma. In suboptimal fracture repair conditions where attachment, migration, proliferation, and differentiation of bone forming cells is hindered, bone grafts play an important role in promoting a milieu in which bone formation is enhanced (Bauer and Muschler, 2000). Autogenous cancellous bone is widely considered to be the most effective graft material, and is used for 50% of the 500,000 bone graft procedures performed annually in the United States (Muschler et al., in press at CORR). Nevertheless, host morbidity and a number of related complications (Bauer and Muschler, 2000; Muschler et al., submitted to JBJS) associated with autografts evidence the need for alternative bone graft materials (Muschler et al., submitted to JBJS) that promote bone healing through osteoconductive, osteoinductive, and osteogenic activity at the fracture site (Muschler et al., in press at CORR). A novel approach to enhance bone healing has been the transplantation of bone marrow derived cells containing connective tissue progenitor cells (CTPs) through their concentration and selective attachment onto an implantable scaffold (Muschler et al., in press at COTT; submitted to JBJS). Successful bone healing requires the presence of a sufficient number of CTPs (Fleming et al., 2000) to adhere, proliferate, and organize extracellular matrix molecules into a functional tissue (Lebaron and Kyriacos, 2000). Mechanical stimuli provided through surface textures (surface topographies) have been reported to influence cell shape, gene expression (Brunnette and Chehroudi, 1999), protein production and deposition (Brunnette and Chehroudi, 1999; Riedel et al., 2001; Walbloomers et al., 1998; von Recum et al., 1996), cell proliferation, migration, differentiation, and survival (Alerts et al., 2001; Chen et al., 1998). Therefore, the incorporation of micro- and nanoscale textures at the cell-scaffold interface might provide an attractive approach to selectively enhance specific and desirable CTP behavior without destabilizing the optimum (and delicate) biochemical conditions during bone fracture healing.

Recent interest has been focused on the use of microfabrication and micromachining technologies to create surfaces that influence cell behavior through surface microtextures comprising posts, holes, and channels. Although cell growth on microtextured surfaces with posts and holes have been reported in literature, including Deutsch et al. (2000), Craighead et al. (1998), Green et al. (1993), Schmidt and von Recum (1992), and Mata et al. (2002), the majority of investigations have concentrated on cell response to channel micro-textures. Brunette (1986a; 1986b) reported in 1986 a hierarchical alignment of fibroblasts cultured on channels of different dimensions. In 1990, Clark et al. (1990) described the relation between kidney and neuronal cells to channel depth and spacing. den Braber et al. (1995a; 1995b; 1996a, 1996b) have reported different degrees of fibroblast morphology, orientation, proliferation, and protein deposition when cultured on various channel textures. Deutsch et al. (2000) reported that topographic features corresponding to cellular dimensions, lead to an enhanced cell attachment, orientation, and a more in vivo-like morphology. Alaerts et al. (2001) described an increased fibroblast orientation on 0.5−10 μm wide channel textures compared to smooth surfaces, while Miller et al. (2001) found 10−20 μm wide channels to be optimal for Schwann cell alignment. van Kooten et al. (1998) reported an increased fibroblast proliferation on 2 and 5 μm wide channels relative to 10 μm wide channels. Other groups such as Singhvi et al. (1994), Curtis and Wilkinson (1998), and Brunette and Chehroudi (1999), have provided reviews of the different behaviors, topographies, and cells that these and other researchers have studied. In many of these investigations (Deutsch et al., 2000; Green et al., 1993; Schmidt and von Recum, 1992; Mata et al., 2002; den Braber 1995a; 1995b; 1996a; 1996b; van Kooten et al., 1998), polydimethylsiloxane (PDMS) was used as the substrate material for cell culture experiments. PDMS is a flexible, biocompatible, highly inert, optically transparent polymeric material that has a low curing temperature and is relatively simple to process and manipulate. It is relatively cheap and commercially available as a two-component mixture consisting of a pre-polymer and a cross-linker.

The present study is designed to advance the application of microtextured surfaces to control and influence the proliferation, migration, and differentiation kinetics of human CTPs, which have relevance to tissue engineering applications involving a variety of connective tissue phenotypes. Specifically, human bone marrow derived cells containing CTPs were cultured on PDMS substrates that were presented as either a smooth surface or textured surface with microchannels. Systematic investigations were then conducted to assess possible effects of surface topography on CTP growth characteristics (attachment, migration, proliferation, alignment, and differentiation) as well as on CTP colony characteristics (cell density, colony area, and colony shape).

