BioMed Research International

BioMed Research International / 2015 / Article

Review Article | Open Access

Volume 2015 |Article ID 409245 | 13 pages |

Ca2+ Signaling in Cytoskeletal Reorganization, Cell Migration, and Cancer Metastasis

Academic Editor: Hiroshi Hasegawa
Received17 Dec 2014
Accepted12 Mar 2015
Published22 Apr 2015


Proper control of Ca2+ signaling is mandatory for effective cell migration, which is critical for embryonic development, wound healing, and cancer metastasis. However, how Ca2+ coordinates structural components and signaling molecules for proper cell motility had remained elusive. With the advance of fluorescent live-cell Ca2+ imaging in recent years, we gradually understand how Ca2+ is regulated spatially and temporally in migrating cells, driving polarization, protrusion, retraction, and adhesion at the right place and right time. Here we give an overview about how cells create local Ca2+ pulses near the leading edge, maintain cytosolic Ca2+ gradient from back to front, and restore Ca2+ depletion for persistent cell motility. Differential roles of Ca2+ in regulating various effectors and the interaction of roles of Ca2+ signaling with other pathways during migration are also discussed. Such information might suggest a new direction to control cancer metastasis by manipulating Ca2+ and its associating signaling molecules in a judicious manner.

1. Introduction

Calcium is one of the most important chemical elements for human beings. At the organismic level, calcium together with other materials composes bone to support our bodies [1]. At the tissue level, the compartmentalization of calcium ions (Ca2+) regulates membrane potentials for proper neuronal [2] and cardiac [3] activities. At the cellular level, increases in Ca2+ trigger a wide variety of physiological processes, including proliferation, death, and migration [4]. Aberrant Ca2+ signaling is therefore not surprising to induce a broad spectrum of diseases in metabolism [1], neuron degeneration [5], immunity [6], and malignancy [7]. However, though tremendous efforts have been exerted, we still do not fully understand how this tiny divalent cation controls our lives.

Such a puzzling situation also exists when we consider Ca2+ signaling in cell migration. As an essential cellular process, cell migration is critical for proper physiological activities, such as embryonic development [8], angiogenesis [9], and immune response [10], and pathological conditions, including immunodeficiency [11], wound healing [12], and cancer metastasis [13]. In either situation, coordination between multiple structural (such as F-actin and focal adhesion) and regulatory (such as Rac1 and Cdc42) components is required for cell migration processes (or modules), including polarization, protrusion, retraction, and adhesion [8]. Since Ca2+ signaling is meticulously controlled temporally and spatially in both local and global manners, it serves as a perfect candidate to regulate cell migration modules. However, although the significant contribution of Ca2+ to cell motility has been well recognized [14], it had remained elusive how Ca2+ was linked to the machinery of cell migration. The advances of live-cell fluorescent imaging for Ca2+ and cell migration in recent years gradually unravel the mystery, but there is still a long way to go.

In the present paper, we will give a brief overview about how Ca2+ signaling is polarized and regulated in migrating cells, its local actions on the cytoskeleton, and its global effect on cell migration and cancer metastasis. The strategies employing Ca2+ signaling to control cell migration and cancer metastasis will also be discussed.

2. History: The Journey to Visualize Ca2+ in Live Moving Cells

The attempt to unravel the roles of Ca2+ in cell migration can be traced back to the late 20th century, when fluorescent probes were invented [15] to monitor intracellular Ca2+ in live cells [16]. Using migrating eosinophils loaded with Ca2+ sensor Fura-2, Brundage et al. revealed that the cytosolic Ca2+ level was lower in the front than the back of the migrating cells. Furthermore, the decrease of regional Ca2+ levels could be used as a marker to predict the cell front before the eosinophil moved [17]. Such a Ca2+ gradient in migrating cells was also confirmed by other research groups [18], though its physiological significance had not been totally understood.

In the meantime, the importance of local Ca2+ signals in migrating cells was also noticed. The use of small molecule inhibitors and Ca2+ channel activators suggested that local Ca2+ in the back of migrating cells regulated retraction and adhesion [19]. Similar approaches were also recruited to indirectly demonstrate the Ca2+ influx in the cell front as the polarity determinant of migrating macrophages [14]. Unfortunately, direct visualization of local Ca2+ signals was not available in those reports due to the limited capabilities of imaging and Ca2+ indicators in early days.

The above problems were gradually resolved in recent years with the advance of technology. First, the utilization of high-sensitive camera for live-cell imaging [20] reduced the power requirement for the light source, which eliminated phototoxicity and improved cell health. A camera with high sensitivity also improved the detection of weak fluorescent signals, which is essential to identify Ca2+ pulses of nanomolar scales [21]. In addition to the camera, the emergence of genetic-encoded Ca2+ indicators (GECIs) [22, 23], which are fluorescent proteins engineered to show differential signals based on their Ca2+-binding statuses, revolutionized Ca2+ imaging. Compared to small molecule Ca2+ indicators, GECIs’ high molecular weights make them less diffusible, enabling the capture of transient local signals. Furthermore, signal peptides could be attached to GECIs so the recombinant proteins could be located to different compartments, facilitating Ca2+ measurements in different organelles. Such tools dramatically improved our knowledge regarding the dynamic and compartmentalized characteristics of Ca2+ signaling.

With the above techniques, “Ca2+ flickers” were observed in the front of migrating cells [18], and their roles in cell motility were directly investigated [24]. Moreover, with the integration of multidisciplinary approaches including fluorescent microscopy, systems biology, and bioinformatics, the spatial role of Ca2+, including the Ca2+ gradient in migrating cells, was also gradually clarified [25]. Our present understanding about Ca2+ signaling in migrating cells is briefly summarized as follows.

3. Ca2+ Transporters Regulating Cell Migration

3.1. Generators of Local Ca2+ Pulses: Inositol Triphosphate (IP3) Receptors and Transient Receptor Potential (TRP) Channels (Figure 1)

For a polarized cell to move efficiently, its front has to coordinate activities of protrusion, retraction, and adhesion [8]. The forward movement starts with protrusion, which requires actin polymerization in lamellipodia and filopodia, the foremost structure of a migrating cell [8, 13, 26]. At the end of protrusion, the cell front slightly retracts and adheres [27] to the extracellular matrix. Those actions occur in lamella, the structure located behind lamellipodia. Lamella recruits myosin to contract and dissemble F-actin in a treadmill-like manner and to form nascent focal adhesion complexes in a dynamic manner [28]. After a successful adhesion, another cycle of protrusion begins with actin polymerization from the newly established cell-matrix adhesion complexes. Such protrusion-slight retraction-adhesion cycles are repeated so the cell front would move in a caterpillar-like manner.

For the above actions to proceed and persist, the structural components, actin and myosin, are regulated in a cyclic manner. For actin regulation, activities of small GTPases, Rac, RhoA, and Cdc42 [29], and protein kinase A [30] are oscillatory in the cell front for efficient protrusion. For myosin regulation, small local Ca2+ signals are also pulsatile in the junction of lamellipodia and lamella [24]. Those pulse signals regulate the activities of myosin light chain kinase (MLCK) and myosin II, which are responsible for efficient retraction and adhesion [31, 32]. Importantly, due to the extremely high affinity between Ca2+-calmodulin complexes and MLCK [33], small local Ca2+ pulses in nanomolar scales are sufficient to trigger significant myosin activities.

The critical roles of local Ca2+ pulses in migrating cells raise the question where those Ca2+ signals come from. In a classical signaling model, most intracellular Ca2+ signals originate from endoplasmic reticulum (ER) through inositol triphosphate (IP3) receptors [34, 35], which are activated by IP3 generated via receptor-tyrosine kinase- (RTK-) phospholipase C (PLC) signaling cascades. It is therefore reasonable to assume that local Ca2+ pulses are also generated from internal Ca2+ storage, that is, the ER. In an in vitro experiment, when Ca2+ chelator EGTA was added to the extracellular space, local Ca2+ pulses were not immediately eliminated from the migrating cells [24], supporting the above hypothesis. Moreover, pan-RTK inhibitors that quenched the activities of RTK-PLC-IP3 signaling cascades reduced local Ca2+ pulses efficiently in moving cells [25]. The observation of enriched RTK and PLC activities at the leading edge of migrating cells was also compatible with the accumulation of local Ca2+ pulses in the cell front [25]. Therefore, polarized RTK-PLC-IP3 signaling enhances the ER in the cell front to release local Ca2+ pulses, which are responsible for cyclic moving activities in the cell front.

