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. Author manuscript; available in PMC: 2015 Oct 1.
Published in final edited form as: J Mech Behav Biomed Mater. 2013 Dec 4;38:198–208. doi: 10.1016/j.jmbbm.2013.11.008

Hydrogels with Differential and Patterned Mechanics to Study Stiffness-Mediated Myofibroblastic Differentiation of Hepatic Stellate Cells

Murat Guvendiren 1, Maryna Perepelyuk 2, Rebecca G Wells 2,*, Jason A Burdick 1,*
PMCID: PMC4045659  NIHMSID: NIHMS551911  PMID: 24361340

Abstract

The differentiation of hepatic stellate cells (HSCs) into myofbroblasts is a key event in liver fibrosis. Due to the local stiffening of the extracellular matrix (ECM) during fibrosis, it is of great interest to develop mimics that can be used to investigate the cellular response to changes in mechanics. Here, we used a step-wise hydrogel crosslinking technique, where macromolecules are crosslinked using a sequence of addition then UV light-mediated radical crosslinking, to generate hydrogels with tunable stiffness. Freshly isolated HSCs remained rounded with lipid droplets and high levels of PPARγ expression on soft substrates (E~2 kPa); however, HSCs spread, lost their lipid droplets, and expressed high levels of α-smooth muscle actin (α-SMA) and type I collagen on stiff substrates (E~ 24 kPa). Similarly, fully differentiated cells reverted to a quiescent state when plated on soft substrates. Stiffness-induced differentiation of HSCs was enhanced in the presence of exogenous TGF-β1, a dominant signal in fibrosis. When the UV-induced secondary crosslinking was restricted with a photomask to spatially control mechanics, HSCs responded based on the local hydrogel stiffness, although they remained quiescent on stiff substrates if the stiff feature size was not sufficient to allow cell spreading. This hydrogel system permits the investigation of HSC response to materials with diverse levels and spatially heterogeneous mechanical properties.

1. Introduction

The differentiation of resident pericytes and fibroblasts to fibrogenic myofibroblasts is a common response to injury in all tissues, including the pancreas, liver, lung and kidney (Desmouliere et al., 2003; Thannickal et al., 2004; Ding et al., 2005). However, the persistence of myofibroblasts leads to excessive ECM deposition and contraction, resulting in fibrosis and further loss of organ function.

Hepatic stellate cells (HSCs) are one of the major sources of hepatic myofibroblasts (Benyon and Arthur, 2001; Perepelyuk et al., 2013). HSCs are a heterogenous group of cells comprising ~15% of the total resident liver cells and are major storage units for retinoids. In response to injury, quiescent HSCs differentiate into highly fibrogenic, proliferative, contractile and migratory myofibroblasts, a process also referred to as activation (Li and Friedman, 1999; Mann and Smart, 2002; Mann and Mann, 2008). The myofibroblastic differentiation of quiescent HSCs is characterized by the loss of vitamin A storing lipid droplets, de novo expression of α-smooth muscle actin (α-SMA), deposition of type I collagen, and production of profibrogenic cytokines (Friedman, 1999; Pinzani and Marra, 2001) such as transforming growth factor-β1 (TGF-β1), platelet-derived growth factor (PDGF) and fibroblast growth factor (FGF). To better understand fibrosis, there is growing interest in the influence of local mechanical properties on the differentiation of HSCs to myofibroblasts.

Anchorage-dependent cells sense the mechanics of their surroundings by pulling and pushing on the ECM, and in response, generate intracellular signals, a process known as mechanotransduction (Ingber, 2006). Matrix mechanics is a key parameter in regulating a range of cell behaviors such as focal adhesion formation, contraction, migration, proliferation and differentiation (Discher et al., 2005; Discher et al., 2009; Guvendiren and Burdick, 2012). Recent evidence also suggests that matrix stiffening, which is generally regarded as an outcome of disease, may be a contributing factor in the development of fibrosis and tumor formation (Georges et al., 2007; Levental et al., 2009; Liu et al., 2010), both of which involve significant mechanical changes at the cellular level as well as in whole tissue. Not surprisingly, there is growing interest in determining the effects of liver stiffness on cell behavior in chronic liver disease. Both noninvasive (transient elastography and magnetic resonance elastography) and invasive (rheology) measurements of liver stiffness illustrate an increase in stiffness with progression of fibrosis (10 to 100 fold compared to healthy tissue) (Foucher et al., 2006; Georges et al., 2007; Yin et al., 2007a; Yin et al., 2007b; Perepelyuk et al., 2013), and measurements of liver stiffness are widely used clinically to assess liver fibrosis.

