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Kozak JA, Putney JW Jr., editors. Calcium Entry Channels in Non-Excitable Cells. Boca Raton (FL): CRC Press/Taylor & Francis; 2018. doi: 10.1201/9781315152592-12

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Calcium Entry Channels in Non-Excitable Cells.

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Chapter 12 Regulation and Role of Store-Operated Ca2+ Entry in Cellular Proliferation

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12.1. Introduction

Ca2+ is a ubiquitous intracellular messenger that transduces a variety of cellular responses downstream of the activation of G-protein-coupled or tyrosine kinase receptors. Depending on the agonist and cellular context, Ca2+ can mediate different responses in the same cell [1]. The specific cellular response transduced downstream of the particular Ca2+ transient is encoded in the spatial and temporal dynamics of the Ca2+ signal, leading to the activation of a subset of Ca2+-dependent effectors and the ensuing cellular response. As such, the duration, amplitude, frequency, and spatial localization of Ca2+ signals encode targeted signals that activate Ca2+-sensitive effectors to define a particular cellular response. To generate and fine-tune those Ca2+ signals, cells use two main Ca2+ sources: entry of extracellular Ca2+ and Ca2+ release from intracellular stores. The primary intracellular Ca2+ store is the endoplasmic reticulum (ER), which can concentrate Ca2+ in the hundreds of μM range [2]. In contrast, cytoplasmic Ca2+ concentration is kept at rest at extremely low levels (∼100 nM or lower), thus providing a low-noise background for detection of complex Ca2+ dynamics [3].

The Ca2+-signaling machinery includes Ca2+ entry and extrusion pathways in the plasma membrane (PM), ER membrane Ca2+ release channels, and Ca2+ reuptake ATPases within the ER membrane [4]. These Ca2+ transport pathways, in addition to intracellular Ca2+ buffers and Ca2+ uptake and release through other intracellular organelles, primarily the mitochondria, combine to shape highly tuned and dynamic Ca2+ transients that regulate cellular functions [5].

Under physiological conditions in non-excitable cells, Ca2+ transients are typically initiated downstream of agonist stimulation through the activation of the PLC-IP3 signal transduction cascade, which leads to the opening of intracellular Ca2+ channel inositol 1,4,5-trisphosphate receptors (IP3Rs) to release Ca2+ from intracellular stores [6]. Ca2+ release depletes the stores and activates a Ca2+ influx pathway in the PM termed store-operated Ca2+ entry (SOCE). SOCE is mediated by two key players: ER transmembrane Ca2+ sensors represented by the STIM family of proteins and PM Ca2+ channels of the Orai family that link directly to STIMs (see Chapters 1 through 3). The N-terminus of STIM1 faces the ER lumen and consists of two EF-hand domains that detect luminal Ca2+ concentration. The loss of STIM1 Ca2+ binding upon store depletion leads to conformational changes in the protein and its aggregation into clusters that translocate and stabilize into ER-PM junctions with very close apposition (∼20 nm) [7]. STIM1 within these ER-PM junctions binds to and recruits Orai1 through a diffusional trap mechanism, resulting in opening Orai1 channels and Ca2+ entry [8]. As such, the STIM-Orai clusters at ER-PM junctions define a specific microdomain at ER-PM junctions that also include the ER Ca-ATPase (SERCA) [9,10].

The tightly regulated remodeling of the Ca2+-signaling machinery upon store depletion allows for specific Ca2+ signaling in the midrange between Ca2+ microdomains and global Ca2+ waves [10] (see Chapter 5). Spatially, Ca2+ signaling can occur in localized spatially restricted elementary Ca2+ release events that activate effectors located in the immediate proximity of the Ca2+ channel. Alternatively, Ca2+ signals/waves occur/spread through the entire cell resulting in a global spatially unrestricted signal.

We have recently described a SOCE-dependent Ca2+-signaling modularity that signals in the midrange between these two spatial extremes [10]. Store depletion downstream of receptor activation and IP3 generation results in a localized Ca2+ entry point source at the SOCE clusters that induces Ca2+ entry into the cytoplasm, which is readily taken up into the ER lumen through SERCA activity only to be released again through open IP3Rs distally to the SOCE entry site and gate Ca2+-activated Cl- channels as downstream Ca2+ effectors. This mechanism, referred to as “Ca2+ teleporting,” allows for specific activation of Ca2+ effectors that are distant from the point source Ca2+ channel without inducing a global Ca2+ wave, thus providing a novel module in the Ca2+-signaling repertoire. A cartoon summary of Ca2+ teleporting is found in Figure 12.1.

Figure 12.1. Cartoon illustrating Ca2+ teleporting that mediates midrange Ca2+ signaling.

Figure 12.1

Cartoon illustrating Ca2+ teleporting that mediates midrange Ca2+ signaling. Receptor activation generates IP3 that releases Ca2+ from stores. Following store depletion, the SOCE machinery forms spatially localized clusters that support Ca2+ entry. Ca (more...)

The relationship between Ca2+ signaling and cellular proliferation is complex with Ca2+ transients detected at various stages of the cell cycle [2]. These transients are thought to activate a multitude of Ca2+ effectors downstream of the initial Ca2+ signal, which were shown to be important for cellular proliferation, including, for example, calmodulin (CaM) and Ca2+-CaM-dependent protein kinase II (CaMKII).

However, there are a few cases where Ca2+ signals have been shown directly to be critical for cell cycle progression, including in mitosis for nuclear envelope breakdown and for chromosome disjunction [11]. In contrast, nuclear envelope breakdown during meiosis in vertebrate oocytes occurs independently of Ca2+, but Ca2+ is required for the completion of meiosis I in vertebrate oocytes [12–14]. Interestingly, multiple Ca2+ signaling pathways are modified during M-phase of the cell cycle, with the best defined example being Xenopus oocyte maturation [15].

Several Ca2+ influx pathways have been implicated in cell proliferation and cell cycle progression, including TRP channels, voltage-gated Ca2+ channels (CaV), purinergic P2X receptors, ionotropic glutamate receptors, and SOCE [16]. Blockers of voltage-gated Ca2+ channels were shown to slow down cell growth, arguing for a role for these channels in cell cycle progression [16–18]. Experimental manipulation of the expression levels of members of the TRPC, TRPV, and TRPM families of cation channels, which are Ca2+ permeable, was linked to cell proliferation with differential effects depending on the particular channel studied [16]. However, some of these studies are difficult to interpret because channel knockdown or overexpression could have significantly broader effects on Ca2+ signaling than affecting Ca2+ influx through the specific channel in question, as it may lead to changes in expression of other Ca2+-signaling pathways as a compensatory mechanism. Furthermore, the majority of TRP channels conduct cations with some Ca2+ permeability and are not Ca2+ selective like Orai1 or CaV channels, with the exception of TRPV5 and TRPV6 (see Chapter 13). Hence, changes in their expression is likely to affect the ionic balance across the cell membrane with effects on resting membrane potential, which may in turn affect cell proliferation.

The relationship between SOCE and cell proliferation is an intimate one that goes beyond the well-recognized roles of Ca2+ signaling in cellular growth and proliferation. SOCE is dramatically downregulated during the division phase of the cell cycle through mechanisms that have not been fully elucidated. This is in line with the significant remodeling of the Ca2+-signaling machinery during M-phase, which has been well characterized during oocyte meiosis. Furthermore, there is mounting evidence from multiple neoplasms for an important role for SOCE in metastasis.

This chapter presents a brief overview of our current knowledge as to the mechanisms regulating SOCE during cell cycle from cellular proliferation to metastasis with an emphasis on SOCE regulation during cell division (mitosis and meiosis).

12.2. Role of Ca2+ Signals in Cellular Proliferation

The requirement for Ca2+ in cell proliferation has been known for decades with the optimum extracellular Ca2+ concentration to support proliferation being in the 0.5–1 mM range [16]. However, the requirement for Ca2+ during cell growth and proliferation is vague and difficult to define given the broad involvement of Ca2+ signaling in many aspects of cellular physiology and signaling. Also, bifurcating signaling pathways such as G-protein-coupled receptor activation can induce cell proliferation together with a Ca2+ signal without any direct link between Ca2+ and cell growth. Nonetheless, Ca2+ influx appears to support cellular proliferation since immortalized/neoplastic cells do not require as much Ca2+ in the extracellular medium as primary cells, and this is of importance for cell cycle studies and in the design of anticancer drugs.

Ca2+ release from intracellular stores through IP3Rs or ryanodine receptors (RyR) has been implicated in proliferation. IP3R activation is involved in stimulating the proliferation of stem cells, kidney cells, pancreatic beta cells, breast cancer, and vascular smooth muscle cells [19–26]. RyR are more likely to be involved in the regulation of cellular differentiation [25]. The activation of IP3Rs also stimulates the proliferation of breast cancer cells [26]. Interestingly, breast cancer cells rely for their survival and proliferation on Ca2+ released via IP3Rs to fuel mitochondrial respiration, making this Ca2+-dependent cross talk between the ER and mitochondria a potential target for drug development [27].

Further strengthening a role for Ca2+ in proliferation, several Ca2+-dependent downstream effectors have been implicated in cellular proliferation. A primary intracellular Ca2+-binding effector, the ubiquitous cellular Ca2+-sensing protein CaM, is required for cell cycle progression and proliferation [28,29]. Furthermore, CaM levels are highly regulated during the cell cycle with an increase at the G1/S transition [30]. Several effectors downstream of CaM have been identified, including CaM kinases I and II that interfere with various steps of the cell cycle, and CaM-dependent protein phosphatase calcineurin [31–33]. In T cells, calcineurin dephosphorylates and activates the nuclear factor of activated T cells (NFAT), inducing its nuclear translocation, where it stimulates gene transcription in support of cell proliferation [34] (see Chapter 5). Several other effectors have been identified downstream of Ca2+ signals and the activation of CaM-dependent phosphorylation/dephosphorylation processes such as NF-κB and the cAMP response element [33].

Ca2+ influx through multiple pathways has also been implicated in cell proliferation. CaV channels, particularly T-type channels, have been described as regulating the proliferation of several cancer cells [35], and in the case of TRP channels, most knockdown experiments revealed that they are also required for cell proliferation [16]. SOCE is now probably the most extensively studied Ca2+ pathway involved in cell proliferation. Beyond the immune response, NFAT stimulation by SOCE is involved in the proliferation of other cell types such as neural progenitor cells [36,37], osteoblasts [38], kidney cells [39], and endothelial and endothelial progenitor cells involved in angiogenesis in cancer patients [40,41].

