Measuring Cytoplasmic Stiffness of Fibroblasts as a Function of

Nov 9, 2018 - The cytoplasmic stiffness of cells plays a significant role during cell migration. As a cell migrates, differences in cytoplasmic proper...
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Measuring Cytoplasmic Stiffness of Fibroblasts as a Function of Location and Substrate Rigidity Using Atomic Force Microscopy Andrew J. Ford†,‡ and Padmavathy Rajagopalan*,†,§ Department of Chemical Engineering and §ICTAS Center for Systems Biology of Engineered Tissues, Virginia Tech, Blacksburg, Virgina 24061, United States

ACS Biomater. Sci. Eng. Downloaded from pubs.acs.org by 181.215.39.47 on 11/10/18. For personal use only.

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ABSTRACT: The cytoplasmic stiffness of cells plays a significant role during cell migration. As a cell migrates, differences in cytoplasmic properties occur that subsequently modulate migratory behavior. The properties of the substrate to which cells are adherent also play a role. To accurately measure the cytoplasmic stiffness of cells, we provide detailed instructions on how to assemble hydrogels that exhibit different elastic moduli, culturing cells on these substrates followed by a step-by-step process to measure and analyze the cytoplasmic properties of fibroblasts. In this study, we have measured the elastic moduli of cells at different locations to demonstrate how this property varies as a function of where the measurement is performed. The degree of anisotropy measured by the difference between cytoplasmic stiffness at the two edges of the cell also varied as a function of the elasticity of their underlying substrates. Larger differences in cytoplasmic stiffness between the leading and trailing edges were observed on substrates with a higher elastic modulus. The methods reported in this study can provide information on cellular properties, specifically, how the elastic modulus of cells can be probed and analyzed in vitro. KEYWORDS: Young’s modulus, cytoplasmic properties, cell migration

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INTRODUCTION

protrusions at the leading edge. Integrin-mediated adhesions to the cell substrate are then formed on these protrusions, after which the cell contracts followed by detachment from the substrate at the rear of the cell, resulting in forward motion.32,33 The initiating step in directed cell migration, which is polarization, results in a loss in symmetry wherein the front and rear of the cell are approximately aligned with the direction of locomotion.34 Asymmetric guidance cues promote the activation of cell receptors that induce signals leading to the generation of protrusions in the front and rear of the cell.35 In most cell types, actin polymerization plays a role in the formation of cellular protrusions or lamellipodia.34 The temporal regulation of the actin cytoskeleton through the class of motor proteins known as myosins (specifically myosin II) coupled with cell−substrate adhesions maintain the driving forces needed for migration.36,37

Cell migration plays an important role in many physiological processes throughout an organism’s lifetime. These processes include gastrulation,1 the development of different organs in the body,2 wound healing,3 disease,4 cancer progression,5 and the immune response.6,7 Several directional cues exist in vivo that influence a cell’s decision to migrate in a particular direction. These directional cues can be mechanical,8,9 electrical,10,11 optical,12 or chemical13,14 in nature. The effects of varying substrate elasticity or the concentration of adhesive ligands have been widely studied in the literature.15−24 Cells are capable of sensing changes in the concentration of chemicals or mechanical properties in their microenvironment and respond accordingly by altering their direction of migration.8,19,22,25−30 Cell migration may be influenced by gradients of soluble chemoattractants (chemotaxis), immobilized molecules (haptotaxis), and the modulus of the underlying matrix (durotaxis).24 To migrate, cells must exhibit a certain degree of anisotropy resulting in a well-defined leading and trailing edge.31 This polarization of the cell leads to the formation of cellular © XXXX American Chemical Society

Received: August 28, 2018 Accepted: October 16, 2018

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DOI: 10.1021/acsbiomaterials.8b01019 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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measure the elastic moduli of Balb/c 3T3 fibroblasts on soft hydrogels with variable stiffness. These studies enable us to make correlations between substrate rigidity and cellular elasticity.

