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.

1. 1. Balasubramanian L
   2. Ahmed A
   3. Lo CM
   4. Sham JS
   5. Yip KP

   (2007) Integrin-mediated mechanotransduction in renal vascular smooth muscle cells: activation of calcium sparks

   American Journal of Physiology Regulatory, Integrative and Comparative Physiology 293:R1586–R1594.

   https://doi.org/10.1152/ajpregu.00025.2007

   • Google Scholar
2. 1. Barritt G
   2. Rychkov G

   (2005) TRPs as mechanosensitive channels

   Nature Cell Biology 7:105–107.

   https://doi.org/10.1038/ncb0205-105

   • Google Scholar
3. 1. Belmonte S
   2. Morad M

   (2008) Shear fluid-induced Ca2+ release and the role of mitochondria in rat cardiac myocytes

   Annals of the New York Academy of Sciences 1123:58–63.

   https://doi.org/10.1196/annals.1420.007

   • Google Scholar
4. 1. Berbey C
   2. Weiss N
   3. Legrand C
   4. Allard B

   (2009) Transient receptor potential canonical type 1 (TRPC1) operates as a sarcoplasmic reticulum calcium leak channel in skeletal muscle

   Journal of Biological Chemistry 284:36387–36394.

   https://doi.org/10.1074/jbc.M109.073221

   • Google Scholar
5. 1. Berns MW

   (2007)

   Optical tweezers: tethers, wavelengths, and heat

   Methods in Cell Biology 82:457–466.

   • Google Scholar
6. 1. Berridge MJ
   2. Lipp P
   3. Bootman MD

   (2000) The versatility and universality of calcium signalling

   Nature Reviews Molecular Cell Biology 1:11–21.

   https://doi.org/10.1038/35036035

   • Google Scholar
7. 1. Bidaux G
   2. Flourakis M
   3. Thebault S
   4. Zholos A
   5. Beck B
   6. Gkika D
   7. Roudbaraki M
   8. Bonnal JL
   9. Mauroy B
   10. Shuba Y
   11. Skryma R
   12. Prevarskaya N

   (2007) Prostate cell differentiation status determines transient receptor potential melastatin member 8 channel subcellular localization and function

   The Journal of Clinical Investigation 117:1647–1657.

   https://doi.org/10.1172/JCI30168

   • Google Scholar
8. 1. Bootman MD
   2. Collins TJ
   3. Mackenzie L
   4. Roderick HL
   5. Berridge MJ
   6. Peppiatt CM

   (2002) 2-aminoethoxydiphenyl borate (2-APB) is a reliable blocker of store-operated Ca2+ entry but an inconsistent inhibitor of InsP3-induced Ca2+ release

   FASEB Journal 16:1145–1150.

   https://doi.org/10.1096/fj.02-0037rev

   • Google Scholar
9. 1. Botvinick EL
   2. Wang Y

   (2007) Laser tweezers in the study of mechanobiology in live cells

   Methods in Cell Biology 82:497–523.

   https://doi.org/10.1016/S0091-679X(06)82018-2

   • Google Scholar
10. 1. Bourguignon LY
    2. Jin HT
    3. Iida N
    4. Brandt NR
    5. Zhang SH

    (1993)

    The involvement of ankyrin in the regulation of inositol 1,4,5-Trisphosphate receptor-mediated Internal Ca2+ release from Ca2+ storage-Vesicles in Mouse t-lymphoma cells

    Journal of Biological Chemistry 268:7290–7297.

    • Google Scholar
11. 1. Cantiello HF
    2. Montalbetti N
    3. Li Q
    4. Chen XZ

    (2007) The cytoskeletal connection to ion channels as a potential mechanosensory mechanism: lessons from polycystin-2 (TRPP2)

    Current Topics in Membranes 59:233–296.

    https://doi.org/10.1016/S1063-5823(06)59010-6

    • Google Scholar
12. 1. Cheng Y
    2. Liu A
    3. Guo Y
    4. Qi X

    (2004)

    Computational models for the helix tilt angle

    Journal of Chemical Information and Computer Sciences 44:1952–1960.