12.3. Remodeling of the Ca2+-Signaling Machinery during Meiosis

As discussed earlier, not only are Ca2+ signals important for cellular proliferation and cell cycle progression, but Ca2+ transport pathways are themselves modulated during the cell cycle. This regulation of Ca2+ signaling during the cell cycle is presumed to support its progression although direct evidence for this remains scarce. An important case study is the remodeling of Ca2+-signaling pathways during Xenopus oocyte meiosis, where multiple Ca2+-signaling modules are modified to prepare the egg for fertilization and the egg-to-embryo transition.

Vertebrate oocytes arrest at prophase of meiosis I for prolonged periods of time [42,43]. Before oocytes acquire the competency to be fertilized, they undergo a maturation period during which they progress to metaphase of meiosis II in a process termed “oocyte maturation.” Mature oocytes, typically referred to as eggs, complete meiosis at fertilization and transition to the mitotic cell cycle. Egg activation is highly Ca2+ dependent as Ca2+ mediates critical steps to initiate development, including the block of polyspermy and the completion of meiosis II to ensure a proper egg-to-embryo transition. In order to be effective, these events need to occur in a chronological fashion since polyspermy needs to be prevented before the completion of meiosis to ensure zygote viability. Throughout phylogeny in all sexually reproducing species investigated to date, Ca2+ has been shown to encode the egg-to-embryo transition through a species-specific Ca2+ transient at fertilization with well-defined spatial, temporal, and amplitude dynamics [44–47]. Eggs acquire the ability to produce this fertilization-specific Ca2+ transient only after oocyte maturation, owing to dramatic remodeling of the Ca2+-signaling machinery during oocyte maturation that affects multiple Ca2+ transport proteins such as IP3R, PMCA, and Orai/STIM [15].

The fertilization-specific Ca2+ signal in Xenopus is distinguished by a sustained elevated Ca2+ plateau that lasts for several minutes following a local Ca2+ rise at the site of sperm entry that spreads slowly (∼9 μm/s), in the form of a Ca2+ wave, across the entire egg [48–51]. In contrast, Ca2+ transients in oocytes tend to have a saltatory mode of propagation with a more rapid wave speed (∼20 μm/s) [52–54]. Given that the frog oocytes express only the type 1 IP3R isoform and no RyR [55], these changes in Ca2+ release during oocyte maturation are thus due to changes in the regulation of the IP3R during maturation as discussed in more detail in the following text. IP3-dependent Ca2+ release plays a central role in mediating the Ca2+ transient at fertilization in vertebrate eggs.

In mammals, the slow oscillations that are maintained for long periods of time depend on Ca2+ influx from the extracellular space, presumably through the SOCE pathway [56–58]. Fertilization activates phospholipase Cγ (PLCγ) in Xenopus or delivers PLCζ in mammalian eggs, which increases IP3 production and gates IP3Rs to release Ca2+ from the ER [59–62]. It is clear that IP3-dependent Ca2+ release properties are modulated during oocyte maturation when an increase in IP3-dependent Ca2+-release sensitivity is noted and conserved among different species [54,63–67].

IP3-dependent Ca2+ release is sensitized during Xenopus oocyte maturation [54]. Threshold IP3 concentrations lead to small and spatially separate elementary Ca2+ release events (puffs) in oocytes [68]; whereas in eggs, similar IP3 concentrations lead to larger consolidated events referred to as single release events (SREs) (Figure 12.2b) [54]. These larger release events are likely due to the clustering of the smaller elementary Ca2+ puffs [54,69–71]. The ER remodels during oocyte maturation forming large patches that are enriched in IP3Rs as compared to the neighboring reticular ER (Figure 12.2a) [72]. Interestingly, we found that this clustering sensitizes IP3Rs within ER patches, to respond to lower IP3 concentrations as compared to IP3Rs that localize to the reticular ER [72]. This sensitization appears to be due to increased Ca2+-dependent cooperativity at subthreshold IP3 concentrations due to the physical clustering of IP3Rs within ER patches, because IP3Rs freely exchange between the two ER compartments (i.e., patches and reticular ER) [72]. A similar sensitization of the IP3R is observed during mitosis [73] and has been suggested to depend on cdk1 phosphorylation of IP3R [74].

Image

Figure 12.2

Clustering of elementary Ca2+ release events during oocyte maturation.

Sensitization of IP3-dependent Ca2+ release during oocyte maturation takes place simultaneously with the activation of multiple kinase cascades that drive this differentiation pathway. This argued for a potential role for phosphorylation of IP3R in modulating its sensitivity in both Xenopus and mouse eggs, especially that IP3R was found to be phosphorylated specifically during oocyte maturation at maturation-promoting factor (MPF)/mitogen-activated protein kinase (MAPK) conserved sites [54,75,76]. However, direct evidence showing that phosphorylation at these residues modulates IP3 sensitivity of the channels is lacking. Furthermore, IP3R sensitization could also be related to the number of functional IP3Rs, given their gating cooperativity. In that context, the number of functional IP3Rs increases during Xenopus oocyte maturation (without a significant change in total IP3R protein pool) as they translocate from annulate lamellae to the ER [77,78]. Annulate lamellae represent an oocyte-specific vesicular compartment to which IP3Rs localize but are silenced presumably through protein-protein interactions [77,78].

Two additional transport pathways important in defining Ca2+ transient in the frog oocyte are the PM Ca2+ ATPase (PMCA) that possesses high affinity for Ca2+ and functions to decrease Ca2+ levels back to the resting state [79,80] and the SERCA pump that ensures Ca2+ sequestration into the ER lumen [81]. PMCA, which localizes to the cell membrane in immature Xenopus oocytes, is internalized into an intracellular vesicular pool during oocyte maturation [71]. Coupled to the continuous and constant SERCA-dependent Ca2+ reuptake and the increased sensitivity of IP3-dependent Ca2+ release, PMCA internalization contributes to the sustained Ca2+ plateau after the slow rising Ca2+ phase at fertilization [15,71].

Finally, SOCE completely inactivates in the mature Xenopus egg compared to immature oocytes [82,83]. The regulation of SOCE during meiosis is covered in more detail in the next section.

12.4. SOCE Inactivation during M-Phase

Immature Xenopus oocytes possess a robust SOC current that has similar biophysical properties to CRAC, the prototypical SOC channel activity originally characterized in immune cells [82,84,85] (see Chapter 1). However, during oocyte maturation, SOCE is inactivated completely: in mature eggs, SOCE is no longer activated when stores are depleted [82]. The inhibition of SOCE requires the activation of maturation-promoting factor (MPF, composed of CDK1 and cyclin B) [83]. This was shown by measuring both the SOC current and the levels of activation of multiple kinases that drive oocyte maturation at the single-oocyte level, to allow for direct correlation between the activity of various kinases and SOCE at the single-cell level [83]. Therefore, the fertilization-specific Ca2+ transient in Xenopus eggs is generated without contribution from SOCE. Xenopus eggs respond to sperm entry with a single sweeping Ca2+ transient that lasts several minutes [49,86]. This Ca2+ signal encodes subsequent events associated with egg activation in the following order: (1) fast block to polyspermy due to gating of Ca2+-activated Cl- channels that depolarize the cell membrane; (2) slow block to polyspermy due to cortical granule fusion; and (3) completion of meiosis due to calcineurin and CaMKII activation [15,87,88]. SOCE inactivation is likely to contribute to shaping the dynamics of the fertilization-specific Ca2+ signal and therefore promote the egg-to-embryo transition.

Interestingly, SOCE is downregulated but not completely inactivated during mammalian oocyte meiosis as store depletion in metaphase II eggs induces Ca2+ entry, which supports Ca2+ oscillations following fertilization [89]. In contrast to Xenopus eggs, mammalian oocytes respond at fertilization with multiple Ca2+ oscillations that can last for hours [57]. Maintenance of these Ca2+ oscillations depends on Ca2+ influx through SOCE, presumably to refill Ca2+ stores and provide a continuous Ca2+ source for the Ca2+ release waves. There is, hence, a correlation between the occurrence of SOCE during meiosis and the ability of the oocyte to support Ca2+ oscillations at fertilization. Some reports have argued that SOCE amplitude increases during oocyte maturation in both mouse and pig [90,91]. In contrast, we and others have shown that SOCE is downregulated but not completely inhibited during mouse oocyte meiosis, and that downregulated SOCE is required for the egg-to-embryo transition [92,93]. The reasons for these discrepancies remain unclear. Nonetheless, collectively the data suggest that SOCE downregulation during vertebrate oocyte maturation represents an important determinant of the remodeling of Ca2+ signaling in preparation for fertilization.

Furthermore, similar to what is observed in meiosis of frog oocytes, SOCE is also inhibited during mitosis of mammalian cells. In the late 1980s, Volpi and Berlin showed that histamine stimulation during interphase in HeLa cells produced an initial Ca2+ rise owing to Ca2+ release from internal stores, followed by an elevated plateau due to Ca2+ influx from the extracellular space [94]. By contrast, histamine stimulation in mitotic cells resulted only in the Ca2+ release phase, arguing that Ca2+ influx is inhibited during mitosis. This observation was made around the time when the initial ideas regarding SOCE were being formulated. The same group later argued that SOCE inhibition during mitosis occurs due to uncoupling of store depletion from SOCE, as thapsigargin (an agent that causes store depletion by blocking SERCA) activated SOCE in interphase but not mitotic cells [95]. More recent studies confirmed that SOCE is inactivated during mitosis in RBL-2H3, HeLa, and HEK293 cells [96–98].

Investigating SOCE levels throughout the cell cycle showed that there is a slight enhancement of SOCE during the G1 and S phases, and dramatic downregulation during M-phase [98]. Consistent with this cell cycle-dependent modulation of SOCE activity, SOCE has been shown to control the G1/S transition but is not necessary during S-phase or the G2/M transition [99]. Furthermore, SOCE has emerged as an important player in cell proliferation, yet the mechanisms by which it controls distinct phases of the cell cycle remain elusive. Recent studies show that inactivation of SOCE by silencing STIM1 in smooth muscle cells, cervical and breast cancer cells significantly inhibited proliferation by slowing down cell cycle progression [21,100]. This is discussed in more detail in Section 12.6.

Collectively, most current evidence argues that SOCE downregulation during M-phase is conserved and as such could be physiologically significant. Although this has not been directly addressed experimentally, one can speculate that tight regulation of Ca2+ signaling is required during M-phase owing to its important functional role at multiple steps throughout the process including nuclear-envelope breakdown, anaphase onset, and cell cleavage. Hence, SOCE inactivation might represent a safety mechanism that prevents sporadic Ca2+ signals from occurring during cell division, which may disrupt its normal progression.