Cytoplasmic stiffness is often used as a measure of the mechanical properties of the cell. The stiffness of the cytoplasm, which includes all cellular material inside the plasma membrane, with the exception of the nucleus, is primarily controlled by the properties of the cytoskeletal network. The three principal components of the cytoskeleton are actin filaments, microtubules, and intermediate filaments. Any changes to the arrangement, concentration or activation of the cytoskeleton can alter the physiological processes and functions of cells. A number of methods have been used for measuring cytoplasmic stiffness and changes to the mechanical properties of cells such as optical tweezers,38−40 magnetic twisting cytometry,41−43 micropipette aspiration,44,45 and atomic force microscopy (AFM).46,47 Optical tweezers and magnetic twisting cytometry monitor changes in ligand-coated micrometer-sized beads that are endocytosed by cells. These methods allow mechanical loads to be applied to intracellular structures via specific receptors. Optical tweezers apply forces to a single bead attached to an individual cell. The bead displacement is visualized and correlated to cell stiffness.40 Magnetic twisting cytometry applies a torque to a large number of ferromagnetic beads bound to a population of cells, and the average bead rotation is measured and an apparent stiffness can be determined.43 Although these methods allow mechanical loads to be selectively applied to intracellular structures via receptors, they also rely on the affinity and binding of foreign particles to the cells. Micropipette aspiration and AFM induce deformations to the cell by applying forces directly to the cell surface, which can be quantified and correlated to the force applied. In micropipette aspiration, the forces range from 10 pN to 1 × 104 nN. This technique is reliant on significant training and skill on the part of the experimentalist but it has relatively low resolution.48 AFM is an attractive option for measuring cytoplasmic stiffness due to the ability to obtain resolution at the nanometer scale. Furthermore, measurements can be obtained under dry or hydrated conditions. When cellular cytoplasmic stiffness measurements are conducted under hydration, the environment is closer to the hydrated states of many tissues. Modern AFM instruments can be coupled with sophisticated optical microscopes enabling measurements on precise locations within a cell. AFM has been utilized for more than two decades to image and investigate the properties of living cells.49,50 The mechanical properties of a wide variety of cells such as erythrocytes, endothelial cells, macrophages, osteoblasts, cardiac cells have been investigated using AFM.41,46,49−51 The elastic modulus (denoted as YM) of cells ranging from 100 Pa to 100 kPa have been reported in the literature.52 Moreover, it has been shown that multiple factors can affect the YM of cells. These include the origin of the cell within the body, disease and aging, cell differentiation, and the properties of the substrate to which a cell is adhered.53 Cellular processes such as division and migration, as well as the region of the cell that is being measured, can also result in differing modulus values.54−58 These factors can affect the organization of the cytoskeleton and subsequently cell stiffness.55,56,58 Cell polarization and the redistribution of cytoskeletal components are key steps in cell migration. Therefore, we seek to investigate whether a certain degree of anisotropy in cytoplasmic modulus could arise due to changes in substrate rigidity. In this report, we describe detailed methods to

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METHOD 1: CASTING POLYACRYLAMIDE (PAAM) HYDROGELS Materials. Reagents and Supplies. • Collagen type I−FITC conjugate from bovine skin (Sigma, St. Louis, MO, cat no: C4361) • 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer (Sigma, St. Louis, MO, cat no: H3375) • Irgacure 2959 (1-[4-(2-hydroxyethoxy)-phenyl]-2-hydroxy-2-methyl-1-propane-1-one) (Sigma, St. Louis, MO, cat no: 410896) • (3-Aminopropyl) triethoxysilane (3-APTES) (Sigma, St. Louis, MO, cat no: A3648) • Glutaraldehyde (25%v/v) (Electron Microscopy Sciences, 16210) • Acrylamide (40% v/v), (Bio-Rad, 1610140) • Bis-acrylamide (2% v/v) (Bio-Rad, 1610143) • Ammonium persulfate (APS) (Bio-Rad, 1610700) • Sulfo-SDA (sulfo-NHS-diazirine) (sulfosuccinimidyl 4,4′-azipentanoate) (Thermo Fisher Scientific, Waltham, MA, cat no: 26173) • 25 mm diameter glass coverslips (Thermo Fisher Scientific, Waltham, MA, cat no: 50-143-784) • 18 mm diameter glass coverslips (Thermo Fisher Scientific, Waltham, MA, cat no: 50-143-783) • Deionized (DI) water • RainX glass water repellent • Cotton swabs (Thermo Fisher Scientific, Waltham, MA, cat no: 14−959−102) • Sodium hydroxide (Sigma, St. Louis, MO, cat no: 415413) • Kimwipes (Thermo Fisher Scientific, Waltham, MA, cat no: 06−666−11C) Equipment. • Clean tweezers, razor blades, propane torch, mechanical pipet, and tips • Spectroline high-intensity longwave UV lamps: SB-100P (λ = 365 nm) (Thermo Fisher Scientific, Waltham, MA, cat no: 11−992−135) • UVP Mineralight Shortwave UV Quartz Display Lamp (λ = 254 nm) (Thermo Fisher Scientific, Waltham, MA, cat no: UVP95002607) • Gray scale masks (Benchmark Technologies, Lynnfield, MA) • Fisherbrand open-air rocking shaker (Thermo Fisher Scientific, Waltham, MA, cat no: 02−217−765) • Thermo Scientific digital dry bath/block heater (Thermo Fisher Scientific, Waltham, MA, cat no: 88− 870−001) • Cole-Parmer SS ultrasonic cleaner, heater/mechanical timer; 0.75 gal, 115 V (Cole-Parmer, Vernon Hills, IL, cat no: EW-08895−02) Procedure. Overview. • Step 1: Coverslip activation • Step 2: Preparation of hydrogel solution • Step 3: Hydrogel photopolymerization

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Figure 1. (A) Grayscale photomasks were used to vary the incident UV light to control the degree of cross-linking and hydrogel stiffness. (B) FITC-labeled Type 1 collagen is photoconjugated to the hydrogels using N-hydroxyl succinimide (NHS) and a hetero bifunctional cross-linker [sulfo-succinimidyl-diazirine (SDA)] through exposure to UV light at λ = 365 nm.