In addition to RTK, the readers may wonder about the potential roles of G protein-coupled receptors (GPCRs) on local Ca2+ pulses during cell migration. As the major pathway to activate PLC, GPCRs coupled to the subunit [36] trigger the cleavage of phosphatidylinositol (4,5)-bisphosphate (PIP2) by PLC to generate diacylglycerol (DAG) and IP3, which subsequently releases Ca2+ from the ER as Ca2+ pulses and spikes [37]. Indeed, Ca2+ oscillations induced by GPCR pathways have been observed in various cell types [38, 39]. However, the GPCRs coupled to and PLC include serotonergic, adrenergic, muscarinic, glutamatergic, and histamine receptors [37], most of which do not directly affect cell migration. In contrast, growth factors contributing to cell migration, such as fibroblast growth factor (FGF) [40], epidermal growth factor (EGF) [41], and vascular endothelial growth factor (VEGF) [42], activate RTK signaling pathways. Therefore, it is more likely for the RTK rather than GPCR pathway to be responsible for local Ca2+ pulses in migrating cells. Nonetheless, more studies are required to clarify this important question.

The readers may also be curious how nonpulsatile RTK-PLC signaling generates oscillatory local Ca2+ pulses. In fact, IP3-induced Ca2+ oscillation has been reported repeatedly in various physiological circumstances [34, 38, 43, 44]. One possibility is that RTK signaling sensitizes IP3 receptors in the front ER but does not directly open those Ca2+ channels. Alternatively, the cyclic Ca2+ channel opening could be triggered by Ca2+-induced Ca2+ release (CICR), which is the activation of IP3 receptors by small changes of local Ca2+ levels [45, 46]. In the protruding cell front, the change of membrane tension may open stretch-activated transient receptor potential (TRP) channels [47], offering the required CICR. Indeed, TRP channels have been extensively reported as major contributors for cell migration and cancer metastasis [48]. Specifically, TRPM7 has been revealed to enhance cancer cell metastasis [49, 50], by mediating Ca2+ influx [51], altering Ca2+ flickers [18, 52], and regulating cell-matrix adhesion [53]. Therefore, oscillatory small Ca2+ pulses in the migrating cell front are probably the integrated results of polarized RTK signaling interacting with pulsatile membrane stretch and TRP channel opening, to release Ca2+ periodically from the front ER.

3.2. Maintainers of Basal Ca2+ Levels and Gradient: Sarcoplasmic/Endoplasmic Reticulum Ca2+-ATPase (SERCA) and Plasma Membrane Ca2+-ATPase (PMCA) (Figure 2)

The fact that tiny cyclic Ca2+ signals induce significant changes of cell motility implies that the basal cytosolic Ca2+ level, especially that at the front of migrating cells, has to be extremely low, so the cell migration machinery, specifically myosin and focal adhesion complexes, can promptly respond to small Ca2+ changes. To achieve the above goal, the migrating cells meticulously utilize two types of Ca2+-ATPase pumps, sarcoplasmic/endoplasmic reticulum Ca2+-ATPase (SERCA) and plasma membrane Ca2+-ATPase (PMCA).

3.2.1. SERCA Pumps Cytosolic Ca2+ into the ER

SERCA is transmembranously located at the ER, continuously pumping cytosolic Ca2+ into the ER lumen with fast speed and high affinity [4]. Although their activities are slightly regulated by phospholamban and protein kinase A (PKA) [54], these pumps maintain the internal Ca2+ storage with high fidelity. Once SERCA is inactivated, the ER luminal Ca2+ leaks out to the cytoplasm in no time [55]. The resulting high cytosolic Ca2+ will saturate MLCK and induce persistent contraction of myosin [25], rendering front protrusion not possible. Furthermore, SERCA dysfunction dramatically reduces the ER luminal Ca2+, disabling further Ca2+ signaling through IP3 receptors. Hence, SERCA is essential for physiological and pathological cell migration. It is therefore not surprising to see aberrant SERCA expressions in cancer progression, invasion, and metastasis [56, 57].

3.2.2. SERCA Is Not Responsible for the Cytosolic Ca2+ Gradient in Migrating Cells

Since SERCA continuously and efficiently removes Ca2+ out of the cytoplasm into the ER, it is convenient to hypothesize that cytosolic Ca2+ gradient in migrating cells results from differential SERCA activities. If the SERCA activity was higher in the front than in the back, more Ca2+ in the front cytosol would be pumped into the front ER, resulting in the low-in-front, high-in-back Ca2+ gradient in the cytoplasm and a reverse (high-in-front, low-in-back) gradient in the ER. However, blocking SERCA activities with small molecule inhibitors caused a paradoxical increase of Ca2+ gradient, in addition to the global increase of cytosolic Ca2+ [25]. Monitoring intra-ER Ca2+ with the T1ER FRET probe [58] also revealed a low-in-front, high-in-back Ca2+ gradient when the cell moved [25]. Therefore, though a pivotal molecule keeps the cytosol Ca2+ free at the basal status, SERCA does not contribute to the Ca2+ gradient in migrating cells.

3.2.3. Differential PMCA Activities Keep [Ca2+] in the Front Lower Than the Back during Cell Migration

Based on the above data, Ca2+ pumps at the plasma membrane might be better candidates for the Ca2+ gradient during cell migration. Similar to SERCA, PMCA also continuously removes cytosolic Ca2+, by pumping it to the extracellular space [4, 59]. Unlike SERCA, recent evidence revealed that PMCA inhibitors and siRNA reduced Ca2+ gradient and cell motility during cell migration [25]. Direct measurement of Ca2+ efflux through plasma membrane also demonstrated an enhancement of PMCA activity by 30–50% in the front of migrating cells [25]. Hence, differential PMCA activities might account for the Ca2+ gradient during cell migration.

It is still not totally understood how cells adjust local PMCA activities to make them high in the front and low in the back. Several modulators have been demonstrated to regulate PMCA, including calmodulin [60], PKA [61], and calpain [62]. Whether those proteins could be spatially regulated inside the cells remains elusive. In addition, PMCA was enriched in the front plasmalemma of moving cells [25], suggesting that its differential distribution might account for the well-recognized front-low, back-high Ca2+ gradient during cell migration. Still, how PMCA is accumulated in the cell front requires further investigation.

3.3. Maintainers of Ca2+ Homeostasis during Migration: Store-Operated Ca2+ (SOC) Influx (Figure 3)

SOC influx is an essential process to maintain internal Ca2+ storage [63] for IP3 receptor-based Ca2+ signaling, during which the luminal ER Ca2+ is evacuated. After IP3-induced Ca2+ release, although Ca2+ can be recycled back to the ER through SERCA, a significant amount of cytosolic Ca2+ will be pumped out of the cell through PMCA, resulting in the depletion of internal Ca2+ storage. To rescue this, low luminal Ca2+ activates STIM1 [55, 64], which is a membranous protein located at the ER and transported to the cell periphery by microtubules [65, 66]. Active STIM1 will be translocated to the ER-plasma membrane junction [67], opening the Ca2+ influx channel ORAI1 [68, 69]. Ca2+ homeostasis could therefore be maintained during active signaling processes including cell migration.

Since the identification of STIM1 and ORAI1 as the major players of SOC influx, numerous reports have emerged confirming their significant roles in cell migration and cancer metastasis (Tables 1 and 2). Although it is reasonable for those Ca2+-regulatory molecules to affect cell migration, the molecular mechanism is still not totally clear. Recent experimental evidence implied that STIM1 helped the turnover of cell-matrix adhesion complexes [7, 25], so SOC influx may assist cell migration by maintaining local Ca2+ pulses in the front of migrating cells. In a moving cell, local Ca2+ pulses near its leading edge result in the depletion of Ca2+ in its front ER. Such depletion subsequently activates STIM1 at the cell front. Compatible with the above assumption, more STIM1 was translocated to the ER-plasma membrane junction in the cell front compared to its back during cell migration [25]. Moreover, in addition to the ER and plasma membrane, STIM1 is also colocalized with EB1 [65, 66], the cargo protein located at the plus ends of microtubules. Further experiments revealed that STIM1 was actively transported to the front ER assisting cell migration [25]. Therefore, STIM1 together with other Ca2+ channels is meticulously regulated in a spatial manner maintaining cell polarity and motility.