12.5. Mechanisms Regulating SOCE Inactivation during M-Phase

Aside from the role of Maturation Promoting Factor (MPF, Cdk1) in SOCE inhibition, very little was known regarding the mechanistic regulation of SOCE inactivation during M-phase. It has also been argued that SOCE inhibition during mitosis in COS-7 cells is the result of the microtubule-network remodeling that accompanies mitosis [101]. Laser scanning confocal microscopy to monitor cytosolic Ca2+ dynamics revealed that SOCE was progressively inhibited in mitosis and became virtually absent during metaphase [101]. Russa and colleagues used various cytoskeletal modifying drugs and immunofluorescence to assess the contribution of microtubule and actin filaments to SOCE. Nocodazole treatment caused microtubule reorganization and retraction from the cell periphery that mimicked the natural mitotic microtubule remodeling that was also accompanied by SOCE inhibition. Short exposure to paclitaxel, a microtubule-stabilizing drug, strengthened SOCE, whereas long exposure resulted in microtubule disruption and SOCE inhibition. Actin-modifying drugs (cytochalasin D, calyculin A) did not affect SOCE. These findings indicate that mitotic microtubule remodeling plays a significant role in the inhibition of SOCE during mitosis. However, recent studies investigating the behavior of STIM1 and Orai1 during M-phase have provided additional insights [96,97,102–104].

STIM1 is a phosphoprotein, as revealed in large-scale mass spectrometry studies with different findings as to the specific phosphorylated residues from immunoprecipitated STIM1, presumably due to the different cell types used with lack of careful control of the cell cycle stage [105]. Smyth and colleagues reported that during mitosis of HeLa and HEK293 cells, which were treated with 1.67 μM nocodazole for 12–16 h, STIM1 fails to move to peripheral junctions to form puncta and interact with Orai1[97]. Furthermore, during mitosis, STIM1 becomes phosphorylated at multiple sites identified by mass spectrometry. Phosphorylation of STIM1 could also be detected by an anti-phospho-Ser/Thr-Pro MPM-2 antibody [106] in this study. STIM1 contains 10 minimal MPF-MAPK consensus sites (S/T-P), all located in the far C-terminus (Figure 12.3). Removal of these 10 residues by truncation at amino acid 482 abolished MPM-2 recognition of mitotic STIM1 [97]. The resulting truncated protein, when coexpressed with Orai1 in mitotic cells, partially rescues SOCE as measured by Ca2+ imaging and SOC current recording by whole-cell patch-clamp technique. In addition, alanine substitution at two residues (Ser486 and Ser668) was sufficient to partially rescue SOCE, although to a lesser extent than the 482 deletion mutant [97]. Cotransfection with an Orai1 construct was necessary in these experiments because truncated STIM1 was not able to rescue SOCE in mitotic cells unless Orai1 was coexpressed. In fact, Smyth and colleagues reported that HEK293 cells expressing the truncated STIM1 did expand at a slightly but significantly slower rate, but no other significant alterations to the mitotic process were observed [97].

Figure 12.3. The molecular domains of human STIM1.

Figure 12.3

The molecular domains of human STIM1. ER STIM1 contains a luminal and a cytosolic domain. Indicated are the locations in the sequence of the N-terminal signal peptide (SP), a Ca2+-binding canonical EF-hand domain (cEF), a non-Ca2+-binding hidden EF-hand (more...)

In a subsequent study, a STIM1 mutant retaining the full-length C-terminus, but lacking the 10 putative phosphorylation sites by alanine substitution (STIM1-10A), was able to rescue SOCE in mitotic cells expressing only endogenous Orai1 [96]. This would suggest that phosphorylation of STIM1 is the major underlying mechanism for shutting down SOCE during mitosis. When the cellular localization of STIM1-10A was examined by confocal microscopy in mitotic cells, the results were striking. STIM1-10A was incapable of dissociating from EB1 and accumulated in the spindle area. Mutation of the TRIP EB1-binding domain to TRNN rescued appropriate partitioning of STIM1 to the cell periphery. However, STIM1-10A supported SOCE in mitosis is likely not due to restoration of the EB1 interaction by STIM1-10A, because STIM1 activation of SOCE is independent of its EB1 interaction [107]. However, ER mislocalization driven by STIM-10A did not cause obvious mitotic defects. In addition, the phosphomimetic STIM1-10E mutant failed to inhibit SOCE activation in interphase cells [96].

Our group studied the mechanisms regulating SOCE inactivation during Xenopus oocyte meiosis with disparate results from what is observed in mitosis [103]. SOCE inactivation during meiosis is dependent on the kinase cascade that drives oocyte maturation, where activation of MPF was shown to be necessary and sufficient to inactivate SOCE in Xenopus oocytes [82,83]. Overexpression of human STIM1 and Orai1 by injection of in vitro transcribed mRNA into Xenopus oocytes greatly enhances SOCE, yet even this current induced by exogenous expression is inactivated during meiosis (Figure 12.4c) [103]. Associated with this inhibition of SOCE, STIM1 fails to cluster following Ca2+ store depletion (Figure 12.4b). Furthermore, meiosis was associated with inhibition of SOCE mediated by constitutively active STIM1 mutants [103], arguing that this inhibition is an active process. STIM1 is phosphorylated at multiple sites during meiosis as shown by a mobility shift on SDS-PAGE and by mass spectrometry. However, mutagenesis of all possible phosphorylation sites to alanines failed to rescue either STIM1 clustering or SOCE activation [103]. Interestingly, STIM1 clustering inhibition during meiosis required activation of MPF and was independent of the activity of the MAPK cascade, consistent with the requirement for MPF activation to inhibit endogenous SOCE during meiosis [83,103]. Our data suggest that STIM1 phosphorylation is not responsible for STIM1 clustering inhibition or SOCE inactivation during M-phase. Thus, while there are similarities between the regulation of STIM1 in meiosis and mitosis, there appear to be some differences as well. Indeed, it is likely that there is much more to be learned about the causes and consequences of STIM1 phosphorylation. More complex and sophisticated assays or models may be necessary to fully understand STIM1 phosphorylation.

Figure 12.4. Meiosis is associated with inhibition of STIM1 clustering and Orai1 internalization.

Figure 12.4

Meiosis is associated with inhibition of STIM1 clustering and Orai1 internalization. Oocyte were injected with mCherry-STIM1 and GFP-Orai1 (a) and matured into eggs (meiosis) (b). Images show the distribution of Orai1 and STIM1 before and after store (more...)

In a follow-up study, substitution of the regulatory region of STIM1, which contains the 10 MPF-MAPK putative phosphorylation sites, with GFP-rescued clustering of STIM1 into puncta during meiosis, but still did not rescue SOCE [102]. These data argue that SOCE inactivation during meiosis is not only due to inhibition of STIM1 clustering in response to store depletion. Indeed, SOCE inactivation during meiosis is also associated with the removal of Orai1 from the cell membrane into an endosomal compartment (Figure 12.4b) [103,104]. Orai1 is enriched in the cell membrane of immature oocytes (Figure 12.4a) and continuously recycles between the cell membrane and an endosomal compartment through a Rho- and Rab5-dependent pathway [104]. However, in eggs, Orai1 is internalized into an endosomal compartment, which requires the activities of dynamin and caveolin in addition to Rab5. Orai1 possesses a consensus caveolin-binding domain in its N-terminal cytoplasmic region that when mutated inhibits the ability of Orai1 to be internalized during meiosis, supporting the role of caveolin-mediated endocytosis in Orai1 internalization [104]. Whether Orai1 trafficking is regulated during mitosis in a similar fashion to meiosis remains unclear (Figure 12.5). Nonetheless, even without the regulation of STIM1 by phosphorylation, Orai1 internalization would be sufficient to inhibit SOCE during M-phase, thus bringing into question the role of STIM1 phosphorylation.

Figure 12.5. Working model of SOCE inactivation during M-phase.

Figure 12.5

Working model of SOCE inactivation during M-phase. (a) Events mediating STIM1Orai1 coupling during interphase. Orai1 has been shown to recycle between an endosomal compartment (Endo) and the cell membrane in Xenopus oocytes; however, it is not known whether (more...)

An important difference between mitosis and meiosis studies is the experimental approaches used as they may affect both the results and interpretations. Xenopus oocyte meiosis provides an important advantage in that oocytes are physiologically arrested at prophase I and eggs at metaphase II of meiosis, as discussed earlier. This removes the need for pharmacological or other experimental interventions to arrest cells in M-phase. M-phase is a transient phase that involves dramatic remodeling of multiple aspects of cellular physiology as the cell prepares to divide. Interventions such as nocodazole treatment may modify physiological processes in ways that are not always predictable. Such treatments, however, are necessary to allow cell synchronization in mitosis. Hence, future research should focus on the regulation of SOCE and STIM1 phosphorylation in mitosis under more physiological conditions, ideally in primary cells. Nonetheless, understanding SOCE inhibition during M-phase will undoubtedly provide important clues regarding the basic mechanisms controlling SOCE activation and regulation.

12.6. SOCE and Cancer

The level of Ca2+-signaling remodeling in cancer cells is remarkable and is associated with dysregulation of several Ca2+ channels and pumps [108,109]. The expression levels of essential components of SOCE, including members of the STIM and Orai families (Orai1, Orai3, STIM1, and STIM2), are modulated in several types of tumors. SOCE affects hallmarks of cancer progression, including cell cycle progression, escaping apoptosis, tumorigenesis, metastasis, angiogenesis, and tumor immunity. The differential expression of these components seems to depend on the cancer type and tumor stage (reviewed in [110–116]).