• Step 4: Photoconjugation of collagen to hydrogel Step 1: Coverslip Activation. Glass coverslips are silanized and activated with glutaraldehyde to promote adhesion between the PAAM hydrogel and the glass surface. 1. Remove any debris from 25 mm coverslips by streaming air across each coverslip surface. 2. While holding 25 mm glass coverslips with tweezers, pass each coverslip briefly through an oxidizing flame to remove hydrocarbon residues. 3. Let the coverslips cool for 5 min. 4. Coat each coverslip with 0.1N NaOH using a cotton swab. Air-dry coverslips in a chemical safety hood for 20−30 min. 5. Coat each coverslip with a thin layer of 3-APTES using a cotton swab and allow coverslips to sit for 10 min. 6. Place each coverslip in an individual well of a 6-well plate and wash the coverslips in DI water 3 times for 15 min each rinse. 7. Remove coverslips from the 6-well plate using tweezers and place on a Kimwipe, ensuring that the treated faces upward. Allow coverslips to dry at room temperature for 1 h. 8. Place each coverslip in individual wells of a new 6-well plate. 9. Make 1 mL of an 8% glutaraldehyde solution in DI water per coverslip. Add 1 mL of glutaraldehyde solution to each well and leave it for overnight on a rocking shaker. Note: Glutaraldehyde solution should be made f resh prior to coverslip activation. Do not try to save or reuse glutaraldehyde after coverslip activation. 10. Remove the glutaraldehyde solution from each well and discard it in a suitable waste container. 11. Wash the coverslips in DI water 3 times for 15 min each rinse. 12. Remove the coverslips from each well of the well plate and place on a Kimwipe, treated side up. Allow coverslips to dry at room temperature for a minimum of 2 h before further use. Step 2: Preparation of Hydrogel Solution.

1. Prepare 1 mL of 10% w/v ammonium persulfate solution in DI water in a micro centrifuge tube. 2. Prepare 1 mL of 3% w/v Irgacure 2959 solution in DI water in a micro centrifuge tube. Note: Irgacure 2959 has low solubility in water at room temperature. The solution should be heated to 80 °C for 30 min. 3. Prepare the hydrogel solution by mixing 250 μL of 40% v/v acrylamide, 360 μL of 2% v/v bis-acrylamide, 5 μL of 10% w/v ammonium persulfate solution, and 400 μL of 3% w/v Irgacure 2959. This will result in final concentrations of 10% v/v acrylamide and 0.72% v/v bis-acrylamide in the hydrogel solution. Step 3: Hydrogel Photopolymerization. PAAM hydrogels were prepared by exposing the hydrogel solution to ultraviolent light (UV) at a wavelength (λ) of 254 nm. Grayscale photomasks were used to control the amount of incident UV light exposed to the hydrogel solution, thereby controlling the degree of cross-linking and hydrogel stiffness (Figure 1A). 1. Remove any debris from 18 mm coverslips by streaming air across each coverslip surface. 2. Coat one side of the 18 mm coverslips with Rain-X using a cotton swab to make the coverslip surface hydrophobic. Allow the coverslips to dry for 15 min. 3. Turn on the UV lamp (λ = 254 nm) at least 15 min before use. 4. Pipette 30 μL drop of the hydrogel solution onto a 25 mm activated coverslip. 5. Gently place an 18 mm coverslip on top of the drop, Rain-X side down. 6. Place this coverslip sandwich on a stand with 18 mm on top and 25 mm coverslips on bottom. 7. Place the photomask above the coverslips making sure that the mask does not touch the coverslip surface. For this study, grayscale photomasks allowing either 30 or 80% of incident UV light were used to generate PAAM hydrogels with two different elastic moduli. 8. Position the UV lamp (λ = 254 nm) directly above the photomask and expose the hydrogel solution to UV light for 1 h. Note: As UV light can be harmful, ensure that appropriate steps are taken to prevent UV exposure. C