Gene(s)/Protein(s)Cell typeHighlightTarget(s)Reference

ORAI1Esophageal squamous cell carcinoma (ESCC) ORAI1 controls intracellular Ca2+ oscillationsN.A.[105]

ORAI1 and STIM1Clear cell renal cell carcinoma (ccRCC)ORAI1 and STIM1 regulate cell proliferation and migrationN.A.[106]

ORAI1 and STIM2Melanoma cell linesORAI1 and STIM2 control melanoma growth and invasion in opposite mannersN.A.[107]

ORAI1Breast cancer cellscAMP-PKA pathway decreases SK3 channel and SK3-ORAI1 complex activities, reducing Ca2+ entry and cancer cell migrationcAMP, PKA[108]

STIM1Breast cancer cell line MDA-MB-435sTargeting SK3-ORAI1 in lipid rafts may inhibit bone metastasisSK3[109]

STIM1Cervical cancer cell lines (SiHa, HT-3, CaSki, and HeLa)HDAC6 may disrupt STIM1-mediated SOC influx and block malignant cell behaviorHDAC6[110]

ORAI1 and STIM1Glioblastoma multiforme (GBM)STIM1 and ORAI1 affect the invasion of GBM cellsN.A.[111]

ORAI1Human T cell leukemia line, Jurkat cellMonoclonal antibodies against ORAI1 reduce SOC influx, NFAT transcription, and cytokine releaseN.A.[112]

ORAI1Human prostate cancer (PCa) cellBisphenol A pretreatment enhances SOC influx and ORAI1 protein in LNCaP cells; it also induces PCa cells migrationN.A.[113]

STIM1Cervical cancer cellSTIM1 regulates actomyosin reorganization and contractile forces to control cell migrationActomyosin[114]

STIM1Hepatocellular carcinoma and hepatocyte cell linesSTIM1 level predicts prognosis in patients of liver cancerN.A.[115]

STIM1Human epidermoid carcinoma A431 cellsSTIM1 regulates SOC influx, cell proliferation, and tumorigenicity N.A.[116]

STIM1Cervical cancer SiHa and CaSki cell linesSTIM1 regulates cervical cancer growth, migration, and angiogenesisFocal adhesion, Pyk2[7]

ORAI1 and STIM1MDA-MB-231 human breast cancer cellsBlocking STIM1 or ORAI1 using RNA interference or small molecule inhibitors decreased tumor metastasis in animal modelsFocal adhesion[82]

Gene(s)/Protein(s)Cell typeHighlightTarget(s)Reference

STIM1Endothelial progenitor cells (EPCs)STIM1 affects EPCs proliferation and migration after vascular injury by regulating Ca2+ levelsN.A.[117]

ORAI1HEK293Selective activation of NFAT by ORAI1NFAT[118]

STIM1Endothelial leader cellsCells employ an integrated and polarized Ca2+ signalling system for directed cell migrationPLC pathway[25]

ORAI1KeratinocytesORAI1-mediated Ca2+ entry enhances the turnover of focal adhesion through PKCβ, calpain, and focal adhesion kinasePKC pathway[119]

ORAI1 and STIM1Retinal pigment epithelial cells (ARPE-19 cell line)STIM1, ORAI1, ERK 1/2, and Akt determine EGF-mediated cell growthMAPK pathway[120]

STIM1HEK293STIM regulates focal adhesion dynamicsFocal adhesion[121]

ORAI1 and STIM1Airway smooth muscle cell (ASMC)STIM1 or ORAI1 controls PDGF-mediated ASMC proliferation and chemotactic migrationN.A.[122]

ORAI1 and STIM1ASMCSTIM1 and ORAI1 control PDGF-induced cell migration and Ca2+ influxN.A.[123]

STIM1Intestinal epithelial cell (IEC) Polyamines control TRPC1-mediated Ca2+ signaling and cell migration via differential STIM1 and STIM2 levelsTRPC1[124]

ORAI1 and STIM1Vascular smooth muscle cells (VSMC)STIM1- and ORAI1-mediated SOC influx regulates angiotensin II-induced VSMC proliferationN.A.[125]

STIM1EPCsSTIM1 regulates the proliferation and migration of EPCsN.A.[126]

ORAI1 and STIM1VSMCSTIM1 and ORAI1 regulate PDGF-mediated Ca2+ entry and migration in VSMCN.A.[127]

ORAI1 and STIM1VSMCKnockdown of STIM1 and ORAI1, but not STIM2, Orai2, or Orai3, reduces VSMC proliferation and migrationN.A.[128]

4. Ca2+ Effectors for Cell Migration (Figure 4)

As described above, intracellular Ca2+ is regulated locally and globally for effective cytoskeletal remodeling, cell migration, and cancer metastasis. Ca2+ pulses and spikes occur at the right place and right time, activating numerous downstream structural and signaling targets, which have been investigated separately over the past decades. The clarification of Ca2+ signaling in recent years has dramatically improved our understanding about how those components are regulated temporally and spatially in migrating cells. However, such advancement has revealed more questions than answers. More efforts are required to resolve those problems in the future.

4.1. Signaling-Related Targets
4.1.1. Protein Kinase C (PKC)

PKC is a typical downstream target of Ca2+ in receptor tyrosine kinase signaling pathways, during which the growth factor binds to the receptor and activates its tyrosine kinase through dimerization and autophosphorylation [70]. The resulting activation of PLC generates diacylglycerol (DAG) and IP3, which subsequently induces Ca2+ release from the ER. DAG and Ca2+ then bind separately to the C1 and C2 domains of classical PKC (PKCα, β, and γ) [71]. Depending on the substrate, classical PKC regulates a wide variety of physiological processes, including cell migration [72]. The action could be direct via phosphorylation or indirect through transcriptional activation.

The classical PKC family has direct and significant impact on cell migration. PKCα is enriched in the front of migrating cells [14]. It directly phosphorylates Rho GTPases and multiple components of focal adhesion complexes, regulating the remodeling of cell-matrix adhesion (see [73] for a more comprehensive review). PKCβ phosphorylates the heavy chains of myosin II, inhibiting myosin contraction and facilitating the process of directional determination in migrating cells [7477]. How these PKCs respond to spatiotemporal Ca2+ signaling and coordinate for effective moving activities requires further investigation.

Besides classical PKCs, atypical PKCs [70] also regulate the polarity of migrating cells. Unlike classical PKCs, those PKCs do not require DAG or Ca2+ for activation [70]. Together with Rho GTPases [78, 79], these PKCs might be actively involved in the dynamic processes of cell protrusion and adhesion [78, 80]. How these actions synchronize with the Ca2+ dynamics during cell migration also awaits more research in the future.

4.1.2. Rho GTPases

Rho GTPases, including Rac1, RhoA, and Cdc42, have been known as the key components for the regulation of actin dynamics [81]. It is therefore not surprising to see their active involvement in cell migration. Spatially, in a simplified model, these GTPases are enriched at specific structures of a migrating cell, Rac1 in lamellipodia, RhoA around focal adhesion complexes, and Cdc42 near filopodia [8]. Temporally, activities of these GTPases are pulsatile and also synchronized to the cyclic lamellipodial activities in the front of migrating cells [29]. Therefore, Rho GTPases, similar to Ca2+ [24], exert actions at the right place and right time for proper actin remodeling and efficient cell migration.

Although the present data reveals no evidence of direct binding between Ca2+ and Rho GTPases, it is reasonable to expect their mutual interactions considering their perfect coordination during cell migration [24, 29, 30]. Such speculation is supported by the observation that blocking Ca2+ influx at the leading edges of polarized macrophages resulted in the disassembly of actin filaments and lamellipodia activities [14]. The facts that constitutively active Rac1 fully rescued the effects of SOC influx inhibition in migrating breast cancer cells [82] also indicate the regulatory role of Ca2+ on Rho GTPases. Moreover, the transamidation of Rac1 was shown to be dependent on intracellular Ca2+ and calmodulin in rat cortical cells, suggesting the biochemical link between Rho GTPases and Ca2+ signaling [83]. Hopefully more studies will be conducted in the near future to clarify the mechanism of how Ca2+ interacts with Rho GTPases.

4.2. Cytoskeleton-Related Targets
4.2.1. Myosin II

As mentioned above, local Ca2+ pulses at the junction of lamellipodia and lamella activate MLCK [24], which subsequently phosphorylates myosin light chain and triggers myosin contraction. It is worth noticing that the affinity between MLCK and myosin-calmodulin is extremely high, with the dissociation constant () of about 1 nM [33]. Therefore, a slight increase of local Ca2+ concentration is sufficient to induce significant activation of MLCK and subsequent contraction of myosin II. Moreover, the high sensitivity of MLCK to Ca2+ implies that the front cytoplasm has to be free of Ca2+ at the basal status, so MLCK can be inactive at baseline but respond to small rises of Ca2+ promptly. Such design justifies the physiological importance of the front-low, back-high Ca2+ gradient in migrating cells.

In cell migration, the immediate effect of myosin contraction is the retraction of actin bundles, which not only facilitates the disassembly of F-actin at lamella but also allows the protruding front to attach to the extracellular matrix [28, 31]. In addition, myosin contraction also stabilizes nascent focal adhesion complexes in the front of migrating cells [32, 84]. This is probably because these contractions apply traction force on the complexes through actin bundles binding to them. Such force subsequently induces remodeling and stabilization of the components in focal adhesion. Therefore, through MLCK and myosin II, local Ca2+ pulses are tightly linked to the oscillatory dynamics of cell protrusion, retraction, and adhesion.