    • Google Scholar
13. 1. Clapham DE

    (2007) Calcium signaling

    Cell 131:1047–1058.

    https://doi.org/10.1016/j.cell.2007.11.028

    • Google Scholar
14. 1. Clark K
    2. Langeslag M
    3. van Leeuwen B
    4. Ran L
    5. Ryazanov AG
    6. Figdor CG
    7. Moolenaar WH
    8. Jalink K
    9. van Leeuwen FN

    (2006) TRPM7, a novel regulator of actomyosin contractility and cell adhesion

    The EMBO Journal 25:290–301.

    https://doi.org/10.1038/sj.emboj.7600931

    • Google Scholar
15. 1. del Rio A
    2. Perez-Jimenez R
    3. Liu R
    4. Roca-Cusachs P
    5. Fernandez JM
    6. Sheetz MP

    (2009) Stretching single talin rod molecules activates vinculin binding

    Science 323:638–641.

    https://doi.org/10.1126/science.116291

    • Google Scholar
16. 1. Deng X
    2. Wang Y
    3. Zhou Y
    4. Soboloff J
    5. Gill DL

    (2009) STIM and Orai: dynamic intermembrane coupling to control cellular calcium signals

    The Journal of Biological Chemistry 284:22501–22505.

    https://doi.org/10.1074/jbc.R109.018655

    • Google Scholar
17. 1. Diaz F
    2. Bourguignon LY

    (2000) Selective down-regulation of IP(3)receptor subtypes by caspases and calpain during TNF alpha -induced apoptosis of human T-lymphoma cells

    Cell Calcium 27:315–328.

    https://doi.org/10.1054/ceca.2000.0126

    • Google Scholar
18. 1. Fajmut A
    2. Brumen M

    (2008) MLC-kinase/phosphatase control of Ca2+ signal transduction in airway smooth muscles

    Journal of Theoretical Biology 252:474–481.

    https://doi.org/10.1016/j.jtbi.2007.10.005

    • Google Scholar
19. 1. Fukatsu K
    2. Bannai H
    3. Zhang SB
    4. Nakamura H
    5. Inoue T
    6. Mikoshiba K

    (2004) Lateral diffusion of inositol 1,4,5-trisphosphate receptor type 1 is regulated by actin filaments and 4.1N in neuronal Dendrites

    Journal of Biological Chemistry 279:48976–48982.

    https://doi.org/10.1074/jbc.M408364200

    • Google Scholar
20. 1. Gallego-Sandin S
    2. Rodriguez-Garcia A
    3. Alonso MT
    4. Garcia-Sancho J

    (2009) The endoplasmic reticulum of dorsal root ganglion neurons contains functional TRPV1 channels

    Journal of Biological Chemistry 284:32591–32601.

    https://doi.org/10.1074/jbc.M109.019687

    • Google Scholar
21. 1. Gui P
    2. Chao JT
    3. Wu X
    4. Yang Y
    5. Davis GE
    6. Davis MJ

    (2010) Coordinated regulation of vascular Ca2+ and K+ channels by integrin signaling

    Advances in Experimental Medicine and Biology 674:69–79.

    https://doi.org/10.1007/978-1-4419-6066-5_7

    • Google Scholar
22. 1. Hamill OP
    2. Martinac B

    (2001)

    Molecular basis of mechanotransduction in living cells

    Physiological Reviews 81:685–740.

    • Google Scholar
23. 1. Hayakawa K
    2. Tatsumi H
    3. Sokabe M

    (2008) Actin stress fibers transmit and focus force to activate mechanosensitive channels

    Journal of Cell Science 121:496–503.

    https://doi.org/10.1242/jcs.022053

    • Google Scholar
24. 1. He WQ
    2. Peng YJ
    3. Zhang WC
    4. Lv N
    5. Tang J
    6. Chen C
    7. Zhang CH
    8. Gao S
    9. Chen HQ
    10. Zhi G
    11. Feil R
    12. Kamm KE
    13. Stull JT
    14. Gao X
    15. Zhu MS

    (2008) Myosin light chain kinase is central to smooth muscle contraction and required for gastrointestinal motility in mice

    Gastroenterology 135:610–620.

    https://doi.org/10.1053/j.gastro.2008.05.032

    • Google Scholar
25. 1. Herring BP
    2. El-Mounayri O
    3. Gallagher PJ
    4. Yin F
    5. Zhou J

    (2006) Regulation of myosin light chain kinase and telokin expression in smooth muscle tissues

    American Journal of Physiology Cell Physiology 291:C817–C827.

    https://doi.org/10.1152/ajpcell.00198.2006

    • Google Scholar
26. 1. Himmel HM
    2. Whorton AR
    3. Strauss HC

    (1993) Intracellular calcium, currents, and stimulus-response coupling in endothelial cells

