Cells Sensing Mechanical Cues: Stiffness Influences the Lifetime of

Cells Sensing Mechanical Cues: Stiffness Influences the Lifetime of Cell−Extracellular Matrix Interactions by Affecting the Loading Rate Li Jiang,†,# Zhenglong Sun,†,‡,# Xiaofei Chen,†,§ Jing Li,† Yue Xu,† Yan Zu,† Jiliang Hu,† Dong Han,*,∥ and Chun Yang*,† †

Institute of Biomechanics and Medical Engineering, School of Aerospace, Tsinghua University, Beijing 100084, People’s Republic of China ‡ Suzhou Institute of Biomedical Engineering and Technology, China Academy of Science, Suzhou 215163, People’s Republic of China § Beijing Institute of Aerospace Systems Engineering, Beijing 100076, People’s Republic of China ∥ National Center for Nanoscience and Technology, Beijing 100190, People’s Republic of China S Supporting Information *

ABSTRACT: The question of how cells sense substrate mechanical cues has gained increasing attention among biologists. By introducing contour-based data analysis to single-cell force spectroscopy, we identified a loading-rate threshold for the integrin α2β1−DGEA bond beyond which a dramatic increase in bond lifetime was observed. On the basis of mechanical cues (elasticity or topography), the effective spring constant of substrates k is mapped to the loading rate r under actomyosin pulling speed v, which, in turn, affects the lifetime of the integrin−ligand bond. Additionally, downregulating v with a low-dose blebbistatin treatment promotes the neuronal lineage specification of mesenchymal stem cells on osteogenic stiff substrates. Thus, sensing of the loading rate is central to how cells sense mechanical cues that affect cell−extracellular matrix interactions and stem cell differentiation. KEYWORDS: single molecule force spectroscopy, single cell force spectroscopy, loading rate, integrin, stem cells differentiation, substrate stiffness minimum free energy12/stochastic-elastic modeling frameworks13 have also been devoted to this issue. Despite these advancements, the mechanism of stiffness sensing remains unsolved. Among candidate stiffness sensors, integrins are known to undergo conformational changes in response to mechanical stimuli and are positioned atop the mechanical sensing pathway. Integrins adhere the cells to the ECM by binding to ECM proteins, e.g., collagen, laminin, fibronectin, vitronectin, etc. We previously demonstrated that soft ECM induces neurogenic differentiation of bone marrow mesenchymal stem cells (BMMSCs) cultured on collagen I-coated soft poly-

I

t has become increasingly apparent that every tissue has a characteristic ‘stiffness phenotype’. All cells in tissues and organs are exposed to extracellular matrix (ECM) stiffness and are specifically tuned to the stiffness of the particular tissue in which the cells reside.1,2 ECM stiffness, being a mechanical property, exerts its effects on a variety of cell behaviors such as proliferation, differentiation, apoptosis, organization, and migration and has thus been intensively studied in recent years.3−8 These phenomena suggest that mechanical cues are crucial factors in the design of implantable materials. How do single cells sense their mechanical environment? A simple mechanism may underlie this puzzling problem. Previous reports have demonstrated that ECM stiffness regulates cell functions through its impact on the contraction force of actomyosin fibers, the subcellular allocation of integrin, and the PI3K pathways.9−11 Modeling efforts based on © 2015 American Chemical Society

Received: May 25, 2015 Accepted: December 23, 2015 Published: December 23, 2015 207

DOI: 10.1021/acsnano.5b03157 ACS Nano 2016, 10, 207−217

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Figure 1. F−D curve analysis of the integrin α2β1−DGEA bond in vitro. (a) Representative F−D curves recorded using integrin α2β1-modified AFM tips on a PDMS substrate modified with DGEA or BSA. The F−D curves on soft substrates (5.1 × 103 Pa) showed the same trend as those on stiff substrates (2.72 × 106 Pa). (b) The binding probabilities of the integrin α2β1−DGEA/BSA bond on soft PDMS (orange) and stiff PDMS (green) (mean ± SEM, n = 3). (c) Distribution of 420 (on the soft PDMS) and 464 (on the stiff PDMS) rupture forces of the integrin α2β1−DGEA bond at a CRS of 1 μm/s after a contact force of ∼500 pN. The solid line represents the unimodal log-normal distribution fitting of the distribution histogram. The peak value of the rupture forces was determined as the maximum of the unimodal lognormal distribution.