4.2.2. Actin

Besides myosin, Ca2+ also affects the dynamics of actin, the major component of cytoskeleton [85, 86]. Although Ca2+ does not directly bind to actin, it affects the activities of multiple actin regulators. First of all, Ca2+ activates protein kinase C and calmodulin-dependent kinases, both of which interact with actin affecting its dynamics [8789]. Secondly, as also described above, Ca2+ signaling regulates the Rho GTPases [14], which are mandatory for the formation of actin bundles for lamellipodia, focal adhesion complexes, and filopodia [8], the major components for cell migration. In addition, the F-actin severing protein cofilin [90, 91] also depends on the cytosolic Ca2+ for its proper activity. Moreover, myosin, as one the major actin regulators, is totally dependent on Ca2+ for its proper activity [24]. Therefore, though not a direct regulator, Ca2+ modulates actin dynamics through multiple signaling pathways and structural molecules.

4.3. Adhesion-Related Targets
4.3.1. Calpain

In addition to kinase activities and physical force, Ca2+ also affects cell migration through protein cleavage and degradation. Calpain, as a Ca2+-dependent intracellular protease [92, 93], is involved in the turnover of stable focal adhesion complexes, probably at the rear end of migrating cells. Calpain has been revealed to cleave several components of the focal adhesion complex, including talin [94], paxillin [95], and focal adhesion kinases [96], compatible with previous reports showing that Ca2+ influx at the back of migrating cells facilitated retraction and detachment at their rear ends [97]. Beside focal adhesion, calpain also degrades PMCA [62]. Since there is an inverse correlation between the front-back gradients of Ca2+ and PMCA in migrating cells [25], decreased amount of PMCA in the cell back may result from the higher Ca2+ level and higher calpain activity in the back than in the front. However, such speculation requires more experimental data to be validated.

4.3.2. Pyk2 and Other Molecules

In addition to calpain, several adhesion-related proteins are also regulated by Ca2+, including Pyk2, plectin, and matrix metallopeptidases.

As a cytoplasmic protein tyrosine kinase, Pyk2 is activated by intracellular Ca2+ and protein kinase C [98]. It regulates the activities of focal adhesion kinase and GRB2, affecting focal adhesion complexes [99] and the MAP kinase signaling pathway [98]. In human cervical cancer cells, aberrant SOC influx changes focal adhesion dynamics through Pyk2 dysregulation [7].

Ca2+ also regulates the conveyance of integrin-based signaling into the cytoskeleton, with its interaction with plectin, the bridge between integrin complexes and actin filaments. Recent biochemical and biophysical evidence indicated that the binding of plectin 1a with Ca2+ effectively decreased its interactions with integrin β and with F-actin, decoupling cell-matrix adhesion with cytoskeletal structures [100, 101]. We may speculate that, with proper temporal and spatial Ca2+ regulation, cells could determine how many environmental signals would be conducted into the cells for cytoskeleton modification. More studies are required to clarify the above hypothesis.

Furthermore, matrix metallopeptidases (MMP), as facilitating factors for cancer metastasis, are also regulated by intracellular Ca2+. In prostate cancer, increased expression of TRPV2 elevated cytosolic Ca2+ levels, which enhanced MMP9 expression and cancer cell aggressiveness [102]. Further investigation in melanoma cells revealed that increased intracellular Ca2+ induced the binding of Ca2+-modulating cyclophilin ligand to basigin, stimulating the production of MMP [103]. Therefore, Ca2+ not only modulates the outside-in (integrin to actin) signaling but also regulates the inside-out (Ca2+ to MMP) signaling for cell migration and cancer metastasis.

5. Future: Interactions between Ca2+ and Other Signaling Pathways

Regarding the complicated temporal and spatial regulation of Ca2+ signaling in migrating cells, we would expect extensive interactions between Ca2+ and other signaling modules during cell migration. Indeed, though still preliminary, recent work has revealed potential cross talk between Ca2+ and other pathways controlling cell motility. These findings will shed new light on our pilgrimage toward a panoramic view of cell migration machinery.

5.1. Interactions between SOC Influx and Cell-Matrix Adhesion

In the present model, SOC influx maintains Ca2+ storage in the ER, which releases local Ca2+ pulses to enhance the formation of nascent focal adhesion complexes [25]. Therefore, the inhibition of SOC influx should weaken cell-matrix adhesion. Interestingly, STIM1, the Ca2+ sensor for the activation of the SOC influx, had been reported as an oncogene [82] or a tumor suppressor gene [104] by different groups. Furthermore, although most recent research suggested a positive role of STIM1 on cancer cell motility (Table 1), other reports revealed the opposite results in primary cells (Table 2). Therefore, effects of SOC influx on cell migration might vary under different circumstances.

One possible explanation of the confusing results uses the interaction between Ca2+ and basal cell-matrix adhesion. Primary cells are usually well attached to the matrix, so further enhancing their adhesion capability might trap them in the matrix and deter them from moving forward. In contrast, metastatic cancer cells often have weak cell-matrix adhesion, so strengthening their attachment to the matrix facilitates the completion of cell migration cycles. Indeed, recent evidence suggested that, in an in vitro cell migration assay [25], SOC influx might increase or decrease the motility of the same cell type depending on concentrations of fibronectin for the cells to attach. Though further explorations are required to validate the present data, the combination of SOC influx inhibition and cell-matrix adhesion blockage might be a novel approach to prevent cancer metastasis.

5.2. Coordination between the Oscillations of Ca2+ and Rho GTPases

Previous reports have revealed the oscillatory activities of Rho GTPases in the front of migrating cells, including Rac1, RhoA, and Cdc42 [29, 30]. These molecules regulate actin dynamics and coordinate with the pulsatile lamellipodial activities. Since the oscillation of local Ca2+ pulses synchronize with the retraction phases of lamellipodial cycles [24], there probably exists cross talk between Ca2+ signaling and Rho GTPases. Clarifying how these molecules are regulated to coordinate with each other will dramatically improve our understanding of lamellipodia and help developing better strategies to control physiological and pathological cell migration.

5.3. Link between Ca2+, RTK, and Lipid Signaling

The meticulous spatial control of Ca2+ signaling in migrating cells, together with the enrichment of RTK, phosphatidylinositol (3,4,5)-triphosphate (PIP3), and DAG in the cell front [25], reveals the complicated nature of the migration polarity machinery. How these signaling pathways act together to determine the direction for cells to move remains elusive and requires more research. In addition, understanding how nonpulsatile RTK and lipid signaling exert effects on oscillatory Ca2+ pulses will improve our knowledge about the spatial and temporal regulation of signal transduction inside the cells. Such information will further enhance our capability to develop novel strategies targeting pathological processes and manipulating diseases.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.