12.6.1. STIM1 and Orai1 Role in Cell Progression, Proliferation, and Cell Death of Cancer Cells

In the past few years, several reports supported a role for SOCE key players, STIM1 and Orai1, in cell cycle progression, proliferation, apoptosis, and during tumor development and progression. Sabbioni and colleagues were the first to report STIM1 (initially known as GOK) deletion in human rhabdomyosarcoma and rhabdoid tumor cell lines, RD and G401, and to show that increasing levels of STIM1 caused growth arrest in these cells [117]. Feng et al. showed that Orai1 knockdown in MCF-7 cells suppressed cell proliferation measured colorimetrically, using colony formation assays, and inhibited tumor formation in nude mice [118]. Expression of two nonfunctional Orai1 mutants L273S and R91W triggers apoptosis resistance in prostate LNCaP cells measured by the TUNEL technique and by Hoechst staining. The rescue of Orai1 function by overexpression of wild-type (WT) Orai1 increased apoptosis levels [119]. In another study, Kondratska and colleagues showed high levels of Orai1 and STIM1 expression in the pancreatic adenocarcinoma Panc1 cell line. siRNA-mediated downregulation of Orai1 and/or STIM1-intensified apoptosis-induced thermotherapy [120]. Similarly, Orai1 overexpression in A549 lung cancer cells decreased SOCE, arrested cells in G0/G1 phase, induced p21 expression, decreased ERK1/2 and Akt phosphorylation, and inhibited EGF-induced proliferation [121]. Treating MDA-MB-231 cells with SOCE inhibitor SKF96365 blocked TGFβ-induced cell proliferation and cell cycle arrest measured by colony formation assay and flow cytometry, respectively. The effect of TGF on proliferation was shown to be mediated by a decrease in STIM1 levels [122].

In cervical cancer tissues from patients with early stage cervical cancer, Chen et al. reported elevated levels of STIM1 protein compared to noncancerous tissues, as analyzed by immunoblotting. On the other hand, siRNA-mediated STIM1 knockdown in cervical cancer SiHa cells inhibits cell proliferation by arresting the cell cycle at S and G2/M phases as determined by flow cytometry of PI-stained cells. This effect of STIM1 on cell cycle progression was suggested to be due to STIM1-dependent modulation of p21 and Cdc25C involved in G2/M checkpoint progression [21].

In another study, DU145 and PC3 prostate cancer cells stably expressing STIM1 and Orai1 showed slower growth rate determined by cell growth curve measured by counting cell number over time [123]. The induction of STIM1 and Orai1 levels, and subsequently SOCE, was accompanied by an increase in the percentage of cells in the G0/G1 phase and decrease in the G2/M phase. The effect of STIM1 and Orai1 overexpression on cell cycle progression was accompanied by altered expression of cell cycle regulatory proteins, cyclin E2, cyclin D1, Wee1, and Myt1. Moreover, cells overexpressing STIM1 and/or Orai1 promoted cell senescence and induced expression of apoptosis inhibitors (DcR2, XIAP, and Bcl2). In another study, Bcl2 overexpression in human prostate cancer LNCaP cells inhibited SOCE and promoted apoptosis resistance [124,125]. Dubois and colleagues showed that heteromeric channels formed by Orai1-Orai3 are regulated by arachidonic acid, and independent of intracellular stores content, to promote cell proliferation and apoptosis resistance of prostate cancer cells. The effect on proliferation was linked to NFAT activation and expression of cyclin D1 controlling G1/S transition [126].

In mouse melanoma cells, silencing of STIM1 caused a reduction in cell growth and increased cell death [127]. Also, pharmacological inhibition of Orai1, or siRNA-mediated silencing of Orai1 and/or STIM2, caused melanoma cells to grow faster but reduced their invading potential. In a similar study, Umemura et al. showed that inhibition of SOCE by inhibitor YM58483 or siRNA knockdown of STIM1 suppresses proliferation and invasion of melanoma cells [128]. In malignant B16BL6 melanoma cells, mitochondrial Ca2+ uptake is coupled to SOCE, which promotes PKB activity favoring cell survival [129]. In a recent study, Hooper and colleagues reported very small SOCE in invasive melanoma depending on PKC-mediated phosphorylation of Orai1 [130]. Collectively, these studies show a correlation between SOCE levels and cancer cell proliferation and the ability to resist apoptosis.

12.6.2. SOCE in Cell Motility, Metastasis, and Tumor Microenvironment

Yang et al. reported that serum-induced migration of human MDA-MB-231 breast cancer cells with STIM1 or Orai1 levels knocked down by siRNA was decreased by 60%–85%, measured by Boyden chamber assay without affecting cell proliferation [100]. The results from these in vitro studies were replicated in mouse models where tumor cells expressing luciferase reporter gene were injected into immune-deficient mice through the tail vein [131,132]. The metastasis of injected cells with STIM1 and Orai1 siRNA to the lungs was much less than the control siRNA-treated cells.

Analysis of cancer tissues from clear-cell renal-cell carcinoma (ccRCC) showed increased expression of Orai1 and STIM1 protein levels in comparison to adjacent normal tissues [133]. Pharmacologic inhibition of SOCE using SKF96365 or 2-APB commonly used blockers, and siRNA-mediated knockdown of Orai1 or STIM1, decreased the motility of ccRCC cell lines, Caki1 and ACHN, independent of proliferation, as measured by wound-healing assay in the presence of the antiproliferative drug Mitomycin C.

In cervical cancer tissues from patients with early stage cervical cancer, the levels of STIM1 were positively correlated with tumor size and metastasis to the lymph nodes [21]. Similarly, injection of cervical cancer cells selected for high STIM1 expression significantly enhanced tumor growth, local spread, and angiogenesis, whereas cells with short hairpin RNAs (shRNAs) targeting human STIM1 showed a significant decrease in tumor growth and angiogenesis.

In human prostate cancer tissues, STIM1 and Orai1 were expressed at significantly lower levels in hyperplasia and tumor tissues at advanced stages. However, the expression of STIM1 was higher in tumors with earlier histological grade than in hyperplasia tissues, suggesting that STIM1 can play a dual role depending on the cancer stage, where it favors malignant transformation of prostate cells at early stages and prohibit tumor growth at advanced stages [123]. The expression levels of STIM1 and Orai1 in the hyperplasic human prostate cell line BPH-1 were lower as compared to the metastatic LNCaP, DU145, and PC3 cell lines with a significantly greater SOCE. Overexpression of Orai1 and STIM1 in prostate cancer cell lines DU145 and PC3 promoted cell migration as tested using the wound-healing and Transwell assays. Furthermore, in vivo, immunodeficient mice (SCID) injected with DU145 cells expressing STIM1-YFP or Orai1-YFP showed retardation in tumor growth and decrease in expression of E-cadherin, demonstrating that STIM1 enhances epithelial-mesenchymal transition (EMT).

Colorectal cancer tissue collected from patients showed high expression of STIM1, with significant correlation between STIM1 levels, tumor growth, invasion depth, and metastasis to lymph nodes. STIM1 knockdown by 50%–90% using shRNA or SOCE inhibition using 2-APB and SKF96365 in three colon cancer cell lines (DLD-1, HCT116, and SW480) strongly inhibited cell motility, and this depends on COX-2 expression and prostaglandin synthesis [134].

Studies by Trebak and colleagues showed that in breast cancer MCF7 cells, SOCE is mediated by STIM1 and Orai3, rather than the ubiquitous STIM1-Orai1 pair [135,136]. Induction of estrogen-receptor-positive breast cancer MCF7 cells using breast growth factors activated Orai3-dependent Ca2+ influx [137]. Orai3 knockdown resulted in a significant decrease in tumor cell invasion measured using Boyden chamber invasion. In addition, Orai3 knockdown with shRNA-encoding lentiviruses significantly reduced the number of MCF7 colonies on agar. In primary glioblastoma cell lines, STIM1 and Orai1 are the key components of SOCE. Interestingly, using siRNA approaches to knockdown Orai3 in MCF7, or STIM1 and Orai1 in glioblastoma cells showed strong reduction in Matrigel invasion of these cells compared to nonmalignant primary astrocytes [138].

Collectively, these studies strongly argue for a ubiquitous role for Ca2+ influx through SOCE in mediating tumor cell metastasis in different cancers. This could be due to the role of SOCE in regulating the cytoskeleton in a polarized fashion in migrating cells, which is important for cell motility [139]. Given that over 90% of breast cancer deaths are associated with metastasis rather than growth of the primary tumor, SOCE may represent an attractive anticancer therapeutic target.

12.6.3. Orai1 and STIM1 in Antitumor Immunity

In addition to the role of SOCE in regulation metastasis, Orai1 and STIM1 have been implicated in regulating immune function in the context of its antitumor activity. The study by Xu et al. using Transwell migration assay reported higher recruitment of human leukemic monocyte macrophage cell line U937 when using culture supernatant from STIM1-knockdown DU145 and PC3 cells and decreased migration of U937 when using supernatant from DU145 and PC3 cells overexpressing STIM1 [123]. Real-time RT-PCR showed decreased expression of cytokines in U937 cells incubated with medium from DU145 and PC3 cells overexpressing STIM1 and/Orai1. In addition, the markers for EMT transition, VEGFA and MMP9 decreased in U937 macrophages, suggesting that STIM1 and Orai1 overexpression, and hence enhanced SOCE activity in prostate cancer cells, hinders EMT in macrophages and their recruitment to tumor sites.

Using Stim1fl/flStim2fl/fl Cd4Cre (DKO) mice with CD4+ and CD8+ T cells lacking SOCE [140], Weidinger et al. examined injecting B16-OVA melanoma cells and MC-38 colon carcinoma cells in DKO and WT mice with depleted immunosuppressive Treg cells. DKO mice failed to control tumor growth suggesting that SOCE in CD8+ T cells mediated by the STIM family favors antitumor immunity [141]. The loss of SOCE in DKO did not compromise the priming or migration of tumor antigen-specific CD8+ T cells, but it inhibited the ability of CTLs to control tumor engraftment and growth. The cytotoxicity of SOCE-deficient CTLs from DKO mice against cocultured EG7-OVA cells was diminished, and the ability of these cells to produce IFN-γ and TNF-α upon stimulation was also reduced, compared to wild-type CTLs.

Although several studies investigating the role of SOCE in cancer have been carried out over the past few years, much remains to be elucidated regarding the mechanisms by which SOCE affects cancer development. This complexity is due to the fact that SOCE regulation is likely to be cancer type, as well as stage specific. Nonetheless, accumulating evidence suggests that SOCE plays a critical role in cancer cell proliferation, metastasis, and antitumor immunity.