ECM and myosin II light chain activation, respectively. We thus proposed that cells sense ECM mechanical cues via loading-rate sensing. Consistent with this model, reducing actomyosin pulling in BMMSCs with a low-dose blebbistatin treatment enhanced integrin internalization and led to neurogenic lineage specification on an osteogenic stiff substrate. This study provides novel insights into the mechanism underlying myosin II-dependent elasticity/topographic sensing by cells and offers a new approach to manipulating lineage specification in mesenchymal stem cells.

acrylamide gels by promoting the caveolae/raft-dependent endocytosis of integrin β1.14 As the rupture of integrin−ligand bonds can trigger caveolin-mediated endocytosis,12,13 these findings suggest that mechanical cues from the substrate may affect the kinetic properties of integrin-ECM protein complexes. In most substrate stiffness studies, gels are typically coated with collagen I as an ECM mimic. Cell adhesion to collagen I is mostly mediated by three integrin types, namely, α1β1, α2β1, and α11β1,15,16 of which integrin α2β1 is expressed in stem cells, osteoblastic cells and epithelial cells, and displays a higher affinity for collagen I than do other integrins.17,18 The present study therefore focused on the effect of ECM stiffness on the kinetic properties of the integrin α2β1-collagen I bond. As a complex large molecule, collagen I contains multiple binding sites for different types of integrins, as well as other membrane proteins, thus is not appropriate for use in the single-molecule force spectroscopy (SMFS)/single-cell force spectroscopy (SCFS). To rule out unexpected interactions between cells and collagen, we used DGEA, the specific integrin α2β1-binding sequence in collagen I,19 as a model peptide to evaluate integrin α2β1−ligand binding properties on the cell surface. Integrin−ligand bonds typically involve nanoscale interactions, in which the adhesion strength ranges from tens to hundreds of piconewtons. Atomic force microscopy (AFM)based approaches are the most frequently used nanotechnologies for detecting piconewton-level forces in biological systems ranging from microscale to nanoscale and has been widely used to study the kinetic properties of receptor−ligand bonds.20−23 By gently conjugating molecules or a cell to the cantilever, researchers can use AFM as a powerful approach to evaluate the interactions/bonds between transmembrane and ECM proteins on a molecular level or in living cells.24,25 With AFM assays, the loading rate, i.e., the changing speed of the force forged on the bond, and the rupture force can be quantitatively evaluated; and the dynamic properties of the bond can thus be approached. By using SMFS/SCFS, we observed that increasing the loading rate of the integrin α2β1− DGEA bond on the cell membrane not only promoted the most probable rupture force to increase, but also dramatically enhanced bond lifetime. The loading rate of the integrin− ligand bond is determined by the effective spring constant of the integrin−ECM system (k) and the actomyosin pulling speed (v), which are affected by the elasticity/topography of the

RESULTS Substrate Stiffness Promotes the Rupture Force of the Integrin α2β1−DGEA Bond in SMFS. To address whether the model peptide recapitulates the activity of the native ECM, we observed the cells adhesion features of DGEAcoated substrates. The cells did not adhere to polyacrylamide gels alone, whereas BMMSCs spread properly on DGEAcoated gels (Figure S1). Because the DGEA peptide is a welldocumented integrin α2β1 binding site on collagen I, these results suggested that the DGEA modification is appropriate for cell culture on compliant substrates. Rupture force is an important indicator of receptor−ligand bond kinetic properties.20,21,26,20,21,26 We therefore investigated the rupture force of the integrin α2β1−DGEA bond on compliant substrates both in vitro and in vivo using SMFS/ SCFS. For in vitro SMFS, polydimethylsiloxane (PDMS) was used instead of polyacrylamide gels as a soft/rigid (5.1 × 103 Pa/2.72 × 106 Pa) substrate to avoid strong interfacial interactions and to stabilize the force−distance curves (F−D curves). The system consisted of an AFM tip conjugated to integrin α2β1 via PEG chains and DGEA-modified PDMS substrates, with BSAmodified PDMS serving as a control. Representative F−D curves for integrin α2β1−DGEA/BSA are shown in Figure 1a. The first peak in the F−D curves is assigned to the nonspecific interaction between the AFM tip and the PDMS substrate. The second peak, which appears about 20−40 nm away from the first peak, denotes the rupture force of the integrin α2β1− DGEA bond because the typical length of a PEG-chain crosslinker is 20−40 nm.27 On the basis of Poisson statistics, ∼85% of unbinding events represent the rupture of a single bond28,29 when the binding probability (e.g., the proportion of curves 208


