Abstract Macrophages participate in immunity, tissue repair and tissue homeostasis. Activation of Toll-like receptors (TLRs) by conserved exogenous or endogenous structures initiates signaling cascades that result in the release of cytokines such as tumor necrosis factor α (TNFα). Extracellular substrate stiffness is known to regulate functions of non-immune cells through a process called mechanotransduction, yet less is known about how physical cues affect macrophage function or TLR signaling. To investigate this question, we cultured murine primary bone marrow-derived macrophages (BMMs) and RAW264.7 cells on fibronectin-coated polyacrylamide (PA) gels of defined stiffnesses (1, 20 and 150 kPa) that approximate the physical properties of physiologic tissues. BMMs on all gels were smaller and more circular than those on rigid glass. Macrophages on intermediate stiffness 20 kPa PA gels were slightly larger and less circular than those on either 1 or 150 kPa. Secretion of the pro-inflammatory cytokine, TNFα, in response to stimulation of TLR4 and TLR9 was increased in macrophages grown on soft gels versus more rigid gels, particularly for BMMs. Inhibition of the rho-associated coiled-coil kinase 1/2 (ROCK1/2), key mediators in cell contractility and mechanotransduction, enhanced release of TNFα in response to stimulation of TLR4. ROCK1/2 inhibition enhanced phosphorylation of the TLR downstream signaling molecules, p38, ERK1/2 and NFκB. Our data indicate that physical cues from the extracellular environment regulate macrophage morphology and TLR signaling. These findings have important implications in the regulation of macrophage function in diseased tissues and offer a novel pharmacological target for the manipulation of macrophage function in vivo. cell signaling, inflammation, innate immunity, mechanosignaling, mechanotransduction Introduction Toll-like receptors (TLRs) detect microbial structures and induce multiple signaling pathways, including those that promote the release of pro-inflammatory cytokines (1). Inappropriate or excessive release of potent inflammatory cytokines negatively affects tissue homeostasis, thus TLR signaling is tightly regulated (2). Surface-associated TLRs (e.g. TLR4, TLR2) directly bind extracellular ligand (3); however, other TLRs (e.g. TLR3, TLR7, TLR8, TLR9) are confined to the endosome and require ligand uptake for receptor activation (4–6). Ligand engagement is restricted to specific cellular compartments (4–10) through regulation of intracellular trafficking (8, 11, 12), proteolytic cleavage (2, 13–23) and post-translational modifications (24–29) of TLRs. Regulatory chaperones (e.g. gp96) provide an additional layer of control (11, 30–33). The extent to which biochemical or biophysical cues from the extracellular environment additionally regulate TLR signaling is not well understood. Substrate stiffness, hydrostatic pressure and flow velocity are features of the extracellular environment that are conveyed to cells by biophysical signaling. Physical inputs are translated into biochemical signals by cells via a process known as mechanotransduction (34–37). Classic mechanotransduction pathways are activated when integrins attach to the extracellular matrix (ECM), and initiate the rapid assembly of adhesion complexes (38–40). These complexes then, in turn, activate downstream signaling kinases such as rho-associated coiled-coil kinase 1/2 (ROCK1/2), cdc42, and the small GTPase, Rac, that regulate actin cytoskeletal rearrangements, phagocytosis and formation of lamellipodia, respectively (41–43). Stiffness of a material is measured by Young’s modulus of elasticity, and expressed in pascals (Pa). In vivo tissue stiffness ranges from very soft brain (<1 kPa) to very stiff bone (1 GPa) (44), which contrasts with traditional plastic and glass cell culture surfaces that are >2 GPa (45). The advent of in vitro growth substrates that can be ‘tuned’ to specific, physiologically relevant, stiffnesses has been critical to our understanding of the importance of biophysical signals and mechanotransduction in modulating cellular functions (46). A pivotal study demonstrated that growth substrate stiffness regulates the differentiation of mesenchymal stem cells grown under the same cytokine and chemokine growth conditions (34). Study of somatic cells and stem cells has significantly advanced our understanding of the basic mechanisms and functional consequences of mechanotransduction (34, 47–49); however, much less is known about how mechanotransduction regulates immune cell function. Here, we show that macrophages respond to different growth surface stiffnesses by regulating inflammatory potential in response to TLR stimulation. In unstimulated cells, macrophage area decreased and circularity increased as growth surface stiffness decreased. TLR signaling and release of the pro-inflammatory cytokine, tumor necrosis factor (TNFα), increased as growth surface stiffness decreased. Inhibition of ROCK1/2 enhanced TLR4 signaling. Our data demonstrate that TLR signaling is modulated by physical cues from the extracellular environment and by a key mechanotransduction kinase. This novel regulatory mechanism of TLR signaling represents a potential target for the pharmacologic regulation of autoimmune- and sepsis-mediated inflammatory pathology. Methods Reagents and antibodies CpG DNAs were synthesized by Eurofins MWG Operon, Inc. (Louisville, KY, USA) or Invivogen (ODN 2395, Class C CpG oligonucleotide) (San Diego, CA, USA). Lipopolysaccharide (LPS 0111:B4) and ROCK1/2 inhibitor (Y-27632) were from Sigma (St. Louis, MO, USA). Alexa Fluor 488 phalloidin and ProLong Gold Anti-Fade with 4′,6-diamidino-2-phenylindole, dihydrochloride (DAPI) were from Molecular Probes/ThermoFisher Scientific (Waltham, MA, USA). Rabbit monoclonal antibodies against murine p38, ERK1/2, JNK/SAPK, ROCK1 and phosphorylated p38 (T180/Y182), ERK1/2 (T202/Y204) and JNK/SAPK (T183/Y185), NFκB (S536) were from Cell Signaling Technology (Danvers, MA, USA). The