    Hypertension 21:112–127.

    https://doi.org/10.1161/01.HYP.21.1.112

    • Google Scholar
27. 1. Horner VL
    2. Wolfner MF

    (2008) Mechanical stimulation by osmotic and hydrostatic pressure activates Drosophila oocytes in vitro in a calcium-dependent manner

    Developmental Biology 316:100–109.

    https://doi.org/10.1016/j.ydbio.2008.01.014

    • Google Scholar
28. 1. Hu S
    2. Chen J
    3. Fabry B
    4. Numaguchi Y
    5. Gouldstone A
    6. Ingber DE
    7. Fredberg JJ
    8. Butler JP
    9. Wang N

    (2003) Intracellular stress tomography reveals stress focusing and structural anisotropy in cytoskeleton of living cells

    American Journal of Physiology Cell Physiology 285:C1082–C1090.

    https://doi.org/10.1152/ajpcell.00159.2003

    • Google Scholar
29. 1. Johnson CP
    2. Tang HY
    3. Carag C
    4. Speicher DW
    5. Discher DE

    (2007) Forced unfolding of proteins within cells

    Science 317:663–666.

    https://doi.org/10.1126/science.1139857

    • Google Scholar
30. 1. Joseph SK
    2. Samanta S

    (1993)

    Detergent solubility of the inositol trisphosphate receptor in rat-brain membranes. Evidence for association of the receptor with ankyrin

    Journal of Biological Chemistry 268:6477–6486.

    • Google Scholar
31. 1. Kaazempur Mofrad MR
    2. Abdul-Rahim NA
    3. Karcher H
    4. Mack PJ
    5. Yap B
    6. Kamm RD

    (2005) Exploring the molecular basis for mechanosensation, signal transduction, and cytoskeletal remodeling

    Acta Biomaterialia 1:281–293.

    https://doi.org/10.1016/j.actbio.2005.02.008

    • Google Scholar
32. 1. Kawano S
    2. Shoji S
    3. Ichinose S
    4. Yamagata K
    5. Tagami M
    6. Hiraoka M

    (2002) Characterization of Ca(2+) signaling pathways in human mesenchymal stem cells

    Cell Calcium 32:165–174.

    https://doi.org/10.1016/S0143416002001240

    • Google Scholar
33. 1. Khan SA
    2. Skaf MW
    3. Harrison RW
    4. Lee K
    5. Minhas KM
    6. Kumar A
    7. Fradley M
    8. Shoukas AA
    9. Berkowitz DE
    10. Hare JM

    (2003) Nitric oxide regulation of myocardial contractility and calcium cycling: independent impact of neuronal and endothelial nitric oxide synthases

    Circulation Research 92:1322–1329.

    https://doi.org/10.1161/01.RES.0000078171.52542.9E

    • Google Scholar
34. 1. Kim T
    2. Im W

    (2010) Revisiting hydrophobic mismatch with free energy simulation studies of transmembrane helix tilt and rotation

    Biophysical Journal 99:175–183.

    https://doi.org/10.1016/j.bpj.2010.04.015

    • Google Scholar
35. 1. Kim TJ
    2. Seong J
    3. Ouyang M
    4. Sun J
    5. Lu S
    6. Hong JP
    7. Wang N
    8. Wang Y

    (2009) Substrate rigidity regulates Ca2+ oscillation via RhoA pathway in stem cells

    Journal of Cellular Physiology 218:285–293.

    https://doi.org/10.1002/jcp.21598

    • Google Scholar
36. 1. Koulen P
    2. Cai YQ
    3. Geng L
    4. Maeda Y
    5. Nishimura S
    6. Witzgall R
    7. Ehrlich BE
    8. Somlo S

    (2002) Polycystin-2 is an intracellular calcium release channel

    Nature Cell Biology 4:191–197.

    https://doi.org/10.1038/ncb754

    • Google Scholar
37. 1. Landsberg JW
    2. Yuan JX

    (2004)

    Calcium and TRP channels in pulmonary vascular smooth muscle cell proliferation

    News in Physiological Sciences 19:44–50.