Materials and Methods

Experimental design

Human bone marrow derived CTPs were cultured on two types of PDMS substrates comprised of Smooth PDMS (unpatterned) surfaces and PDMS Channel textures with curved cross-sections that were 11 μm high, 45 μm wide, and separated by 5 μm wide ridges. The textured PDMS substrates were realized by microfabrication and micro-machining technologies. Fresh bone marrow derived cells were plated on the substrates, cultured for 9 days, fixed and analyzed using phase contrast, fluorescent, and scanning electron microscopy. Differentiation toward an osteoblastic phenotype was assessed using in situ staining for alkaline phosphatase activity. Each experiment was repeated three times and the results were compared to those from corresponding glass tissue culture dishes that served as controls.

Substrate preparation

The PDMS substrates were manufactured by Soft Lithography (Figure 1) as described by Xia and Whitesides (1998). Briefly, an 11 μm-thick layer of photoresist (AZ-9260, AZ Electronic Materials, Somerville, NJ) was coated on top of a standard 100 mm diameter, (100)-oriented silicon wafer. Photolithography was then performed to transfer straight channel patterns from a photomask to the coated photoresist. The reflow of the photoresist during the final bake step (115 °C for 30 minutes) of the photolithography process resulted in rounding of the edges of photoresist patterns. This patterned photoresist on the silicon wafer constituted the master for subsequent PDMS molding. The patterned master was coated with 1H,1H,2H,2H-perfluorodecyltrichlorosilane (Lancaster, Pelham, NH) to facilitate release of the cast PDMS after curing. The liquid PDMS pre-polymer and cross-linker (Sylgard 184) (Dow Corning, Midland, MI) components were mixed in a ratio of 10:1, degassed for 7 minutes, and poured uniformly on top of the patterned master. After additional degassing for 12 minutes, the PDMS on the patterned master was cured at 65 °C for 3 hours and at room temperature (∼ 25 °C) for 21 hours. The cured PDMS cast was released from the master and sectioned into 1 cm × 1 cm samples. Representative samples were inspected for defects by scanning electron microscopy (SEM) (JSM-5310, JEOL USA, Peabody, MA) before and after sterilization in ethanol as described in the next section. The resulting PDMS substrates were textured with curved channels that were nominally 11 μm high and 45 μm wide. The microchannels were separated by ridges that were flat and 5 μm wide at the top and offset at maximum of 50° from the channel wall (Figure 2). In addition to PDMS Channel textures, substrates comprising Smooth PDMS surfaces were also produced by performing the above-mentioned procedure on an unpatterned silicon wafer. Glass slides (Lab-Tek Chamber Slide System) (Nalge Nunc International, Naperville, IL) that are regularly employed for tissue culture applications served as control surfaces. The PDMS substrates were placed inside the 2 cm × 2cm standard tissue culture dishes (Lab-Tek Chamber Slide System).

Fig. 1.

Fabrication of microtextured polydimethylsiloxane (PDMS) substrates by Soft Lithography. The cross-sectional schematic diagrams depict: (a) starting substrate, which is a 100 mm diameter, 500 μm thick, (100)-oriented silicon (Si) wafer; (b) thick photoresist coating patterned by photolithography; (c) rounding of pattern edges due to reflow of photoresist during baking; (d) molding of PDMS by casting onto patterned master after coating with a fluorinated alkyltrichlorosilane (R-SiCl₃) to facilitate mold release; and (e) release of PDMS cast from master.

Fig. 2.

Scanning electron microscope (SEM) images of PDMS Channel textures: (a) after release from patterned master; and (b) edge-on view to show channel cross-section. The tangent at the intersection of the ridge with the channel wall is offset at an angle of 50°.

Cell culture

Bone marrow aspirates were harvested with informed consent from patients immediately prior to elective orthopaedic procedures as described by Muschler et al. (1997). Briefly, a 2 ml sample of bone marrow was aspirated from the anterior iliac crest into 1 ml of saline containing 1000 units of heparin (Vector, Burlingame, CA). The heparinized marrow sample was suspended into 20 ml of Heparinized Carrier Media (α-MEM + 2 units/ml Na-heparin; Gibco, Grand Island, NY) and centrifuged at 1500 rpm (400 × g) for 10 minutes. The buffy coat was removed, resuspended in 20 ml of 0.3% BSA-MEM (Gibco), and the number of nucleated cells was counted. All PDMS substrates and control surfaces were sterilized for 30 minutes with 70% ethanol (Fisher Scientific, Fair Lawn, NJ). Cells were then plated on Day 0 at a density of 125,000 cells/cm² and cultured for 9 days in α-MEM media (Gibco #11900-073) with 10% Fetal Bovine Serum (Whittaker, Walkersville, MD) plus Dexamethasone (Sigma-Aldrich #D-1756), which was used to enhance osteoblastic expression. The media was removed and replaced on days 1, 2, 5, and 7.