antibody against phosphorylated ROCK1 (T455/S456) was from Abcam (Cambridge, MA, USA), the antibody against p65 NFκB was from Santa Cruz Biotechnology (Dallas, TX, USA) and the antibody against α-tubulin (DM1A) was from ThermoFisher. Cell culture media and additives were from Corning CellGro (Tewksbury, MA, USA). Cell culture Murine RAW264.7 macrophages (American Type Culture Collection, Rockville, MD, USA) were grown in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% heat-inactivated low endotoxin fetal bovine serum (v/v) (VWR Life Science Seradigm, Radnor, PA, USA), sodium pyruvate (1 mM), 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (10 mM), l-glutamine (2 mM), and penicillin (100 U ml−1) and streptomycin (100 µg ml−1) (complete DMEM). As needed, RAW264.7 macrophages were detached with mechanical disruption. Murine primary bone marrow-derived macrophages (BMMs) were generated from the femurs and tibias from 8–12-week-old male C57BL/6 mice (Jackson Laboratory, Bar Harbor, ME, USA). Bone marrow cells were plated at 0.8 × 106 cells ml−1 in 10-cm non-treated tissue culture dishes in complete DMEM supplemented with 10% L-cell conditioned media (v/v). All cells were cultured at 37°C in a humidified incubator with 5% CO2. BMMs were detached with 0.25% trypsin (Corning CellGro), as needed. Cell viability was >95% (trypan blue exclusion) and cells were enumerated with a hemacytometer. All animal experiments were approved by Cornell University’s Institutional Animal Care and Use Committee (Animal Welfare Assurance A3347-01). Cornell University is accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International. Preparation of polyacrylamide gels Polyacrylamide (PA) gels of uniform thickness were prepared by polymerizing the gels between two differently treated 22-mm glass coverslips. To adhere the PA gel to the bottom coverslip, this coverslip was pretreated with 1% polyethyleneimine for 10 min, followed by 30-min incubation in 0.1% glutaraldehyde, and three 5-min washes in 1× phosphate-buffered saline (PBS). The second (top) coverslip was lightly coated with Rain-X® (Houston, TX, USA) to create a hydrophobic surface. Excess reagent was removed by buffing with a lint-free wipe. PA mixtures were prepared with stiffness-specific ratios of bisacrylamide:acrylamide, along with HEPES (14 mM), tetramethylethylenediamine (0.0054%) and ammonium persulfate (0.05%) as previously described (46). Once gels were fully polymerized in a vacuum chamber, the top hydrophobic coverslip was removed and the newly exposed gel surface was coated with sulfa-SANPAH [sulfosuccinimidyl 6-(4′-azido-2′-nitrophenylamino)hexanoate] (0.2 mg ml−1, Sigma), a compound that contains an amine-reactive N-hydroxysuccinimide (NHS) ester and a photoactivatable nitrophenyl azide, and cross-linked with ultraviolet light for 10 min. Three washes in 50 mM HEPES (pH 8.0) removed excess reagent, and the gels were incubated with fibronectin (20 µg ml−1, Corning, Bedford, MA, USA) overnight at 4°C. Excess fibronectin was removed with three washes in molecular grade water, and the gels were equilibrated in appropriate media at 37°C for 1 h prior to cell attachment. Attachment kinetics BMMs and RAW264.7 cells were detached from traditional tissue culture surfaces, and incubated in suspension for 30 min prior to being transferred to new tissue culture wells. Cells were allowed to adhere undisturbed for 0.5, 1, 2, 4, 6 or 18 h, after which the media were replaced with fresh media with or without TLR ligand. For the 0-h attachment time point, the TLR ligand was added to the cells as the cells were transferred into the new tissue culture well. Supernatant was collected after 6 h. Immunofluorescent staining and microscopy Cells on fibronectin-coated glass or gel coverslips were rinsed in 1× PBS, fixed in 3% paraformaldehyde in PBS, permeabilized in 0.1% Triton-X and blocked with 1% bovine serum albumin (BSA; ThermoFisher) in PBS. Filamentous actin was stained with Alexa Fluor 488 phalloidin (165 nM, ThermoFisher) in 1% BSA (w/v) in PBS. Coverslips and PA gels were mounted onto glass slides with Prolong Anti-Fade with DAPI (ThermoFisher). Slides were imaged with an Axio Imager M1 microscope (Zeiss, Thornwood, NY, USA) and an Axiocam MRm (Zeiss). Confocal images were obtained using a Leica TCS SP5 spectral confocal microscope (Buffalo Grove, IL, USA). Morphometric analysis Images from a minimum of 100 cells from each condition were collected with the 63× objective. Using ImageJ software (open source), cell perimeters were outlined and the surface area, circularity and aspect ratio of each cell were calculated. Circularity is mathematically represented by the equation: 4π (area) perimeter−2, where a perfect circle has a value of 1 and an infinitely elongated polygon has a value of 0. To calculate the aspect ratio of each cell, the software first determines the fitted ellipse of each cell and then calculates the ratio of major to minor axes [(major axis)/(minor axis)]. Measurement of cytokines TNFα and interleukin-10 (IL-10) in the supernatants were measured by enzyme-linked immunosorbent assay (ELISA) (BioLegend, San Diego, CA, USA) according to the manufacturer’s instructions. RNA purification and quantitative real-time polymerase chain reaction Total RNA was isolated and purified with TRIzol reagent (Life Technologies) according to the manufacturer’s instructions. RNA was quantified by spectrophotometry (Quawell Q3000, Palo Alto, CA, USA). Residual genomic DNA was digested with DNase I (Invitrogen), and cDNA was synthesized using Invitrogen reverse transcription reagents according to the manufacturer’s instructions on a thermocycler (BioRad My Cycler, Hercules, CA, USA). Quantitative real-time polymerase chain reaction (PCR) was performed in triplicate using Power SYBR Green master mix reagent (Applied Biosystems by Life Technologies) according to the manufacturer’s instructions. Primers for murine TLR9 (forward: 5′-CCTGGCTAATGGTGTGAAG-3’; reverse: 