    • Google Scholar
38. 1. Lehtonen JY
    2. Kinnunen PK

    (1995) Phospholipase A2 as a mechanosensor

    Biophysical Journal 68:1888–1894.

    https://doi.org/10.1016/S0006-3495(95)80366-8

    • Google Scholar
39. 1. Lu S
    2. Kim TJ
    3. Chen CE
    4. Ouyang M
    5. Seong J
    6. Liao X
    7. Wang Y

    (2011) Computational analysis of the spatiotemporal coordination of polarized PI3K and Rac1 activities in micro-patterned live cells

    PLOS ONE 6:e21293.

    https://doi.org/10.1371/journal.pone.0021293

    • Google Scholar
40. 1. Maeda Y
    2. Nitanai Y
    3. Oda T

    (2007) From the crystal structure of troponin to the mechanism of calcium regulation of muscle contraction

    Advances in Experimental Medicine and Biology 592:37–46.

    https://doi.org/10.1007/978-4-431-38453-3_5

    • Google Scholar
41. 1. Matsumoto H
    2. Baron CB
    3. Coburn RF

    (1995)

    Smooth muscle stretch-activated phospholipase C activity

    The American Journal of Physiology 268:C458–C465.

    • Google Scholar
42. 1. Matthews BD
    2. Overby DR
    3. Mannix R
    4. Ingber DE

    (2006) Cellular adaptation to mechanical stress: role of integrins, Rho, cytoskeletal tension and mechanosensitive ion channels

    Journal of Cell Science 119:508–518.

    https://doi.org/10.1242/jcs.02760

    • Google Scholar
43. 1. Matthews BD
    2. Thodeti CK
    3. Ingber DE

    (2007) Activation of mechanosensitive ion channels by forces transmitted through integrins and the cytoskeleton

    Mechanosensitive Ion Channels, Part A 58:59–85.

    https://doi.org/10.1016/S1063-5823(06)58003-2

    • Google Scholar
44. 1. McKinsey TA
    2. Zhang CL
    3. Olson EN

    (2002) MEF2: a calcium-dependent regulator of cell division, differentiation and death

    Trends in Biochemical Sciences 27:40–47.

    https://doi.org/10.1016/S0968-0004(01)02031-X

    • Google Scholar
45. 1. Missiaen L
    2. De Smedt H
    3. Parys JB
    4. Sienaert I
    5. Vanlingen S
    6. Droogmans G
    7. Nilius B
    8. Casteels R

    (1996) Hypotonically induced calcium release from intracellular calcium stores

    The Journal of Biological Chemistry 271:4601–4604.

    https://doi.org/10.1074/jbc.271.21.12287

    • Google Scholar
46. 1. Miyawaki A
    2. Llopis J
    3. Heim R
    4. McCaffery JM
    5. Adams JA
    6. Ikura M
    7. Tsien RY

    (1997) Fluorescent indicators for Ca2+ based on green fluorescent proteins and calmodulin

    Nature 388:882–887.

    https://doi.org/10.1038/42264

    • Google Scholar
47. 1. Moe PC
    2. Blount P
    3. Kung C

    (1998) Functional and structural conservation in the mechanosensitive channel MscL implicates elements crucial for mechanosensation

    Molecular Microbiology 28:583–592.

    https://doi.org/10.1046/j.1365-2958.1998.00821.x

    • Google Scholar
48. 1. Morris CE

    (2011) Voltage-gated channel mechanosensitivity: fact or friction?

    Frontiers in Physiology 2:25.

    https://doi.org/10.3389/fphys.2011.00025

    • Google Scholar
49. 1. Numata T
    2. Shimizu T
    3. Okada Y

    (2007a) Direct mechano-stress sensitivity of TRPM7 channel

    Cellular Physiology and Biochemistry 19:1–8.

    https://doi.org/10.1159/000099187

    • Google Scholar
50. 1. Numata T
    2. Shimizu T
    3. Okada Y

    (2007b) TRPM7 is a stretch- and swelling-activated cation channel involved in volume regulation in human epithelial cells

    American Journal of Physiology Cell Physiology 292:C460–C467.

    https://doi.org/10.1152/ajpcell.00367.2006

    • Google Scholar
51. 1. Olsson MC
    2. Patel JR
    3. Fitzsimons DP
    4. Walker JW
    5. Moss RL

    (2004) Basal myosin light chain phosphorylation is a determinant of Ca2+ sensitivity of force and activation dependence of the kinetics of myocardial force development

    American Journal of Physiology Heart and Circulatory Physiology 287:H2712–H2718.

