Advances in Intracellular Calcium Signaling Reveal Untapped Targets for Cancer Therapy

Sharma, Aarushi; Ramena, Grace T.; Elble, Randolph C.

doi:10.3390/biomedicines9091077

Open AccessReview

Advances in Intracellular Calcium Signaling Reveal Untapped Targets for Cancer Therapy

by

Aarushi Sharma

¹

,

Grace T. Ramena

² and

Randolph C. Elble

^1,*

¹

Department of Pharmacology and Simmons Cancer Institute, Southern Illinois University School of Medicine, Springfield, IL 62702, USA

²

Department of Aquaculture, University of Arkansas, Pine Bluff, AR 71601, USA

^*

Author to whom correspondence should be addressed.

Biomedicines 2021, 9(9), 1077; https://doi.org/10.3390/biomedicines9091077

Submission received: 14 June 2021 / Revised: 15 July 2021 / Accepted: 18 July 2021 / Published: 24 August 2021

(This article belongs to the Special Issue Calcium Signaling and Its Dysregulation in Cancer)

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Abstract

:

Intracellular Ca²⁺ distribution is a tightly regulated process. Numerous Ca²⁺ chelating, storage, and transport mechanisms are required to maintain normal cellular physiology. Ca²⁺-binding proteins, mainly calmodulin and calbindins, sequester free intracellular Ca²⁺ ions and apportion or transport them to signaling hubs needing the cations. Ca²⁺ channels, ATP-driven pumps, and exchangers assist the binding proteins in transferring the ions to and from appropriate cellular compartments. Some, such as the endoplasmic reticulum, mitochondria, and lysosomes, act as Ca²⁺ repositories. Cellular Ca²⁺ homeostasis is inefficient without the active contribution of these organelles. Moreover, certain key cellular processes also rely on inter-organellar Ca²⁺ signaling. This review attempts to encapsulate the structure, function, and regulation of major intracellular Ca²⁺ buffers, sensors, channels, and signaling molecules before highlighting how cancer cells manipulate them to survive and thrive. The spotlight is then shifted to the slow pace of translating such research findings into anticancer therapeutics. We use the PubMed database to highlight current clinical studies that target intracellular Ca²⁺ signaling. Drug repurposing and improving the delivery of small molecule therapeutics are further discussed as promising strategies for speeding therapeutic development in this area.

Keywords:

SOCE; ORAI; STIM; STIMATE; SERCA; PNCA; calmodulin; TRP; IP3R; MCU; VGCC

Graphical Abstract

1. Introduction

Ca²⁺ is the quintessential ion central to numerous cellular homeostasis and physiological functions. Low hydration energy, high polarizability, relative flexibility of coordination sites and bond length, and large concentration gradient across cellular membranes (100 nM intracellular to 2 mM extracellular) due to low intracellular levels make it the ion of choice at the core of cellular signaling in prokaryotes and eukaryotes alike [1,2]. The mechanisms adopted by cells for intracellular Ca²⁺ buffering involve sequestration by special proteins [3,4] (Figure 1). Some of these proteins exist in the soluble or non-membranous parts of the cytoplasm within or outside organelles that serve as repositories for Ca²⁺ ions [3]. Such proteins sequester cytosolic Ca²⁺ upon sensing an increase in its levels and participate in relaying the associated cellular messages. Other proteins that work as intracellular Ca²⁺ buffers exist in the lipid bilayers, plasma membrane, or organelle membranes, like pumps or transporters. Apart from these proteins, intracellular Ca²⁺ is regulated by inter-organellar transport and the influx of Ca²⁺ ions from extracellular space [5]. In this review, we provide an overview of key components and the associated major mechanisms of intracellular Ca²⁺ regulation under physiological conditions. It is followed by delineating how these proteins and pathways are manipulated by cancerous cells during tumorigenesis and progression.

Intracellular Ca²⁺ levels are managed through binding to special proteins or sequestration within different cellular compartments. The three main ways by which intracellular Ca²⁺ is buffered are depicted above—soluble or unbound proteins that are found in the non-membranous parts of a cell (cytosol or inside organelles), membrane-bound or intramembranous proteins (generally Ca²⁺ channels (like TRP), ATP-driven pumps (SERCA or PMCA), and ion exchangers (NCX)), and organellar compartments such as endoplasmic reticulum (ER), mitochondria, acidic vesicles (mainly lysosomes and Golgi bodies) or organellar junctions (endoplasmic reticulum-plasma membrane (ER-PM), endoplasmic reticulum-mitochondria, or endoplasmic reticulum-lysosomes) [3,4,5]. The major players regulating inter-organellar Ca²⁺ are mentioned in parenthesis within the cellular organelles section. IP₃R, inositol-3,4,5-triphosphate; NCX, sodium-Ca²⁺ exchanger; ORAI1 (or CRACM1), Ca²⁺ release activated modulator 1; PMCA, plasma-membrane Ca²⁺ ATPase; SERCA, sarco-endoplasmic reticulum Ca²⁺ ATPase; STIM1, stromal interaction molecule 1; SOCE (store-operated Ca²⁺ entry); TPC1/2, two-pore channel; TRP, transient receptor potential; VDAC, voltage-dependent anion channel.