5’-CAAAGCAGTCCCAAGAGAG-3′) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (forward: 5′-TCCCACTCTTCCACCTTC-3′; reverse: 5′-ACCACCCTGTT GCTGTA-3′) were used at a final concentration of 580 nM, and cDNA was used at 20 ng per reaction. Reactions were run on an Applied Biosystems 7500 Fast Real-Time PCR System (Life Technologies) in standard mode. Cycle parameters were: 1 cycle at 50°C for 2 min, followed by 1 cycle at 95°C for 10 min, then 45 cycles at 95°C for 15 s and 60°C for 1 min. Expression data were obtained in the form of threshold cycle (Ct) values, and relative gene expression was calculated with the ΔΔCt method, using GAPDH as the reference housekeeping gene. Flow cytometry RAW264.7 cells and BMMs were plated on fibronectin-coated gels and glass for 24 h. Cells were incubated with or without 1 µM Cy3-labeled CpG DNA for 1 h at 37°C. Cells were detached and single cell suspensions were prepared in 1% BSA (w/v) in PBS with 0.1% sodium azide (w/v). Data were acquired with a BD FACSCantoTM II Flow Cytometer (BD Biosystems, San Jose, CA, USA) and analyzed using FlowJo software (FlowJo, LLC, Ashland, OR, USA). Cell viability assay RAW264.7 cells were treated with Y-27632 (2.5, 5, 10 µM) for 24 h. Viability was measured with the Cell Signaling kit-8 (Sigma) according to the manufacturer’s instructions and absorbance was read using a BioTek PowerWave XS microplate reader (Winooski, VT, USA). Protein immunoblotting Protein lysates were collected into β-mercaptoethanol reduced Laemmli loading dye and boiled prior to resolving by sodium dodecylsulfate–polyacrylamide gel electrophoresis. Proteins were transferred to nitrocellulose, blocked in 4% nonfat milk (w/v) in Tris-buffered saline with 0.1% Tween 20 (v/v) (TBST) and probed sequentially with antibody against phosphorylated and total p38, ERK1/2, SAPK/JNK, NFκB, followed by α-tubulin in 5% BSA in TBST. Films of blots were scanned and densitometry of scanned images was performed with ImageJ software. Statistical analysis Data were graphed and analyzed by the statistical tests described in the figure legends using GraphPad by Prism (version 7) software (La Jolla, CA, USA). Experimental replicate numbers are indicated in the figure legends. Results Macrophage adhesion and morphology are regulated by substrate surface mechanics Macrophage shape has been correlated with functional activity. Macrophages treated with LPS and interferon γ (IFNγ) to induce the classically activated (M1) phenotype exhibit a rounded shape. Induction of the alternatively activated (M2) phenotype by IL-4 and IL-13 causes macrophages to become elongated (50). When forced into an elongated shape using micropatterned growth surfaces, macrophage expression of arginase is increased, suggesting a more M2-like phenotype (50). Growth surface mechanics also regulate functions of a number of different cell types, including T cells and B cells (34, 51–56). We first compared the morphology of BMMs grown on fibronectin-coated glass to BMMs grown on fibronectin-coated 20 kPa PA gels which approximate the stiffness of skeletal muscle (57). BMMs on glass had a larger surface area than BMMs on 20 kPa PA gels (Fig. 1A). Circularity was lower in BMMs on glass compared to BMMs on 20 kPa PA gels (Fig. 1B). The marked morphometric differences in BMMs grown on glass versus 20 kPa PA gels illustrate the role that the physical extracellular environment plays in regulating cytoskeletal arrangement and basic morphologic features. Fig. 1. View largeDownload slide Substrate stiffness regulates macrophage morphology. BMMs were plated on fibronectin (fn)-coated glass or 20 kPa gels (A, B) or on fn-coated 1, 20 and 150 kPa gels (C, D) for 24 h. Cells were fixed and stained with Alexa Fluor 488 phalloidin (filamentous actin stain) and 4′,6-diamidino-2-phenylindole, dihydrochloride. Epifluorescent images of at least 100 cells from each condition were analyzed by ImageJ. Each dot represents morphometric data from an individual cell. (A, C) Cell surface area (in μm2). (B, D) Cell circularity in arbitrary units (A.U.). (E–H) BMMs were plated on fn-coated gels of indicated stiffnesses for 24 h and treated with vehicle (-) or 3 µM CpG DNA (+). Surface area and circularity were quantified as in (A) and (B), respectively. Representative fluorescent images from a minimum of three independent experiments are shown on the right. Scale bars = 10 μm. Note that the images of macrophages on glass are shown at lower magnification, but that the scale bar is still 10 µm. Mean ± SD from two independent experiments is shown. Data were analyzed by one-way ANOVA. *P < 0.05, ***P < 0.001, ****P < 0.0001. Fig. 1. View largeDownload slide Substrate stiffness regulates macrophage morphology. BMMs were plated on fibronectin (fn)-coated glass or 20 kPa gels (A, B) or on fn-coated 1, 20 and 150 kPa gels (C, D) for 24 h. Cells were fixed and stained with Alexa Fluor 488 phalloidin (filamentous actin stain) and 4′,6-diamidino-2-phenylindole, dihydrochloride. Epifluorescent images of at least 100 cells from each condition were analyzed by ImageJ. Each dot represents morphometric data from an individual cell. (A, C) Cell surface area (in μm2). (B, D) Cell circularity in arbitrary units (A.U.). (E–H) BMMs were plated on fn-coated gels of indicated stiffnesses for 24 h and treated with vehicle (-) or 3 µM CpG DNA (+). Surface area and circularity were quantified as in (A) and (B), respectively. Representative fluorescent images from a minimum of three independent experiments are shown on the right. Scale bars = 10 μm. Note that the images of macrophages on glass are shown at lower magnification, but that the scale bar is still 10 µm. Mean ± SD from two independent experiments is shown. Data were analyzed by one-way ANOVA. *P < 0.05, ***P < 0.001, ****P < 0.0001. We next asked whether BMMs exhibited morphologic differences when grown on fibronectin-coated PA gels that model physiologically and pathologically relevant differences in tissue stiffness. The softest 1 kPa PA gels approximate the stiffness of adipose