Matrix stiffness-induced myofibroblastic differentiation of HSCs has also been confirmed with in vitro models. When cultured on stiff substrates (tissue culture plastic, or hydrogels with elastic modulus E > ~3kPa), freshly isolated quiescent HSCs lose their lipid droplets and express α-SMA, whereas they remain quiescent on soft substrates (E< ~3kPa)(Friedman et al., 1989; Gaca et al., 2002; Li et al., 2007; Olsen et al., 2011). Despite this observation, current in vitro models are limited. For example, in all of these studies the substrate modulus was uniform, preventing study of the mechanical heterogeneity typical of fibrosis (i.e., fibrous septa, with nodules). The severity and progression of liver fibrosis are correlated with nodule size, septal width and fibrosis area in such a way that small nodules, thick septa and large fibrosis areas are indications of severe cirrhosis (Kage et al., 1997; Garcia-Tsao et al., 2010; Bedossa et al., 2013). For instance, in rodents, the septal width is in the range of 20 to 80 microns for severe fibrosis (Patsenker et al., 2009). Liver biopsy of patients revealed that fibrous septae ranged from ~38 to ~1150 microns, such that ~38 to ~800 micron thickness denoted severe fibrosis, and ~78 to ~1150 microns corresponded to cirrhosis (Kage et al., 1997; Caballero et al., 2001; Nagula et al., 2006; Sethasine et al., 2012). Therefore, it is important to develop model hydrogel substrates that enable precise control of matrix mechanics with spatial control to investigate the activation of HSCs and other myofibroblast precursors.

For the studies reported here, we use hydrogels based on hyaluronic acid (HA), a natural component of the ECM involved in many biological processes (Burdick and Prestwich, 2011). HA can be chemically modified in a variety of ways to obtain hydrogels with tunable properties, such as degradation and mechanics (Marklein and Burdick, 2010; Burdick and Prestwich, 2011). In this work, we used HA macromers modified with methacrylates, which can react with both thiols and radicals for crosslinking reactions. By using a previously employed sequential crosslinking method (i.e., Michael-type addition reaction followed by UV induced radical polymerization) (Khetan and Burdick, 2010), hydrogels with elastic moduli mimicking that of healthy or fibrotic liver tissue were fabricated via Michael-type addition reaction alone or by secondary crosslinking via UV exposure of the initially formed hydrogel, respectively, to investigate HSC activation in both uniform and spatially controlled systems.

2. Materials and methods

2.1. Fabrication of hydrogel substrates for cell culture

Hydrogels were fabricated from methacrylated hyaluronic acid (MeHA), which was synthesized following a previously described procedure (Burdick et al., 2005). Briefly, sodium hyaluronate (Lifecore, 59 kDa) was dissolved in deionized water (1 wt%) and reacted with methacrylic anhydride (MA, Sigma-Aldrich; 2.4 ml MA per gram of HA) at pH 8.0 on ice for 8 h. This was followed by overnight incubation at 4°C, and further reaction with MA (1.2 ml MA per gram of HA) at pH 8.0 on ice for 4 h. The macromer solution was then dialyzed in deionized water (SpectraPor, molecular weight cutoff, 6–8 kDa) at room temperature for 4 days and lyophilized. The macromer exhibited methacrylate modification of each HA repeat unit via 1H NMR (Bruker).

Prior to the formation of hydrogels, an RGD adhesion moiety was coupled to MeHA macromers via a Michael-type addition reaction by incubating the GCGYGRGDSPG oligopeptide (GenScript) in 3 wt% solution of MeHA in PBS buffer containing 0.2 M triethanolamine (at pH 9, for 45 min at room temperature), such that the final concentration of RGD was 1 mM. Soft hydrogel (E ~2.1 kPa) substrates were fabricated using Michael-type addition crosslinking via introduction of dithiothreitol (DTT, 18% of methacylates) to the RGD-coupled MeHA solution. This solution (110 µl) was pipetted into a poly(dimethylsiloxane) mold (Sylgard 184® Silicone Elastomer Kit, Dow Corning, ~150 µm thick), covered with a methacrylated glass slide (Guvendiren et al., 2009), and incubated for 2 h at room temperature to obtain a hydrogel film. To fabricate stiff hydrogel (E ~24 kPa) substrates, soft substrates were equilibrated in PBS containing I2959 photoinitiator (0.05 wt%) for 30 min and exposed to ultraviolet light (Ominicure S1000 Spotcure, 10 mW cm−2) for 2 min. Hydrogels with patterned mechanics were fabricated by exposing the soft hydrogels to UV light through a photomask. In this study, transparent circles on a dark background, with diameters equal to 1000, 200, 50 and 25 microns, and transparent stripes with 500 micron width, were used as photomasks.