References

1.
Berridge, M. J., Lipp, P., and Bootman, M. D. (2000) The versatility and universality of calcium signalling. Nature Reviews. Molecular Cell Biology 1, 11-21. [PubMed: 11413485]
2.
Demaurex, N. and Frieden, M. (2003) Measurements of the free luminal ER Ca2+ concentration with targeted “cameleon” fluorescent proteins. Cell Calcium 34, 109-119. [PubMed: 12810053]
3.
Berridge, M. J. (2012) Calcium signalling remodelling and disease. Biochemical Society Transactions 40, 297-309. [PubMed: 22435804]
4.
Clapham, D. E. (1995) Calcium signaling. Cell 80, 259-268. [PubMed: 7834745]
5.
Berridge, M. J., Bootman, M. D., and Roderick, H. L. (2003) Calcium signalling: Dynamics, homeostasis and remodelling. Nature Reviews. Molecular Cell Biology 4, 517-529. [PubMed: 12838335]
6.
Clapham, D. E. (2007) Calcium signaling. Cell 131, 1047-1058. [PubMed: 18083096]
7.
Wu, M. M., Buchanan, J., Luik, R. M., and Lewis, R. S. (2006) Ca2+ store depletion causes STIM1 to accumulate in ER regions closely associated with the plasma membrane. Journal of Cell Biology 174, 803-813. [PMC free article: PMC2064335] [PubMed: 16966422]
8.
Hogan, P. G., Lewis, R. S., and Rao, A. (2010) Molecular basis of calcium signaling in lymphocytes: STIM and ORAI. Annual Review of Immunology 28, 491-533. [PMC free article: PMC2861828] [PubMed: 20307213]
9.
Hogan, P. G. (2015) The STIM1-ORAI1 microdomain. Cell Calcium 58, 357-367. [PMC free article: PMC4564343] [PubMed: 26215475]
10.
Courjaret, R. and Machaca, K. (2014) Mid-range Ca2+ signalling mediated by functional coupling between store-operated Ca2+ entry and IP3-dependent Ca2+ release. Nature Communications 5, 3916. [PubMed: 24867608]
11.
Arredouani, A., Yu, F., Sun, L., and Machaca, K. (2010) Regulation of store-operated Ca2+ entry during the cell cycle. Journal of Cell Science 123, 2155-2162. [PMC free article: PMC2886739] [PubMed: 20554894]
12.
Sun, L. and Machaca, K. (2004) Ca2+ cyt negatively regulates the initiation of oocyte maturation. Journal of Cell Biology 165, 63-75. [PMC free article: PMC1289150] [PubMed: 15067021]
13.
Sun, L., Hodeify, R., Haun, S., Charlesworth, A., MacNicol, A. M., Ponnappan, S., Ponnappan, U., Prigent, C., and Machaca, K. (2008) Ca2+ homeostasis regulates Xenopus oocyte maturation. Biology of Reproduction 78, 726-735. [PMC free article: PMC2587222] [PubMed: 18094360]
14.
Tombes, R. M., Simerly, C., Borisy, G. G., and Schatten, G. (1992) Meiosis, egg activation, and nuclear envelope breakdown are differentially reliant on Ca2+, whereas germinal vesicle breakdown is Ca2+ independent in the mouse oocyte. Journal of Cell Biology 117, 799-811. [PMC free article: PMC2289470] [PubMed: 1577859]
15.
Machaca, K. (2007) Ca2+ signaling differentiation during oocyte maturation. Journal of Cellular Physiology 213, 331-340. [PubMed: 17620315]
16.
Capiod, T. (2013) The need for calcium channels in cell proliferation. Recent Patents on Anti-Cancer Drug Discovery 8, 4-17. [PubMed: 22519598]
17.
Nel, A. E., Dirienzo, W., Stefanini, G. F., Wooten, M. W., Canonica, G. W., Lattanze, G. R., Stevenson, H. C., Miller, P., Fudenberg, H. H., and Galbraith, R. M. (1986) Inhibition of T3 mediated T-cell proliferation by Ca2+-channel blockers and inhibitors of Ca2+/phospholipid-dependent kinase. Scandinavian Journal of Immunology 24, 283-290. [PubMed: 3489285]
18.
Taylor, J. T., Huang, L., Pottle, J. E., Liu, K., Yang, Y., Zeng, X., Keyser, B. M., Agrawal, K. C., Hansen, J. B., and Li, M. (2008) Selective blockade of T-type Ca2+ channels suppresses human breast cancer cell proliferation. Cancer Letters 267, 116-124. [PubMed: 18455293]
19.
Aguiari, G., Trimi, V., Bogo, M., Mangolini, A., Szabadkai, G., Pinton, P., Witzgall, R., et al. (2008) Novel role for polycystin-1 in modulating cell proliferation through calcium oscillations in kidney cells. Cell Proliferation 41, 554-573. [PMC free article: PMC2440503] [PubMed: 18422703]
20.
Resende, R. R., Adhikari, A., da Costa, J. L., Lorencon, E., Ladeira, M. S., Guatimosim, S., Kihara, A. H., and Ladeira, L. O. (2010) Influence of spontaneous calcium events on cell-cycle progression in embryonal carcinoma and adult stem cells. Biochimica et Biophysica Acta 1803, 246-260. [PubMed: 19958796]
21.
Chen, Y. F., Chiu, W. T., Chen, Y. T., Lin, P. Y., Huang, H. J., Chou, C. Y., Chang, H. C., Tang, M. J., and Shen, M. R. (2011) 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 108, 15225-15230. [PMC free article: PMC3174613] [PubMed: 21876174]
22.
Stanimirovic, D. B., Ball, R., Mealing, G., Morley, P., and Durkin, J. P. (1995) The role of intracellular calcium and protein kinase C in endothelin-stimulated proliferation of rat type I astrocytes. Glia 15, 119-130. [PubMed: 8567063]
23.
Shawl, A. I., Park, K. H., and Kim, U. H. (2009) Insulin receptor signaling for the proliferation of pancreatic beta-cells: Involvement of Ca2+ second messengers, IP3, NAADP and cADPR. Islets 1, 216-223. [PubMed: 21099275]
24.
Cheng, D., Zhu, X., Barchiesi, F., Gillespie, D. G., Dubey, R. K., and Jackson, E. K. (2011) Receptor for activated protein kinase C1 regulates cell proliferation by modulating calcium signaling. Hypertension 58, 689-695. [PMC free article: PMC3174333] [PubMed: 21844488]
25.
Forostyak, O., Forostyak, S., Kortus, S., Sykova, E., Verkhratsky, A., and Dayanithi, G. (2016) Physiology of Ca2+ signalling in stem cells of different origins and differentiation stages. Cell Calcium 59, 57-66. [PubMed: 26905828]
26.
Szatkowski, C., Parys, J. B., Ouadid-Ahidouch, H., and Matifat, F. (2010) Inositol 1,4,5-trisphosphate-induced Ca2+ signalling is involved in estradiol-induced breast cancer epithelial cell growth. Molecular Cancer 9, 156. [PMC free article: PMC2906470] [PubMed: 20565939]
27.
Cardenas, C., Muller, M., McNeal, A., Lovy, A., Jana, F., Bustos, G., Urra, F., et al. (2016) Selective vulnerability of cancer cells by inhibition of Ca transfer from endoplasmic reticulum to mitochondria. Cell Reports 15, 219-220. [PubMed: 27050774]
28.
Rasmussen, C. D. and Means, A. R. (1989) Calmodulin is required for cell-cycle progression during G1 and mitosis. EMBO Journal 8, 73-82. [PMC free article: PMC400774] [PubMed: 2469574]
29.
Hidaka, H., Sasaki, Y., Tanaka, T., Endo, T., Ohno, S., Fujii, Y., and Nagata, T. (1981) N-(6-aminohexyl)-5-chloro-1-naphthalenesulfonamide, a calmodulin antagonist, inhibits cell proliferation. Proceedings of the National Academy of Sciences of the United States of America 78, 4354-4357. [PMC free article: PMC319788] [PubMed: 6945588]
30.
Kahl, C. R. and Means, A. (2003) Regulation of cell cycle progression by calcium/calmodulin-dependent pathways. Endocrine Reviews 24, 719-736. [PubMed: 14671000]
31.
Skelding, K. A., Rostas, J. A., and Verrills, N. M. (2011) Controlling the cell cycle: The role of calcium/calmodulin-stimulated protein kinases I and II. Cell Cycle 10, 631-639. [PubMed: 21301225]
32.
Courjaret, R. and Machaca, K. (2012) STIM and Orai in cellular proliferation and division. Frontiers in Bioscience 4, 331-341. [PubMed: 22201875]
33.
Pinto, M. C., Kihara, A. H., Goulart, V. A., Tonelli, F. M., Gomes, K. N., Ulrich, H., and Resende, R. R. (2015) Calcium signaling and cell proliferation. Cellular Signalling 27, 2139-2149. [PubMed: 26275497]
34.
Shaw, P. J. and Feske, S. (2012) Regulation of lymphocyte function by ORAI and STIM proteins in infection and autoimmunity. The Journal of Physiology 590, 4157-4167. [PMC free article: PMC3473275] [PubMed: 22615435]
35.
Dziegielewska, B., Gray, L. S., and Dziegielewski, J. (2014) T-type calcium channels blockers as new tools in cancer therapies. Pflügers Archiv 466, 801-810. [PubMed: 24449277]
36.
Somasundaram, A., Shum, A. K., McBride, H. J., Kessler, J. A., Feske, S., Miller, R. J., and Prakriya, M. (2014) Store-operated CRAC channels regulate gene expression and proliferation in neural progenitor cells. The Journal of Neuroscience 34, 9107-9123. [PMC free article: PMC4078087] [PubMed: 24990931]
37.
Li, M., Chen, C., Zhou, Z., Xu, S., and Yu, Z. (2012) A TRPC1-mediated increase in store-operated Ca2+ entry is required for the proliferation of adult hippocampal neural progenitor cells. Cell Calcium 51, 486-496. [PubMed: 22579301]
38.
Hu, F., Pan, L., Zhang, K., Xing, F., Wang, X., Lee, I., Zhang, X., and Xu, J. (2014) Elevation of extracellular Ca2+ induces store-operated calcium entry via calcium-sensing receptors: A pathway contributes to the proliferation of osteoblasts. PLoS One 9, e107217. [PMC free article: PMC4177836] [PubMed: 25254954]
39.
Madsen, C. P., Klausen, T. K., Fabian, A., Hansen, B. J., Pedersen, S. F., and Hoffmann, E. K. (2012) On the role of TRPC1 in control of Ca2+ influx, cell volume, and cell cycle. American Journal of Physiology. Cell Physiology 303, C625-C634. [PubMed: 22744003]
40.
Kito, H., Yamamura, H., Suzuki, Y., Yamamura, H., Ohya, S., Asai, K., and Imaizumi, Y. (2015) Regulation of store-operated Ca2+ entry activity by cell cycle dependent up-regulation of Orai2 in brain capillary endothelial cells. Biochemical and Biophysical Research Communications 459, 457-462. [PubMed: 25748572]