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Figure 2. F−D curve analysis of the integrin α2β1−DGEA bond at the cell−ECM interface. (a) Optical image of the SCFS operating system, showing a BMMSC at the end of the concanavalin-A-functionalized AFM cantilever. The picture is taken when the experiment is just completed and a trypan blue staining was carried out. Scale bar: 50 μm. (b) Representative F−D curves recorded using MSC-modified AFM tips on DGEA- or BSA-modified polyacrylamide gels. The F−D curves on soft substrates (500 Pa) showed the same trend as those on stiff substrates (1 × 105 Pa). (c) Binding probabilities of single BMMSC-DGEA/BSA bonds on soft polyacrylamide gel (orange) and stiff polyacrylamide gel (green) (mean ± SEM, n = 3). (d) Distribution of 136 (on the soft polyacrylamide gel) and 214 (on the stiff polyacrylamide gel) rupture forces of the integrin α2β1−DGEA bond. The peak value was determined as mentioned before.

displaying specific binding events, Figure 1b) is 28.0% ± 3.0%. Histogram analysis of 300−400 observed specific unbinding events demonstrated a log-normal distribution of the rupture force for the integrin α2β1−DGEA bond, in which the peak value of the rupture force was 83 and 119 pN for soft and stiff gels, respectively, at a cantilever retraction speed (CRS) of 1 μm/s (Figure 1c). Integrin dimers can undergo conformational changes upon activation that may have a major impact on ligand binding and signaling activities.30 According to the results from Miller et al.,31 integrin α2β1 binds ligands via the I-domain in the α subunit, but switching I-domain from a closed, inactive conformation to an open, active conformation is regulated by the I-like domain of the β subunit. TC-I 15 is a small molecule that docks with the integrin α2β1 I-like domain and can lock the heterodimer into the inactive conformation. To test the influence of integrin conformation on rupture forces, we applied TC-I 15 to inhibit the activation of integrin α2β1. Integrin α2β1-immobilized AFM tips were incubated for 2 h with 20 nM TC-I 15 (4527, Tocris Bioscience), and SMFS was then performed in 20 nM TC-I 15 buffer. Treatment with TC-I 15 significantly reduced the rupture force of the integrin α2β1− DGEA bond from 94.9 to 58.7 pN on a soft PDMS substrate; and from 130 to 77.3 pN on a stiff substrate (Figure S3a). Integrin inhibition also reduce the binding frequency of the integrin α2β1−DGEA bond to 17% ± 2% on soft PDMS; and 18% ± 3% on stiff PDMS, showing a significant effect (p < 0.01) of TC-I 15 treatment. These results suggested that the exposure of the I-domain of integrin α2 contributes to the high binding strength of the integrin α2β1−DGEA bond.

Substrate Stiffness Promotes the Rupture Force of the Integrin−DGEA Bond in SCFS. Given that integrin α2β1 function is regulated by cytosolic factors and the membrane bilayer, an in vivo system is needed to evaluate the rupture force of the integrin α2β1−DGEA bond in living cells. We therefore conjugated BMMSCs to the end of the cantilever as a cellular model to study the rupture force of the integrin α2β1−DGEA bond on substrates with different elasticity. Independent groups have demonstrated that soft gels whose elasticity mimics that of the brain (500 Pa to ∼1 kPa) induce the neurogenic differentiation of BMMSCs, whereas comparatively rigid matrices that mimic collagenous bone (25−100 kPa) induce osteogenic differentiation. On the basis of these reports,7,14,32 polyacrylamide gels of 500 Pa (neurogenic) and 1 × 105 Pa (osteogenic) were used substrates and functionalized with the DGEA peptide. The rupture forces of integrin the α2β1−DGEA bond on the cell−ECM interface was detected when the single cell-functionalized AFM tip was brought into and out of contact with the DGEA-modified substrate. Optical images of the cell during the force measurement were captured every 10 min. The cell on the cantilever remained an intact sphere shape, and its integrity showed no obvious changes during the experiment (Figure 2a); Trypan blue staining demonstrated that the cell was still alive after the experiment (Figure S2). Typical F−D curves are shown in Figure 2b. The only peak represents the interaction between the DGEA peptide and integrin α2β1 on the cell surface. The specificity of the integrin α2β1−DGEA bond was confirmed, as significantly fewer interactions were detected in the control group (BSA-modified substrate, Figure 2c). The low binding probability of 20.0% ± 3.0% ensured that 89% of unbinding events represented the 209