tissue, the intermediate stiffness 20 kPa PA gels approximate the stiffness of skeletal muscle and the highest stiffness 150 kPa PA gels model fibrotic tissue (44). We observed that unstimulated BMMs grown on 20 kPa gels had a larger surface area and were less circular than BMMs grown on either the soft 1 kPa PA gels or stiff 150 kPa PA gels (Fig. 1C and D). There was no difference in surface area between BMMs grown on 1 kPa PA gels or 150 kPa PA gels; however, BMMs on 150 kPa PA gels were more circular than BMMs on 1 kPa PA gels. The aspect ratio, which is the ratio of the largest diameter of the cell to its smallest orthogonal diameter, was slightly larger in BMMs on 20 kPa PA gels than those on 1 or 150 kPa PA gels, which were not significantly different from each other (Supplementary Figure 1A). We conclude that in two-dimensional culture, physiologically relevant differences in stiffness regulate cell morphology of unstimulated BMMs. Stimulation of TLR9 by CpG DNA induces significant changes in macrophage morphology (28). We found that CpG DNA-stimulated BMMs on fibronectin-coated glass had a larger surface area than unstimulated BMMs; however, they did not have a statistically significant difference in circularity (Fig. 1E). We next asked whether macrophages on physiologically relevant stiffness surfaces also exhibited CpG DNA-stimulated changes in morphology. In response to CpG DNA, the surface area of BMMs on the softest 1 kPa PA gels did not change (Fig. 1F); however, CpG DNA did induce an increase in surface area of BMMs on the intermediate (20 kPa, Fig. 1G) and high (150 kPa, Fig. 1H) stiffness PA gels. CpG DNA stimulation decreased the circularity of BMMs on gels of each stiffness, but the most pronounced effect was seen in BMMs on the 150 kPa PA gels (Fig. 1E–H). The aspect ratio was unchanged by CpG DNA stimulation in BMMs on 1 and 20 kPa PA gels, but significantly increased in BMMs on 150 kPa PA gels and decreased on glass (Supplementary Figure 1B and E). These findings indicate that stimulation of TLR9 signaling by CpG DNA induces morphologic changes in BMMs, and that the stiffness of the underlying growth substrate regulates the morphology of both unstimulated and stimulated BMMs. Duration of cell attachment regulates spontaneous and TLR-induced TNFα secretion In cell culture experiments, cells are routinely detached from and subsequently allowed to reattach to growth surfaces. Convention dictates that there should be an interval between reattachment and use in any assay to allow the cells to ‘equilibrate’. Since the processes of attachment and reattachment disrupt cellular interactions with the extracellular environment and attachment to stiff surfaces will induce mechanotransduction with unknown effects on inflammatory signaling, we next asked whether macrophage TLR-mediated inflammatory cytokine release is disrupted by detachment and reattachment. BMMs and RAW264.7 cells were detached from untreated plastic or standard tissue culture plates, respectively, and then allowed to adhere to a new set of standard tissue culture wells for different lengths of time prior to stimulation for 6 h with TLR ligands (Fig. 2A). In BMMs, CpG DNA-induced secretion of TNFα decreased over the first 2 h of attachment, and then increased at 4 h and remained stable up to 18 h (Fig. 2B). The BMM response to LPS, a TLR4 ligand, was very similar except there was little initial decrease in LPS-induced secretion of TNFα, with an increase by 2 h that remained high for 6 h prior to reducing slightly at 18 h (Fig. 2C). In RAW264.7 macrophages, there was very little change in the level of TNFα induced by CpG DNA for the first 4 h (Fig. 2D), but responsiveness began to increase at 6 h of attachment and reached a maximum at 18 h (Fig. 2D). Maximal response to LPS in RAW264.7 cells also occurred at 18 h of attachment time (Fig. 2E). Fig. 2. View largeDownload slide The magnitude of TLR-induced TNFα secretion is regulated by time of attachment to tissue culture substrates. (A) Experimental design schematic. Briefly, macrophages were allowed to attach to tissue culture plates for the indicated times (white bars), and media were changed (black bars) before cells were stimulated for 6 h (gray bars). Supernatants were analyzed for TNFα by ELISA. (B) BMMs were stimulated with 3 µM CpG DNA (+). (C) BMMs were stimulated with 100 ng ml−1 LPS (+). (D) RAW264.7 cells were stimulated with 3 µM CpG DNA (+). (E) RAW264.7 cells were stimulated with 100 ng ml−1 LPS (+). Data are representative of three independent experiments each with a minimum of three biologic replicates. Mean ± SD are shown and results were analyzed by one-way ANOVA, *P < 0.05, **P < 0.01, ****P < 0.0001. Selected statistical comparisons are shown. Full statistical analysis results are available in Supplementary Figure 2D. Fig. 2. View largeDownload slide The magnitude of TLR-induced TNFα secretion is regulated by time of attachment to tissue culture substrates. (A) Experimental design schematic. Briefly, macrophages were allowed to attach to tissue culture plates for the indicated times (white bars), and media were changed (black bars) before cells were stimulated for 6 h (gray bars). Supernatants were analyzed for TNFα by ELISA. (B) BMMs were stimulated with 3 µM CpG DNA (+). (C) BMMs were stimulated with 100 ng ml−1 LPS (+). (D) RAW264.7 cells were stimulated with 3 µM CpG DNA (+). (E) RAW264.7 cells were stimulated with 100 ng ml−1 LPS (+). Data are representative of three independent experiments each with a minimum of three biologic replicates. Mean ± SD are shown and results were analyzed by one-way ANOVA, *P < 0.05, **P < 0.01, ****P < 0.0001. Selected statistical comparisons are shown. Full statistical analysis results are available in Supplementary Figure 2D. Strikingly, unstimulated BMMs (and to a lesser extent, RAW264.7 cells) released detectable TNFα during the first hour of attachment (Fig. 2B and expanded in Supplementary Figure 2A–C). These data suggest that during the process of