Engineered Fibrous Networks To Investigate the Influence of Fiber

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Engineered fibrous networks to investigate the influence of fiber mechanics on myofibroblast differentiation Matthew Davidson, KwangHoon Song, Mu Huan Lee, Jessica Llewellyn, Yu Du, Brendon Baker, Rebecca G Wells, and Jason A. Burdick ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.8b01276 • Publication Date (Web): 11 Mar 2019 Downloaded from http://pubs.acs.org on March 16, 2019

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ACS Biomaterials Science & Engineering

Engineered fibrous networks to investigate the influence of fiber mechanics on myofibroblast differentiation Matthew D. Davidson‡∥, Kwang Hoon Song‡, Mu-Huan Lee‡, Jessica Llewellyn§∥, Yu Du§∥, Brendon M. Baker⊥, Rebecca G. Wells§‡∥, and Jason A. Burdick‡∥* ‡Department of Bioengineering, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States §Department of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States ∥ NSF Science and Technology Center for Engineering Mechanobiology, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States ⊥Department of Biomedical Engineering, University of Michigan, Ann Arbor, Michigan 48109, United States

*Corresponding author: Jason A. Burdick Department of Bioengineering, University of Pennsylvania, 210 South 33rd Street, Philadelphia, PA 19104, USA E-mail: [email protected]

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ACS Biomaterials Science & Engineering 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract Tissue fibrosis is a leading cause of mortality and is characterized by excessive protein deposition and altered tissue mechanical properties. In pathological fibrosis, as well as cancer related fibrosis, tissue pericytes and fibroblasts transition from a quiescent to a myofibroblastic phenotype. In vitro models are needed to better understand how these cells are influenced by their local microenvironment. Here, we developed a fibrous network platform to mimic the structure of extracellular matrix, where fibers consist of crosslinked hyaluronic acid hydrogels with controlled crosslink density and mechanical properties. As a model myofibroblast precursor, primary hepatic stellate cells were seeded onto fibers with either low (soft) or high (stiff) crosslink density either directly after isolation (quiescent) or following pre-culture on tissue culture plates (activated). In general, both quiescent and activated cells showed an increase in spreading, alpha smooth muscle actin expression, and the formation of multicellular clusters on soft fibers when compared to stiff fibers. Further, inhibition of alpha smooth muscle actin decreased activation of cells on soft fibers. This is likely due to fiber recruitment in soft fibers that increased local fiber density, whereas stiff fibers resisted recruitment. This work emphasizes the importance of substrate topography on cell-material interactions and shows that tunable fibrous hydrogels are a relevant culture platform for studying fibrosis and mechanotransduction in disease.

Keywords: electrospinning, hydrogel, hepatic stellate cells, myofibroblasts, fibrosis


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1. Introduction Fibrosis is a leading cause of mortality and is a consequence of aberrant healing when tissues are chronically injured or stressed. Fibrosis occurs in multiple organs including the lung, heart, kidney and liver, and may eventually lead to organ failure.1 Additionally, fibrosis often accompanies cancer and is emerging as an important factor in metastasis.2 The differentiation of tissue pericytes and fibroblasts into a fibrogenic myofibroblast phenotype is a central feature of fibrosis and their persistence and excessive extracellular matrix (ECM) deposition contributes to tissue stiffening.3 Importantly, mechanisms in tissue fibrosis and cancer-associated fibrosis are thought to be conserved.4 Within the liver, hepatic stellate cells, or stellate cells, are pericytes that are the major source of myofibroblasts.5,6 Although stellate cells normally aid in homeostasis in a quiescent state, their chronic activation or myofibroblastic differentiation plays a role in liver fibrosis and they are the major contributors to the liver microenvironment that leads to the development of hepatocellular carcionoma. In order to develop new therapies, it is critically important to understand the factors that contribute to stellate cell activation,7 including the biophysical cues within the microenvironment8,9 as well as soluble factors that contribute to changes in liver tissue stiffness.9,10,11 The ECM surrounding cells has tissue-specific stiffness, porosity, and non-linear strain stiffening behaviors that influence cellular functions including contraction, migration, and differentiation.12,13 Importantly, local ECM stiffness has emerged as a key driver of tumor metastasis and fibrosis progression, rather than solely being a consequence of disease.14,15 Initial studies using hydrogel substrates that mimic a healthy or fibrotic liver stiffness showed that stellate cells are highly mechanosensitive and differentiate towards a myofibroblast phenotype in stiff environments;8,16,17 similar relationships have been observed for myofibroblast precursors in other organs. Dynamic stiffening and softening hydrogel substrates have further shown that stellate cells become activated rapidly (within hours) after stiffening,17 or regress to an intermediate activated state with softening.18 Recently, viscoelastic hydrogels have highlighted the ability of substrate viscosity to shield stellate cells from substrate stiffness and attenuate activation.19 Clearly, local ECM mechanics contribute to stellate cell activation; however, cells in the liver reside in a fibrous ECM and in contrast, published studies on ECM mechanics have all been performed with cells seeded atop hydrogels lacking topography. In fibrous environments such as type I collagen gels, cells remodel the matrix to increase adhesion sites and locally stiffen their environment by increasing matrix density, which could