  1. R. Masuyama, “Role of local vitamin D signaling and cellular calcium transport system in bone homeostasis,” Journal of Bone and Mineral Metabolism, vol. 32, no. 1, pp. 1–9, 2014. View at: Publisher Site | Google Scholar
  2. I. Calin-Jageman and A. Lee, “Cav1 L-type Ca2+ channel signaling complexes in neurons,” Journal of Neurochemistry, vol. 105, no. 3, pp. 573–583, 2008. View at: Publisher Site | Google Scholar
  3. X. Zhao, D. Yamazaki, S. Kakizawa, Z. Pan, H. Takeshima, and J. Ma, “Molecular architecture of Ca2+ signaling control in muscle and heart cells,” Channels, vol. 5, no. 5, pp. 391–396, 2011. View at: Publisher Site | Google Scholar
  4. D. E. Clapham, “Calcium Signaling,” Cell, vol. 131, no. 6, pp. 1047–1058, 2007. View at: Publisher Site | Google Scholar
  5. F. M. LaFerla, “Calcium dyshomeostasis and intracellular signalling in Alzheimer's disease,” Nature Reviews Neuroscience, vol. 3, no. 11, pp. 862–872, 2002. View at: Publisher Site | Google Scholar
  6. P. J. Shaw and S. Feske, “Physiological and pathophysiological functions of SOCE in the immune system,” Frontiers in Bioscience (Elite Edition), vol. 4, no. 6, pp. 2253–2268, 2012. View at: Google Scholar
  7. Y.-F. Chen, W.-T. Chiu, Y.-T. Chen et al., “Calcium store sensor stromal-interaction molecule 1-dependent signaling plays an important role in cervical cancer growth, migration, and angiogenesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 108, no. 37, pp. 15225–15230, 2011. View at: Publisher Site | Google Scholar
  8. A. J. Ridley, M. A. Schwartz, K. Burridge et al., “Cell migration: integrating signals from front to back,” Science, vol. 302, no. 5651, pp. 1704–1709, 2003. View at: Publisher Site | Google Scholar
  9. L. Lamalice, F. Le Boeuf, and J. Huot, “Endothelial cell migration during angiogenesis,” Circulation Research, vol. 100, no. 6, pp. 782–794, 2007. View at: Publisher Site | Google Scholar
  10. B. A. Imhof and D. Dunon, “Basic mechanism of leukocyte migration,” Hormone and Metabolic Research, vol. 29, no. 12, pp. 614–621, 1997. View at: Publisher Site | Google Scholar
  11. S. Hanna and A. Etzioni, “Leukocyte adhesion deficiencies,” Annals of the New York Academy of Sciences, vol. 1250, no. 1, pp. 50–55, 2012. View at: Publisher Site | Google Scholar
  12. S. Barrientos, O. Stojadinovic, M. S. Golinko, H. Brem, and M. Tomic-Canic, “Growth factors and cytokines in wound healing,” Wound Repair and Regeneration, vol. 16, no. 5, pp. 585–601, 2008. View at: Publisher Site | Google Scholar
  13. M. Vicente-Manzanares and A. R. Horwitz, “Cell migration: an overview,” Methods in Molecular Biology, vol. 769, pp. 1–24, 2011. View at: Publisher Site | Google Scholar
  14. J. H. Evans and J. J. Falke, “Ca2+ influx is an essential component of the positive-feedback loop that maintains leading-edge structure and activity in macrophages,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 41, pp. 16176–16181, 2007. View at: Publisher Site | Google Scholar
  15. G. Grynkiewicz, M. Poenie, and R. Y. Tsien, “A new generation of Ca2+ indicators with greatly improved fluorescence properties,” The Journal of Biological Chemistry, vol. 260, no. 6, pp. 3440–3450, 1985. View at: Google Scholar
  16. R. Y. Tsien, T. J. Rink, and M. Poenie, “Measurement of cytosolic free Ca2+ in individual small cells using fluorescence microscopy with dual excitation wavelengths,” Cell Calcium, vol. 6, no. 1-2, pp. 145–157, 1985. View at: Publisher Site | Google Scholar
  17. R. A. Brundage, K. E. Fogarty, R. A. Tuft, and F. S. Fay, “Calcium gradients underlying polarization and chemotaxis of eosinophils,” Science, vol. 254, no. 5032, pp. 703–706, 1991. View at: Publisher Site | Google Scholar
  18. C. Wei, X. Wang, M. Chen, K. Ouyang, L.-S. Song, and H. Cheng, “Calcium flickers steer cell migration,” Nature, vol. 457, no. 7231, pp. 901–905, 2009. View at: Publisher Site | Google Scholar
  19. J. T. H. Mandeville, R. N. Ghosh, and F. R. Maxfield, “Intracellular calcium levels correlate with speed and persistent forward motion in migrating neutrophils,” Biophysical Journal, vol. 68, no. 4, pp. 1207–1217, 1995. View at: Publisher Site | Google Scholar
  20. J. Elf, G.-W. Li, and X. S. Xie, “Probing transcription factor dynamics at the single-molecule level in a living cell,” Science, vol. 316, no. 5828, pp. 1191–1194, 2007. View at: Publisher Site | Google Scholar
  21. K. L. Rogers, J.-R. Martin, O. Renaud et al., “Electron-multiplying charge-coupled detector-based bioluminescence recording of single-cell Ca2+,” Journal of Biomedical Optics, vol. 13, no. 3, Article ID 031211, 2008. View at: Publisher Site | Google Scholar
  22. M. Mank and O. Griesbeck, “Genetically encoded calcium indicators,” Chemical Reviews, vol. 108, no. 5, pp. 1550–1564, 2008. View at: Publisher Site | Google Scholar
  23. T.-W. Chen, T. J. Wardill, Y. Sun et al., “Ultrasensitive fluorescent proteins for imaging neuronal activity,” Nature, vol. 499, no. 7458, pp. 295–300, 2013. View at: Publisher Site | Google Scholar
  24. F.-C. Tsai and T. Meyer, “Ca2+ pulses control local cycles of lamellipodia retraction and adhesion along the front of migrating cells,” Current Biology, vol. 22, no. 9, pp. 837–842, 2012. View at: Publisher Site | Google Scholar
  25. F.-C. Tsai, A. Seki, H. W. Yang et al., “A polarized Ca2+, diacylglycerol and STIM1 signalling system regulates directed cell migration,” Nature Cell Biology, vol. 16, no. 2, pp. 133–144, 2014. View at: Publisher Site | Google Scholar
  26. J. V. Small, T. Stradal, E. Vignal, and K. Rottner, “The lamellipodium: where motility begins,” Trends in Cell Biology, vol. 12, no. 3, pp. 112–120, 2002. View at: Publisher Site | Google Scholar
  27. D. J. Webb, J. T. Parsons, and A. F. Horwitz, “Adhesion assembly, disassembly and turnover in migrating cells—over and over and over again,” Nature Cell Biology, vol. 4, no. 4, pp. E97–E100, 2002. View at: Publisher Site | Google Scholar
  28. D. T. Burnette, S. Manley, P. Sengupta et al., “A role for actin arcs in the leading-edge advance of migrating cells,” Nature Cell Biology, vol. 13, no. 4, pp. 371–382, 2011. View at: Publisher Site | Google Scholar
  29. M. MacHacek, L. Hodgson, C. Welch et al., “Coordination of Rho GTPase activities during cell protrusion,” Nature, vol. 461, no. 7260, pp. 99–103, 2009. View at: Publisher Site | Google Scholar
  30. E. Tkachenko, M. Sabouri-Ghomi, O. Pertz et al., “Protein kinase A governs a RhoA-RhoGDI protrusion-retraction pacemaker in migrating cells,” Nature Cell Biology, vol. 13, no. 6, pp. 660–667, 2011. View at: Publisher Site | Google Scholar
  31. G. Giannone, B. J. Dubin-Thaler, H.-G. Döbereiner, N. Kieffer, A. R. Bresnick, and M. P. Sheetz, “Periodic lamellipodial contractions correlate with rearward actin waves,” Cell, vol. 116, no. 3, pp. 431–443, 2004. View at: Publisher Site | Google Scholar
  32. G. Giannone, B. J. Dubin-Thaler, O. Rossier et al., “Lamellipodial actin mechanically links myosin activity with adhesion-site formation,” Cell, vol. 128, no. 3, pp. 561–575, 2007. View at: Publisher Site | Google Scholar
  33. R. Kasturi, C. Vasulka, and J. D. Johnson, “Ca2+, caldesmon, and myosin light chain kinase exchange with calmodulin,” The Journal of Biological Chemistry, vol. 268, no. 11, pp. 7958–7964, 1993. View at: Google Scholar
  34. M. J. Berridge, “Inositol trisphosphate and calcium signalling,” Nature, vol. 361, no. 6410, pp. 315–325, 1993. View at: Publisher Site | Google Scholar
  35. C. W. Taylor, Taufiq-Ur-Rahman, and E. Pantazaka, “Targeting and clustering of IP3 receptors: key determinants of spatially organized Ca2+ signals,” Chaos, vol. 19, no. 3, Article ID 037102, 2009. View at: Publisher Site | Google Scholar
  36. K. Qin, C. Dong, G. Wu, and N. A. Lambert, “Inactive-state preassembly of Gq-coupled receptors and Gq heterotrimers,” Nature Chemical Biology, vol. 7, no. 10, pp. 740–747, 2011. View at: Publisher Site | Google Scholar