detachment and reattachment, unstimulated macrophages spontaneously release TNFα. However, the spontaneous release rapidly dropped to very low levels within 2 h of attachment. After 2 h, the cells produce TNFα that is at or below the detection limit of the ELISA. We conclude that detachment results in the spontaneous release of TNFα at low levels that persists during the first hours of attachment, and that macrophages require at least 4 h of attachment to achieve a ‘steady state’ of responsiveness. Detachment and reattachment of macrophages to the underlying growth surfaces alters their response to TLR stimulation. Response to both CpG DNA and LPS were maximal in RAW264.7 cells at 18 h, as was BMM response to CpG DNA. Although the BMM response to LPS was lower at 18 h than at 4 h, the optimal adaptation and response were generally at 18 h. Thus, we chose to perform all subsequent experiments using macrophages that had equilibrated to the defined growth surface for 18 h. Cytokine secretion is regulated by growth substrate stiffness We next asked whether changing the stiffness of the growth substrate could, by itself, affect the ability of macrophages to respond to TLR9 and TLR4 stimulation. BMMs plated on fibronectin-coated 1 kPa PA gels secreted more TNFα in response to CpG DNA than BMM plated on either fibronectin-coated 20 or 150 kPa PA gels (Fig. 3A). Similarly, BMMs plated on 1 kPa gels secreted more TNFα in response to LPS (Fig. 3B). BMMs on 1 kPa PA gels also secreted more of the anti-inflammatory cytokine, IL-10, in response to CpG DNA (Fig. 3C). There was no significant difference in TNFα, or IL-10, released from BMMs on either 20 or 150 kPa PA gels. We conclude that TLR-mediated cytokine responses in BMMs are regulated by physiologically relevant differences in extracellular substrate stiffness. The absence of a difference in cytokine response by BMMs on 20 and 150 kPa PA gels indicates that the mechanoregulation of TLR signaling is not linear, but that there is a specific threshold for surface stiffness to regulate macrophage TLR response. Fig. 3. View largeDownload slide TLR-induced cytokine release by macrophages is regulated by substrate stiffness. (A–C) BMMs were plated on fibronectin (fn)-coated 1, 20 and 150 kPa PA gels for 18 h. (A) BMMs were stimulated for 24 h with media (-) or 3 µM CpG DNA (+). (B) BMMs were stimulated with media (-) or 100 ng ml−1 LPS (+) and secreted TNFα was quantified by ELISA. (C) As in (A) except secreted IL-10 was quantified by ELISA. (D–F) As in (A–C) except RAW264.7 cells were plated on fn-coated PA gels of the indicated stiffness and the attached cells were mounted in 4′,6-diamidino-2-phenylindole, dihydrochloride mounting medium and nuclei were quantified by ImageJ to normalize cytokine secretion per million cells. Data are representative of a minimum of three independent experiments, each with three biologic replicates. Mean ± SD is shown, and results were analyzed by one-way ANOVA. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. Fig. 3. View largeDownload slide TLR-induced cytokine release by macrophages is regulated by substrate stiffness. (A–C) BMMs were plated on fibronectin (fn)-coated 1, 20 and 150 kPa PA gels for 18 h. (A) BMMs were stimulated for 24 h with media (-) or 3 µM CpG DNA (+). (B) BMMs were stimulated with media (-) or 100 ng ml−1 LPS (+) and secreted TNFα was quantified by ELISA. (C) As in (A) except secreted IL-10 was quantified by ELISA. (D–F) As in (A–C) except RAW264.7 cells were plated on fn-coated PA gels of the indicated stiffness and the attached cells were mounted in 4′,6-diamidino-2-phenylindole, dihydrochloride mounting medium and nuclei were quantified by ImageJ to normalize cytokine secretion per million cells. Data are representative of a minimum of three independent experiments, each with three biologic replicates. Mean ± SD is shown, and results were analyzed by one-way ANOVA. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. Unlike BMMs, which adhered similarly to the different stiffness gels, RAW264.7 macrophages exhibited lower adhesion on soft gels (Supplementary Figure 3). Differential stiffness-dependent cell adhesion has been previously reported in other cell types (55, 58, 59). The differences in adherence between the two cells suggest a difference in attachment formation, and we hypothesized that TLR signaling in RAW264.7 macrophages would be less sensitive to differences in extracellular substrate stiffness than BMMs. Cytokine production by the RAW264.7 was normalized to the number of cells attached to the gels. Unlike BMMs, RAW264.7 macrophages plated on 1, 20 and 150 kPa PA gels responded similarly to CpG DNA (Fig. 3D). RAW264.7 showed a stiffness-dependent, biphasic LPS-induced TNFα secretion with more TNFα released on soft 1 kPa and stiff 150 kPa PA gels and a lower response on the intermediate stiffness 20 kPa PA gels (Fig. 3E). A similar biphasic response was observed in release of IL-10 in response to CpG DNA (Fig. 3F). Together, we conclude that TLR4- and TLR9-mediated secretion of pro- and anti-inflammatory cytokines in both the RAW264.7 cell line and primary BMMs is regulated by the physical stiffness of the underlying substrate, but that the two macrophage models differ in their responsiveness to the underlying stiffness. PA gel stiffness does not regulate mRNA expression of TLR9, uptake of CpG DNA or secreted factors Regulation of TLR9 signaling is complex, with multiple potential points for modulation by stiffness-dependent signals (2). Therefore, we next asked whether TLR9 mRNA expression was affected by stiffness. We collected total RNA from BMMs plated for 24 h on fibronectin-coated 1, 20 and 150 kPa gels and glass and performed quantitative real-time PCR for TLR9 and the housekeeping gene GAPDH. TLR9 mRNA expression was decreased in BMMs on 1 kPa PA gels compared to glass, but no significant differences in expression were observed in BMMs on each of the three stiffness PA gels (Supplementary Figure 4A). It is important