play important roles in stellate cell activation.20,21,22 Stellate cells have extensive cellular processes in vivo23 and when cultured on fibrous matrices,24,25 but these are lost during longterm culture on plastic or atop hydrogels; this may have important implications for studying their interactions with the matrix as well as signaling interactions with other cells. At the tissue level, the fibrous structure of the ECM along with its non-linear mechanics enables long-range force transmission between cells through fiber alignment.26,27 This long range force transmission could explain how cells communicate location and form multicellular clusters on fibrous matrices24,25 and during fibrosis.28 During liver injury and fibrosis progression, extensive fibrin deposition29 and fibrillar ECM synthesis occurs within the hepatic sinusoids and near fibrotic septa, respectively, and fibrillar ECM mechanics are altered via dynamic changes in enzymes that degrade30 and crosslink31 ECM. Thus, understanding how quiescent and myofibroblastic stellate cells respond to fibrillar matrices with varied mechanics is critical towards our understanding of stellate cell activation. Difficulties in tuning the properties of natural ECM without changing variables such as biochemical composition limit their use in studying the role of fiber mechanics on stellate cell activation. Recently, we designed electrospun synthetic hyaluronic acid (HA)-based fibrous hydrogels that are stable over time, closely mimic the microarchitecture and mechanics of fibrous ECMs, and influence mesenchymal stromal cell behavior.32 Such electrospun fibrous hydrogels have been used to investigate the role of local fiber recruitment on cell spreading, proliferation and focal adhesion signaling.33,34 Importantly, these hydrogels recapitulate the network properties of the ECM, which are largely lost with typical hydrogel substrates that lack such topography.35 In this study, we report a modified suspended fibrous hydrogel system where the mechanics of individual fibers can be altered while maintaining other features of the fibrous matrix. Using hepatic stellate cells as a model system, we show that this system enables study of the contribution of fiber mechanics to myofibroblast differentiation and collective cell behavior.

2. Materials and Methods 2.1. Material synthesis Methacrylated hyaluronic acid (MeHA) was synthesized as previously described.32 Briefly, 2 grams of HA sodium salt (Lifecore, 76 kDa) was dissolved in 200 mL DIH2O to achieve a 1% w/v solution. While mixing and maintaining a pH of ~8.5-9, 0.45 mL or 1.5 mL methacrylic


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anhydride (Sigma) was added dropwise to this solution, to achieve low or high modified MeHA. The solution was maintained on ice at a pH of ~8.5-9 for 8 hours, mixed for an additional 12 hours at room temperature, dialyzed for 7 days, and lyophilized. The final methacrylate modification was ~30 or ~97%, as quantified by 1H NMR (Bruker, 360 MHZ) (Figure S1). RGD (GCGYGRGDSPG, Genescript) and 5(6)-Carboxyfluorescein (FAM) (GCDDD-FAM) peptides were conjugated to MeHA via Michael addition between thiols on cysteines and methacrylates. Briefly, MeHA was dissolved in pH 8 triethanolamine buffer (Sigma) at a final concentration of 2% w/v. RGD (2mM) and FAM (0.5mM) peptides were added and the reaction was carried out for 12 hours at 37°C, dialyzed for 5 days, and lyophilized. Peptides were synthesized via standard solid supported Fmoc chemistry and the sequence was confirmed via MALDI. The spectra

for

the

FAM

peptide

EEEDGRQIKIWFPNRRMKWKK)17

(GCDDD-FAM)36

and

αSMA

blocking

peptide

(Ac-

have previously been published.

2.2. Fibrous network fabrication Solutions for electrospinning included 4% w/v MeHA, 3.5% w/v polyethylene oxide (Sigma), and 0.05% Irgacure 2959 dissolved in DIH2O.

Electrospinning was performed as

previously described,32 with constant needle (+26-27 kV), collector (-3-4 kV), and deflector (+6-8 kV) voltages and varied needle-to-collector distances and polymer extrusion rates. Electrospun fibers were collected on foil, methacrylated glass coverslips, or fabricated polydimethylsiloxane (PDMS) (SYLGARD 184, DOW) wells. Glass coverslips and PDMS were methacrylated as previously described.32,33 For indentation mechanical testing, ~1.2 ml of polymer solution was electrospun to achieve a final thickness of ~130 µm. For suspended fiber mats, ~185 µl of polymer solution was electrospun onto PDMS wells to achieve a final thickness of ~17 µm. After collection, fibers were purged under nitrogen and crosslinked with 15 mW/cm2 ultraviolet light (320-390 nm, Omnicure S1500 UV Spot Cure Systems) for 20 minutes. 2.3. Fiber characterization Fiber diameters were measured from scanning electron microscopy (SEM, FEI Quanta 600 ESEM, University of Pennsylvania Singh Center for Nanotechnology) images and confocal microscopy (Leica SP5) images using ImageJ (NIH). To measure hydrated fiber diameters, a super resolution technique was employed (Figure S2). Briefly, fluorescence line intensity profiles were acquired perpendicular to the long axis of fibers to encompass the entire fiber width and the full width at half maximum fluorescence intensity was used to approximate fiber diameters.