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Figure 3. Analysis of the rupture forces and loading rates of the integrin α2β1−DGEA bond at varying cantilever retracting speeds. (a) The distribution of the rupture forces of the integrin α2β1−DGEA bond on soft polyacrylamide gel (500 Pa) or stiff polyacrylamide gel (105Pa) at CRSs of 1 μm/s, 10 μm/s and 32 μm/s. Approximately 200 F−D curves displaying specific binding events were acquired to draw each histogram. The average of each group in the classical method (b), the scatter plot (c) and the contour (d) of the distribution of the F−D curves in the loading rate-rupture force coordinate system. The solid lines represent the line fitting. (e) The distribution of the loading rate of the integrin α2β1−DGEA bond on soft polyacrylamide gel (500 Pa, orange) or stiff polyacrylamide (105 Pa, green) at a CRS of 32 μm/s. The loading rates were calculated by r = keff × vc as described previously,20 where keff is the effective spring constant derived from each rupture event on the F−D curves (see Supporting Information Part 8) and vc is the CRS.

reduced the binding frequency to an equivalent level to that of the nonadhesion control group (the cell-BSA group, as mentioned above), and the rare binding events that occurred showed a random distribution in rupture forces (Figure S5a,b). On the other hand, integrin α1 and α11 siRNA transfections, as well as the control siRNA transfection, had no effect on the rupture force distribution (Figure S5a,b) and the binding frequency (Figure S5c,d) of BMMSCs to DGEA-coated polyacrylamide gels. These results demonstrated the key role of integrin α2β1 in the interaction of BMMSCs with the DGEAcoated polyacrylamide gels. Taken together, these results demonstrate that the integrin α2β1−DGEA bond is weaker on a softer substrate than on a stiffer substrate both in vitro and on a living cell surface. In addition, we carried out an SCFS assay using rat osteosarcoma cells, which showed a similar dual-slope property of the rupture force and loading rate of the integrin α2β1−DGEA bond (Figure S6). These results suggested that the loading rate determination of the α2β1−DGEA bond is not cell-type specific.

dissociation of a single bond. Histogram analysis of the 150− 200 observed unbinding events demonstrated a unimodal lognormal distribution of the rupture force of the integrin α2β1− DGEA bond, in which the peak value is 94.9 pN and 130 pN on soft and stiff gels, respectively, at a CRS of 1 μm/s (Figure 2d). Integrin α1β1, α2β1, and α11β1 have all been demonstrated as collagen I receptors. To rule out the possible impact of integrins α1β1 and α11β, we performed siRNA knockdowns of integrin α1, α2, and α11 and repeated the SMFS experiments at a CRS of 1 μm/s. By counting the FAM-siRNA-labeled cells (Figure S4a,c,e), we observed that more than 95% of the cells were transfected with integrin α1, α2, or α11 siRNA, and Western blotting (Figure S4b,d,e) showed that siRNA transfection dramatically reduced integrin α1, α2, or α11 protein levels. Cells labeled with FAM were conjugated to the cantilever under a fluorescence microscope to perform the experiments. As shown in Figure S5c,d, integrin α2 siRNA transfection 210


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ACS Nano High Loading Rate of the Integrin α2β1−DGEA Bond Dramatically Increases Bond Lifetime. The in vitro and in vivo experiments both indicate that integrin α2β1−DGEA bonds are more easily disrupted on soft substrates than on the stiff substrates. Inspired by the Bell model,33,34 we proposed that substrate elasticity affects the rupture force of ligand−receptor bonds by manipulating the loading rate r,14 e.g., fpeak =