41.
Lodola, F., Laforenza, U., Bonetti, E., Lim, D., Dragoni, S., Bottino, C., Ong, H. L., et al. (2012) Store-operated Ca2+ entry is remodelled and controls in vitro angiogenesis in endothelial progenitor cells isolated from tumoral patients. PLoS One 7, e42541. [PMC free article: PMC3458053] [PubMed: 23049731]
42.
Smith, L. D. (1989) The induction of oocyte maturation: Transmembrane signaling events and regulation of the cell cycle. Development 107, 685-699. [PubMed: 2698799]
43.
Miyazaki, S. (1995) Calcium signalling during mammalian fertilization. Ciba Foundation Symposium 188, 235-251. [PubMed: 7587620]
44.
Antoine, A. F., Faure, J. E., Cordeiro, S., Dumas, C., Rougier, M., and Feijo, J. A. (2000) A calcium influx is triggered and propagates in the zygote as a wavefront during in vitro fertilization of flowering plants. Proceedings of the National Academy of Sciences of the United States of America 97, 10643-10648. [PMC free article: PMC27078] [PubMed: 10973479]
45.
Miyazaki, S., Shirakawa, H., Nakada, K., and Honda, Y. (1993) Essential role of the inositol 1,4,5-trisphosphate receptor/Ca2+ release channel in Ca2+ waves and Ca2+ oscillations at fertilization of mammalian eggs. Developmental Biology 158, 62-78. [PubMed: 8392472]
46.
Runft, L. L., Jaffe, L. A., and Mehlmann, L. (2002) Egg activation at fertilization: Where it all begins. Developmental Biology 245, 237-254. [PubMed: 11977978]
47.
Stricker, S. A. (1999) Comparative biology of calcium signaling during fertilization and egg activation in animals. Developmental Biology 211, 157-176. [PubMed: 10395780]
48.
Busa, W. B., Ferguson, J. E., Joseph, S. K., Williamson, J. R., and Nuccitelli, R. (1985) Activation of frog (Xenopus laevis) eggs by inositol trisphosphate. I. Characterization of Ca2+ release from intracellular stores. Journal of Cell Biology 101, 677-682. [PMC free article: PMC2113661] [PubMed: 3874873]
49.
Fontanilla, R. A. and Nuccitelli, R. (1998) Characterization of the sperm-induced calcium wave in Xenopus eggs using confocal microscopy. Biophysical Journal 75, 2079-2087. [PMC free article: PMC1299880] [PubMed: 9746550]
50.
Kline, D. and Nuccitelli, R. (1985) The wave of activation current in the Xenopus egg. Developmental Biology 111, 471-487. [PubMed: 3840102]
51.
Nuccitelli, R., Yim, D. L., and Smart, T. (1993) The sperm-induced Ca2+ wave following fertilization of the Xenopus egg requires the production of Ins(1, 4, 5)P3. Developmental Biology 158, 200-212. [PubMed: 7687224]
52.
Lechleiter, J. D. and Clapham, D. E. (1992) Molecular mechanisms of intracellular calcium excitability in X. laevis oocytes. Cell 69, 283-294. [PubMed: 1568248]
53.
Callamaras, N., Marchant, J. S., Sun, X. P., and Parker, I. (1998) Activation and co-ordination of InsP3-mediated elementary Ca2+ events during global Ca2+ signals in Xenopus oocytes. The Journal of Physiology 509(Pt 1), 81-91. [PMC free article: PMC2230929] [PubMed: 9547383]
54.
Machaca, K. (2004) Increased sensitivity and clustering of elementary Ca2+ release events during oocyte maturation. Developmental Biology 275, 170-182. [PubMed: 15464580]
55.
Parys, J. B. and Bezprozvanny, I. (1995) The inositol trisphosphate receptor of Xenopus oocytes. Cell Calcium 18, 353-363. [PubMed: 8581964]
56.
Igusa, Y. and Miyazaki, S. (1983) Effects of altered extracellular and intracellular calcium concentration on hyperpolarizing responses of the hamster egg. Journal of Physiology 340, 611-632. [PMC free article: PMC1199230] [PubMed: 6887062]
57.
Kline, D. and Kline, J. T. (1992) Thapsigargin activates a calcium influx pathway in the unfertilized mouse egg and suppresses repetitive calcium transients in the fertilized egg. Journal of Biological Chemistry 267, 17624-17630. [PubMed: 1387638]
58.
Mohri, T., Shirakawa, H., Oda, S., Sato, M. S., Mikoshiba, K., and Miyazaki, S. (2001) Analysis of Mn2+/Ca2+ influx and release during Ca2+ oscillations in mouse eggs injected with sperm extract. Cell Calcium 29, 311-325. [PubMed: 11292388]
59.
Sato, K., Iwao, Y., Fujimura, T., Tamaki, I., Ogawa, K., Iwasaki, T., Tokmakov, A. A., Hatano, O., and Fukami, Y. (1999) Evidence for the involvement of a Src-related tyrosine kinase in Xenopus egg activation. Developmental Biology 209, 308-320. [PubMed: 10328923]
60.
Sato, K., Tokmakov, A. A., Iwasaki, T., and Fukami, Y. (2000) Tyrosine kinase-dependent activation of phospholipase Cgamma is required for calcium transient in Xenopus egg fertilization. Developmental Biology 224, 453-469. [PubMed: 10926780]
61.
Tokmakov, A. A., Sato, K. I., Iwasaki, T., and Fukami, Y. (2002) Src kinase induces calcium release in Xenopus egg extracts via PLCgamma and IP3-dependent mechanism. Cell Calcium 32, 11-20. [PubMed: 12127058]
62.
Saunders, C. M., Larman, M. G., Parrington, J., Cox, L. J., Royse, J., Blayney, L. M., Swann, K., and Lai, F. A. (2002) PLCzeta: A sperm-specific trigger of Ca2+ oscillations in eggs and embryo development. Development 129, 3533-3544. [PubMed: 12117804]
63.
Chiba, K., Kado, R. T., and Jaffe, L. A. (1990) Development of calcium release mechanisms during starfish oocyte maturation. Developmental Biology 140, 300-306. [PubMed: 2373255]
64.
Mehlmann, L. and Kline, D. (1994) Regulation of intracellular calcium in the mouse egg: Calcium release in response to sperm or inositol trisphosphate is enhanced after meiotic maturation. Biology of Reproduction 51, 1088-1098. [PubMed: 7888488]
65.
Fujiwara, T., Nakada, K., Shirakawa, H., and Miyazaki, S. (1993) Development of inositol trisphosphate-induced calcium release mechanism during maturation of hamster oocytes. Developmental Biology 156, 69-79. [PubMed: 8383620]
66.
Jones, K. T., Carroll, J., and Whittingham, D. G. (1994) Ionomycin, thapsigargin, ryanodine, and sperm induced Ca2+ release increase during meiotic maturation of mouse oocytes. Journal of Biological Chemistry 270, 6671-6677. [PubMed: 7896808]
67.
Terasaki, M., Runft, L. L., and Hand, A. R. (2001) Changes in organization of the endoplasmic reticulum during Xenopus oocyte maturation and activation. Molecular Biology of the Cell 12, 1103-1116. [PMC free article: PMC32290] [PubMed: 11294910]
68.
Parker, I., Choi, J., and Yao, Y. (1996) Elementary events of InsP 3-induced Ca2+ liberation in Xenopus oocytes: Hot spots, puffs and blips. Cell Calcium 20, 105-121. [PubMed: 8889202]
69.
Parys, J. B., McPherson, S. M., Mathews, L., Campbell, K. P., and Longo, F. J. (1994) Presence of inositol 1,4,5-trisphosphate receptor, calreticulin, and calsequestrin in eggs of sea urchins and Xenopus laevis. Developmental Biology 161, 466-476. [PubMed: 8313995]
70.
Kume, S., Yamamoto, A., Inoue, T., Muto, A., Okano, H., and Mikoshiba, K. (1997) Developmental expression of the inositol 1,4,5-trisphosphate receptor and structural changes in the endoplasmic reticulum during oogenesis and meiotic maturation of Xenopus laevis. Developmental Biology 182, 228-239. [PubMed: 9070324]
71.
El Jouni, W., Jang, B., Haun, S., and Machaca, K. (2005) Calcium signaling differentiation during Xenopus oocyte maturation. Developmental Biology 288, 514-525. [PubMed: 16330019]
72.
Sun, L., Yu, F., Ullah, A., Hubrack, S., Daalis, A., Jung, P., and Machaca, K. (2011) Endoplasmic reticulum remodeling tunes IP3-dependent Ca2+ release sensitivity. PLoS One 6, e27928. [PMC free article: PMC3227640] [PubMed: 22140486]
73.
Malathi, K., Kohyama, S., Ho, M., Soghoian, D., Li, X., Silane, M., Berenstein, A., and Jayaraman, T. (2003) Inositol 1,4,5-trisphosphate receptor (type 1) phosphorylation and modulation by Cdc2. Journal of Cellular Biochemistry 90, 1186-1196. [PubMed: 14635192]
74.
Li, X., Malathi, K., Krizanova, O., Ondrias, K., Sperber, K., Ablamunits, V., and Jayaraman, T. (2005) Cdc2/cyclin B1 interacts with and modulates inositol 1,4,5-trisphosphate receptor (type 1) functions. Journal of Immunology 175, 6205-6210. [PubMed: 16237118]
75.
Sun, L., Haun, S., Jones, R. C., Edmondson, R. D., and Machaca, K. (2009) Kinase-dependent regulation of IP3-dependent Ca2+ release during oocyte maturation. Journal of Biological Chemistry 284, 20184-20196. [PMC free article: PMC2740445] [PubMed: 19473987]
76.
Lee, B., Vermassen, E., Yoon, S. Y., Vanderheyden, V., Ito, J., Alfandari, D., De Smedt, H., Parys, J. B., and Fissore, R. A. (2006) Phosphorylation of IP3R1 and the regulation of [Ca2+]i responses at fertilization: A role for the MAP kinase pathway. Development 133, 4355-4365. [PMC free article: PMC2909192] [PubMed: 17038520]
77.
Boulware, M. J. and Marchant, J. S. (2005) IP3 receptor activity is differentially regulated in endoplasmic reticulum subdomains during oocyte maturation. Current Biology 15, 765-770. [PubMed: 15854911]
78.
Boulware, M. J. and Marchant, J. S. (2008) Nuclear pore disassembly from endoplasmic reticulum membranes promotes Ca2+ signalling competency. The Journal of Physiology 586, 2873-2888. [PMC free article: PMC2517208] [PubMed: 18450775]