Fiber fluorescence intensity and fibrous network average pore areas were also measured using ImageJ. Single fiber mechanics were measured as previously described.32,33 Briefly, 125 µm PDMS troughs were fabricated using a 3D printed mold (Protolabs), methacrylated, and fibers were aligned perpendicular to the troughs by electrospinning onto a rotating mandrel collector moving at 10m/s. To enable visualization of fibers under bright field microscopy, fibers were hydrated in a 3% w/v solution of black RIT dye in PBS for 24 hours. 3-point bending tests of single fibers that spanned the troughs were performed using atomic force microscopy (Asylum MFP-3D, University of Pennsylvania Singh Center for Nanotechnology) with a calibrated cantilever (0.06 N/m) (µmasch) with a 22.5 µm polystyrene bead tip. Force and displacement measurements were used to calculate the Young’s modulus of single fibers.37 To measure the Young’s moduli of thick fiber mats, fibers were sandwiched between two pieces of PDMS containing 4 mm wells. Indentation testing (shaft loaded blister tester, Penn Center for Musculoskeletal Disorders core facility) was then performed using a 500 µm cylindrical tip and a 250 g load cell. The Young’s moduli were approximated using equations from Baker et al.33 2.4. Hepatic stellate cell isolation and culture Primary hepatic stellate cells were isolated from Sprague-Dawley rat livers as previously described.38 All studies involving animals were approved by the University of Pennsylvania Institutional Animal Care and Use Committee and followed the guidelines set out by the US Public Health Service Policy on the Humane Care and Use of Laboratory Animals. Rat livers were perfused and then digested in situ with a 0.4% pronase (Roche Diagnostics) and 0.04% collagenase II (Worthington) solution. The digested liver was then diluted in minimal essential media (MEM) and filtered through cheesecloth. This produced a cell suspension that was then washed twice in 0.002% DNase solution (Worthington). Stellate cells were subsequently isolated by density gradient centrifugation over a 9% Nycodenz (Sigma) solution at 1400x gravity for 25 minutes, then washed with MEM and stored on ice until seeding. Prior to seeding hepatic stellate cells on suspended fiber substrates, fibers were hydrated for 24 hours in PBS, washed 3x and then sterilized via a germicidal lamp for 15 minutes. Fiber substrates were then incubated in cell culture medium (phenol red free αMEM (Gibco), 10% v/v fetal bovine serum (Sigma) and 2% v/v penicillin streptomycin (Corning)) for 30 minutes. Cells were seeded on fibers at a density of 1.5 x 105 cells/cm2. Media was


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replaced every 3 days. For α-SMA blocking studies, the peptide was added to culture medium daily at a concentration of 25 μg/mL. 2.5. Cell imaging, staining, and quantification Cells were fixed at stated times in culture using warmed 4% paraformaldehyde (ThermoFisher). For immunolabeling with anti-αSMA antibodies (mouse monoclonal antiαSMA clone 1A4, Sigma), cells were first permeabilized in 0.1% Triton X-100 (Sigma) solution in PBS for 20 minutes at room temperature, then blocked with a 1% w/v bovine serum albumin (BSA)(Sigma) solution dissolved in PBS for 1 hour, and incubated with primary antibodies (1:500) in the blocking solution overnight at 4 °C. Cell-fiber substrates were then washed 3x with PBS and incubated with secondary antibodies (1:200) (AF 635, Goat anti-mouse IgG, Invitrogen) in blocking solution for 2 hours at room temperature. For the last 15 minutes of secondary antibody incubation, Hoechst 33342 (ThermoFisher) was added (50 µg/ml) to the solution. Finally, the cell-fiber substrates were washed 3x with PBS and stored in the dark until imaging. Labeled cells were then imaged using a confocal microscope (Nikon TE2000) and average fluorescence intensities of αSMA per cell were quantified on maximum z-projections using ImageJ. For live cell imaging, the cell membrane was labeled with CellMask deep red membrane dye (1:1000) (ThermoFisher) for 15 minutes in PBS containing 1% FBS, and then transferred to a custom imaging chamber in cell culture medium. Imaging was carried out on a spinning disk confocal microscope (Nikon TE2000) housed in a CO2- and temperaturecontrolled environmental chamber. Z stacks were acquired every 20 minutes for 15-20 hours. 2.6. Statistical analysis Student’s t-tests and One-way ANOVAs with Tukey post hoc tests were performed to determine significance, which was indicated by p1500) for soft and stiff fibers. Error bars represent s.e.m. Figure 2. Mechanical properties of electrospun fibers. (a) Three point bending analysis of single fibers using AFM to determine Young’s moduli of hydrated fibers (n=11). (b) Indentation of thick fiber mats to determine Young’s moduli of hydrated fibers (n=4). *p25 cells), as well as (g) normalized fiber recruitment per cell (n≥20). *p

Engineered Fibrous Networks To Investigate the Influence of Fiber

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