to note that we observed more cytokine production in response to TLR9 stimulation in BMMs on 1 kPa gels, yet slightly lower TLR9 expression. Thus, the reduced TLR9 expression does not account for increased cytokine production observed in BMMs on 1 kPa PA gels. We next asked whether decreased substrate stiffness, which inhibited cytoskeletal rearrangements in response to CpG DNA, enhanced the uptake of CpG DNA to drive the increased release of TNFα by BMMs on soft 1 kPa gels versus the stiff 20 and 150 kPA gels. BMMs attached to fibronectin-coated 1, 20 and 150 kPa gels were incubated with Cy3-labeled CpG DNA, and detached to measure uptake by flow cytometry. There was no difference in median fluorescent intensity in BMMs incubated with Cy3-CpG DNA on any stiffness gel (Supplementary Figure 4B). Although CpG DNA induced RAW264.7 macrophages to secrete similar levels of TNFα regardless of whether they were attached to 1, 20 or 150 kPa gels, TNFα secretion by macrophages on any of the three stiffness gels was greater than by macrophages on glass (Supplementary Figure 4C). The same was true for BMMs (Supplementary Figure 4D). Both RAW264.7 and BMMs took up slightly more CpG DNA on glass than on 20 kPa (Supplementary Figure 4E and F). Thus, decreased responsiveness to TLR9 stimulation by macrophages on glass is not due to decreased uptake of CpG DNA. We conclude that the enhanced release of TNFα in response to CpG DNA stimulation by BMMs on 1 kPa PA gels is not due to an increase in TLR9 mRNA expression or increased ligand uptake. We next asked whether TLR9 signaling was indirectly regulated through secretion of soluble factors acting in an autocrine or paracrine fashion. RAW264.7 macrophages were plated on fibronectin-coated glass or 20 kPa gels for 24 h. Conditioned media from these cells were added to a second set of RAW264.7 macrophages plated on glass, and the cells were stimulated with CpG DNA for 24 h (Supplementary Figure 4G). There was no difference in TNFα secretion by macrophages on glass treated with either conditioned media (Supplementary Figure 4H). Similarly, RAW 264.7 macrophages on 20 kPa gels treated with each conditioned medium (Supplementary Figure 4I) did not differ in CpG DNA-induced TNFα secretion (Supplementary Figure 4J). We conclude that stiffness-dependent differences in TLR signaling are not due to production of secreted factors. ROCK inhibits TLR signaling through inactivation of p38 In fibroblasts and endothelial cells, increases in extracellular stiffness induce multiple mechanotransduction signaling cascades including Rho and its downstream effector ROCK (60, 61). In RAW264.7 cells plated on fibronectin-coated glass, ROCK1 was expressed at significantly higher levels and was constitutively phosphorylated, compared to cells plated on 1, 20 or 150 kPa gels (Fig. 4A). We observed several bands detected by the ROCK1 antibody (Supplementary Figure 5A), and the phosphorylated band corresponded to a lower molecular weight fragment and not full-length ROCK1, which is predicted to be 160 kDa. To investigate whether TLR signaling was regulated by the Rho/ROCK mechanotransduction pathway, we cultured RAW264.7 cells on tissue culture plastic and blocked ROCK1/2 with the small inhibitor, Y-27632 (ROCKi). Inhibition of ROCK1/2 did not affect viability (Supplementary Figure 5B), but enhanced TNFα secretion induced by various concentrations of LPS. The enhancing effect was more pronounced at higher concentrations of LPS (Fig. 4B). ROCK1 was barely detectable in BMMs, even when the cells were plated on fibronectin-coated glass (Supplementary Figure 5A), and ROCK1/2 inhibitor did not significantly change LPS-induced TNFα production by BMMs (Supplementary Figure 5C). Since ROCK1/2 inhibition augments TLR signaling, we conclude that when ROCK1/2 are normally active in cells, these kinases negatively regulate macrophage TLR-induced signaling. Fig. 4. View largeDownload slide ROCK enhances TNFα release by RAW264.7 cells. (A) RAW264.7 cells were plated on fibronectin-coated 1, 20 and 150 kPa PA gels or glass for 24 h. Cells were lysed in reduced sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS–PAGE) loading dye. Lysates were boiled, resolved by SDS–PAGE and analyzed by sequential immunoblotting for phosphorylated ROCK (top), total ROCK1 (middle) and α-tubulin (α-tub, bottom). (B) RAW264.7 cells were treated with the ROCK1/2 inhibitor, Y-27632 (10 µM), or vehicle control (media) for 1 h, followed by a 6-h incubation with LPS at the indicated concentrations (n = 4). TNFα in the supernatants was measured by ELISA. Mean ± SD is shown and results were analyzed by one-way ANOVA with Sidak’s multiple comparisons test ****P < 0.0001. Data are representative of three independent experiments. Fig. 4. View largeDownload slide ROCK enhances TNFα release by RAW264.7 cells. (A) RAW264.7 cells were plated on fibronectin-coated 1, 20 and 150 kPa PA gels or glass for 24 h. Cells were lysed in reduced sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS–PAGE) loading dye. Lysates were boiled, resolved by SDS–PAGE and analyzed by sequential immunoblotting for phosphorylated ROCK (top), total ROCK1 (middle) and α-tubulin (α-tub, bottom). (B) RAW264.7 cells were treated with the ROCK1/2 inhibitor, Y-27632 (10 µM), or vehicle control (media) for 1 h, followed by a 6-h incubation with LPS at the indicated concentrations (n = 4). TNFα in the supernatants was measured by ELISA. Mean ± SD is shown and results were analyzed by one-way ANOVA with Sidak’s multiple comparisons test ****P < 0.0001. Data are representative of three independent experiments. TLR signaling requires the activity of many kinases including the MAP kinases: p38, ERK1/2 and JNK (2). Therefore, we asked whether inhibition of ROCK1/2 would enhance the LPS-induced phosphorylation, and thus activation, of these kinases in RAW264.7 cells. LPS induced p38 phosphorylation in untreated RAW264.7 cells at 15 min, which remained high at 30 min, and was reduced to near background levels