79.
Guerini, D., Coletto, L., and Carafoli, E. (2005) Exporting calcium from cells. Cell Calcium 38, 281-289. [PubMed: 16102821]
80.
Strehler, E. E., Filoteo, A. G., Penniston, J. T., and Caride, A. J. (2007) Plasma-membrane Ca2+ pumps: Structural diversity as the basis for functional versatility. Biochemical Society Transactions 35, 919-922. [PMC free article: PMC2276580] [PubMed: 17956246]
81.
Strehler, E. E. and Treiman, M. (2004) Calcium pumps of plasma membrane and cell interior. Current Molecular Medicine 4, 323-335. [PubMed: 15101689]
82.
Machaca, K. and Haun, S. (2000) Store-operated calcium entry inactivates at the germinal vesicle breakdown stage of Xenopus meiosis. Journal of Biological Chemistry 275, 38710-38715. [PMC free article: PMC1201341] [PubMed: 10991950]
83.
Machaca, K. and Haun, S. (2002) Induction of maturation-promoting factor during Xenopus oocyte maturation uncouples Ca2+ store depletion from store-operated Ca2+ entry. Journal of Cell Biology 156, 75-85. [PMC free article: PMC1307503] [PubMed: 11781335]
84.
Hartzell, H. C. (1996) Activation of different Cl currents in Xenopus oocytes by Ca liberated from stores and by capacitative Ca influx. Journal of General Physiology 108, 157-175. [PMC free article: PMC2229319] [PubMed: 8882861]
85.
Yao, Y. and Tsien, R. Y. (1997) Calcium current activated by depletion of calcium stores in Xenopus oocytes. Journal of General Physiology 109, 703-715. [PMC free article: PMC2217046] [PubMed: 9222897]
86.
Busa, W. B. and Nuccitelli, R. (1985) An elevated free cytosolic Ca2+ wave follows fertilization in eggs of the frog, Xenopus laevis. Journal of Cell Biology 100, 1325-1329. [PMC free article: PMC2113751] [PubMed: 3980584]
87.
Liu, J. and Maller, J. L. (2005) Calcium elevation at fertilization coordinates phosphorylation of XErp1/Emi2 by Plx1 and CaMK II to release metaphase arrest by cytostatic factor. Current Biology 15, 1458-1468. [PubMed: 16040245]
88.
Mochida, S. and Hunt, T. (2007) Calcineurin is required to release Xenopus egg extracts from meiotic M phase. Nature 449, 336-340. [PubMed: 17882219]
89.
Martin-Romero, F. J., Lopez-Guerrero, A. M., Alvarez, I. S., and Pozo-Guisado, E. (2012) Role of store-operated calcium entry during meiotic progression and fertilization of mammalian oocytes. International Review of Cell and Molecular Biology 295, 291-328. [PubMed: 22449493]
90.
Gomez-Fernandez, C., Lopez-Guerrero, A. M., Pozo-Guisado, E., Alvarez, I. S., and Martin-Romero, F. J. (2012) Calcium signaling in mouse oocyte maturation: The roles of STIM1, ORAI1 and SOCE. Molecular Human Reproduction 18, 194-203. [PubMed: 22053056]
91.
Wang, C., Lee, K., Gajdocsi, E., Papp, A. B., and Machaty, Z. (2012) Orai1 mediates store-operated Ca2+ entry during fertilization in mammalian oocytes. Developmental Biology 365, 414-423. [PubMed: 22445508]
92.
Lee, B., Palermo, G., and Machaca, K. (2013) Downregulation of store-operated Ca2+ entry during mammalian meiosis is required for the egg-to-embryo transition. Journal of Cell Science 126, 1672-1681. [PubMed: 23424198]
93.
Cheon, B., Lee, H. C., Wakai, T., and Fissore, R. A. (2013) Ca2+ influx and the store-operated Ca2+ entry pathway undergo regulation during mouse oocyte maturation. Molecular Biology of the Cell 24, 1396-1410. [PMC free article: PMC3639051] [PubMed: 23468522]
94.
Volpi, M. and Berlin, R. D. (1988) Intracellular elevations of free calcium induced by activation of histamine H1 receptors in interphase and mitotic HeLa cells: Hormone signal transduction is altered during mitosis. Journal of Cell Biology 107, 2533-2539. [PMC free article: PMC2115666] [PubMed: 3204119]
95.
Preston, S. F., Sha'afi, R. I., and Berlin, R. D. (1991) Regulation of Ca2+ influx during mitosis: Ca2+ influx and depletion of intracellular Ca2+ stores are coupled in interphase but not mitosis. Cell Regulation 2, 915-925. [PMC free article: PMC361890] [PubMed: 1809398]
96.
Smyth, J. T., Beg, A. M., Wu, S., Putney, J. W., Jr., and Rusan, N. M. (2012) Phosphoregulation of STIM1 leads to exclusion of the endoplasmic reticulum from the mitotic spindle. Current Biology 22, 1487-1493. [PMC free article: PMC3427412] [PubMed: 22748319]
97.
Smyth, J. T., Petranka, J. G., Boyles, R. R., DeHaven, W. I., Fukushima, M., Johnson, K. L., Williams, J. G., and Putney, J. W., Jr. (2009) Phosphorylation of STIM1 underlies suppression of store-operated calcium entry during mitosis. Nature Cell Biology 11, 1465-1472. [PMC free article: PMC3552519] [PubMed: 19881501]
98.
Tani, D., Monteilh-Zoller, M. K., Fleig, A., and Penner, R. (2007) Cell cycle-dependent regulation of store-operated ICRAC and Mg2+-nucleotide-regulated MagNuM (TRPM7) currents. Cell Calcium 41, 249-260. [PMC free article: PMC5663638] [PubMed: 17064762]
99.
Chen, Y. W., Chen, Y. F., Chen, Y. T., Chiu, W. T., and Shen, M. R. (2016) The STIM1-Orai1 pathway of store-operated Ca2+ entry controls the checkpoint in cell cycle G1/S transition. Scientific Reports 6, 22142. [PMC free article: PMC4768259] [PubMed: 26917047]
100.
Yang, S., Zhang, J. J., and Huang, X. Y. (2009) Orai1 and STIM1 are critical for breast tumor cell migration and metastasis. Cancer Cell 15, 124-134. [PubMed: 19185847]
101.
Russa, A. D., Ishikita, N., Masu, K., Akutsu, H., Saino, T., and Satoh, Y. (2008) Microtubule remodeling mediates the inhibition of store-operated calcium entry (SOCE) during mitosis in COS-7 cells. Archives of Histology and Cytology 71, 249-263. [PubMed: 19359807]
102.
Yu, F., Sun, L., Courjaret, R., and Machaca, K. (2011) Role of the STIM1 C-terminal domain in STIM1 clustering. Journal of Biological Chemistry 286, 8375-8384. [PMC free article: PMC3048722] [PubMed: 21220431]
103.
Yu, F., Sun, L., and Machaca, K. (2009) Orai1 internalization and STIM1 clustering inhibition modulate SOCE inactivation during meiosis. Proceedings of the National Academy of Sciences of the United States of America 106, 17401-17406. [PMC free article: PMC2765092] [PubMed: 19805124]
104.
Yu, F., Sun, L., and Machaca, K. (2010) Constitutive recycling of the store-operated Ca2+ channel Orai1 and its internalization during meiosis. Journal of Cell Biology 191, 523-535. [PMC free article: PMC3003315] [PubMed: 21041445]
105.
Pozo-Guisado, E. and Martin-Romero, F. J. (2013) The regulation of STIM1 by phosphorylation. Communicative & Integrative Biology 6, e26283. [PMC free article: PMC3914909] [PubMed: 24505502]
106.
Westendorf, J. M., Rao, P. N., and Gerace, L. (1994) Cloning of cDNAs for M-phase phosphoproteins recognized by the MPM2 monoclonal antibody and determination of the phosphorylated epitope. Proceedings of the National Academy of Sciences of the United States of America 91, 714-718. [PMC free article: PMC43019] [PubMed: 8290587]
107.
Grigoriev, I., Gouveia, S. M., van, D. V., Demmers, J., Smyth, J. T., Honnappa, S., Splinter, D., et al. (2008) STIM1 is a MT-plus-end-tracking protein involved in remodeling of the ER. Current Biology 18, 177-182. [PMC free article: PMC2600655] [PubMed: 18249114]
108.
Monteith, G. R., Davis, F. M., and Roberts-Thomson, S. J. (2012) Calcium channels and pumps in cancer: Changes and consequences. The Journal of Biological Chemistry 287, 31666-31673. [PMC free article: PMC3442501] [PubMed: 22822055]
109.
Prevarskaya, N., Ouadid-Ahidouch, H., Skryma, R., and Shuba, Y. (2014) Remodelling of Ca2+ transport in cancer: How it contributes to cancer hallmarks?Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 369, 20130097. [PMC free article: PMC3917351] [PubMed: 24493745]
110.
Xie, J., Pan, H., Yao, J., Zhou, Y., and Han, W. (2016) SOCE and cancer: Recent progress and new perspectives. International Journal of Cancer 138, 2067-2077. [PMC free article: PMC4764496] [PubMed: 26355642]
111.
Wong, H. S. and Chang, W. C. (2015) Correlation of clinical features and genetic profiles of stromal interaction molecule 1 (STIM1) in colorectal cancers. Oncotarget 6, 42169-42182. [PMC free article: PMC4747217] [PubMed: 26543234]
112.
Stanisz, H., Vultur, A., Herlyn, M., Roesch, A., and Bogeski, I. (2016) The role of Orai/STIM calcium channels in melanocytes and melanoma. Journal of Physiology 594, 2825-2835. [PMC free article: PMC4887671] [PubMed: 26864956]
113.
Vashisht, A., Trebak, M., and Motiani, R. K. (2015) STIM and Orai proteins as novel targets for cancer therapy. A review in the theme: Cell and molecular processes in cancer metastasis. American Journal of Physiology. Cell Physiology 309, C457-C469. [PMC free article: PMC4593768] [PubMed: 26017146]
114.
Vanden Abeele, F., Shuba, Y., Roudbaraki, M., Lemonnier, L., Vanoverberghe, K., Mariot, P., Skryma, R., and Prevarskaya, N. (2003) Store-operated Ca2+ channels in prostate cancer epithelial cells: Function, regulation, and role in carcinogenesis. Cell Calcium 33, 357-373. [PubMed: 12765682]
115.
Fiorio Pla, A., Kondratska, K., and Prevarskaya, N. (2016) STIM and ORAI proteins: Crucial roles in hallmarks of cancer. American Journal of Physiology. Cell Physiology 310, C509-C519. [PubMed: 26791491]
