Cells receive many signals from their environment, for example, when they are compressed or pulled about by neighboring cells. Information about these ‘mechanical stimuli’ can be transmitted within the cell to trigger changes in gene expression and cell behavior.

When a cell receives a mechanical stimulus, it can activate the release of calcium ions from storage compartments within the cell, including from a compartment called the endoplasmic reticulum. Calcium ions can also enter the cell from outside via channels located in the membrane that surrounds the cell (the plasma membrane).

Kim et al. investigated how mechanical forces are transmitted in a type of human cell called mesenchymal stem cells using optical tweezers to apply a gentle force to the outside of a cell. These tweezers use a laser to attract tiny objects, in this case a bead attached to proteins in the cell's outer membrane. The cell's response to this mechanical stimulation was measured using a sensor protein that fluoresces a different color when it binds to calcium ions.

With this set-up, Kim et al. found that mesenchymal stem cells are able to transmit mechanical forces to different depths within the cell. The forces can travel deep to trigger the release of calcium ions from the endoplasmic reticulum. This process involves a network of protein fibers that criss-cross to support the structure of a cell—called the cytoskeleton—and also requires proteins that are associated with the cytoskeleton to contract. However, calcium ion entry through the plasma membrane due to a mechanical force does not require these contractile proteins—only the cytoskeleton is involved.

These results demonstrate that the transmission of mechanical signals to different depths within mesenchymal stem cells involves different components. Future work should shed light on how these mechanical signals control gene expression and the development of mesenchymal stem cells.

Mechanical factors are known to play crucial roles in both development and tissue regeneration from stem cells. However, it remains unclear how these factors, such as mechanical forces, are converted into biochemical signals in stem cells to regulate regeneration processes. Calcium ion (Ca²⁺) as one of the most important biochemical signals, is involved in many cellular processes, including muscle contraction, differentiation, proliferation, gene expression, and apoptosis (Berridge et al., 2000; West et al., 2001; McKinsey et al., 2002; Landsberg and Yuan, 2004; Clapham, 2007; Maeda et al., 2007; Rong and Distelhorst, 2008). Various mechanical stimulations can affect cytosolic Ca²⁺ signals as well as Ca²⁺ dynamics in organelles or subcellular compartments, such as mitochondria and focal adhesion sites (Balasubramanian et al., 2007; Belmonte and Morad, 2008; Hayakawa et al., 2008; Horner and Wolfner, 2008). To apply mechanical force precisely, we utilize optical laser tweezers to generate force in a bead coupled to the cell surface through the ligation of adhesion receptors and transmit it into the cell at subcellular locations to trigger signal transduction (Berns, 2007; Botvinick and Wang, 2007).

Most cells mobilize their Ca²⁺ signals via the Ca²⁺ entry across the plasma membrane and/or the Ca²⁺ release from intracellular stores such as endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) (Wehrens et al., 2005; Clapham, 2007). Biochemical signals, such as inositol 1,4,5-trisphosphate (IP₃), are known to regulate Ca²⁺ release from ER. But direct regulation of ER Ca²⁺ signals by mechanical force is unknown. The human mesenchymal stem cells (HMSCs) displaying Ca²⁺ oscillations provide a model system to study that (Kawano et al., 2002; Sun et al., 2007). As Ca²⁺ influx at the plasma membrane and release from ER are the only two sources for Ca²⁺ oscillations in HMSCs (Kawano et al., 2002; Kim et al., 2009), we dissected the effect of mechanical force on each process by monitoring the calcium signals at subcellular locations. To visualize Ca²⁺ signal with high spatiotemporal resolutions, we employed a fluorescence resonance energy transfer (FRET)-based Ca²⁺ biosensor and its variants anchored at subcellular organelles (Miyawaki et al., 1997; Ouyang et al., 2008). Here, we combined optical laser tweezers to deliver local mechanical force and FRET biosensors to investigate how mechanical force regulates Ca²⁺ signals at different subcellular locations in HMSCs.