2. Intracellular Ca²⁺ Buffers in Normal Cells

2.1. Soluble and Unbound Intracellular Proteins: Calmodulin, Calbindin, and Calretinin

The non-membranous proteins inside a cell can act as both Ca²⁺ sensors and buffers [2,3]. Most of these proteins have EF-hand motif(s) that allows Ca²⁺ ions to bind and trigger changes in protein folding, thereby influencing downstream or linked cellular pathways [3,4]. Calmodulin (CaM) is one of the best-studied and ubiquitously expressed Ca²⁺-sensing proteins known to play a key role in intracellular Ca²⁺ homeostasis [3]. As the prototype for intracellular Ca²⁺ sensors, its 148 amino acid structure is comprised of two Ca²⁺-binding sites, each with two EF-hand motifs: N- and C-termini alpha-helices with a Ca²⁺ coordination loop in between providing affinity for Ca²⁺ ion docking and sequestration [6]. The ability of CaM to transmit a change in free intracellular Ca²⁺ levels into modulation of a cellular response comes through its Ca²⁺-dependent structural flexibility. CaM can exist in a Ca²⁺-free closed conformational state (Apo-CaM), a semi-open (Ca2-CaM), or an open state (Holo-CaM or Ca4-CaM) after Ca²⁺-binding [6,7,8] (Figure 2). The latter two conformational states expose hydrophobic residues of this protein, thus allowing it to bind to target or effector molecules and acting as a fast-acting intermediary between change in intracellular Ca²⁺ and cellular processes. Differential Ca²⁺-binding on the two lobes of CaM makes fast buffering of a wide range of free intracellular Ca²⁺ possible for this protein. Analysis of CaM kinetics by Faas et al. has revealed that the N-lobe of CaM acts as the first site for Ca²⁺-binding during a massive increase in intracellular Ca²⁺ levels (>100 mM) in a nanodomain [9], whereas the C-lobe, having a higher affinity for Ca²⁺ than the N-lobe, captures Ca²⁺ in its EF-hand motifs when the Ca²⁺ concentration in the pool of intracellular fluid is 1–10 mM and both the motifs have Ca²⁺-bound to them. The presence of methionine residues in its lobes and plasticity of the central linker in its structure also provides CaM with properties to function as an adaptor protein in intracellular Ca²⁺ signaling [10]. CaM can bind to several targets or effector molecules over a variable distance and in multiple orientations to mediate change in intracellular Ca²⁺ signaling. Some major effector proteins that are regulated by CaM binding and are relevant for Ca²⁺ homeostasis include ORAI, EGFR, PI3K, IQGAP, and connexins [1].

Calbindin D-28k is another Ca²⁺-binding protein with six EF-hand motifs that buffers and transports free cytosolic Ca²⁺ but, unlike calmodulin, does not act as a linker or adaptor protein in shaping intracellular Ca²⁺ signaling [11,12]. Additionally, its expression is limited to a few cell types such as mammalian kidney ductal cells, intestinal absorptive epithelia, and neurons. Calretinin or calbindin D-29k, with 58% homology to calbindin D-28k, acts both as a nonlinear Ca²⁺ buffer and sensor predominantly in the neurons [13,14]. Expressed in the kidney and duodenum epithelial cells, calbindin D-9k or S100G is a monomer comprised of two EF motifs [15]. With no known binding partners, it is only considered a Ca²⁺ buffer. ER (endoplasmic reticulum) molecular chaperones, calreticulin (in the lumen), and calnexin (on the membrane) are also known to be Ca²⁺ buffers [5,16].

Physiological relevance: CaM is required for spatial and temporal regulation of [Ca²⁺]i as evident by its role in modulation (activation or inactivation) of Ca²⁺ pumps (such as PMCA and SERCA) and Ca²⁺ channels (such as CaV1.3, TRPV5 and 6, ORAI) [17,18,19]. CaM also acts via serine/threonine kinases known as Calmodulin-activated Kinases (CaMKs) to influence cellular processes like proliferation (for example, centrosome duplication at G1/S or anaphase to metaphase transition via CaMKII) [20]. Calbindin D-28k acts as a Ca²⁺ buffer proximal to Ca²⁺ channels like TRPV5 and maintains a steep gradient for ion entry [21].