kBT kT γ = a(ln r ) + b ln r + B ln γ γ k 0kBT

system over loading rates ranging from 102 to 104 pN/s.20 On the other hand, for the high-slope line where f = 511.8 log10r − 1810.8, we obtained koff = 0.0104 s−1, 1/koff= 96.53 s and xu = 0.42 Å. We also plotted the experimental data from the groups on soft and stiff gels separately; both of the scatter plots from the soft- and stiff-gel groups show a similar dual-slope property (Figure S7). The low-slope and high-slope lines intersect at a loading rate of 5888 ± 103.5 pN/s (Figure 3c). These results suggest that a loading rate higher than 5888 pN/s results in a significantly longer lifetime (96.53 s) for the integrin α2β1−DGEA bond compared with the lower loading rate (2.6594 s). At the CRS of 32 μm/s, 89% of the adhesive events on the stiff substrate fall into the long lifetime region (>5888 pN/s), while only 67% of the adhesive events on the soft substrate fall into this region (Figure 3e). At the CRS of 10 μm/s, 76% and 68% of the adhesive events on stiff and soft substrate fall into the long lifetime region, respectively. This result indicates that stiff substrates induce much longer bond lifetimes, and hence, a much lower possibility of bond disassociation compared with the soft substrate. At the CRS of 1 μm/s, however, none of the adhesive events fall into the long lifetime region, suggesting that when the pulling speed on the integrin α2β1−DGEA bond is low, the bond will not respond to substrate elasticity. By introducing a three springs-piston model (Supporting Information Part 7), we estimated the real pulling speed of actin−myosin on integrin, which is on the same order of magnitude as in living cells, ensuring that the experimental data were collected within the normal physiological range. A Loading-Rate-Sensing Model. In adhesive cells, the loading rate of the integrin−ligand bond is calculated by multiplying the effective spring constant of the system (k) by the actomyosin pulling speed (v):

(1)

and r=

1 dw 1/ks + 1/k t dt

(2)

where f peak represents the most probable rupture force of the integrin α2β1−DGEA bonds, k0 is the dissociation rate constant in the absence of applied force, kB is the Boltzmann constant, T is the absolute temperature, γ is the distance from the energy minimum to the barrier, a and b are constant parameters that can be derived from the experiment, kt and ks are the spring constants of the cantilever−cell-integrin−ligand system and the substrate, and w is the total deformation of the system.14 Models from independent groups35−37 also proposed that substrate elasticity influences the loading rate on integrins, thus affecting bond kinetic properties. To verify whether the loading rate is determinative, we investigated the rupture force of cell-surface integrin α2β1− DGEA complexes on the same substrate under different CRSs, as varying the CRS has been widely used to vary the receptor− ligand bond loading rate in SCFS.20,27 The rupture force values in each F−D curve are shown in Figure 3a, and a unimodal lognormal fit demonstrated that the peak values were 121 pN on soft (500 Pa) and 168 pN on stiff (1 × 105 Pa) substrates at a CRS of 10 μm/s and 169 pN on soft (500 Pa) and 225N on stiff (1 × 105 Pa) substrate at a CRS of 32 μm/s. These results indicate that the rupture force of the integrin α2β1−DGEA bond increases with the CRS, as well as with substrate stiffness. To gain insight into the impact of loading rate on the bond rupture force, we introduced a novel analysis approach here, which is quite different from the previous data analytical method. The more classical data analytic method plots samples under the same CRS as a single spot according to the average of the loading rate and rupture force.16 However, when the loading rate data scatters in a broad region in each sample, this method may direct a significant misleading on the curve fitting. For example, by doing average upon each group of data, the data from 6 groups in the present work converges into a linear line (Figure 3b). However, when calculating the loading rate from each unbinding event containing F−D curve and plotting each curve as a dot in the effective loading rate-rupture force coordinates (Figure 3c), we observe a significant none linear trend of the these data. The resulting scatter plot was then used to construct a contour (see AFM Data Analysis in Supporting Information), as shown in Figure 3d, which demonstrated a dual-slope property and can be fitted by two lines with different slopes (Figure 3d). This analysis enables the calculation of integrin α2β1−DGEA bond-specific parameters such as the unstressed dissociation rate koff, the lifetime 1/koff, and the barrier width xu.33 For the low-slope line fitting where f = 14.1 log10r + 65.3, we obtained koff = 0.3760 s−1, 1/koff = 2.6594 s and xu = 15.21 Å, which are similar to the results reported by Anna Taubenberger et al.20 for the integrin α2β1−collagen I