DOI: 10.1021/acsbiomaterials.8b01019 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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METHOD 2: MEASURING CYTOPLASMIC STIFFNESS OF FIBROBLASTS ON PAAM HYDROGELS Materials. Reagents and Supplies. • Fisherbrand Petri dishes with clear lid (Thermo Fisher Scientific, Waltham, MA, cat no: FB0875713) • Phosphate buffered saline (PBS) (Invitrogen Life Technologies, Carlsbad, CA, cat no: 10010023) • Dulbecco’s modified Eagle medium (Invitrogen Life Technologies, Carlsbad, CA, cat no: 12100046) • Penicillin-streptomycin (Invitrogen Life Technologies, Carlsbad, CA, cat no: 15140148) • Bovine calf serum (BCS) (Thermo Fisher Scientific, Waltham, MA, cat no: SH3007203) • Pyramidal SiN cantilever tips (Bruker AFM Probes, Camarillo, CA, cat no: DNP-10) • T75 culture flasks (Thermo Fisher Scientific, Waltham, MA, cat no: 10−126−11) • Murine Balb/c 3T3 cells (Balb/3T3 clone A31, American Type Culture Collection, Manassas, VA cat no: ATCC CCL-163) • 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer (Sigma, St. Louis, MO, cat no: H3375) • Trypsin-EDTA (0.25%), phenol red (Thermo Fisher Scientific, Waltham, MA, cat no: 25200056) Equipment. • Veeco BioScope II AFM (Veeco, Santa Barbara CA) • Eclipse TE2000U microscope (Nikon Instruments Inc., Melville, NY) • Hemocytometer (Thermo Fisher Scientific, Waltham, MA, cat no: 02−671−51B) • Fisherbrand accuSpin 1/1R benchtop centrifuge (Thermo Fisher Scientific, Waltham, MA, cat no: 75−400− 102) • In-VitroCell ES NU-5800 direct heat 7 ft3 (200L) CO2 Incubator (Nuaire, Plymouth, MN, cat no: NU-5800) Procedure. Overview. • Step 1: Fibroblast culture and seeding onto PAAM Hydrogels • Step 2: Setup of AFM • Step 3: Obtaining force−distance measurements on fibroblasts • Step 4: Analysis of force−distance curves Step 1: Fibroblast Culture and Seeding onto PAAM Hydrogels. 1. Maintain Balb/c 3T3 fibroblasts in T-75 flasks at 37 °C under a humidified atmosphere at 5% CO2. Fibroblasts should be cultured in DMEM supplemented with 10% v/v BCS and 2% v/v penicillin-streptomycin. 2. Remove 6-well plate containing FITC labeled collagen conjugated PAAM hydrogels from 4 °C. Transfer a hydrogel to a 50 mm Petri dish in a sterile biosafety cabinet and add 2 mL of 50 mM HEPES. Hydrogels can be stored at 4 °C. 3. Sterilize the FITC-labeled collagen conjugated PAAM hydrogels under germicidal UV light in the biosafety cabinet for 1 h. While the hydrogel is sterilizing, prepare fibroblasts for seeding.

9. Following photopolymerization, turn off the UV light, and gently remove the top 18 mm coverslip from the hydrogel using a razor blade and tweezers. 10. Place each hydrogel in a 6-well plate. 11. Rinse the gels (6 times) with 50 mM HEPES for a minimum of 15 min each rinse to remove any unpolymerized monomers. 12. Store the gels for a maximum of 1 week at 4 °C until further use. Step 4: Photoconjugation of Collagen to Hydrogel. To promote the adhesion of cells to the PAAM hydrogels, we photoconjugated FITC-labeled type 1 collagen to the hydrogels using N-hydroxyl succinimide (NHS) and a hetero bifunctional cross-linker {sulfo-succinimidyl-diazirine (SDA)} through exposure to UV light at λ = 365 nm (Figure 1B). 1. Dissolve 4 mg/mL of FITC-labeled collagen in 0.02 M acetic acid by sonicating for 30 min to generate a stock solution. Store this FITC-labeled collagen stock solution at 4 °C wrapped in aluminum foil to prevent exposure to light. 2. Prepare a 1 mL aliquot of FITC-labeled collagen by diluting the stock solution with 50 mM HEPES to the desired concentration. Sonicate this diluted solution an additional 30 min. For this study, a 250 μg/mL solution was prepared. 3. Prepare a 10 mL of 0.2 mg/mL solution of sulpho-NHSdiazirine in DI water. 4. Add the sulpho-NHS-diazirine to the FITC labeled collagen at a 10:1 molar ratio. Allow the mixture to react at 4 °C for 8 h. This solution can be stored at 4 °C until further use and can be further diluted in 50 mm HEPES to a desired concentration. For this study, a final concentration of 50 μg/mL collagen was used. 5. Remove the 6-well plate containing polymerized PAAM hydrogels from refrigeration. Remove one gel from the 6-well plate using tweezers and wick away the buffer from the side of the gel using a Kimwipe. Allow the gel to sit at room temperature for 1 min. 6. Carefully, place 60 μL of the sulpho-NHS-diazirine conjugated, FITC-labeled collagen on top of the hydrogel. Place a square 25 mm coverslip on top of the gel to spread the collagen solution. 7. Place the UV lamp (λ = 365 nm) above the gel and expose to UV light for 10 min at room temperature. 8. Following exposure to UV light, turn off the lamp and carefully remove the top coverslip using tweezers. 9. Transfer the coverslip to a new 6-well plate. 10. Repeat steps 5−9 for any remaining hydrogels. Rinse the gels 6 times with 50 mM HEPES for a minimum of 15 min each rinse. 11. Visualize conjugation of FITC labeled collagen to PAAM hydrogels through fluorescence microscopy. 12. Gels can be stored in 50 mM HEPES up to 1 week at 4 °C until further use. Anticipated Results. PAAM hydrogels were polymerized using photomasks with gray scales of 30% or 80% according to previously established methods.59 The use of these photomasks resulted in hydrogels with elastic moduli of 46.7 ± 1.2 kPa or 126.7 ± 1.9 kPa (n = 3 gels, minimum 3 locations per gel), as determined by AFM, for gels polymerized using the 30 or 80% masks, respectively. D