116.
Bergmeier, W., Weidinger, C., Zee, I., and Feske, S. (2013) Emerging roles of store-operated Ca2+ entry through STIM and ORAI proteins in immunity, hemostasis and cancer. Channels (Austin, TX) 7, 379-391. [PMC free article: PMC3913761] [PubMed: 23511024]
117.
Sabbioni, S., Barbanti-Brodano, G., Croce, C. M., and Negrini, M. (1997) GOK: A gene at 11p15 involved in rhabdomyosarcoma and rhabdoid tumor development. Cancer Research 57, 4493-4497. [PubMed: 9377559]
118.
Feng, M., Grice, D. M., Faddy, H. M., Nguyen, N., Leitch, S., Wang, Y., Muend, S., et al. (2010) Store-independent activation of Orai1 by SPCA2 in mammary tumors. Cell 143, 84-98. [PMC free article: PMC2950964] [PubMed: 20887894]
119.
Flourakis, M., Lehen’kyi, V., Beck, B., Raphael, M., Vandenberghe, M., Abeele, F. V., Roudbaraki, M., et al. (2010) Orai1 contributes to the establishment of an apoptosis-resistant phenotype in prostate cancer cells. Cell Death Disease 1, e75. [PMC free article: PMC3032347] [PubMed: 21364678]
120.
Kondratska, K., Kondratskyi, A., Yassine, M., Lemonnier, L., Lepage, G., Morabito, A., Skryma, R., and Prevarskaya, N. (2014) Orai1 and STIM1 mediate SOCE and contribute to apoptotic resistance of pancreatic adenocarcinoma. Biochimica et Biophysica Acta 1843, 2263-2269. [PubMed: 24583265]
121.
Hou, M. F., Kuo, H. C., Li, J. H., Wang, Y. S., Chang, C. C., Chen, K. C., Chen, W. C., Chiu, C. C., Yang, S., and Chang, W. C. (2011) Orai1/CRACM1 overexpression suppresses cell proliferation via attenuation of the store-operated calcium influx-mediated signalling pathway in A549 lung cancer cells. Biochimica et Biophysica Acta 1810, 1278-1284. [PubMed: 21782006]
122.
Cheng, H., Wang, S., and Feng, R. (2016) STIM1 plays an important role in TGF-beta-induced suppression of breast cancer cell proliferation. Oncotarget 7, 16866-16878. [PMC free article: PMC4941356] [PubMed: 26919241]
123.
Xu, Y., Zhang, S., Niu, H., Ye, Y., Hu, F., Chen, S., Li, X., et al. (2015) STIM1 accelerates cell senescence in a remodeled microenvironment but enhances the epithelial-to-mesenchymal transition in prostate cancer. Scientific Reports 5, 11754. [PMC free article: PMC4530453] [PubMed: 26257076]
124.
Vanden Abeele, F., Skryma, R., Shuba, Y., Van Coppenolle, F., Slomianny, C., Roudbaraki, M., Mauroy, B., Wuytack, F., and Prevarskaya, N. (2002) Bcl-2-dependent modulation of Ca2+ homeostasis and store-operated channels in prostate cancer cells. Cancer Cell 1, 169-179. [PubMed: 12086875]
125.
Skryma, R., Mariot, P., Bourhis, X. L., Coppenolle, F. V., Shuba, Y., Abeele, F. V., Legrand, G., Humez, S., Boilly, B., and Prevarskaya, N. (2000) Store depletion and store-operated Ca2+ current in human prostate cancer LNCaP cells: Involvement in apoptosis. The Journal of Physiology 527(Pt 1), 71-83. [PMC free article: PMC2270062] [PubMed: 10944171]
126.
Dubois, C., Vanden Abeele, F., Lehen’kyi, V., Gkika, D., Guarmit, B., Lepage, G., Slomianny, C., et al. (2014) Remodeling of channel-forming ORAI proteins determines an oncogenic switch in prostate cancer. Cancer Cell 26, 19-32. [PubMed: 24954132]
127.
Stanisz, H., Stark, A., Kilch, T., Schwarz, E. C., Muller, C. S., Peinelt, C., Hoth, M., Niemeyer, B. A., Vogt, T., and Bogeski, I. (2012) ORAI1 Ca2+ channels control endothelin-1-induced mitogenesis and melanogenesis in primary human melanocytes. The Journal of Investigative Dermatology 132, 1443-1451. [PubMed: 22318387]
128.
Umemura, M., Baljinnyam, E., Feske, S., De Lorenzo, M. S., Xie, L. H., Feng, X., Oda, K., et al. (2014) Store-operated Ca2+ entry (SOCE) regulates melanoma proliferation and cell migration. PLoS One 9, e89292. [PMC free article: PMC3931742] [PubMed: 24586666]
129.
Feldman, B., Fedida-Metula, S., Nita, J., Sekler, I., and Fishman, D. (2010) Coupling of mitochondria to store-operated Ca2+-signaling sustains constitutive activation of protein kinase B/Akt and augments survival of malignant melanoma cells. Cell Calcium 47, 525-537. [PubMed: 20605628]
130.
Hooper, R., Zhang, X., Webster, M., Go, C., Kedra, J., Marchbank, K., Gill, D. L., Weeraratna, A. T., Trebak, M., and Soboloff, J. (2015) Novel protein kinase C-mediated control of Orai1 function in invasive melanoma. Molecular and Cellular Biology 35, 2790-2798. [PMC free article: PMC4508319] [PubMed: 26055321]
131.
Minn, A. J., Kang, Y., Serganova, I., Gupta, G. P., Giri, D. D., Doubrovin, M., Ponomarev, V., Gerald, W. L., Blasberg, R., and Massague, J. (2005) Distinct organ-specific metastatic potential of individual breast cancer cells and primary tumors. The Journal of Clinical Investigation 115, 44-55. [PMC free article: PMC539194] [PubMed: 15630443]
132.
Ponomarev, V., Doubrovin, M., Serganova, I., Vider, J., Shavrin, A., Beresten, T., Ivanova, A., et al. (2004) A novel triple-modality reporter gene for whole-body fluorescent, bioluminescent, and nuclear noninvasive imaging. European Journal of Nuclear Medicine and Molecular Imaging 31, 740-751. [PubMed: 15014901]
133.
Kim, J. H., Lkhagvadorj, S., Lee, M. R., Hwang, K. H., Chung, H. C., Jung, J. H., Cha, S. K., and Eom, M. (2014) Orai1 and STIM1 are critical for cell migration and proliferation of clear cell renal cell carcinoma. Biochemical and Biophysical Research Communications 448, 76-82. [PubMed: 24755083]
134.
Wang, J. Y., Sun, J., Huang, M. Y., Wang, Y. S., Hou, M. F., Sun, Y., He, H., et al. (2015) STIM1 overexpression promotes colorectal cancer progression, cell motility and COX-2 expression. Oncogene 34, 4358-4367. [PMC free article: PMC4426254] [PubMed: 25381814]
135.
Motiani, R. K., Abdullaev, I. F., and Trebak, M. (2010) A novel native store-operated calcium channel encoded by Orai3: Selective requirement of Orai3 versus Orai1 in estrogen receptor-positive versus estrogen receptor-negative breast cancer cells. Journal of Biological Chemistry 285, 19173-19183. [PMC free article: PMC2885196] [PubMed: 20395295]
136.
Zhang, W., Zhang, X., Gonzalez-Cobos, J. C., Stolwijk, J. A., Matrougui, K., and Trebak, M. (2015) Leukotriene-C4 synthase, a critical enzyme in the activation of store-independent Orai1/Orai3 channels, is required for neointimal hyperplasia. The Journal of Biological Chemistry 290, 5015-5027. [PMC free article: PMC4335238] [PubMed: 25540197]
137.
Motiani, R. K., Zhang, X., Harmon, K. E., Keller, R. S., Matrougui, K., Bennett, J. A., and Trebak, M. (2013) Orai3 is an estrogen receptor alpha-regulated Ca2+ channel that promotes tumorigenesis. The FASEB Journal 27, 63-75. [PMC free article: PMC3528310] [PubMed: 22993197]
138.
Motiani, R. K., Hyzinski-Garcia, M. C., Zhang, X., Henkel, M. M., Abdullaev, I. F., Kuo, Y. H., Matrougui, K., Mongin, A. A., and Trebak, M. (2013) STIM1 and Orai1 mediate CRAC channel activity and are essential for human glioblastoma invasion. Pflügers Archiv 465, 1249-1260. [PMC free article: PMC3748246] [PubMed: 23515871]
139.
Tsai, F. C., Seki, A., Yang, H. W., Hayer, A., Carrasco, S., Malmersjo, S., and Meyer, T. (2014) A polarized Ca2+, diacylglycerol and STIM1 signalling system regulates directed cell migration. Nature Cell Biology 16, 133-144. [PMC free article: PMC3953390] [PubMed: 24463606]
140.
Oh-hora, M., Yamashita, M., Hogan, P. G., Sharma, S., Lamperti, E., Chung, W., Prakriya, M., Feske, S., and Rao, A. (2008) Dual functions for the endoplasmic reticulum calcium sensors STIM1 and STIM2 in T cell activation and tolerance. Nature Immunology 9, 432-443. [PMC free article: PMC2737533] [PubMed: 18327260]
141.
Weidinger, C., Shaw, P. J., and Feske, S. (2013) STIM1 and STIM2-mediated Ca2+ influx regulates antitumour immunity by CD8(+) T cells. EMBO Molecular Medicine 5, 1311-1321. [PMC free article: PMC3799488] [PubMed: 23922331]
142.
Courjaret, R., Dib, M., and Machaca, K. (2017) Store-operated Ca2+ entry in oocytes modulate the dynamics of IP3-dependent Ca2+ release from oscillatory to tonic. Journal of Cellular Physiology 232, 1095-1103. [PubMed: 27504787]

Rawad Hodeify

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Fang Yu

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Raphael Courjaret

Department of Physiology and Biophysics

Weill-Cornell Medical College

Doha, Qatar

Nancy Nader

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Maya Dib

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Lu Sun

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Ethel Adap

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Satanay Hubrack

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

Khaled Machaca

Department of Physiology and Biophysics

Weill Cornell Medicine Qatar

Doha, Qatar

© 2017 by Taylor & Francis Group, LLC.

This work is licensed under a Creative Commons Attribution-NonCommercial-NoDerivs 3.0 Unported License. To view a copy of this license, visit http://creativecommons.org/licenses/by-nc-nd/3.0/

Bookshelf ID: NBK531436PMID: 30299656DOI: 10.1201/9781315152592-12

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