Cell fixation and staining

On Day 9, the cultures were fixed and permeabilized by placing the substrates in acetone : methanol in a 1 :1 ratio for 10 minutes. Afterwards, cells were stained with 6-diamidino-2-phenylindole dihydrochloride hydrate (DAPI) (Vector), a nuclear fluorescent stain (Deutsch, 2000), for 6 minutes at 25 °C and subsequently washed three times with phosphate buffer. After imaging nuclei using DAPI staining, cells were again stained in situ for alkaline phosphatase (ALP), a marker for osteoblastic differentiation (Muschler et al., 1997; Majors et al., 1997; Muschler et al., 2001), using 0.9 mM Napthol ASMX phosphate and 1.8 mM Fast Red-TR Salt for 30 minutes at 37 °C. This staining forms an insoluble Napthol-Fast Red complex that precipitates in regions where cells express ALP activity. The precipitate was observed using autofluorescence.

SEM fixation

SEM was used to observe cell morphology. In order to assess the possibility of cell damage due to cell fixation (den Braber et al., 1995a), two different procedures were used to prepare the cells for SEM observation. On Day 9, the media was removed and the plated substrates were immersed in a solution containing 2% glutaraldehyde (Electron Microscopy Sciences, Fort Washington, PA), 3% sucrose (Sigma-Aldrich Co., Irvine, United Kingdom), and 0.1 M phosphate buffer (Baxter, Deerfield, IL) at 4 °C and 7.4 pH. After 1 hour, substrates were rinsed twice with the phosphate buffer for 30 minutes at 4 °C and washed with deionized (DI) water for 5 minutes. Dehydration was achieved by placing the plated substrates in 50% ethanol (in DI) for 15 minutes and replacing it every 15 minutes while increasing the concentration of ethanol to 60, 70, 80, 90, and finally 100% (Aaper Alcohol and Chemical Co.). The liquid ethanol was removed using critical point drying. The second fixation protocol followed the same steps, except that dehydrated cells were immersed for 5 minutes in hexamethyldisilazane (HMDS)(Nation, 1983) instead of using critical point drying (den Braber et al., 1995a).

Cell culture analysis

A phase contrast microscope (Olympus CK2) (Olympus Optical Co., Japan) was used for daily observation of the cells. Cells were fixed on Day 9, stained with DAPI and viewed under a fluorescent microscope (Olympus BX50F) (Olympus Optical Co.) to study different cell parameters: cell attachment, by counting the number of colonies that are formed; cell proliferation, by counting the number of cells within a single colony; cell migration, by studying the distribution of cells within the colony; and cell differentiation, by assay of markers of the osteoblastic phenotype. Furthermore, the following colony parameters were also quantified: colony area, length of longest axis of the colony (MAXL), and length of shortest axis of the colony (MINL) (Figure 3(a)). A colony was defined as a cluster of eight or more cells (Muschler et al., 2001). Colony area, MAXL, MINL, and the resulting colony aspect ratio (AR = MAXL/MINL) were quantified using the computer software Image-Pro Plus (Media Cybernetics, Inc., Silver Spring, MD). In order to account for random variations between and within experiments, statistical significance was defined at the 95% confidence interval using analysis of variance (ANOVA) test performed in Microsoft Excel (Microsoft Corp., Redmond, WA). Cell alignment was quantified by photographing five randomly chosen fields of vision of CTPs on PDMS Channel textures, and measuring the angle between the longest axis of the cell and the channel axis (Walbloomers et al., 1998; Clark et al., 1990; den Braber et al., 1995b)(Figure 3(b)). The morphology of cells cultured on the PDMS substrates and control surfaces was also examined using SEM. Finally, cells were stained in situ for ALP and viewed using a fluorescent microscope to verify differentiation into osteoblastic phenotype.

Fig. 3.

Schematic diagrams depicting: (a) a single colony of cells and colony parameters: longest axis of the colony (MAXL), shortest axis of the colony (MINL), and colony aspect ratio (AR); and (b) a pair of channels illustrating alignment of cells with respect to channel axis.