DOI: 10.1021/acsbiomaterials.8b01019 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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4. Aspirate the cell culture media from a fibroblast containing T-75 flask. Wash the cells by gently dispensing 10 mL of PBS buffer using a pipet into the cell culture flask. Gently swirl the flask making sure that the buffer reaches every corner of the flask. Aspirate the PBS from the flask. Repeat this step 2 more times. 5. Pipette 5 mL of trypsin into the flask before placing it in the incubator at 37 °C for 3−5 min. 6. After 3 min, examine the flask under a microscope to ensure cells have begun to detach. If not, gently tap the underside of the flask to dislodge attached cells. Place the flask back in the incubator for an additional minute if the cells have not detached fully. 7. Transfer the flask back to the biosafety cabinet and add 15 mL of cell culture media in order to neutralize the trypsin. 8. Spread the media in the flask by gently pipetting using a 10 mL pipet to ensure all cells from the flask have come in contact with the medium. 9. Transfer the entire volume of media from the flask to a 50 mL Falcon tube. 10. Centrifuge at 120g for 5 min at room temperature. 11. Carefully, transfer the tube back to the biosafety cabinet without disturbing the cell pellet. 12. Aspirate the supernatant without disturbing the cell pellet using a Pasteur pipet. 13. Resuspend the cell pellet by gently pipetting 5 mL of warm media up and down into the tube until the pellet is no longer visible. 14. Count the number of cells using a hemocytometer according to the manufacturer’s instructions. 15. Aspirate the 50 mM HEPES from Petri dish containing the sterilized FITC-labeled, collagen-conjugated PAAM hydrogel. 16. Carefully add the volume of cell suspension corresponding to 4 × 105 cells to the top of hydrogel using a micropipette, spreading the cell suspension over the surface of the gel. 17. Bring the volume of media in the Petri dish up to 3 mL by gently pipetting media to the side of the dish. Ensure that the top surface of the hydrogel is submerged in media. 18. Transfer the Petri dish to an incubator and allow cells to adhere for 8 h at 37 °C. Step 2: Setup of AFM. Described below is the setup of a Veeco Bioscope II AFM for obtaining force−distance measurements of fibroblasts seeded on PAAM hydrogels in a liquid environment. A general schematic of an AFM setup is shown in Figure 2A. The use of an AFM coupled with an optical microscope allows the user to obtain cytoplasmic stiffness measurements at precise locations along the cell body (Figure 2B). The anisotropy in the elastic modulus of a cell can be measured by obtaining force−distance measurements at the leading and trailing edges of fibroblasts (Figure 2C). 1. Turn on the heated stage of the Bioscope II. Make sure the controller is set to 37 °C. 2. Insert a cantilever into the tip holder and then slide the tip holder onto the four prongs of the Bioscope II head. For taking AFM measurements on cells, a cantilever with an unsharpened tip and low spring constant is desired. Here, DNP-10 silicon nitride AFM probes with a spring constant of 0.06 N/m were used.

Figure 2. (A) Schematic of an AFM setup to measure the elastic modulus of cells. (B) Use of an AFM coupled with an optical microscope enables the measurement of elastic moduli at precise locations along the cell body. (C) Cellular cytoplasmic stiffness can be measured by obtaining force−distance measurements at the leading and trailing edges of fibroblasts.