    https://doi.org/10.1152/ajpheart.01067.2003

    • Google Scholar
52. 1. Orr AW
    2. Helmke BP
    3. Blackman BR
    4. Schwartz MA

    (2006) Mechanisms of mechanotransduction

    Developmental Cell 10:11–20.

    https://doi.org/10.1016/j.devcel.2005.12.006

    • Google Scholar
53. 1. Ouyang M
    2. Sun J
    3. Chien S
    4. Wang Y

    (2008) Determination of hierarchical relationship of Src and Rac at subcellular locations with FRET biosensors

    Proceedings of the National Academy of Sciences of USA 105:14353–14358.

    https://doi.org/10.1073/pnas.0807537105

    • Google Scholar
54. 1. Palmer AE
    2. Giacomello M
    3. Kortemme T
    4. Hires SA
    5. Lev-Ram V
    6. Baker D
    7. Tsien RY

    (2006) Ca2+ indicators based on computationally redesigned calmodulin-peptide pairs

    Chemistry & Biology 13:521–530.

    https://doi.org/10.1016/j.chembiol.2006.03.007

    • Google Scholar
55. 1. Perozo E
    2. Rees DC

    (2003) Structure and mechanism in prokaryotic mechanosensitive channels

    Current Opinion in Structural Biology 13:432–442.

    https://doi.org/10.1016/S0959-440X(03)00106-4

    • Google Scholar
56. 1. Pinsky DJ
    2. Patton S
    3. Mesaros S
    4. Brovkovych V
    5. Kubaszewski E
    6. Grunfeld S
    7. Malinski T

    (1997) Mechanical transduction of nitric oxide synthesis in the beating heart

    Circulation Research 81:372–379.

    https://doi.org/10.1161/01.RES.81.3.372

    • Google Scholar
57. 1. Rath AL
    2. Bonewald LF
    3. Ling J
    4. Jiang JX
    5. Van Dyke ME
    6. Nicolella DP

    (2010) Correlation of cell strain in single osteocytes with intracellular calcium, but not intracellular nitric oxide, in response to fluid flow

    Journal of Biomechanics 43:1560–1564.

    https://doi.org/10.1016/j.jbiomech.2010.01.030

    • Google Scholar
58. 1. Rong Y
    2. Distelhorst CW

    (2008) Bcl-2 protein family members: versatile regulators of calcium signaling in cell survival and apoptosis

    Annual Review of Physiology 70:73–91.

    https://doi.org/10.1146/annurev.physiol.70.021507.105852

    • Google Scholar
59. 1. Rosenblatt N
    2. Hu S
    3. Suki B
    4. Wang N
    5. Stamenović D

    (2007) Contributions of the active and passive components of the cytoskeletal prestress to stiffening of airway smooth muscle cells

    Annals of Biomedical Engineering 35:224–234.

    https://doi.org/10.1007/s10439-006-9228-z

    • Google Scholar
60. 1. Schug ZT
    2. da Fonseca PC
    3. Bhanumathy CD
    4. Wagner L II
    5. Zhang X
    6. Bailey B
    7. Morris EP
    8. Yule DI
    9. Joseph SK

    (2008) Molecular characterization of the inositol 1,4,5-trisphosphate receptor pore-forming segment

    The Journal of Biological Chemistry 283:2939–2948.

    https://doi.org/10.1074/jbc.M706645200

    • Google Scholar
61. 1. Schwartz MA
    2. DeSimone DW

    (2008) Cell adhesion receptors in mechanotransduction

    Current Opinion in Cell Biology 20:551–556.

    https://doi.org/10.1016/j.ceb.2008.05.005

    • Google Scholar
62. 1. Seong J
    2. Ouyang M
    3. Kim T
    4. Sun J
    5. Wen PC
    6. Lu S
    7. Zhuo Y
    8. Llewellyn NM
    9. Schlaepfer DD
    10. Guan JL
    11. Chien S
    12. Wang Y

    (2011) Detection of focal adhesion kinase activation at membrane microdomains by fluorescence resonance energy transfer

    Nature Communications 2:406.

    https://doi.org/10.1038/ncomms1414

    • Google Scholar
63. 1. Su LT
    2. Agapito MA
    3. Li M
    4. Simonson WT
    5. Huttenlocher A
    6. Habas R
    7. Yue L
    8. Runnels LW

    (2006) TRPM7 regulates cell adhesion by controlling the calcium-dependent protease calpain