  37. B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, and P. Walter, Molecular Biology of the Cell, Garland Science, 4th edition, 2002.
  38. R. Jacob, J. E. Merritt, T. J. Hallam, and T. J. Rink, “Repetitive spikes in cytoplasmic calcium evoked by histamine in human endothelial cells,” Nature, vol. 335, no. 6185, pp. 40–45, 1988. View at: Publisher Site | Google Scholar
  39. L. Giri, A. K. Patel, W. K. A. Karunarathne, V. Kalyanaraman, K. V. Venkatesh, and N. Gautam, “A G-protein subunit translocation embedded network motif underlies GPCR regulation of calcium oscillations,” Biophysical Journal, vol. 107, no. 1, pp. 242–254, 2014. View at: Publisher Site | Google Scholar
  40. E. G. Levin, “Cancer therapy through control of cell migration,” Current Cancer Drug Targets, vol. 5, no. 7, pp. 505–518, 2005. View at: Publisher Site | Google Scholar
  41. J. Grahovac and A. Wells, “Matrikine and matricellular regulators of EGF receptor signaling on cancer cell migration and invasion,” Laboratory Investigation, vol. 94, no. 1, pp. 31–40, 2014. View at: Publisher Site | Google Scholar
  42. A. Angelucci and M. Bologna, “Targeting vascular cell migration as a strategy for blocking angiogenesis: the central role of focal adhesion protein tyrosine kinase family,” Current Pharmaceutical Design, vol. 13, no. 21, pp. 2129–2145, 2007. View at: Publisher Site | Google Scholar
  43. O. H. Petersen, “Stimulus-secretion coupling: cytoplasmic calcium signals and the control of ion channels in exocrine acinar cells,” Journal of Physiology, vol. 448, pp. 1–51, 1992. View at: Publisher Site | Google Scholar
  44. P. Thorn, A. M. Lawrie, P. M. Smith, D. V. Gallacher, and O. H. Petersen, “Local and global cytosolic Ca2+ oscillations in exocrine cells evoked by agonists and inositol trisphosphate,” Cell, vol. 74, no. 4, pp. 661–668, 1993. View at: Publisher Site | Google Scholar
  45. E. A. Finch, T. J. Turner, and S. M. Goldin, “Calcium as a coagonist of inositol 1,4,5-trisphosphate-induced calcium release,” Science, vol. 252, no. 5004, pp. 443–446, 1991. View at: Publisher Site | Google Scholar
  46. J. Keizer and L. Levine, “Ryanodine receptor adaptation and Ca2+(−)induced Ca2+ release- dependent Ca2+ oscillations,” Biophysical Journal, vol. 71, no. 6, pp. 3477–3487, 1996. View at: Publisher Site | Google Scholar
  47. A. Di and A. B. Malik, “TRP channels and the control of vascular function,” Current Opinion in Pharmacology, vol. 10, no. 2, pp. 127–132, 2010. View at: Publisher Site | Google Scholar
  48. N. Nielsen, O. Lindemann, and A. Schwab, “TRP channels and STIM/ORAI proteins: sensors and effectors of cancer and stroma cell migration,” British Journal of Pharmacology, vol. 171, no. 24, pp. 5524–5540, 2014. View at: Publisher Site | Google Scholar
  49. H. Gao, X. Chen, X. Du, B. Guan, Y. Liu, and H. Zhang, “EGF enhances the migration of cancer cells by up-regulation of TRPM7,” Cell Calcium, vol. 50, no. 6, pp. 559–568, 2011. View at: Publisher Site | Google Scholar
  50. J. Middelbeek, A. J. Kuipers, L. Henneman et al., “TRPM7 is required for breast tumor cell metastasis,” Cancer Research, vol. 72, no. 16, pp. 4250–4261, 2012. View at: Publisher Site | Google Scholar
  51. J.-P. Chen, Y. Luan, C.-X. You, X.-H. Chen, R.-C. Luo, and R. Li, “TRPM7 regulates the migration of human nasopharyngeal carcinoma cell by mediating Ca2+ influx,” Cell Calcium, vol. 47, no. 5, pp. 425–432, 2010. View at: Publisher Site | Google Scholar
  52. B. Roy, T. Das, D. Mishra, T. K. Maiti, and S. Chakraborty, “Oscillatory shear stress induced calcium flickers in osteoblast cells,” Integrative Biology, vol. 6, no. 3, pp. 289–299, 2014. View at: Publisher Site | Google Scholar
  53. L.-T. Su, M. A. Agapito, M. Li et al., “TRPM7 regulates cell adhesion by controlling the calcium-dependent protease calpain,” The Journal of Biological Chemistry, vol. 281, no. 16, pp. 11260–11270, 2006. View at: Publisher Site | Google Scholar
  54. D. H. MacLennan and E. G. Kranias, “Phospholamban: a crucial regulator of cardiac contractility,” Nature Reviews Molecular Cell Biology, vol. 4, no. 7, pp. 566–577, 2003. View at: Publisher Site | Google Scholar
  55. J. Liou, M. L. Kim, D. H. Won et al., “STIM is a Ca2+ sensor essential for Ca2+-store- depletion-triggered Ca2+ influx,” Current Biology, vol. 15, no. 13, pp. 1235–1241, 2005. View at: Publisher Site | Google Scholar
  56. W.-F. Gou, Z.-F. Niu, S. Zhao, Y. Takano, and H.-C. Zheng, “Aberrant SERCA3 expression during the colorectal adenoma-adenocarcinoma sequence,” Oncology Reports, vol. 31, no. 1, pp. 232–240, 2014. View at: Publisher Site | Google Scholar
  57. F.-Y. Chung, S.-R. Lin, C.-Y. Lu et al., “Sarco/endoplasmic reticulum calcium-ATPase 2 expression as a tumor marker in colorectal cancer,” The American Journal of Surgical Pathology, vol. 30, no. 8, pp. 969–974, 2006. View at: Publisher Site | Google Scholar
  58. E. Abell, R. Ahrends, S. Bandara, B. O. Park, and M. N. Teruel, “Parallel adaptive feedback enhances reliability of the Ca2+ signaling system,” Proceedings of the National Academy of Sciences of the United States of America, vol. 108, no. 35, pp. 14485–14490, 2011. View at: Publisher Site | Google Scholar
  59. E. E. Strehler and M. Treiman, “Calcium pumps of plasma membrane and cell interior,” Current Molecular Medicine, vol. 4, no. 3, pp. 323–335, 2004. View at: Publisher Site | Google Scholar
  60. P. James, M. Maeda, R. Fischer et al., “Identification and primary structure of a calmodulin binding domain of the Ca2+ pump of human erythrocytes,” The Journal of Biological Chemistry, vol. 263, no. 6, pp. 2905–2910, 1988. View at: Google Scholar
  61. P. H. James, M. Pruschy, T. E. Vorherr, J. T. Penniston, and E. Carafoli, “Primary structure of the cAMP-dependent phosphorylation site of the plasma membrane calcium pump,” Biochemistry, vol. 28, no. 10, pp. 4253–4258, 1989. View at: Publisher Site | Google Scholar
  62. P. James, T. Vorherr, J. Krebs et al., “Modulation of erythrocyte Ca2+-ATPase by selective calpain cleavage of the calmodulin-binding domain,” The Journal of Biological Chemistry, vol. 264, no. 14, pp. 8289–8296, 1989. View at: Google Scholar
  63. J. W. Putney Jr., “Capacitative calcium entry: sensing the calcium stores,” Journal of Cell Biology, vol. 169, no. 3, pp. 381–382, 2005. View at: Publisher Site | Google Scholar
  64. O. Brandman, J. Liou, W. S. Park, and T. Meyer, “STIM2 is a feedback regulator that stabilizes basal cytosolic and endoplasmic reticulum Ca2+ levels,” Cell, vol. 131, no. 7, pp. 1327–1339, 2007. View at: Publisher Site | Google Scholar
  65. I. Grigoriev, S. M. Gouveia, B. van der Vaart et al., “STIM1 is a MT-plus-end-tracking protein involved in remodeling of the ER,” Current Biology, vol. 18, no. 3, pp. 177–182, 2008. View at: Publisher Site | Google Scholar
  66. S. Honnappa, S. M. Gouveia, A. Weisbrich et al., “An EB1-binding motif acts as a microtubule tip localization signal,” Cell, vol. 138, no. 2, pp. 366–376, 2009. View at: Publisher Site | Google Scholar
  67. J. Liou, M. Fivaz, T. Inoue, and T. Meyer, “Live-cell imaging reveals sequential oligomerization and local plasma membrane targeting of stromal interaction molecule 1 after Ca2+ store depletion,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 22, pp. 9301–9306, 2007. View at: Google Scholar
  68. S. Feske, Y. Gwack, M. Prakriya et al., “A mutation in Orai1 causes immune deficiency by abrogating CRAC channel function,” Nature, vol. 441, no. 7090, pp. 179–185, 2006. View at: Publisher Site | Google Scholar
  69. M. Prakriya, S. Feske, Y. Gwack, S. Srikanth, A. Rao, and P. G. Hogan, “Orai1 is an essential pore subunit of the CRAC channel,” Nature, vol. 443, no. 7108, pp. 230–233, 2006. View at: Publisher Site | Google Scholar