3. Turn on Bioscope II and open the NanoScope software. In the NanoScope software, double-click on the yellow microscope icon in the top left corner to connect the software to the AFM. In the window that appears, select “Use Default Parameters”. 4. For obtaining force−distance curves to determine a cells elastic modulus, check the boxes for “Scan Parameter List,” “Ramp Plot,” “Ramp Parameter List,” and “Navigate” in the window that appears next. 5. Initialize the piezo motors by clicking “Wake Up” in the next window. 6. In the “Scan Parameter List,” click on the “Other” tab and make sure the Bioscope II is in contact mode. 7. Turn on the Nikon TE2000U microscope interfaced with the Bioscope II. 8. Place the Bioscope II head onto the EasyAlign. Using the knobs on the side of the Bioscope II head, align the laser onto tip of the cantilever. The laser should appear as a spot on the apex of the cantilever. For a DNP-10 cantilever, the sum signal displayed on the Bioscope II controller should be 6−8 V. 9. Place the Petri dish containing the cell-containing PAAM hydrogel onto the heated stage of the Bioscope II. 10. Carefully place the Bioscope II head on top of the Petri dish. The cantilever should not yet be in contact with the cell culture media. 11. In the “Navigate” window in the NanoScope software, press the Z-down button until the cantilever comes into contact with the media. At the point of contact, the sum signal displayed on the Bioscope II controller should drop to roughly 0.15 V. This occurs due to refraction of the laser as it passes into the media and requires laser realignment onto the cantilever tip. 12. Realign laser by shifting it to the left using the laser align knobs on the Bioscope II head. The sum should return to 6−8 V. E

DOI: 10.1021/acsbiomaterials.8b01019 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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of the cantilever during the ramp, as to ensure the cantilever is in contact with the cell. The resulting plot should appear as a flat line leading into a smooth upward curve as the cantilever indents into the cell. 10. Save a minimum of two curves per location of the cell indented by clicking “Capture.” 11. While the cantilever is still in contact with the cell, obtain an image of the cell being indented using the Nikon TE2000U microscope. This will provide a reference later for whether the force−distance curve obtained corresponds to the leading edge, trailing edge, or nuclear region. 12. Click “Withdraw” in the NanoScope software and move the cantilever to a new position on cell or to another cell. 13. Once force−distance measurements have been obtained for a sufficient number of cells, move the cantilever to a region above the glass coverslip or Petri dish. 14. Engage the cantilever and in “Ramp Parameter Settings” adjust the “Z Scan Size” to 2 μm. 15. Click the “Continuous Ramp” icon and adjust the “Z Scan Start” until the cantilever comes into contact with the surface. The resulting graph should appear as a flat line leading to a sharp linear increase. In the plot window, click and drag the two boundaries of the plot to either side of the flat, sloped region. Click “Update Force Sensitivity” and record the value reported. For the DNP10 cantilevers and a sum of 6−8 V, this value should be 15−25 nm/V. This value is the sensitivity of the cantilever and will be used to convert the signal from the photodetector into the force applied to the cells by the cantilever. 16. Withdraw the cantilever and raise the piezo to the maximum height. Remove the sample from the microscope and dispose of it in an appropriate biohazardous waste container. 17. In the NanoScope software, export the force−distance curves as .txt files. 18. Shut down the Bioscope II, Nikon TE2000U, and NanoScope software. Step 4: Analysis of Force−Distance Curves. The elastic modulus of a cell at a given indented location is obtained by fitting the raw data to a modified Hertz cone model (eqs 1 and 2).61