    The Journal of Biological Chemistry 281:11260–11270.

    https://doi.org/10.1074/jbc.M512885200

    • Google Scholar
64. 1. Sun S
    2. Liu Y
    3. Lipsky S
    4. Cho M

    (2007) Physical manipulation of calcium oscillations facilitates osteodifferentiation of human mesenchymal stem cells

    FASEB Journal 21:1472–1480.

    https://doi.org/10.1096/fj.06-7153com

    • Google Scholar
65. 1. Tanimura A
    2. Morita T
    3. Nezu A
    4. Shitara A
    5. Hashimoto N
    6. Tojyo Y

    (2009) Use of fluorescence resonance energy transfer-based biosensors for the Quantitative analysis of inositol 1,4,5-Trisphosphate dynamics in calcium oscillations

    The Journal of Biological Chemistry 284:8910–8917.

    https://doi.org/10.1074/jbc.M805865200

    • Google Scholar
66. 1. Turvey MR
    2. Fogarty KE
    3. Thorn P

    (2005) Inositol (14,5)-trisphosphate receptor links to filamentous actin are important for generating local Ca2+ signals in pancreatic acinar cells

    Journal of Cell Science 118:971–980.

    https://doi.org/10.1242/jcs.01693

    • Google Scholar
67. 1. Vila Petroff MG
    2. Mattiazzi AR

    (2001) Angiotensin II and cardiac excitation-contraction coupling: questions and controversies

    Heart, Lung & Circulation 10:90–98.

    https://doi.org/10.1046/j.1444-2892.2001.00083.x

    • Google Scholar
68. 1. Walker DS
    2. Ly S
    3. Lockwood KC
    4. Baylis HA

    (2002) A direct interaction between IP3 receptors and myosin II regulates IP3 signaling in C-elegans

    Current Biology 12:951–956.

    https://doi.org/10.1016/S0960-9822(02)00868-0

    • Google Scholar
69. 1. Wang N
    2. Butler JP
    3. Ingber DE

    (1993) Mechanotransduction across the cell-surface and through the cytoskeleton

    Science 260:1124–1127.

    https://doi.org/10.1126/science.7684161

    • Google Scholar
70. 1. Wang Y
    2. Botvinick EL
    3. Zhao Y
    4. Berns MW
    5. Usami S
    6. Tsien RY
    7. Chien S

    (2005) Visualizing the mechanical activation of Src

    Nature 434:1040–1045.

    https://doi.org/10.1038/nature03469

    • Google Scholar
71. 1. Wang Y
    2. Jin G
    3. Miao H
    4. Li JY
    5. Usami S
    6. Chien S

    (2006) Integrins regulate VE-cadherin and catenins: dependence of this regulation on Src, but not on Ras

    Proceedings of the National Academy of Sciences of USA 103:1774–1779.

    https://doi.org/10.1073/pnas.0510774103

    • Google Scholar
72. 1. Wehrens XH
    2. Lehnart SE
    3. Marks AR

    (2005) Intracellular calcium release and cardiac disease

    Annual Review of Physiology 67:69–98.

    https://doi.org/10.1146/annurev.physiol.67.040403.114521

    • Google Scholar
73. 1. Wei C
    2. Wang X
    3. Chen M
    4. Ouyang K
    5. Song LS
    6. Cheng H

    (2009) Calcium flickers steer cell migration

    Nature 457:901–905.

    https://doi.org/10.1038/nature07577

    • Google Scholar
74. 1. West AE
    2. Chen WG
    3. Dalva MB
    4. Dolmetsch RE
    5. Kornhauser JM
    6. Shaywitz AJ
    7. Takasu MA
    8. Tao X
    9. Greenberg ME

    (2001) Calcium regulation of neuronal gene expression

    Proceedings of the National Academy of Sciences of USA 98:11024–11031.

    https://doi.org/10.1073/pnas.191352298

    • Google Scholar
75. 1. Yuan JP
    2. Kiselyov K
    3. Shin DM
    4. Chen J
    5. Shcheynikov N
    6. Kang SH
    7. Dehoff MH
    8. Schwarz MK
    9. Seeburg PH
    10. Muallem S
    11. Worley PF

    (2003) Homer binds TRPC family channels and is required for gating of TRPC1 by IP3 receptors

    Cell 114:777–789.

    https://doi.org/10.1016/S0092-8674(03)00716-5

    • Google Scholar

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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