The mechanical properties of hydrogel substrates were measured by atomic force microscopy (AFM, Veeco Bioscope I) as described previously (Guvendiren et al., 2009). Briefly, force curves from individual points on the surface of the hydrogel (at least 20 points per gel) were obtained using a silicon bead AFM tip (spring constant of 0.06 N/m) and local elastic modulus values were calculated following a Hertz model for small indentations. For patterned substrates, methacrylated rhodamine (MeRho, 10 mM) was used as a contrast agent for confocal microscopy. MeRho was swollen into the hydrogels prior to secondary crosslinking and was covalently attached with UV exposure. During cell staining, DAPI diffuses from the soft regions much faster than stiff regions, which also allows pattern visualization.

For cell studies, hydrogel substrates (equilibrated thickness ~200 µm) were incubated in sterile PBS overnight, sterilized under a germicidal lamp for 2 h and incubated in culture media for 30 min before cell seeding.

2.2. HSC isolation and culture

Hepatic stellate cells (HSCs) were isolated as previously described (Uemura et al., 2005). Briefly, rat liver was digested in situ with 0.4% pronase (Roche Diagnostics) followed by 0.04% collagenase II (Worthington). The liver slurry was diluted with Minimal Essential Media (MEM) (GIBCO) and filtered through cheesecloth. The resulting cell suspension was centrifuged at 1500g for 5 min, resuspended in MEM with 0.002% DNAse (Worthington) and centrifuged at 1500g for 5 min. The wash step with DNAse was repeated once. Cells were then layered over a 9% Nycodenz (Sigma-Aldrich) solution and centrifuged at 1400g for 25 min. HSCs were collected in between layers and washed in MEM. HSCs were cultured in medium 199 (Invitrogen) supplemented with 10% fetal bovine serum (Gemini Bioproducts), 2% penicillin streptomycin and 1% fungizone amphotericin B (Gibco). Freshly isolated HSCs were seeded onto MeHA substrates placed in a 6-well plate or onto the plate itself (tissue culture plate, TCPS) as a control at 5×103 cells per cm2. The culture media was replaced the following day and every 2 days during culture. For treatment studies, transforming growth factor-β1 (TGF-β1, 0.1nM, R&D Systems) was administered in the culture media.

2.3. Cell staining, imaging and quantification

For immunostaining, cells were fixed (4% formalin for 10 min), permeabilized (0.1% triton X-100 for 15 min), and non-specific binding sites were blocked (10% horse serum in PBS for 45 min) at room temperature. Cells were then incubated with primary antibodies against α-SMA (mouse monoclonal anti–α-SMA clone 1A4 Ab; Sigma; 1:400) and peroxisome proliferator-activated receptor-γ (PPAR-γ; rabbit polyclonal anti–PPAR-γ Ab; Abcam; 1:400) in bovine serum albumin (BSA) solution (3% BSA in PBS) overnight at 4 °C. After several washes with the BSA solution, cells were incubated with secondary antibodies (AlexaFluor® 488 donkey anti-rabbit IgG and AlexaFluor® 594 chicken anti-mouse IgG; 1:200; Invitrogen) in BSA solution at room temperature for 2 h. Cells were then washed with PBS and nuclei were stained with DAPI in PBS. Cells were stained for vitamin A-containing lipid droplets using oil red O (ORO, 3 mg per ml in 60% isopropanol). Phase-contrast and fluorescent images of the fixed cells were captured on an Olympus BX51 microscope (B&B Microscopes Limited). For cell area measurements of HSCs during culture, at least 10 bright field images were taken from each hydrogel surface (at least 3 gels for each condition) at each time point using a Zeiss Axiovert 200 inverted microscope (Hitech Instruments, Inc.). NIH ImageJ was used to measure the cell area for each HSC.