We first investigated how force regulates Ca²⁺ release from ER with the FRET-based Ca²⁺ biosensor (Figure 1A) to dissect the effect of mechanical force on each of the two Ca²⁺ mobilization processes (Figure 1B). The experiments were done in the absence of extracellular Ca²⁺ so that there was no Ca²⁺ influx through plasma membrane and the oscillations were mainly from ER release (Figure 1B). When fibronectin (Fn)-coated beads were seeded onto the HMSCs and 300 pN of mechanical force was applied by optical laser tweezers as described previously (Wang et al., 2005), Ca²⁺ oscillations were induced in HMSCs (Figure 1C-F, Video 1) but not in bovine aortic endothelial cells (BAECs) (Figure 1—figure supplement 1). In contrast, laser-tweezer-traction on bovine serum albumin (BSA)-coated beads did not cause any oscillations (Figure 1C–F). These results indicate that without extracellular Ca²⁺, mechanical force can induce Ca²⁺ oscillations by triggering Ca²⁺ release from ER. Further experiments showed that depletion of ER Ca²⁺ by Thapsigargin (TG) or inhibition of Ca²⁺ release from the ER by 2-Amino-ethoxydiphenylborate (2-APB) entirely abolished the force-induced oscillations (Figure 1—figure supplement 2A,B). These results confirmed that the ER Ca²⁺ store is the main source for the force-induced oscillations in HMSCs without extracellular Ca²⁺.

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Two possible mechanisms can regulate Ca²⁺ release from ER upon mechanical laser-tweezer-traction. External force can either 1) transmit deep inside the cell and mechanically alter the channels on ER for Ca²⁺ release (Himmel et al., 1993; Missiaen et al., 1996; Rath et al., 2010); or 2) trigger biochemical signaling cascades to produce IP₃ that diffuses inside to activate IP₃-sensitive Ca²⁺ channels (Lehtonen and Kinnunen, 1995; Matsumoto et al., 1995). To distinguish them, we directly monitored the IP₃ level using a FRET-based biosensor, LIBRAvIIs. Mechanical force did not induce any change of IP₃ while ATP increased IP₃ production in HMSCs (Figure 1—figure supplement 2C), suggesting that the second mechanism is unlikely. Therefore, laser-tweezer-traction should transmit deep inside the cells to mechanically release Ca²⁺ from ER.

Cytoskeleton is known to transmit mechanical forces and conduct mechanotransduction (Hamill and Martinac, 2001; Orr et al., 2006; Schwartz and DeSimone, 2008), so we investigated the role of cytoskeleton and its associated proteins in the force-induced ER Ca²⁺ release. The disruption of cytoskeletal actin filaments by cytochalasin D (Cyto D) or microtubules by nocodazole (Noc) completely eliminated the force-induced Ca²⁺ oscillations (Figure 2A–B). In addition, the inhibition of actomyosin contractility by ML-7 or blebbistatin had the same effect (Figure 2C–D). Thus, the deep penetration and transmission of force inside the cell and the induction of Ca²⁺ release from ER depend on both cytoskeletal support and actomyosin contractility. This matches with the previous reports that long-distance force propagation to the deep cytoplasm depends on cytoskeleton tension (Hu et al., 2003) as well as the pivotal role of myosin light chain kinase (MLCK) and myosin II in regulating force development (Olsson et al., 2004; Herring et al., 2006; Fajmut and Brumen, 2008; He et al., 2008). There are two possible signals in ER that may trigger the Ca²⁺ release in response to mechanical force. 1) IP₃R channels on the ER membrane are mechanosensitive and can be directly opened by transmitted mechanical force. Several lines of evidence supported this hypothesis. First, IP₃R is coupled to cytoskeleton and associated proteins allowing mechanical coupling. A direct binding between IP₃R and myosin II was discovered in C. elegans (Walker et al., 2002). In addition, IP₃R has linkage to actin mediated by an adaptor 4.1N protein (Fukatsu et al., 2004; Turvey et al., 2005). IP₃R also binds to ankyrins, which are adaptor proteins coupled to the spectrin-based cytoskeleton (Bourguignon et al., 1993; Joseph and Samanta, 1993). Second, IP₃R channel has an α-helix bundle at the pore forming region, similar to voltage-gated potassium and calcium channels (Schug et al., 2008), which are generally found to be mechanosensitive (Morris, 2011). The mechanism for their mechanosensitivity is possibly that the α-helix tilt angle tends to change when the membrane thins upon mechanical tension, in order to do proper hydrophobic matching with the interfacial region of the membrane, which leads to channel opening (Cheng et al., 2004; Kim and Im, 2010). Therefore, it is likely that the IP₃R channel is also mechanosensitive. 2) Other mechanosensitive channels on ER, for example, transient receptor potential (TRP) family, may also contribute to this force-induced Ca²⁺ release. A number of TRP channels have been found to express at ER membranes, such as TRPC1 (Berbey et al., 2009), TRPV1 (Gallego-Sandin et al., 2009), TRPM8 (Bidaux et al., 2007), and TRPP2 (Koulen et al., 2002). As some TRP channels have been shown to be mechanosensitive and have linkage to cytoskeleton (Barritt and Rychkov, 2005), it is likely that TRP channels located at ER may mediate, at least in part, the force-induced ER calcium release. Notably, these two possibilities are not mutually exclusive as more than one type of channels can be responsible for the force-induced ER calcium release.