  70. C. Rosse, M. Linch, S. Kermorgant, A. J. M. Cameron, K. Boeckeler, and P. J. Parker, “PKC and the control of localized signal dynamics,” Nature Reviews Molecular Cell Biology, vol. 11, no. 2, pp. 103–112, 2010. View at: Publisher Site | Google Scholar
  71. H. Mellor and P. J. Parker, “The extended protein kinase C superfamily,” Biochemical Journal, vol. 332, part 2, pp. 281–292, 1998. View at: Google Scholar
  72. H. P. Rang, M. M. Dale, J. M. Ritter, R. J. Flower, and G. Henderson, Rang & Dale's Pharmacology, Churchill Livingstone, Edinburgh, UK, 7th edition, 2011.
  73. B. S. Fogh, H. A. B. Multhaupt, and J. R. Couchman, “Protein kinase C, focal adhesions and the regulation of cell migration,” Journal of Histochemistry and Cytochemistry, vol. 62, no. 3, pp. 172–184, 2014. View at: Publisher Site | Google Scholar
  74. N. Murakami, S. S. Singh, V. P. Chauhan, and M. Elzinga, “Phospholipid binding, phosphorylation by protein kinase C, and filament assembly of the COOH terminal heavy chain fragments of nonmuscle myosin II isoforms MIIA and MIIB,” Biochemistry, vol. 34, no. 49, pp. 16046–16055, 1995. View at: Publisher Site | Google Scholar
  75. N. Murakami, V. P. S. Chauhan, and M. Elzinga, “Two nonmuscle myosin II heavy chain isoforms expressed in rabbit brains: filament forming properties, the effects of phosphorylation by protein kinase C and casein kinase II, and location of the phosphorylation sites,” Biochemistry, vol. 37, no. 7, pp. 1989–2003, 1998. View at: Publisher Site | Google Scholar
  76. N. G. Dulyaninova, V. N. Malashkevich, S. C. Almo, and A. R. Bresnick, “Regulation of myosin-IIA assembly and Mts1 binding by heavy chain phosphorylation,” Biochemistry, vol. 44, no. 18, pp. 6867–6876, 2005. View at: Publisher Site | Google Scholar
  77. N. G. Dulyaninova, R. P. House, V. Betapudi, and A. R. Bresnick, “Myosin-IIA heavy-chain phosphorylation regulates the motility of MDA-MB-231 carcinoma cells,” Molecular Biology of the Cell, vol. 18, no. 8, pp. 3144–3155, 2007. View at: Publisher Site | Google Scholar
  78. S. Etienne-Manneville and A. Hall, “Integrin-mediated activation of Cdc42 controls cell polarity in migrating astrocytes through PKCζ,” Cell, vol. 106, no. 4, pp. 489–498, 2001. View at: Publisher Site | Google Scholar
  79. S. Etienne-Manneville and A. Hall, “Cdc42 regulates GSK-3β and adenomatous polyposis coli to control cell polarity,” Nature, vol. 421, no. 6924, pp. 753–756, 2003. View at: Publisher Site | Google Scholar
  80. C. Rosse, E. Formstecher, K. Boeckeler et al., “An aPKC-Exocyst complex controls paxillin phosphorylation and migration through localised JNK1 activation,” PLoS Biology, vol. 7, no. 11, Article ID e1000235, 2009. View at: Publisher Site | Google Scholar
  81. X. R. Bustelo, V. Sauzeau, and I. M. Berenjeno, “GTP-binding proteins of the Rho/Rac family: Regulation, effectors and functions in vivo,” BioEssays, vol. 29, no. 4, pp. 356–370, 2007. View at: Publisher Site | Google Scholar
  82. S. Yang, J. J. Zhang, and X.-Y. Huang, “Orai1 and STIM1 are critical for breast tumor cell migration and metastasis,” Cancer Cell, vol. 15, no. 2, pp. 124–134, 2009. View at: Publisher Site | Google Scholar
  83. Y. Dai, N. L. Dudek, Q. Li, and N. A. Muma, “Phospholipase C, Ca2+, and calmodulin signaling are required for 5-HT2A receptor-mediated transamidation of Rac1 by transglutaminase,” Psychopharmacology, vol. 213, no. 2-3, pp. 403–412, 2011. View at: Publisher Site | Google Scholar
  84. P. Kanchanawong, G. Shtengel, A. M. Pasapera et al., “Nanoscale architecture of integrin-based cell adhesions,” Nature, vol. 468, no. 7323, pp. 580–584, 2010. View at: Publisher Site | Google Scholar
  85. E. Mostafavi, A. A. Nargesi, Z. Ghazizadeh et al., “The degree of resistance of erythrocyte membrane cytoskeletal proteins to supra-physiologic concentrations of calcium: an in vitro study,” The Journal of Membrane Biology, vol. 247, no. 8, pp. 695–701, 2014. View at: Publisher Site | Google Scholar
  86. F. Wang, D.-Z. Liu, H. Xu et al., “Thapsigargin induces apoptosis by impairing cytoskeleton dynamics in human lung adenocarcinoma cells,” The Scientific World Journal, vol. 2014, Article ID 619050, 7 pages, 2014. View at: Publisher Site | Google Scholar
  87. Y. Ohta, E. Nishida, and H. Sakai, “Type II Ca2+/calmodulin-dependent protein kinase binds to actin filaments in a calmodulin-sensitive manner,” FEBS Letters, vol. 208, no. 2, pp. 423–426, 1986. View at: Publisher Site | Google Scholar
  88. C. Larsson, “Protein kinase C and the regulation of the actin cytoskeleton,” Cellular Signalling, vol. 18, no. 3, pp. 276–284, 2006. View at: Publisher Site | Google Scholar
  89. L. Hoffman, M. M. Farley, and M. N. Waxham, “Calcium-calmodulin-dependent protein kinase II isoforms differentially impact the dynamics and structure of the actin cytoskeleton,” Biochemistry, vol. 52, no. 7, pp. 1198–1207, 2013. View at: Publisher Site | Google Scholar
  90. C.-B. Guan, H.-T. Xu, M. Jin, X.-B. Yuan, and M.-M. Poo, “Long-range Ca2+ signaling from growth cone to soma mediates reversal of neuronal migration induced by slit-2,” Cell, vol. 129, no. 2, pp. 385–395, 2007. View at: Publisher Site | Google Scholar
  91. Z.-H. Huang, Y. Wang, Z.-D. Su et al., “Slit-2 repels the migration of olfactory ensheathing cells by triggering Ca2+-dependent cofilin activation and RhoA inhibition,” Journal of Cell Science, vol. 124, no. 2, pp. 186–197, 2011. View at: Publisher Site | Google Scholar
  92. M. Nagano, D. Hoshino, N. Koshikawa, T. Akizawa, and M. Seiki, “Turnover of focal adhesions and cancer cell migration,” International Journal of Cell Biology, vol. 2012, Article ID 310616, 10 pages, 2012. View at: Publisher Site | Google Scholar
  93. A. Glading, D. A. Lauffenburger, and A. Wells, “Cutting to the chase: calpain proteases in cell motility,” Trends in Cell Biology, vol. 12, no. 1, pp. 46–54, 2002. View at: Publisher Site | Google Scholar
  94. S. J. Franco, M. A. Rodgers, B. J. Perrin et al., “Calpain-mediated proteolysis of talin regulates adhesion dynamics,” Nature Cell Biology, vol. 6, no. 10, pp. 977–983, 2004. View at: Publisher Site | Google Scholar
  95. C. L. Cortesio, L. R. Boateng, T. M. Piazza, D. A. Bennin, and A. Huttenlocher, “Calpain-mediated proteolysis of paxillin negatively regulates focal adhesion dynamics and cell migration,” The Journal of Biological Chemistry, vol. 286, no. 12, pp. 9998–10006, 2011. View at: Publisher Site | Google Scholar
  96. K. T. Chan, D. A. Bennin, and A. Huttenlocher, “Regulation of adhesion dynamics by calpain-mediated proteolysis of focal adhesion kinase (FAK),” The Journal of Biological Chemistry, vol. 285, no. 15, pp. 11418–11426, 2010. View at: Publisher Site | Google Scholar
  97. M. D. Sjaastad and W. J. Nelson, “Integrin-mediated calcium signaling and regulation of cell adhesion by intracellular calcium,” BioEssays, vol. 19, no. 1, pp. 47–55, 1997. View at: Publisher Site | Google Scholar
  98. S. Lev, H. Moreno, R. Martinez et al., “Protein tyrosine kinase PYK2 involved in Ca2+-induced regulation of ion channel and MAP kinase functions,” Nature, vol. 376, no. 6543, pp. 737–745, 1995. View at: Publisher Site | Google Scholar
  99. S. Avraham, R. London, Y. Fu et al., “Identification and characterization of a novel related adhesion focal tyrosine kinase (RAFTK) from megakaryocytes and brain,” The Journal of Biological Chemistry, vol. 270, no. 46, pp. 27742–27751, 1995. View at: Publisher Site | Google Scholar
  100. J. Kostan, M. Gregor, G. Walko, and G. Wiche, “Plectin isoform-dependent regulation of keratin-integrin α6β4 anchorage via Ca2+/calmodulin,” The Journal of Biological Chemistry, vol. 284, no. 27, pp. 18525–18536, 2009. View at: Publisher Site | Google Scholar