by 60 min. Pre-treatment with ROCKi also resulted in detectable p38 phosphorylation at 15 min, but phosphorylation continued to increase at 30 min and remained higher than uninhibited cells at 60 min (Fig. 5A, Supplementary Figure 6A). Phosphorylated ERK1/2 peaked at 15 min, and was reduced to near background levels by 60 min. Pre-treatment with ROCKi resulted enhanced ERK1/2 phosphorylation at 30 min and more sustained phosphorylation at 60 min (Fig. 5A, Supplementary Figure 6B and C). In contrast, phosphorylation of JNK was lower overall and equivalent in control and ROCKi-treated cells (Fig. 5A, Supplementary Figure 6D and E). The downstream transcription factor NFκB (p65) exhibited basal phosphorylation, and phosphorylation increased over 60 min following stimulation with LPS (Fig. 5B). Inhibition of ROCK1/2 augmented p65 NFκB phosphorylation (Fig. 5B). Fig. 5. View largeDownload slide Inhibition of ROCK enhances TLR signaling and prolongs TLR-induced p38, ERK 1/2 and NFκB phosphorylation. (A) RAW264.7 cells plated on tissue culture plastic dishes were treated with ROCKi (10 µM) or vehicle control for 1 h, and then treated with LPS (100 ng ml−1) for the indicated times. Cells were lysed in reduced sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS–PAGE) loading dye and boiled. Lysates were resolved by SDS–PAGE and analyzed by sequential immunoblotting for phosphorylated p38 (P-p38) and total p38 (p38) or phosphorylated ERK1/2 (P-ERK1/2) and phosphorylated JNK (P-JNK) and α-tubulin. (B) As in (A) except immunoblots were probed sequentially for phosphorylated p65 NFκB (P-NFκB), total NFκB (tot NFκB) and α-tubulin (α-tub). (C) BMMs plated on tissue culture plastic dishes were treated as in (A) and protein lysates were similarly resolved and immunoblotted for P-p38, total p38 (p38), P-ERK1/2 and α-tubulin (α-tub). (D) As in (B) except with BMMs. Data are representative of two (A) or three (B–D) independent experiments. Fig. 5. View largeDownload slide Inhibition of ROCK enhances TLR signaling and prolongs TLR-induced p38, ERK 1/2 and NFκB phosphorylation. (A) RAW264.7 cells plated on tissue culture plastic dishes were treated with ROCKi (10 µM) or vehicle control for 1 h, and then treated with LPS (100 ng ml−1) for the indicated times. Cells were lysed in reduced sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS–PAGE) loading dye and boiled. Lysates were resolved by SDS–PAGE and analyzed by sequential immunoblotting for phosphorylated p38 (P-p38) and total p38 (p38) or phosphorylated ERK1/2 (P-ERK1/2) and phosphorylated JNK (P-JNK) and α-tubulin. (B) As in (A) except immunoblots were probed sequentially for phosphorylated p65 NFκB (P-NFκB), total NFκB (tot NFκB) and α-tubulin (α-tub). (C) BMMs plated on tissue culture plastic dishes were treated as in (A) and protein lysates were similarly resolved and immunoblotted for P-p38, total p38 (p38), P-ERK1/2 and α-tubulin (α-tub). (D) As in (B) except with BMMs. Data are representative of two (A) or three (B–D) independent experiments. Primary BMMs cells stimulated with LPS also had detectable phosphorylation of p38 at 15 and 30 min and remained increased compared to the background levels even at 60 min. Pre-treatment with ROCKi resulted in earlier p38 phosphorylation at 5 min with a peak at 30 min, and sustained higher levels of phosphorylation at 60 min compared to vehicle control treated cells (Fig. 5C, Supplementary Figure 6F). Pre-treatment of BMMs with ROCKi resulted in detectable phosphorylated ERK1/2 at 5 min (Fig. 5C, Supplementary Figure 6G and H). ERK1 peaked at 15 min, but remained increased through 60 min. ERK2 continued to increase through 60 min. Unlike in RAW264.7 cells, phosphorylation of JNK was not detected at any time point in either vehicle control or ROCKi BMMs (not shown). In BMMs, NFκB p65 was phosphorylated with slower kinetics than in RAW264.7 cells, peaking at 60 min (Fig. 5D). This phosphorylation was slightly increased by ROCKi at 5 and 30 min (Fig. 5D). We conclude that blocking traditional ROCK-dependent pathways augments TLR signaling by enhancing phosphorylation of at least two key kinases, p38 and ERK1/2, and the downstream transcription factor NFκB. Discussion In this study, we have demonstrated that primary and immortalized macrophages sense and respond to the physical mechanics of their growth environment to regulate morphology and TLR signaling. We also demonstrated that the inhibition of ROCK1/2, which interrupts mechanotransduction signaling initiated when cells bind to the growth surface, augmented TLR signaling. ROCK1/2 inhibition both enhanced TNFα production and prolonged activation of TLR downstream kinases p38 and ERK1/2. Thus, our data support a model where physical stiffness of growth surfaces provides mechanical signals that specifically modulate TLR-mediated inflammatory responses. The most profound differences in morphology were observed between macrophages grown on traditional glass surfaces and those grown on PA gels, which are all significantly less stiff than glass. The differences in morphology in unstimulated BMMs were modest, but notable because they occur within the narrow range of physiologically relevant stiffnesses. The morphologic changes in BMMs that occur in response to CpG DNA stimulation were also dependent on stiffness. Stiffness-dependent changes in cell height, cell area and adhesion have been reported in rat alveolar macrophages when attached to glass (>1 MPa) and 40 kPa PA gels coated with collagen type I (62). Human monocyte-derived macrophages have a larger surface area and migrate faster on stiff PA gels (280 kPa) compared to soft PA gels (1–5 kPa) (63). Because macrophage polarization to the anti-inflammatory phenotype is regulated by increased stiffness (64) or simply by being forced into an elongated morphology (50), the relatively modest differences in morphology observed here could have significant consequences on macrophage differentiation within tissues of different stiffness. Our data, and those