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Surprisingly, blocking stretch-activated or store-operated channels at the plasma membrane by Gd³⁺, La³⁺, or streptomycin, but not L-type voltage-operated Ca²⁺ channels by Nifedipine, abolished the mechanical force-induced Ca²⁺ release from ER (Figure 2—figure supplement 1). These results suggest a possible coupling between force-induced Ca²⁺ release at ER and stretch-activated and store-operated channels at the plasma membrane. As TRPM7 is one of the major Ca²⁺ permeable mechanosensitive channels (Wei et al., 2009), we knocked down TRPM7 with targeting small interfering RNA (siRNA) to examine its role in the force-induced Ca²⁺ oscillations. The decreased expression of TRPM7 and the lower percentile of HMSCs with Ca²⁺ oscillations confirmed the effect of siRNA (Figure 2—figure supplement 2A,B) TRPM7 siRNA further abrogated the force-induced oscillations (Figure 2E–F). It is intriguing that the inhibition of TRPM7 at the plasma membrane can block the force transmission into ER in regulating calcium signals. Several possibilities may contribute to the observed results. 1) TRPM7 is functionally coupled to integrin, actomyocin contractility and cytoskeleton. As such, it may mediate and facilitate the force transmission to ER. Indeed, it has been shown that TRPM7 kinase can phosphorylate myosin II heavy chain (Clark et al., 2006) and regulate actomyocin contractility. 2) TRPM7 activity may have some downstream effect on IP₃R in ER. For example, TRPM7 can control the protease calpain (Su et al., 2006), which can regulate IP₃R degradation in ER (Diaz and Bourguignon, 2000). 3) TRPM7 may also be directly coupled to IP₃R in the ER through adaptor proteins. Indeed, another TRP channel, TRPC1 has been shown to directly couple to IP₃R in the ER through an adaptor protein Homer (Yuan et al., 2003). It becomes clear that membrane channels are not isolated entities floating in the plasma membrane. Instead, they are intimately coupled to integrins, cytoskeleton, actomyocin contractility, and ER membrane channels (Cantiello et al., 2007; Matthews et al., 2007; Deng et al., 2009). Therefore, these structural and physical couplings enable membrane channels to participate in direct force transmission to ER.

We further visualized the Ca²⁺ release from ER directly, by generating and employing an improved ER-targeting Ca²⁺ biosensor (D3ER) (Palmer et al., 2006; Ouyang et al., 2008). Mechanical force could directly induce the Ca²⁺ release from ER without extracellular Ca²⁺, as evidenced by the decrease in ER Ca²⁺ concentration (Figure 3A-B, Video 2). The measurements of direct Ca²⁺ release from ER in the presence of inhibitors or siRNA of TRMP7 were also consistent with our observations when cytosolic Ca²⁺ biosensor was used (Figure 3C).

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To gain more insights, we then examined the effect of force on Ca²⁺ influx at the plasma membrane. ER Ca²⁺ release was blocked by an IP₃Rs inhibitor, 2-APB, in the presence of extracellular Ca²⁺ so that the only source of intracellular Ca²⁺ change is the calcium influx from extracellular space (Bootman et al., 2002). Nifedipine did not have any effect on the force-induced Ca²⁺ influx, while Gd³⁺, La³⁺, streptomycin, and TRPM7 siRNA significantly inhibited this Ca²⁺ influx (Figure 4A–B), confirming the involvement of mechanosensitive channels, especially TRPM7, which was shown to be directly activated by a membrane stretch or shear stress in various cell types (Numata et al., 2007a, 2007b; Wei et al., 2009). The disruption of actin filaments or microtubules also inhibited the force-induced Ca²⁺ influx (Figure 4C–D), suggesting that cytoskeletal integrity is essential for the membrane channels to respond to force, consistent with previous reports (Hayakawa et al., 2008). Interestingly, ML-7 or blebbistatin did not affect this force-induced Ca²⁺ influx, suggesting that active actomyosin contractility may not be involved in the mechano-regulation of membrane channels (Figure 4C–D), different from their indispensable roles in the force-induced ER Ca²⁺ release (Figure 4E).