r=

df df dΔ dΔ = × =k× =k×v dt dΔ dt dt

where Δ is the actomyosin pulling distance. The elasticity of the ECM influences the loading rate of integrin−ligand bonds via its impact on the effective spring constant of the system, thereby inducing significant changes in the bond lifetime. To evaluate the impact of substrate elasticity on the loading rate on integrin in adherent cells, we calculated the loading-rate distribution of integrin-ECM protein bonds using actomyosin pulling speeds collected from previous experimental reports (see Supporting Information Part 9). In total, 51% of adhesive events on stiff substrates fall into the long lifetime region (>5888 pN/s), whereas a negligible fraction of adhesive events on soft substrates fall in this region, indicating a dramatic decline in integrin−ECM protein bond lifetime on the cell membrane on neurogenic substrates. Because the disruption of integrin results in caveolin-mediated endocytosis and thus contributes to neurogenic lineage specification,38,39 the present results show a cellular mechanism through which the mechanical properties of the ECM are sensed. From the aforementioned results, we propose that cells sense mechanical cues from the ECM through the loading rate of the integrin−ligand bond. We thus conclude that reducing the pulling speed on this bond may reduce the rupture force as well as the loading rate of the bond and thereby direct stem cells toward neuronal differentiation on stiff substrates. Blebbistatin is a well-documented inhibitor of nonmuscle myosin II (NM II) ATPase activity. Upon the formation of 211


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Figure 4. Blebbistatin treatment affects activated integrin β1 internalization and BMMSC fate determination on stiff substrates. (a) Typical immunocytochemistry images and statistical analyses of activate integrin β1 expression in BMMSCs cultured on osteogenic stiff substrate (E = 105 Pa) in the presence/absence (control) of 5 μM blebbistatin (scale bar, 5 μm; DIO, cell membrane marker; DAPI, DNA marker). P-value indicates differences between control and blebbistatin-treated groups (mean ± SEM; n = 20). (b and c) BMMSCs were cultured on osteogenic stiff substrate (E = 105 Pa) for 7 days in the presence or absence of 5 μM blebbistatin, followed by staining for Runx2, Collagen I, Nestin and NFL. Data are representative of three experiments (scale bar: 50 μm). The fluorescence intensities of Runx2, Collagen I, Nestin and NFL were quantified using Image Pro Plus 6.0. P-values indicate differences between control and blebbistatin treatment on stiff substrate (mean ± SEM; n = 5).

stable cell adhesions on the matrix, NM II will move along actin fibers in an ATP-dependent manner, pulling the integrin− ligand bond toward the cell center. Thus, NM IIs exhibit low speed and low contraction when their ATPase activity is downregulated. To test our loading-rate-sensing theory, we then asked whether treating stem cells with a low-dose blebbistatin treatment would reduce the loading rate of the integrin−ligand bond. Consistent with our model, we found that a low-dose blebbistatin treatment (5 μM) results in a

relatively low loading rate and rupture force on osteogenic stiff substrates (see Supporting Information Part 10). We then used BMMSCs as a model to investigate whether the low-dose blebbistatin treatment can mimic the effect of low substrate elasticity in manipulating stem cell specification. BMMSCs can sense soft polyacrylamide gels (500 Pa) via internalization of activated integrin β1 and be directed toward a neurogenic lineage on soft gels.14 We assayed the internalization of activated integrin β1 and integrin α2, as well as the lineage 212


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Figure 5. Low-dose blebbistatin treatment affects BMMSC fate determination on stiff substrates. BMMSCs were cultured on stiff substrate (E = 105 Pa) in the presence of 5 μM blebbistatin or on soft substrate (E = 500 Pa) in the absence of blebbistatin for 7 days. Neuronal differentiation markers, Nestin, NFL, and MAP2, were then labeled by fluorescence staining. The fluorescence intensities of Nestin, NFL and MAP2 were quantified using Image Pro Plus 6.0. P-values indicate differences between soft and rigid + blebbistatin (mean ± SEM; n = 5).