2.4. Quantitative PCR

Total RNA was extracted with an RNeasy Plus Micro kit (Qiagen) according to manufacturer’s instructions. The RNA template was reverse transcribed with Super Script III Reverse Transcriptase (Invitrogen) and the resulting cDNA was used as a template for qPCR, carried out in a StepOnePlus instrument (Applied Biosystems) with three targets: α-SMA, type I collagen (COL1), and peroxisome proliferator-activated receptor γ (PPARγ). Ribosomal protein S12 (RPS12) was used for normalization. The sequences of the primers were as following: (F)TGACCAGGGAGTTCCTCAAAA, (R)GGCGGTCTCCACTGAGAATAA (PPARγ), (F)TGTGCTGGACTCTGGAGATG, (R)GAAGGAATAGCCACGCTCAG (α-SMA), (F)CATAAAGGGTCATCGTGGCT, (R)TTGAGTCCGTCTTTGCCAG (COL1A1), (F)CCTCGATGACATCCTTGG, (R) GGAAGGCATAGCTGCTGG (RPS12).

2.5. Statistical analysis

The data were processed using KaleidaGraph®. Cell area measurements are depicted as statistical box plots for each condition and time point. Remaining data are presented as mean ± standard deviation. ANOVA with Tukey’s HSD post hoc test of the means was used to make comparisons between groups (n=3 samples per group).

3. Results

3.1. Fabricating hydrogels with tunable stiffness

Hydrogel substrates displaying “soft” (E=2.1±0.7 kPa) and “stiff” (E=23.8±4.6 kPa) elastic moduli were fabricated using MeHA macromers and a multi-step crosslinking process (Figure 1) and assessed with atomic force microscopy. Specifically, after covalent incorporation of a cell adhesive peptide (arginine-glycine-aspartic acid, RGD) to the macromers (Figure 1B) for a final concentration in the hydrogel of 1.0 mM, soft hydrogels were fabricated via Michael-type addition reaction of methacrylates on MeHA with thiols on dithiothreitol (DTT). Stiff hydrogels were then fabricated by secondary crosslinking of the remaining methacrylates in the soft hydrogels by UV-induced radical polymerization in the presence of a photoinitiator (Igracure 2959, I2959, 0.05 vol%), as shown in Figure 1C. Final moduli were tailored via the amount of DTT used and extent of secondary crosslinking.

Figure 1.

Figure 1

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(A) Chemical structure of methacrylated hyaluronic acid (MeHA). (B) Schematic representation of the incorporation of RGD groups onto MeHA macromers to mediate adhesion. (C) Schematic of Michael –type addition polymerization to form “soft” MeHA hydrogels and secondary crosslinking via UV exposure (kinetic chains shown as red dashed lines) to form “stiff” MeHA hydrogels.

3.2. HSC interaction with substrates of varied stiffness

To assess the response of HSCs to hydrogel substrates with varied stiffness, freshly isolated cells were cultured on soft and stiff hydrogels as well as tissue culture plastic (TCPS) and cultured for up to 14 days. The morphology of HSCs was dependent on the substrate mechanical properties, as shown in Figure 2. Specifically, HSCs retained their initial cell area (~300 µm2) when cultured on soft hydrogels for 14 days, whereas HSC area significantly increased over the first 5 days of culture on stiff hydrogels (e.g., 224 to 4286 µm2 from day 1 to 5) and remained constant for the remaining culture period. The same behavior was observed for HSCs cultured on TCPS. Bright field images revealed that the rounded morphology of quiescent HSCs was well preserved on soft substrates; however, on stiff substrates (and TCPS) cell shape changed significantly and intense cellular spreading correlated with a loss of vitamin A-storing lipid droplets (Figure 2B). Transmitted light phase contrast images also demonstrated that the lipid droplets were well preserved after 14 days of culture on soft MeHA substrates and were completely lost on stiff MeHA substrates.

Figure 2.

Figure 2

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(A) Box plots showing cell area distributions of HSCs cultured on MeHA substrates mimicking the stiffness of healthy (soft, E=2.1±0.7 kPa) and fibrotic (stiff, E=23.8±4.6 kPa) liver tissue or on tissue culture plastic (TCPS) controls for up to 14 days. Bright field images (≥10) of cells (n≥30) were used for quantification of cell area at each time point (n=3 substrates for each condition) during culture. (B) Bright field and phase contrast images of HSCs on soft and stiff MeHA substrates after culture for 1 and 14 days. Note that lipid droplets appear white under transmitted light phase contrast mode (Scale bars, 20 µm.).

Immunostaining was performed to determine the effects of stiffness-induced morphological changes on HSC gene expression level, with PPARγ used as an indicator of quiescent HSCs, and α-SMA as characteristic of myofibroblastic differentiation of HSCs (Figure 3A and B). On soft substrates, the majority of the cells stained positive for PPARγ such that the fraction of PPARγ positive cells was 0.78, 0.74 and 0.68 for culture days 4, 7 and 14, respectively. On stiff substrates (and TCPS controls), the fraction of PPARγ positive cells (≤0.10) was significantly lower than for soft substrates and α-SMA stained cells increased significantly from 0.47 at day 4 to 0.79 at day 7, and finally to 0.85 at day 14 (Figure 3B). Note that some cells displayed no markers and some stained positive for both markers. Gene expression at 14 days (Figure 3C) confirmed these results such that PPARγ was significantly up-regulated when HSCs were cultured on soft substrates whereas α-SMA and COL1 were significantly up-regulated for stiff substrates and for TCPS controls.