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Since mechanical stimulation such as stretch affects integrin adhesion and integrin-associated signaling, including Src and focal adhesion kinase (FAK) (Wang et al., 2005; Orr et al., 2006), we investigated whether Src and FAK mediate the force-induced Ca²⁺ signals. Our results indicated that neither PP1, an inhibitor of Src family kinases, nor PF228, an inhibitor of FAK, had a significant effect on the force-induced Ca²⁺ influx or ER release (Figure 4—figure supplement 1). Inhibition of phosphoinositide 3-kinases (PI3Ks) with LY294002 did not have any effect either (Figure 4—figure supplement 1). Although integrin-mediated Src phosphorylation and activation can regulate L-type Ca²⁺ channel functions (Gui et al., 2010), the force-induced Ca²⁺ oscillations were clearly independent of L-type Ca²⁺ channels in HMSCs (Figure 2—figure supplement 1). Mechanical stretch can induce the phosphorylation of FAK and nitric oxide formation in cardiomyocytes (Pinsky et al., 1997; Khan et al., 2003), which can modulate ryanodine receptor 2 (RYR2) and Ca²⁺ release at the SR (Vila Petroff and Mattiazzi, 2001). However, there is minimal expression of RYRs in HMSCs (Kawano et al., 2002), which may explain the lack of FAK involvement in the mechano-regulation of Ca²⁺ signals. Together, the biochemical activities above have no detected involvement in the force-induced Ca²⁺ signals.

Further analysis of the duration between force application and the first Ca²⁺ signal indicated that in the absence of extracellular Ca²⁺, the average delay time of ER calcium release monitored by both cytosolic Ca²⁺ biosensor and ER Ca²⁺ biosensor are around 100 s (Figure 4—figure Supplement 1C). Meanwhile, the average delay time of the calcium influx through plasma membrane in the presence of 2-APB and extracellular calcium is much shorter, around 20 s (Figure 4—figure supplement 1C), indicating faster response to mechanical force stimulation. Although the delay time of ER calcium release is longer than that of calcium influx across the plasma membrane, it does not contradict mechanical signal-mediated mechanism. Several reasons may contribute to longer time delay of ER calcium release. 1) The mechanical coupling machinery may need time for reinforcement to allow sufficient force transfer between the plasma membrane and the ER membrane. It is likely that mechanical force transmits through integrins and cytoskeleton (Wang et al., 1993; Perozo and Rees, 2003) as only Fn-coated, but not BSA-coated beads can induce ER calcium release (Figure 1C–F). However, the discrete network linkage of the existing cytoskeleton at the time of force application may not be sufficient for force focusing and transmission to ER, particularly because the apical integrins bound to the Fn-coated beads had not previously experienced force and therefore may have limited connection with cytoskeleton (Matthews et al., 2006). As force can unfold proteins, change their conformations to expose cryptic binding sites to allow the assembly of new/stronger network linkages (del Rio et al., 2009; Johnson et al., 2007), mechanical force can modulate the structural coupling of molecules and mechanical properties of the cells through cytoskeletal remodeling (Kaazempur Mofrad et al., 2005; Matthews et al., 2006; Rosenblatt et al., 2007). These modulation processes of mechanical coupling and reinforcement may need time to reach the threshold for sufficient mechanotransduction of ER calcium release; 2) the channels on ER membrane could have different kinetics and mechanosensitivity (Moe et al., 1998), therefore they may need larger focused and reinforced force to be developed at the site of ER to reach the threshold for physical opening. All these factors may contribute to the longer delay time for force-induced ER calcium release. Again our model does not rule out biochemical signal-mediated mechanisms. As a matter of fact, biochemical signal-mediated mechanisms, such as protein–protein interaction and cytoskeletal remodeling under mechanical tension are important to mediate mechanical signals as discussed above.

As such, our results have provided molecular insights on how mechanical force triggers intracellular Ca²⁺ oscillations through two mechanisms in HMSCs: Ca²⁺ influx at the plasma membrane and Ca²⁺ release from ER (Figure 1B). Our results showed that the deep penetration and transmission of mechanical force to regulate ER functions is dependent on not only the passive cytoskeletal support of actin filaments and microtubules, but also the active actomyosin contractility controlled by MLCK and myosin II. In contrast, the passive cytoskeletal support, but not active actomyosin contractility, is needed for the mechanotransduction at the plasma membrane levels, including the mechanoactivation of channels and Ca²⁺ influx across the plasma membrane (Figure 4E). These results hence provide direct evidence that the mechanotransduction at different depths of cell body is mediated by differential sets of mechanosensing elements.