13. Using the Nikon TE2000U microscope and a low magnification objective, such as 10x, focus on the surface of the PAAM hydrogel through the eyepiece. Once focused on the surface, bring the microscope slightly out of focus to just above the hydrogel. In the NanoScope software, continue lowering cantilever until it comes into focus on the Nikon TE2000U. Note: Lowering the cantilever too much could cause premature contact between the cantilever and hydrogel and may result in damage to cells. 14. Adjust the dichroic mirror to focus the laser onto the photodetector. Adjust the knobs on the top of the Bioscope II head until the vertical position reads −1 V and the horizontal position reads 0 V on the Bioscope II controller. 15. In the NanoScope software, under “Scan Parameters”, set the “Scan Size” to 0 nm, the “Integral Gain” to 0.6, the “Proportional Gain” to 1.2, and the “Deflection Set point” to 0 V. 16. Under “Ramp Parameter Settings”, set “Channel 1 Data Type” to “Deflection Error,” the “Data Scale” to “Auto Scale,” and the “Scan Rate” to 1 Hz. Step 3: Obtaining Force−Distance Measurements on Fibroblasts. 1. In the NanoScope software, make sure the box next to “Use Joystick” is checked in the “Navigate” window. Using the joystick, position the tip of the cantilever next to a fibroblast of interest above the PAAM hydrogel. The position of the cantilever can be viewed through the Nikon TE2000U microscope. 2. Bring the cantilever into contact with the surface of the PAAM hydrogel by clicking “Engage”. Note the zposition of the piezo at the point of contact with the hydrogel. 3. Withdraw the cantilever from the hydrogel by clicking “Withdraw”. 4. Using the joystick position the cantilever over the region of cell to be indented. For this study, fibroblasts were indented at the leading edge, trailing edge, and nuclear regions of the cells. The leading edge was identified by broad lammellipodia found at the front of the cell in the direction of motion. 5. Bring the cantilever into contact with the fibroblast by clicking “Engage”. 6. Note the z-position of the piezo at the point of contact with the cell. 7. While the cantilever is still in contact with the cell, open the “Ramp Parameter Settings” and “Ramp Plot”. 8. In “Ramp Parameter Settings” set the “Z Scan Size” to 10% of cell’s height. The cell’s height can be determined by taking the difference of the z-positions when in contact with the hydrogel and the cell. The cell is only indented to 10% to avoid any substrate contributions to the elastic modulus from the PAAM hydrogel below. This becomes especially important on thin regions of the cell, where the underlying substrate can artificially inflate modulus measurements.60 9. Click the “Continuous Ramp” icon at the top of the NanoScope software. A plot of the voltage detected by the photodetector versus the piezo height or z position will appear in the “Ramp Plot” window. The “Z Scan Start” can then be changed to adjust the starting position

F = k(d − do) Ä É 2tan α ÅÅÅÅ E ÑÑÑÑ 2 F= Å Ñδ π ÅÅÅÇ 1 − ν 2 ÑÑÑÖ

(1)

(2)

where F = applied force, α = 18°, E = elastic modulus, k = spring constant of the cantilever, υ = Poisson’s ratio (constant = 0.5),53 d = deflection of the cantilever, do = deflection point during contact, and δ = indentation. Described below is the workflow for obtaining the elastic modulus from raw data in Microsoft Excel. 1. Import .txt files from force distance curves into Excel or other software. In a new sheet, copy and paste the photodetector voltage and z-position for one curve. 2. Plot the photodetector voltage (V) in volts vs the zposition (z) in nanometers. 3. Determine where the point of contact between the cantilever and the cell occurred by identifying the point where the deflection begins to increase from a flat F

DOI: 10.1021/acsbiomaterials.8b01019 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Figure 3. (A−C) Fibroblasts exhibit heterogeneous Young’s moduli profiles depending on the position of the cell indented and substrate rigidity. (D) Cytoplasmic stiffness of fibroblasts in the leading edge. (E) Cytoplasmic stiffness of fibroblasts in the trailing edge. (F) Anisotropy in cytoplasmic stiffness as a function of substrate rigidity.

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7. 8.

9. 10. 11. 12. 13. 14. 15.

horizontal line. Enter the z-position at this point of contact in a new cell, which will be denoted as zo. This will serve as a starting value when fitting the curve to the Hertz cone model. Calculate z − zo for all data points in a new column. Calculate deflection (d) by multiplying the photodetector voltage (V) by the sensitivity factor obtained on the rigid glass or Petri dish in a new column. Find the deflection point corresponding to the z-position where contact between the cantilever and cell is made. This is determined as the point where the slope of the curve begins to increase from a flat horizontal line. Take the average deflection of this point and three on either side. Enter this average value into a new cell. This value is do. Calculate d − do for all data points Calculate the force (F) applied to the cell by the cantilever for all data points by multiplying d − do by the cantilever spring constant (k). For this study, a cantilever with a spring constant of 0.06 N/m was used. Calculate indentation (δ) where δ = (z − zo) − (d − do). Plot force (F) vs indentation (δ). Calculate natural log of indentation and natural log of force in new columns. In the force versus indentation graph, adjust the data points selected so that no data points corresponding to a natural log #num error are shown. Fit the force vs indentation data with a power law curve and display the fitted power law equation on the graph. Adjust zo until the power in the equation is 2.0 Use the slope of the power law equation to calculate the elastic modulus where

E = slope

π (1 − υ2) 2tan α

16. Proceed to the next data set. The photodetector voltage and z-position can be copied and pasted into the first two columns of the spreadsheet and steps 3−15 repeated for any remaining force−distance curves. Young’s moduli of multiple curves obtained at the same location can be averaged. 17. Statistical significance and p-values can be calculated by a two-sample t test, assuming unequal variance. Statistical significance between different data sets is determined using ANOVA and compared post hoc using two-tailed t tests while applying the Bonferroni (multiple hypothesis testing) correction. For all statistical testing α = 0.05. All data are reported as mean ± standard deviation; n denotes sample size. Statistical significance is determined by p < 0.05.