Figure 3.

Figure 3

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(A) Fraction of cells stained positive for α-SMA and/or PPARγ, or without any stain (except for DAPI) when cultured on soft, stiff or tissue culture plastic (TCPS) substrates. * P < 0.001 compared to Soft groups. Error bars denote s.d. for 3 samples (n≥50 cells). (B) Fluorescent images of the HSCs on soft, stiff or TCPS substrates, immunostained for PPARγ (green), α-SMA (red), and cell nuclei (DAPI, blue) after 4, 7 and 14 days of culture. (Scale bars, 200 µm.) (C) Relative gene expression (mean fold difference in gene expression normalized to GAPDH and isolated cells prior to culture) of α-SMA, type 1 collagen (COL1) and PPARγ for HSCs cultured for 14 days on soft and stiff MeHA substrates and TCPS. # P < 0.01 compared to Soft group; * P < 0.001 compared to Soft group. Error bars denote s.d.(n=3).

3.3. Biochemically induced changes in HSC differentiation

The presentation of exogenous TGF-β1 did not induce significant morphological changes and differentiation of HSCs when they were cultured on soft MeHA substrates (Figure 4A and B). Although there was a slight increase in cell area and COL1 gene expression in the presence of TGF-β1, PPARγ was significantly up-regulated for HSCs cultured on soft substrates with TGF-β1 (Soft (+)). However, when administered during culture on stiff MeHA substrates (Stiff (+)), TGF-β1 significantly enhanced HSC spreading, such that mean cell area increased from 4020 µm2 in the absence of TGF-β1 (Stiff (−), Figure 4A) to 6300 µm2 in the presence of TGF-β1 (Stiff (+), Figure 4A), and differentiation also increased, as measured by gene expression for α-SMA and COL1 (Figure 4B).

Figure 4.

Figure 4

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(A) Box plots showing cell area distributions of HSCs cultured on soft and stiff MeHA substrates for 14 days in the presence (+) or absence (−) of TGF-β1. (B) Relative gene expression (mean fold difference in gene expression normalized to GAPDH and isolated cells prior to culture) of α-SMA, type 1 collagen (COL1) and PPARγ for the corresponding HSCs. * P < 0.0001 compared to Soft groups in (A), compared to Soft groups and Stiff (−) in (B) (all data from 3 samples, n≥50 cells).

3.4. Hydrogels with patterned mechanical properties and HSC response

To pattern substrate mechanics, a photomask was used to spatially control the UV exposure during the secondary crosslinking process (Figure 5A). Transparent regions on the dark photomask allowed light penetration only through defined regions with well-controlled shapes and sizes. To assess the fidelity of the patterned mechanics, methacrylated rhodamine (MeRho) was introduced into the soft hydrogel prior to secondary crosslinking, which selectively binds to the UV exposed regions (red, Figure 5B) via the radical polymerization. Fluorescent images revealed formation of patterns (circles and stripes) with relatively high fidelity, and AFM indicated a change in mechanics from 2.5±0.6 kPa (outside patterns) to 25.3±5.7 kPa (within patterns) (Figure 5B). Our results showed a partial loss of pattern size during transfer from photomask to hydrogel substrate. As an example, circle patterns changed in diameter (in microns) from photomask to hydrogel: 1000 to 987, 200 to 187, 100 to 85, 50 to 38, and 25 to not observable.

Figure 5.