Results and Discussion

SEM examinations revealed that the PDMS substrates did not exhibit significant geometrical variations with respect to the photoresist/silicon master. Furthermore, a comparison of the PDMS substrates before and after ethanol sterilization did not reveal any apparent pattern degradation. The effect of the SEM fixation procedure on the PDMS substrates was minimal; only slight markings were observed on PDMS surfaces immersed in HMDS. Qualitative SEM examinations revealed that morphologies of cells on the various PDMS substrates were similar for both critical point drying and HMDS-based fixation procedures.

CTPs attached, proliferated, migrated, and differentiated on all PDMS substrates and control surfaces. The number of colonies ranged from 4 to 15 per 500,000 plated cells among different patient donors (separate experiments), and was generally consistent (±2) on all substrates for an individual donor (same experiment). Variations in CTP prevalence between donors and differences in CTP behavior within an individual donor have been identified (Mata et al., 2002; Muschler et al., 1997) and represent an inherent variability of our investigation. Nevertheless, the similarity in colony number suggests that the colony forming efficiency (i.e., the ability to support both attachment and proliferation of CTPs) of both Smooth PDMS and PDMS Channel textures was comparable to the control surface. These results correlate with our previous experiments (Mata et al., 2002).

Table 1 presents results from the cell culture analysis for different cell and colony parameters on PDMS Channel textures, Smooth PDMS, and control surfaces. The mean and standard deviation parameters were derived from three different experiments. The AR values were computed from the corresponding MAXL and MINL (not shown) values. On Day 9, PDMS Channel textures exhibited the highest cell number with a mean of 174 cells per colony. The Smooth PDMS and control surfaces exhibited means of 146 and 114 cells per colony, respectively. However, the differences in cell number among the different substrates were not statistically significant, indicating that the effective proliferation rate is similar for the various surfaces. The similarity in cell number between Smooth PDMS and control surfaces contradict previous reports (Green et al., 1993; Mata et al., 2002) where cell number on smooth PDMS surfaces was lower than on tissue culture dishes. Although the reasons for the discrepancy between our observations and the previous reports are not clear, it is possible that the difference in cell types and corresponding surface affinities (Kapur et al., 1996; Rich and Harris, 1981; Sigurdson et al., 2002; Bruinink and Wintermantel, 2001), along with variations in topography, biochemical stimuli, and quantification methods might be contributing factors.

Table 1.

Quantification of cell and colony parameters

	Cell number	Colony area (mm2)	MAXL (μm)	AR	Cell alignment
PDMS Channels	174±134	0.64±0.55*	3252±1522*	13.72±3.65*	14.44°±15.85°
Smooth PDMS	117±103	0.96±0.62	1265±442	1.57±0.64	—
Control	146±157	1.05±0.98	1319±544	1.51±0.43	—

^*Denotes statistical significance at 95% confidence interval.

The size and shape of cell colonies on the PDMS Channel textures were strikingly different from those on the Smooth PDMS and control surfaces (Figure 4). Areas of cell colonies on Smooth PDMS substrates (0.96 mm²) and control surfaces (1.05 mm²) were comparable, but significantly larger than the areas of the cell colonies on the PDMS Channel textures (0.64 mm²). Furthermore, an examination of cell density within colonies (cell number per unit area) reveals that colonies on PDMS Channel textures (272 cells/mm²) were significantly more dense than those on Smooth PDMS (122 cells/mm²) and control surfaces (139 cells/mm²). Without any kind of cell motility assay such as time-lapse microscopy, it is impossible to conclude that a particular colony shape or size is not due to fusion of multiple colonies; however, these observations collectively suggest that the rate of migration of cells within colonies on PDMS Channel textures was significantly lower than on Smooth PDMS and control surfaces.

Fig. 4.

Phase contrast microscope images of living cells and fluorescent microscope images of cells stained with DAPI on Day 9 to qualitatively show the difference in colony morphology and cell density between: (a) Smooth PDMS surfaces; and (b) PDMS Channel textures. Arrows denote single cells. Hematopoietic cells (round appearance) from the bone marrow aspirate are also observed within the channels.

CTP migration on PDMS Channel textures is highly directional. Colonies on Smooth PDMS substrates and control surfaces were similar and exhibited arbitrary shapes without any preferred orientation. In contrast, colonies on PDMS Channel textures appeared elongated along the direction of the channel axes, with mean and maximum MAXL of 3252 μm and 6689 μm, respectively. The Smooth PDMS substrates exhibited significantly lower mean and maximum MAXL of 1265 μm and 1994 μm. The directionality of the CTP migration is reflected by the higher AR values of colonies on the PDMS Channel textures. Cell colonies on PDMS Channel textures exhibited a mean AR of 13.72, which was significantly higher than the mean AR of the colonies on the Smooth PDMS substrates (1.57) and control surfaces (1.51). These results correlate with those from Brunette (1986a) where epithelial cell colonies cultured on grooved titanium substrates exhibited preferred orientation.