osteoblasts while promoting neurogenic lineage specification on osteogenic substrate (Figure 4b,c). Further analysis indicated that low-dose blebbistatin treatment resulted in neurogenic phenotypes on stiff substrates that were comparable to those obtained on soft substrates (Figure 5). By siRNA knocking down, we observed that the down regulation of integrin α2 or β1 rather than integrin α1 or α11 repressed the neurogenic lineage specification of BMMSCs on stiff substrate induced by low dose blebbistatin (Figure S13). On the other hand, upregulating the actomyosin pulling speed via stimulating cell contraction by caliculin A, a well documented chemical leading to hyperactivation and contraction of myosin, significantly inhibited fluorescence intensity of NFL, MAP2 and Nestin (Figure S14). These results suggest that regulating the actomyosin pulling speed produce a similar effect on integrin

specification of MSCs on neurogenic (500 Pa) and osteogenic (105 Pa) gels after the low-dose (5 μM) blebbistatin treatment. The levels of activated integrin β1 in the whole cell were measured via immunocytochemistry, and the results showed that the treatment markedly enhanced internalization on osteogenic stiff substrates (Figure 4a). Integrin α2 has been found colocalized with activated integrin β1 on the cell membrane without 5 μM blebbistatin treatment (Figure S12b), while the 5 μM blebbistatin treatment repressed their membrane distributions and leaded to dramatic cytoplasmic distributions (Figure S12). Immunofluorescence staining for runt-related transcription factor 2 (Runx 2), collagen I, Nestin, Neuro-filament light protein (NFL) and microtubule-associated protein 2 (MAP2) further showed that blebbistatin treatment blocked the stiff substrate-induced differentiation of MSCs into 213


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ACS Nano internalization, as well as on stem cell lineage specification, with regulating the effective spring constant of the substrate.

DISCUSSION The functional role of cytoskeletal proteins in integrin activation has been discussed in several recent studies.42 Our group and others have demonstrated that ECM stiffness regulates cell function through its impact on the contraction force of actomyosin fibers.9−11 In the present study, we demonstrated that NM II regulates the dynamic properties of the integrin α2β1−DGEA bond.42 However, a large number of proteins, including myosin, talin, filamin, and α-actinin, have been described to directly interact within integrin cytoplasmic domains and regulate the activation of integrin43−47 The influence of other cytoskeletal proteins on the rupture force and the loading rate of the integrin α2β1−DGEA bond, as well as the corresponding effects on cell function, need further investigation. CONCLUSION Mechanical cues from the substrate have been demonstrated to play key roles in regulating numerous cell functions and protein functions, including cell proliferation, cell migration, and cell differentiation.40,41 However, the mechanical sensors responsible for stiffness sensing remain largely unknown. Using an SMFS/SCFS-based approach, we found that increasing the loading rate of the integrin α2β1−ligand bond on the cell membrane not only promoted a larger rupture force but also, and more importantly, dramatically enhanced bond lifetime. Variations in the loading rate (r) are controlled by two factors: the effective spring constant of the integrin−ECM system (k), which represents extracellular mechanical cues such as the elasticity and pore size of the substrate, and the actomyosin pulling speed (v), which corresponds to cell contraction. We have previously demonstrated that the disruption of integrin results in caveolin-mediated endocytosis and thus contributes to neurogenic lineage specification.14 We thus propose that cells sense soft substrate via the loading rate created on the integrin−ligand bond. On the basis of this assumption, we inferred that reducing the pulling speed on the bond also promotes the disruption and endocytosis of integrin, and plays an equal role to that of reducing substrate elasticity in inducing neurogenesis differentiation of cells. By manipulating myosin activation, drug treatments such as a low dose of blebbistatin can also manipulate the lineage specification of stem cells, supporting our prediction based on the loading rate-sensing theory. The loading rate-sensing theory provides a unified physical core underlying myosin II-dependent elasticity/ topographic sensing by cells (Figure 6). The relationship between rupture force and loading rate in rat osteosarcoma cells on soft/stiff gels shows a similar dual-slope property, indicating that the loading rate sensing mechanism is cell-type independent. Although substrate mechanical cues are well-known to regulate cell function, whether cells sense the stiffness or the pore size of the substrate remains controversial.48,49 Britta Trappmann and colleagues have challenged the notion that cells sense ECM elasticity by proposing a particular arrangement of collagen beams on the pores on the substrate.7,48,50 According to their model, the deformation of collagen fibers/ beams varies according to the pore size of the substrate, and thus the mechanical cues sensed by cells are affected by the

Figure 6. Mechanical cues from the external environment and physiological cues from the cell jointly modulate cell functions. (a) A schematic of the cell−integrin−ECM protein system. (b) Mechanical cues and biological factors affect cell function through the loading rate on the integrin−ECM protein bond.