Figure 5

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(A) Schematic showing two-step process to create hydrogels with patterned mechanics. Hydrogels formed via addition reaction were spatially stiffened via a UV light-induced secondary crosslinking process in the presence of a photomask. (B) Fluorescent images of hydrogel surfaces for corresponding photomasks. Methacrylated rhodamine (red) was introduced prior to the secondary crosslinking step to visualize the pattern since it is covalently bound to regions exposed to UV light. (C) Box plots showing cell area distributions of HSCs cultured on soft hydrogels with circular stiff regions with a range of diameter from 50 to 1000 microns. HSC location relative to pattern is denoted as “In” for inside the stiffer region or “Out” for on the soft region. (D) Fraction of HSCs stained positive for PPARγ and/or α-SMA when cultured on hydrogels with patterned stiffness for 14 days. NT refers to cells with no staining. (E) Corresponding fluorescent images of patterns (blue; circle diameter of 100, 200 and 1000 microns from top to bottom) and HSCs (red for α-SMA, green for PPARγ, blue for cell nuclei). (Scale bars, 200 µm.) (F) Fraction of HSCs stained positive for PPARγ and/or α-SMA, or without any stain (NT) cultured on hydrogels with striped patterns (blue indicating stiff regions ~400 µm width, indicated as In). (G) Corresponding fluorescent images of patterns and HSCs as in (E). # P < 0.01 and * P < 0.0001 compared to Out (for 3 samples, n=50 cells).

To investigate the influence of pattern size on HSC morphology and differentiation, soft hydrogels were fabricated with photomasks containing circular patterns of diameter equal to 50, 100, 200 and 1000 microns. For all samples, cell area increased significantly for HSCs that were in the stiffer regions (In, Figure 5C) when compared to HSCs that were on the soft regions (Out, Figure 5C) after 14 days culture. However, when the mean cell area values of the HSCs in the fibrotic regions were compared, the values for 200 and 1000 µm diameter regions were significantly higher than that of 100 and 50 µm. For instance, the mean cell area was 4450±240 and 3720±310 µm2 for 200 and 1000 µm and only 640±380 and 1030±350 µm2 for 50 and 100 µm, respectively. In most cases, there was more than one cell per circular pattern. The imaging and quantification of immunostaining for α-SMA and PPARγ are shown in Figure 5D and E and are consistent with the observed cell area distributions for films, except for 50 and 100 µm patterns. Specifically, HSCs mainly differentiated into myofibroblastic HSCs (indicated by mature α-SMA fibers, red) when they were located inside the stiff regions (see Figure 5E), whereas HSCs mainly stained positive for PPARγ (green) outside the stiff regions (Figure 5D). In the case of 50 and 100 µm patterns however, the majority of the HSCs stained positive for PPARγ independent of their location, suggesting that cell spreading was necessary for myofibroblast differentiation, even on a stiff substrate. We also investigated HSC morphology and differentiation on soft hydrogels with stripe patterns of 400 µm width, similar to the width of fibrous septae in cirrhotic livers, and observed that HSCs spread out and stained positive for α-SMA within the stiffer regions and remained fairly rounded and quiescent in softer regions (Figure 5F and G). Also, there was no differential response observed based on the distance of the cell from the pattern edge.

3.5. HSC response to substrate stiffness after mechanical priming

To investigate the effects of mechanical priming on HSC behavior, myofibroblastic HSCs (obtained after 14 days of culture on TCPS) were reseeded on soft and stiff MeHA hydrogel substrates and TCPS as a control (Figure 6). When reseeded on soft substrates, myofibroblastic HSCs immediately became rounded (at day 15) and remained rounded for an additional 14 days of culture (28 days of total culture) with a mean cell area (equal to ~200 µm2) significantly lower than that of myofibroblastic HSCs (~4200 µm2) prior to reseeding at day 14 (Figure 6B, D). Cells reseeded on soft substrates accumulated lipid droplets and stained positive for ORO and PPARγ after a total culture time of 28 days (including 14 days of mechanical priming on TCPS) (Figure 6C, E). On stiff substrates, HSCs remained spread, increased their mean cell area to 5500 µm2 at day 28 (Figure 6B, D), and stained negative for ORO but positive for α-SMA (Figure 6C, E). Finally, the relative expression of PPARγ significantly increased while that of α-SMA and COL1 significantly decreased for myofibroblastic HSCs when they were cultured on soft substrates for an additional 14 days (Figure 6F). For cells on stiff substrates and TCPS, the relative expression of the fibroblastic genes α-SMA and COL1 remained significantly up-regulated.

Figure 6.