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RESULTS To examine the role of substrate rigidity on cytoplasmic stiffness, fibroblasts were cultured on PAAM hydrogels with Young’s moduli of 46.7 or 126.7 kPa, as well as on highly stiff glass coverslips (∼50 GPa). When cultured on each substrate, fibroblasts exhibited highly heterogeneous Young’s moduli profiles, with values ranging from 1.0 to 27.0 kPa depending on the location of the cell that was indented (Figure 3A−C). It was found that fibroblasts exhibited the highest cytoplasmic stiffness at the leading edge, where Young’s moduli were 4.41 ± 3.52 kPa, 6.28 ± 7.41 kPa, and 10.96 ± 8.56 kPa for fibroblasts on 46.7 kPa PAAM gels, 126.7 kPa PAAM gels, and glass coverslips, respectively (n = 16 fibroblasts on 46.7 kPa PAAM hydrogels, n = 20 fibroblasts on 126.7 kPa PAAM G

DOI: 10.1021/acsbiomaterials.8b01019 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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Methods/Protocols

hydrogels, n = 15 fibroblasts on glass coverslips) (Figure 3D). The corresponding average Young’s moduli at the trailing edge were 3.48 ± 1.77 kPa, 2.38 ± 1.45 kPa, and 4.37 ± 3.06 kPa (Figure 3E). Large standard deviations resulted from cell to cell heterogeneity. Although the average Young’s moduli of the leading edge were statistically insignificant compared to average values at the trailing edge, both the leading and trailing edges were found to be statistically higher than for regions close to the nuclei for all substrates. Young’s moduli in regions close to the nucleus were 2.09 ± 1.07 kPa, 1.89 ± 1.12 kPa, and 2.06 ± 1.55 kPa for cells on 46.7 kPa PAAM gels, 126.7 kPa PAAM gels, and glass coverslips, respectively. Furthermore, when the fold-changes between the leading and trailing edge Young’s moduli for individual cells were analyzed, we observed increasing anisotropy in cytoplasmic stiffness with increasing substrate rigidity (Figure 3F). On average, the stiffness at the leading edge was found to be 1.71 ± 0.95-fold, 2.70 ± 0.83-fold, and 3.48 ± 2.0-fold higher than at the trailing edge for fibroblasts on the 46.7 kPa PAAM gels, 126.7 kPa PAAM gels, and glass coverslips, respectively.

ORCID

Padmavathy Rajagopalan: 0000-0001-9997-4953 Present Address ‡

A.J.F. is currently at 153A Science & Technology Center, Tufts University, 4 Colby St., Medford, MA 02155, United States. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We gratefully acknowledge financial support from the National Science Foundation (NSF DMR-1410341, CBET-1510920). We also acknowledge financial support from the Institute for Critical Technologies and Applied Sciences at Virginia Tech and the Computational Tissue Engineering Graduate Education Program at Virginia Tech.

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SUMMARY In this study, we have provided detailed methods for investigating changes to the cytoplasmic stiffness of fibroblasts when cultured on substrates with varying rigidities. We have shown changes in the cytoplasmic stiffness of fibroblasts depend on substrate stiffness as well as the location of the cell indented. Larger differences in cytoplasmic stiffness between the leading edge and trailing edge were observed on substrates with higher elastic modulus. Directed migration occurs when cells are exposed to asymmetric guidance cues, resulting in the activation of cell receptors that induce polarized signals leading to the generation of a leading and trailing edge.35 In most cell types, actin polymerization and the assembly of other cytoskeletal components are associated with the formation of the cell front.34 The temporal regulation of these cytoskeletal components coupled with cell−substrate adhesions is crucial to migration.36,37 As the cytoskeleton is altered during migration, the cytoplasmic stiffness of cells likely changes as actin and focal adhesions assemble and disassemble. Such cellular changes warrant the further correlation between cytoskeletal proteins such as actin and tubulin and cellular elasticity. Future studies will require the evaluation of cytoplasmic stiffness for a given cell as it is migrating to truly understand how this physical property relates to cell speed and directionality. However, such measurements are extremely difficult to conduct because continuous indentations of the cytoplasm can lead to cell damage or even death. A greater understanding of the complex interplay between cytoskeletal organization, migration, cytoplasmic stiffness, and substrate properties could lead to future clinical applications in disease and wound healing. By isolating the contributions to changes in cytoskeletal stiffness from individual components of the cytoskeleton as well as external migration cues, we will gain a greater knowledge of cell decision making during the migration process.

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AUTHOR INFORMATION

Corresponding Author

*Email: [email protected]. Tel: 1-540-231-4851. Fax: 1-540231-5022 H

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