  101. J.-G. Song, J. Kostan, F. Drepper et al., “Structural insights into Ca2+-calmodulin regulation of plectin 1a-integrin β4 interaction in hemidesmosomes,” Structure, vol. 23, no. 3, pp. 558–570, 2015. View at: Publisher Site | Google Scholar
  102. M. Monet, V. Lehen'kyi, F. Gackiere et al., “Role of cationic channel TRPV2 in promoting prostate cancer migration and progression to androgen resistance,” Cancer Research, vol. 70, no. 3, pp. 1225–1235, 2010. View at: Publisher Site | Google Scholar
  103. T. Long, J. Su, W. Tang et al., “A novel interaction between calcium-modulating cyclophilin ligand and Basigin regulates calcium signaling and matrix metalloproteinase activities in human melanoma cells,” Cancer Letters, vol. 339, no. 1, pp. 93–101, 2013. View at: Publisher Site | Google Scholar
  104. E. Suyama, R. Wadhwa, K. Kaur et al., “Identification of metastasis-related genes in a mouse model using a library of randomized ribozymes,” The Journal of Biological Chemistry, vol. 279, no. 37, pp. 38083–38086, 2004. View at: Publisher Site | Google Scholar
  105. H. Zhu, H. Zhang, F. Jin et al., “Elevated Orai1 expression mediates tumor-promoting intracellular Ca2+ oscillations in human esophageal squamous cell carcinoma,” Oncotarget, vol. 5, pp. 3455–3471, 2014. View at: Google Scholar
  106. J.-H. Kim, S. Lkhagvadorj, M.-R. Lee et al., “Orai1 and STIM1 are critical for cell migration and proliferation of clear cell renal cell carcinoma,” Biochemical and Biophysical Research Communications, vol. 448, no. 1, pp. 76–82, 2014. View at: Publisher Site | Google Scholar
  107. H. Stanisz, S. Saul, C. S. L. Müller et al., “Inverse regulation of melanoma growth and migration by Orai1/STIM2-dependent calcium entry,” Pigment Cell and Melanoma Research, vol. 27, no. 3, pp. 442–453, 2014. View at: Publisher Site | Google Scholar
  108. L. Clarysse, M. Guéguinou, M. Potier-Cartereau et al., “cAMP-PKA inhibition of SK3 channel reduced both Ca2+ entry and cancer cell migration by regulation of SK3-Orai1 complex,” Pflügers Archiv, vol. 466, no. 10, pp. 1921–1932, 2014. View at: Publisher Site | Google Scholar
  109. A. Chantôme, M. Potier-Cartereau, L. Clarysse et al., “Pivotal role of the lipid raft SK3-orai1 complex in human cancer cell migration and bone metastases,” Cancer Research, vol. 73, no. 15, pp. 4852–4861, 2013. View at: Publisher Site | Google Scholar
  110. Y.-T. Chen, Y.-F. Chen, W.-T. Chiu et al., “Microtubule-associated histone deacetylase 6 supports the calcium store sensor STIM1 in mediating malignant cell behaviors,” Cancer Research, vol. 73, no. 14, pp. 4500–4509, 2013. View at: Publisher Site | Google Scholar
  111. R. K. Motiani, M. C. Hyzinski-García, X. Zhang et al., “STIM1 and Orai1 mediate CRAC channel activity and are essential for human glioblastoma invasion,” Pflugers Archiv European Journal of Physiology, vol. 465, no. 9, pp. 1249–1260, 2013. View at: Publisher Site | Google Scholar
  112. M. L. Greenberg, Y. Yu, S. Leverrier, S. L. Zhang, I. Parker, and M. D. Cahalan, “Orai1 function is essential for T cell homing to lymph nodes,” Journal of Immunology, vol. 190, no. 7, pp. 3197–3206, 2013. View at: Publisher Site | Google Scholar
  113. S. Derouiche, M. Warnier, P. Mariot et al., “Bisphenol A stimulates human prostate cancer cell migration via remodelling of calcium signalling,” SpringerPlus, vol. 2, article 54, 2013. View at: Publisher Site | Google Scholar
  114. Y.-T. Chen, Y.-F. Chen, W.-T. Chiu, Y.-K. Wang, H.-C. Chang, and M.-R. Shen, “The ER Ca2+ sensor STIM1 regulates actomyosin contractility of migratory cells,” Journal of Cell Science, vol. 126, no. 5, pp. 1260–1267, 2013. View at: Publisher Site | Google Scholar
  115. N. Yang, Y. Tang, F. Wang et al., “Blockade of store-operated Ca2+ entry inhibits hepatocarcinoma cell migration and invasion by regulating focal adhesion turnover,” Cancer Letters, vol. 330, no. 2, pp. 163–169, 2013. View at: Publisher Site | Google Scholar
  116. J. Yoshida, K. Iwabuchi, T. Matsui, T. Ishibashi, T. Masuoka, and M. Nishio, “Knockdown of stromal interaction molecule 1 (STIM1) suppresses store-operated calcium entry, cell proliferation and tumorigenicity in human epidermoid carcinoma A431 cells,” Biochemical Pharmacology, vol. 84, no. 12, pp. 1592–1603, 2012. View at: Publisher Site | Google Scholar
  117. X.-P. Cong, W.-H. Wang, X. Zhu, C. Jin, L. Liu, and X.-M. Li, “Silence of STIM1 attenuates the proliferation and migration of EPCs after vascular injury and its mechanism,” Asian Pacific Journal of Tropical Medicine, vol. 7, no. 5, pp. 373–377, 2014. View at: Publisher Site | Google Scholar
  118. P. Kar, K. Samanta, H. Kramer, O. Morris, D. Bakowski, and A. B. Parekh, “Dynamic assembly of a membrane signaling complex enables selective activation of NFAT by Orai1,” Current Biology, vol. 24, no. 12, pp. 1361–1368, 2014. View at: Publisher Site | Google Scholar
  119. M. Vandenberghe, M. Raphaël, V. Lehen'kyi et al., “ORAI1 calcium channel orchestrates skin homeostasis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 110, no. 50, pp. E4839–E4848, 2013. View at: Publisher Site | Google Scholar
  120. I.-H. Yang, Y.-T. Tsai, S.-J. Chiu et al., “Involvement of STIM1 and Orai1 in EGF-mediated cell growth in retinal pigment epithelial cells,” Journal of Biomedical Science, vol. 20, article 41, 2013. View at: Publisher Site | Google Scholar
  121. C. Schäfer, G. Rymarczyk, L. Ding, M. T. Kirber, and V. M. Bolotina, “Role of molecular determinants of store-operated Ca2+ entry (Orai1, phospholipase A2 group 6, and STIM1) in focal adhesion formation and cell migration,” The Journal of Biological Chemistry, vol. 287, no. 48, pp. 40745–40757, 2012. View at: Google Scholar
  122. A. M. Spinelli, J. C. González-Cobos, X. Zhang et al., “Airway smooth muscle STIM1 and Orai1 are upregulated in asthmatic mice and mediate PDGF-activated SOCE, CRAC currents, proliferation, and migration,” Pflügers Archiv, vol. 464, no. 5, pp. 481–492, 2012. View at: Publisher Site | Google Scholar
  123. N. Suganuma, S. Ito, H. Aso et al., “STIM1 regulates platelet-derived growth factor-induced migration and Ca2+ influx in human airway smooth muscle cells,” PLoS ONE, vol. 7, no. 9, Article ID e45056, 2012. View at: Publisher Site | Google Scholar
  124. J. N. Rao, N. Rathor, R. Zhuang et al., “Polyamines regulate intestinal epithelial restitution through TRPC1-mediated Ca2+ signaling by differentially modulating STIM1 and STIM2,” The American Journal of Physiology—Cell Physiology, vol. 303, no. 3, pp. C308–C317, 2012. View at: Publisher Site | Google Scholar
  125. R.-W. Guo, L.-X. Yang, M.-Q. Li, X.-H. Pan, B. Liu, and Y.-L. Deng, “Stim1-and Orai1-mediated store-operated calcium entry is critical for angiotensin II-induced vascular smooth muscle cell proliferation,” Cardiovascular Research, vol. 93, no. 2, pp. 360–370, 2012. View at: Publisher Site | Google Scholar
  126. C.-Y. Kuang, Y. Yu, R.-W. Guo et al., “Silencing stromal interaction molecule 1 by RNA interference inhibits the proliferation and migration of endothelial progenitor cells,” Biochemical and Biophysical Research Communications, vol. 398, no. 2, pp. 315–320, 2010. View at: Publisher Site | Google Scholar
  127. J. M. Bisaillon, R. K. Motiani, J. C. Gonzalez-Cobos et al., “Essential role for STIM1/Orai1-mediated calcium influx in PDGF-induced smooth muscle migration,” The American Journal of Physiology—Cell Physiology, vol. 298, no. 5, pp. C993–C1005, 2010. View at: Publisher Site | Google Scholar
  128. M. Potier, J. C. Gonzalez, R. K. Motiani et al., “Evidence for STIM1- and Orai1-dependent store-operated calcium influx through ICRAC in vascular smooth muscle cells: role in proliferation and migration,” The FASEB Journal, vol. 23, no. 8, pp. 2425–2437, 2009. View at: Publisher Site | Google Scholar

Copyright © 2015 Feng-Chiao Tsai et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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