Figure 6

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(A) Bright field and fluorescent images of myofibroblastic HSCs after mechanical priming on tissue culture plastic for 14 days. (B) Bright field images of mechanically primed HSCs further cultured for 14 days (total culture of 28 days) on soft or stiff substrates. (C) Bright field and fluorescent images of HSCs corresponding to (B) after 28 days of culture. HSCs were either stained for oil red O (ORO) or immunostained for PPARγ (green), α-SMA (red) and nuclei (DAPI, blue). (Scale bars, 200 µm) (D) Box plots showing cell area distributions and (E) quantification of immunostaining of HSCs from (B) after 28 days of total culture. (F) Relative gene expression (mean fold difference in gene expression normalized to GAPDH and isolated cells prior to culture) of α-SMA, type 1 collagen (COL1) and PPARγ for HSCs from (B). * P < 0.0001 Soft vs. TCPS, Stiff and TCPS (Day 14), in (D), and in (E) for PPARγ, and TCPS, Stiff and TCPS (Day 14) vs. Soft, for α-SMA. # P < 0.01 Stiff or TCPS (Day 28) vs. TCPS (Day 14), in (D). * P < 0.001 Stiff or TCPS (at Day 28) vs. TCPS (at Day 14) for COL1, and Soft (Day 28) vs Stiff and TCPS (at Day 28), and TCPS (at Day 14), for PPARγ, in (E).

4. Discussion

HSCs are key participants in liver fibrosis, differentiating into myofibroblasts, which are highly fibrogenic, proliferative, contractile and migratory cells. The general process of fibrosis involves local changes in matrix stiffness due to excessive ECM production and organization. There is growing evidence that these two events – matrix stiffening and myofibroblastic differentiation of HSCs – are correlated (Georges et al., 2007; Wells, 2008; Gordon-Walker et al., 2012; Perepelyuk et al., 2013). This is not surprising, as stem cells have been shown to sense the stiffness of their microenvironment, and in response, display changes in cell morphology, cytoskeletal organization, contractility, migratory behavior and gene expression (Discher et al., 2009; Burdick and Murphy, 2012). In liver fibrosis, there are spatial changes in matrix mechanics such that fibrous septa develop and reorganize the liver (Garcia-Tsao et al., 2010). The severity and progression of liver fibrosis correlates with septal width and fibrosis area in such a way that thick septae and large areas of fibrosis are typical of advanced cirrhosis (Bedossa et al., 2013). Thus, it is crucial to develop hydrogels with tunable and spatially controlled stiffness that can be used as in vitro culture systems in order to study the contributions of matrix stiffness and its spatial presentation.

We used a material platform that enables precise and spatial control of matrix stiffness. This was achieved by a two-step crosslinking process through the same reactive group on the HA precursors. The initial gelation was obtained by an addition reaction, which is used to fabricate hydrogels with stiffness (E~2 kPa) mimicking that of healthy tissue, and the hydrogel was then stiffened by secondary crosslinking via a UV light-mediated radical polymerization, mimicking the stiffness of fibrotic tissue (E~25kPa). This technique also allowed us to spatially control the stiffness using a photomask, as UV light was used during secondary crosslinking. Although we used materials based on HA, this sequential crosslinking technique can be applied to any reactive polymer that contains chemical groups that permit both types of reactions.

To demonstrate the utility of our system, we first investigated the changes in morphology and gene expression of HSCs on soft and stiff hydrogels. COL1 is a major component of the hepatic scar, and its production by activated stellate cells has been a major focus due to its role in the pathogenesis of fibrosis. Similarly, mature α-SMA fibers are a major signature of the myofibroblastic phenotype. Therefore, in this study, COL1 and α-SMA were genes of interest to represent myofibroblastic differentiation of HSCs, and PPARγ was used as the indicator of quiescent HSCs. HSCs were found to remain quiescent with elevated levels of PPARγ expression on soft hydrogels (E~2kPa), whereas HSC area significantly increased with loss of vitamin A-storing lipid droplets within 5 days and significant upregulation of α-SMA and COL1 at day 14 when cells were cultured on stiff substrates (E~25Pa). Notably, however, cell spreading was required for differentiation, as cells on stiff but relatively small patterns remained quiescent.

A fraction of the cells displayed no markers and some stained positive for both markers during the course of differentiation. We believe that this is due to the heterogeneous behavior of cells as they down-regulate PPARγ or up-regulate α-SMA in response to matrix stiffness. Our results in Figure 2A show that HSCs respond to matrix stiffness and reach an equilibrium cell area distribution within 5 days on stiff substrates. However, reaching this fibrotic area distribution does not necessarily mean that there is complete myofibloblastic differentiation of all HSCs. In fact, our results show that even though ~90% of the cells lost their lipid droplets (and did not show PPARγ staining) only ~47% of the total cells stained positive for α-SMA. However, the percentage of cells with no markers decreased significantly with culture time (i.e., ~16% of cells on stiff substrates at Day 14. This heterogeneity and extent of differentiation likely changes with respect to culture time on these substrates.