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Author details

1. Tae-Jin Kim

   1. Neuroscience Program, University of Illinois, Urbana-Champaign, Urbana, United States
   2. Beckman Institute for Advanced Science and Technology, University of Illinois, Urbana-Champaign, Urbana, United States

   Contribution

   T-JK, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

2. Chirlmin Joo

   1. Department of Physics, University of Illinois, Urbana-Champaign, Urbana, United States
   2. Kavli Institute of NanoScience and Department of BioNanoScience, Delft University of Technology, Delft, Netherlands

   Contribution

   CJ, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

3. Jihye Seong

   1. Neuroscience Program, University of Illinois, Urbana-Champaign, Urbana, United States
   2. Center for Neuro-Medicine, Korea Institute of Science and Technology, Seoul, Republic of Korea

   Contribution

   JS, Conception and design, Acquisition of data, Analysis and interpretation of data

   Competing interests

   No competing interests declared.

4. Reza Vafabakhsh

   Department of Physics, University of Illinois, Urbana-Champaign, Urbana, United States

   Contribution

   RV, Conception and design, Acquisition of data

   Competing interests

   No competing interests declared.

5. Elliot L Botvinick

   Department of Biomedical Engineering, Beckman Laser Institute, University of California, Irvine, Irvine, United States

   Contribution

   ELB, Conception and design, Analysis and interpretation of data

   Competing interests

   No competing interests declared.

6. Michael W Berns

   Department of Biomedical Engineering, Beckman Laser Institute, University of California, Irvine, Irvine, United States

   Contribution

   MWB, Conception and design, Analysis and interpretation of data

   Competing interests

   No competing interests declared.

7. Amy E Palmer

   Department of Chemistry and Biochemistry, University of Colorado, Boulder, Boulder, United States

   Contribution

   AEP, Analysis and interpretation of data, Contributed unpublished essential data or reagents

   Competing interests

   No competing interests declared.

8. Ning Wang

   Department of Mechanical Science and Engineering, University of Illinois, Urbana-Champaign, Urbana, United States

   Contribution

   NW, Conception and design, Analysis and interpretation of data

   Competing interests

   No competing interests declared.

9. Taekjip Ha

   1. Department of Physics, University of Illinois, Urbana-Champaign, Urbana, United States
   2. Center for Biophysics and Computational Biology, University of Illinois, Urbana-Champaign, Urbana, United States
   3. Institute of Genomic Biology, University of Illinois, Urbana-Champaign, Urbana, United States
   4. Howard Hughes Medical Institute, University of Illinois, Urbana-Champaign, Urbana, United States

   Contribution

   TH, Conception and design, Analysis and interpretation of data

   Competing interests

   TH: Reviewing editor, eLife.

10. Eric Jakobsson

    1. Beckman Institute for Advanced Science and Technology, University of Illinois, Urbana-Champaign, Urbana, United States
    2. Department of Molecular and Integrative Physiology, University of Illinois, Urbana-Champaign, Urbana, United States

    Contribution

    EJ, Analysis and interpretation of data, Drafting or revising the article

    Competing interests

    No competing interests declared.

11. Jie Sun

    1. Beckman Institute for Advanced Science and Technology, University of Illinois, Urbana-Champaign, Urbana, United States
    2. Department of Molecular and Integrative Physiology, University of Illinois, Urbana-Champaign, Urbana, United States

    Contribution

    JS, Conception and design, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    jiesun2@illinois.edu

    Competing interests

    No competing interests declared.

12. Yingxiao Wang

    1. Neuroscience Program, University of Illinois, Urbana-Champaign, Urbana, United States
    2. Beckman Institute for Advanced Science and Technology, University of Illinois, Urbana-Champaign, Urbana, United States
    3. Center for Biophysics and Computational Biology, University of Illinois, Urbana-Champaign, Urbana, United States
    4. Department of Molecular and Integrative Physiology, University of Illinois, Urbana-Champaign, Urbana, United States
    5. Department of Bioengineering, University of Illinois, Urbana-Champaign, Urbana, United States
    6. Department of Bioengineering, Institute of Engineering in Medicine, University of California, San Diego, San Diego, United States

    Contribution

    YW, Conception and design, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    yiw015@eng.ucsd.edu

    Competing interests

    No competing interests declared.

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