of others, support a role for mechanoregulation of TLR signaling. We did not observe significantly altered levels of TLR9 mRNA in macrophages attached to different stiffness surfaces, and we observed less and not more CpG DNA uptake in macrophages on lower stiffness surfaces. Thus, it is unlikely that expression level or ligand access explains the augmented response of macrophages on lower stiffness surfaces to TLR stimuli. Instead, our data show a role for substrate stiffness and the rho/ROCK pathway in regulating TLR signaling. These findings expand upon previous work and underscore the importance of physical cues in determining macrophage functional fate. The attachment of RAW264.7 cells to tissue culture plastic has been shown to activate the cell cycle kinase cdc42, an important mediator in the mechanoregulation of phagocytosis (65). Macrophages on stiff substrates also display increased LPS-induced phagocytosis compared to macrophages on soft substrates (65, 66). Studies have also shown that LPS-induced TNFα production decreased with increased stiffness (65); however, those studies used supraphysiologic concentrations of LPS (1 µg ml−1) and did not detect the subtle, and more physiologically relevant, differences in morphology and inflammatory activity we report here (65). In contrast, studies in the human promonocytic THP-1 cells found that stiffness did not regulate the release of TNFα (58), which is similar to our observations in RAW264.7 cells (Fig. 3). These discrepancies likely reflect cell-specific differences in the sensitivity to mechanosignals and underscore the need to carefully document mechanoregulation in each cell type. While integrins may be mechanosensory receptors that integrate sensation of surface mechanics with regulation of TLR signaling (67–74), several recent studies have implicated mechanosensitive members of the transient receptor potential (TRP) family, including TRPV4 and TRPM7, in the regulation of TLR4 signaling. For example, TRPM7 is critical for LPS-induced Ca++ flux, and TRPM7-deficient mouse macrophages exhibit reduced TLR4 endocytosis as well as both interferon regulatory transcription factor 3- and NFκB-dependent gene up-regulation (75). TRPM7-deficient mice were protected from lethality induced by intraperitoneal injection of LPS (75). Similarly, macrophages from TRPV4-deficient mice had a significantly lower response to LPS than wild-type macrophages (66). In contrast, another group found that in the absence of TRPV4, LPS induced greater macrophage and neutrophil recruitment, due to increased cytokine and chemokine production by TRPV4-deficient airway epithelial cells (76). Thus, additional studies will be required to determine the role of various mechanosensory receptors in TLR signaling. In several experiments, we noted that stiffness induced a biphasic response in macrophages. The morphology of BMMs and cytokine production by RAW264.7 cells in response to LPS were similar on soft (1 kPa) and stiff (150 kPa) gels, but different than those observed on intermediate stiffness (20 kPa) gels (Fig. 3). A similar biphasic phenomenon has been reported in human THP-1 cells treated with phorbol myristate acetate, where maximal release of IL-8 was observed when cells were grown on interpenetrating polymer networks with a stiffness of 9.9 kPa versus 1 or 389 kPa (58). One possible explanation is that macrophage response to an inflammatory stimulus is optimal at ‘average’ tissue stiffness (e.g. 20 kPa PA gels). Thus, conditions that either decrease stiffness (e.g. necrosis) or increase stiffness (e.g. fibrosis) can modulate macrophage responses. Changes in tissue stiffness via ECM remodeling occur in diseases such as cancer, cardiovascular disease and hepatic disease (35, 36, 52, 77–83). Changes in stiffness are not simply a pathologic consequence, but have also been shown to affect disease course. For example, increased tissue stiffness drives the metastatic potential of cancer cells (52, 82). Reducing tissue stiffness through the administration of a lysyl oxidase inhibitor, β-aminopropionitrile, reduced tumor stiffness and breast tumor progression in a mouse model (84). Macrophages play a critical regulatory role in inflammatory and fibrotic diseases; thus reprogramming macrophages in vivo is a potential way to treat diseases and repair tissue (85). Our studies suggest that interrupting macrophage mechanosensing in vivo will enhance TLR-mediated inflammation, and thus may have implications for treatment of chronic inflammatory and fibrotic diseases. Targeted modulation of tissue stiffness offers a novel approach to directly promote favorable macrophage functions and indirectly improve disease outcome via macrophage-mediated activities. In summary, our studies offer new insight into the role of mechanotransduction in regulating macrophage function, and identify specific cellular pathways that could be therapeutically targeted to reprogram macrophages. Funding This work was supported by the Center for Vertebrate Genomics, Cornell University (C.L.); the Cornell-Rochester Collaborative Trans-Institutional Pilot Award Program in Immunity and Infection to C.L.; an American Association of Immunologists Careers in Immunology Fellowship Program to E.G. and National Institutes of Health (T32OD011000) to E.G. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Conflicts of interest statement: the authors declared no conflicts of interest. Acknowledgements We thank Cynthia Reinhart-King for sharing her expertise in the fabrication of PA gels. We thank Siddhartha Sinha for his contributions to the conceptualization of the project. Specific contributions of the authors: E.G., C.H. and C.L. conceived the project and designed experiments; E.G., C.H. and J.C. performed experiments; E.G., C.H. and C.L. analyzed data and wrote the manuscript. 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International Immunology – Oxford University Press
Published: Apr 18, 2018
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