TGF-β1 plays a critical role in mediating liver fibrosis as it stimulates ECM production such as the synthesis and secretion of COL1 by stellate cells. Our results showed that when HSCs were cultured on soft hydrogels, the presence of exogenous TGF-β1 was not enough to alter HSC morphology and α-SMA and COL1 gene expression levels. Therefore, a stiff microenvironment was required for HSCs to differentiate. However, when this requirement was satisfied, TGF-β1 was found to enhance stiffness induced morphological changes and expression levels of fibrogenic genes. These findings were consistent with previous studies (Li et al., 2007; Olsen et al., 2011).

To further demonstrate the utility of the sequential crosslinking approach, we fabricated hydrogels with spatially patterned stiffer regions within a soft hydrogel, to mimic spatial organization during liver fibrosis. For circular fibrotic areas with diameter ≤ 100, although we observed a significant increase in cell area (mean cell area equal to 640±380 and 1030±350 µm2 for 50 and 100 µm) when compared to soft regions (mean cell area ~300 µm2) HSCs remained quiescent. For circular fibrotic areas with diameter ≥ 200, the mean cell area values were much higher (4450±240 and 3720±310 µm2 for 200 and 1000 µm) than that of smaller diameter fibrotic regions as well as soft regions. These HSCs showed mature α-SMA fiber formation and significant levels of COL1. Although the available area was over 5000 µm2 for a 100 µm diameter circle for a single cell, due to the effects of cell crowding the observed mean cell area was much lower (~1030 µm2). These results clearly show that it is possible to block HSC spreading, and hence, differentiation by simply physically limiting the available area for cell spreading. This has not been shown previously for HSCs, but has been well described for mesenchymal stem cells where cytoskeletal organization, contraction and differentiation are strongly correlated with cell shape. In the presence of adipogenic and osteogenic mixed induction media, rounded cells display diffuse actin and vinculin, accumulate lipid droplets, and acquire an adipogenic phenotype. In contrast, spread cells display mature actin fibers and vinculin patches, and acquire an osteogenic phenotype (McBeath et al., 2004; Guvendiren and Burdick, 2010). We believe that both stiffness and cell spreading are required for HSC differentiation; however, since spreading seems to be the result of increased stiffness, it is possible that spreading alone is sufficient.

Finally, we investigated the effects of matrix stiffness on the reversion of myofibroblastic differentiation of HSCs. Our results showed that myofibroblastic HSCs revert to a quiescent state when further cultured on soft substrates for 14 days. However, on stiff substrates, myofibroblastic HSCs displayed enhanced morphological changes and levels of fibrotic genes. These results indicate that myofibroblastic differentiation of HSCs is reversible based simply on substrate stiffness. HSC reversion has recently been studied in detail, and it has been shown that in the setting of fibrosis regression roughly half of myofibroblasts undergo apoptosis, while the remaining cells revert to a quiescent-like but “primed” state (Kisseleva et al., 2012; Troeger et al., 2012). The role of mechanical factors in this process has not been examined but the method we report here could be used to better define the changes that result in loss of myofibroblasts.

5. Conclusions

In summary, we have developed a culture system that can be used to study the effects of matrix stiffness in a time-dependent and spatially-controlled fashion. With this system, we have confirmed the stiffness dependence of HSC differentiation, but our findings are likely to be widely applicable to other fibrosis-related cell systems as well as to other diseases such as cancer. With HSCs, we have demonstrated that the requirement for stiffness cannot be overcome with TGF-β, and importantly that cell spreading in addition to stiffness is necessary. Through the use of patterned substrates, we have shown that even in a relatively small region of increased stiffness (200 micron diameter), cells can become myofibroblastic, with clear differentiation outcomes even at pattern interfaces. This has significant implications for understanding the mechanism of fibrosis, suggesting that mechanical changes leading to myofibroblast differentiation could be highly localized and that there is a mechanical mechanism underlying the self-perpetuation of fibrosis. Additionally, the finding that changes in local mechanics can lead to myofibroblast reversion has important ramifications for antifibrotic therapy, suggesting that targeting the mechanical properties of a diseased organ may be therapeutically effective in preventing progression of fibrosis.

Highlights.

  • A sequential crosslinking approach was used to develop hydrogels with spatially controlled mechanics.

  • Hepatic stellate cells were cultured on hydrogel films with varying mechanical properties.

  • Myofibroblastic differentiation was dependent on mechanics, including in a spatially controlled presentation.

Acknowledgements

We acknowledge funding from a David and Lucile Packard Foundation Fellowship in Science and Engineering, a National Science Foundation CAREER award (to JAB), and the National Institutes of Health (NIH R01 DK-058123, to RGW).

Footnotes

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