Investigation of cell morphology demonstrates that surface topography influences cell size, shape, and orientation. Before spreading, the CTPs generally exhibited a circular shape of approximately 10–12 μm diameter. After attachment and spreading, cell morphology varied between the two topographies. On the Smooth PDMS substrates and control surfaces, cell bodies adopted a broad flattened shape with the longest axis ranging from 40 to 100 μm and exhibited randomly oriented processes with mean lengths up to 80 μm (Figures 4 and 5). In contrast, the cells on PDMS Channel textures tended to attach and spread mostly within and along the channels with a mean alignment of 14.44°. Although the process lengths were comparable to those on cells grown on the smooth surfaces, cell bodies were narrower and were oriented along the channel axis. Similar characteristics of cell alignment in channels have been previously reported (Brunnette and Chehroudi, 1999; Deutsch et al., 2000; Brunnette, 1986a; 1986b; den Braber, 1995a; 1995b; 1996a; 1996b; van Kooten, 1998). Cell shape has been implicated to participate in the regulation of cell differentiation (Chen et al., 1998). However, our qualitative observations revealed similar ALP staining on all the surfaces, which suggested that surface texture did not affect the differentiation into osteoblastic phenotype.

Fig. 5.

SEM image of a cell cultured on: (a) Smooth PDMS substrate adopting a broad flattened shape with randomly oriented processes (mean lengths up to 80 μm); and (b) PDMS Channel texture exhibiting a narrower cell body aligned along channel axis and with similar process lengths to cells on the Smooth PDMS and control.

The majority of CTPs on PDMS Channel textures was located within the channels and aligned to the channel axis with the major processes generally directed towards those of other cells. However, a few cells straddled the ridges between channels, which is consistent with observations reported by Brunnette (1986b). SEM and phase contrast microscope images revealed distortions on ridges at points of attachment by cell processes, which suggests strong adhesion that is likely mediated by either adsorbed serum proteins from the medium to the surface, or by cell attachment factors expressed by CTPs on the PDMS surfaces (den Braber, 1996b).

Most investigations into the effects of microtextured surfaces comprising microchannels for in vitro cell studies have concentrated on individual cell morphology and not colony characteristics. In addition to knowledge of the effects of topography on individual cells, the successful engineering of functional tissues will require understanding how cells behave with their neighbors (Brunnette and Chehroudi, 1999). Therefore, in vitro colony investigations can be clinically relevant in designing and predicting local tissue reactions to bone implants (Brunnette and Chehroudi, 1999). The present study is unique not only in combining CTPs with microfabricated surfaces, but also in quantifying both cell and colony behavior as a consequence of microtopographies. The drastic increase in colony AR and MAXL for the CTPs cultured on PDMS Channel textures are important parameters that should be considered when designing not only bone, but a variety of tissue engineering constructs that use CTPs. These effects may be particularly applicable when cells need to be localized in specific areas or when cell migration needs to be directed.

Conclusion

The effective use of bone marrow derived cells and CTPs for bone healing applications, is dependent upon the creation of a local environment in which CTPs will attach, proliferate, migrate, survive, and differentiate to form bone. Optimizing the efficiency of 2-dimensional and 3-dimensional scaffolds depends upon the design and control of variables that influence cellular behavior. Reproducible patterning of precise and desirable surface microtextures on scaffolds using microfabrication and micromachining technologies is one possible means of controlling CTP behavior through surface topography.

We have successfully created microchannels on PDMS substrates that affected both morphology and alignment of CTPs and their colonies. CTP migration on the surface increased along the channel axis, decreased across the channels and formed higher density colonies that were preferentially elongated along the direction of the channel axes. These results demonstrate a significant response of CTPs to topography and illustrate the practicality in modifying CTP behavior for bone applications through surface microtextures. Knowledge of CTP response to surface stimuli could lead to the incorporation of specific microtextures into surfaces of bone implant materials that would increase the recruitment, migration, and proliferation of cells to achieve a better and faster implant integration. By engineering CTPs and their location within an implant, it might be possible to control local tissue behavior, not only for bone, but for a variety of other tissues.