pore size of the ECM. In fact, both elasticity and topography, such as pore size and pillar length (Figure 7), affect the effective spring constant of the substrate; thus, both parameters contribute to the loading rate of integrin−ligand bonds. Our results indicate that these mechanical cues regulate cell functions via the same principle. To understand the kinetic properties of a specific bond, a linear fit of the average of the measured rupture force and loading rate is frequently performed. However, without careful insight into the trend of the data, fitting can be subtle and misleading. In the present work, we introduced a contour-based dynamic-force spectroscopy analysis to specify the trend in the relationship between the rupture force and the loading rate. With this method, we demonstrated a dual-slope property of the relationship between the rupture force and the loading rate of the integrin−ligand bond in living cells. This new method reveals the dynamic properties of protein−peptide bonds. Whether the dual-lifetime property of other protein−peptide bonds can be observed using this method remains to be investigated. In addition to substrate mechanical cues, several biological factors have been demonstrated to affect the kinetic properties of integrin−ligand bonds. For example, the types and concentrations of divalent cations can regulate integrin conformation and ligand-binding affinity.51−54 Here, we also 214


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Figure 7. Myosin II-induced contractility subject to integrin−ECM protein bond on substrates with different effective spring constants and different stiffness, pore sizes, or pillar lengths.

found that a high concentration of Mn2+ results in a higher rupture force for the integrin α2β1−DGEA bond (Supporting Information Part 14), which is consistent with the results reported by Agnieszka Ligezowska and colleagues.55 The role of cations in regulating the conformation of integrin and the kinetic properties of integrin−ligand bonds should be addressed in further studies. With the use of a specific binding peptide of integrin α2β1, DGEA, the present study focused on the role of integrin α2β1 in stiffness sensing by cells. However, other integrin family subunits, such as α1β1 and α11β1, also play key roles in cell adhesion and cell functions. Whether the lifetime of α1β1 or α11β1 integrin−ligand bonds on the cell-matrix interface depends on the loading rate, along with their role in stiffness sensing, merits additional study. In conclusion, on the basis of SMFS/SCFS and a contourbased analysis of the loading rate and rupture force of the integrin α2β1−DGEA bond, the present study proposed a loading-rate-sensing theory that provides a unified interpretation of the myosin II-dependent sensing of mechanical cues by cells. These results deepen our understanding of the fundamental mechanisms of mechanical sensing by cells and offer an approach to manipulate stem cell specification by adjusting myosin activation.

kinetic property of integrin−DGEA bond, the specificity of integrin α2β1in single cell force spectroscopy, the loading rat sensing model is cell type independent, “big data” analysis of the loading rate and rupture force, rupture force and loading rate analysis on substrate of different stiffness, estimation of pulling speed of actin bundle on the integrin, the effective spring constant estimation on F−D curves, the loading rate distribution of integrin−ECM protein bond in adherent cells, blebbistatin treatment influences the rupture force and loading rate on integrinα2β1−DGEA bond, the distribution of integrin α2 and its colocalization with activated integrin β1, the neurogenic lineage specification of BMMSCs on stiff substrate induced by low dose blebbistatin depended on the integrin α2 or β1, caliculin A repressed the neurogenic lineage specification of BMMSCs on soft substrate, and the addition of Mn2+ increases the rupture force of integrinα2β1−DGEA bond (PDF)

AUTHOR INFORMATION Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected].

METHODS

Author Contributions

AFM tips and substrates were functionalized according to previously established protocols.20,25,27,5620,25,27,55 Cultured primary BMMSCs were derived from Sprague−Dawley rats. Detailed information on methods is described in Supporting Information, Part 15 Materials and Methods. These include substrate preparation and elasticity identification, AFM tip functionalization, SMFS/SCFS, cell culture, immunocytochemical staining, Western blotting and AFM data analysis.

#

L.J. and Z.S. contributed equally to this work.

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS We thank Quanmei Sun, Jiantao Feng, Jing Liu, Xiaoli Shi, Hainan Gao (National Center for Nanoscience and Technology) for excellent technical assistance and Chunyang Xiong (Peking University) for discussion. This work was financially supported by the National Natural Science Foundation of China (Nos. 31170885, 31370939 and 31400799) and Tsinghua University (2011Z02175).

ASSOCIATED CONTENT S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.5b03157. Cells adhere and spread on DGEA-functionalized substrate, the effect of integrin α2β1 activation on the 215


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Cells Sensing Mechanical Cues: Stiffness Influences the Lifetime of

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