2.2. Intramembranous Molecular Buffers: SERCA, PMCA, NCX, and TRP

Intra-membrane Ca²⁺ buffers primarily translocate free Ca²⁺ between domains and organelles. These mainly comprise ion exchangers, channels, and ATP-driven pumps [22]. SERCA or Sarcoendoplasmic Reticulum Ca²⁺ ATPase is an ATP-dependent ion pump known to significantly maintain free cytosolic Ca²⁺ concentration via actively pumping the ion into the endoplasmic reticulum (or sarcoplasmic reticulum in muscle cell). Among the eleven of these P-type ATPase pump isoforms (and variants) recognized so far, SERCA1a and SERCA1b are mainly expressed in adult and neonatal skeletal muscle cells, respectively. SERCA2a is found in cardiomyocytes, while SERCA2b and 2c are expressed ubiquitously. SERCA3 (all the six splice variants) are largely co-expressed with SERCA2b in hematopoietic, endothelial, and epithelial cells. With 85 percent and 75 percent overlap of SERCA2 and SERCA3 primary sequences to that of SERCA1, these isoforms exhibit differential affinity for Ca²⁺; SERCA3 demonstrates a fivefold lower propensity to bind Ca²⁺ than other isoforms [23,24,25]. Regardless, the isoforms share a general structure that includes 10-pass transmembrane helices and three cytoplasmic domain lobes (Figure 3A) [26,27]. Two closely spaced Ca²⁺-binding sites are present on the cytoplasmic side of transmembrane domains. These sites act cooperatively with each other such that the binding of Ca²⁺ ions to site I increases the binding affinity for site II [27]. Once the two Ca²⁺-sites are occupied, the cytoplasmic lobes—the nucleotide-binding (N)-domain followed by phosphorylation (P), and actuator (A)-domains—undergo conformational shifts and translocate Ca²⁺ ions [25,26] (Figure 3B).

Although phospholamban (PLN) has been shown to be a stronger inhibitor of the pump, some studies indicate sarcolipin (SLN) to inhibit SERCA at high Ca²⁺ concentrations [24,28]. Both PLN and SLN are type I transmembrane micropeptides that bind as dephosphorylated monomers (active form) to a groove surrounded by TM2, TM4, TM8, and TM9 of SERCA. SERCA2b being the more widely expressed isoform in non-muscle cells is modulated by other means than PLN and SLN. The inhibitors include another-regulin (ALN; a ubiquitously expressed inhibitor with PLN/SLN key SERCA2b-interacting amino acids), an additional transmembrane helix (TM11), and a cytoplasmic end luminal extension of SERC2b [28] (Table 1).

Figure 3. Sarcoplasmic endoplasmic Ca²⁺-ATPase structure and mechanism of Ca²⁺ ion transport. (A) SERCA pump structure is comprised of Actuator or A domain (for dephosphorylation), transmembrane or M domain (10 helices and two Ca²⁺ ion binding sites between helices 4 and 5), Nucleotide or N domain (source of phosphorylation), and Phosphorylation or P domain (consists of the site of phosphorylation; aspartate 351 residue) [26]. Picture in the inset is a simplified representation of SERCA crystal structure; (B) Numerous steps are involved in the transport of Ca²⁺ via SERCA across the endoplasmic reticulum membrane [28,29,30,31]. As per the above model, the pump cycle begins with SERCA having a high affinity to Ca²⁺ in its physiologically predominant E1-ATP state. After cytosolic Ca²⁺ ions bind to SERCA, the pump transitions into Ca2-E1-P-ADP state utilizing the energy from the transfer of ATP γ-phosphate in the N domain to D351 amino acid residue in P domain. Transport of Ca²⁺ ions to the ER lumen parallels the shift of pump from its high-energy intermediate conformation to the low energy E2-ATP state (with less affinity to Ca²⁺; not shown) triggered by replacement of ADP with ATP. Ca²⁺ ions are then exchanged in the lumen with 2–3 protons compensating for the loss of positive charge. Followed by dephosphorylation at D351. In a Ca²⁺-free resting stage, the γ-phosphate of ATP in N-domain is masked from phosphorylation site (P domain Asp³⁵¹) as well as the dephosphorylation site (A-domain TGES sequence) [25,32]. When two cytosolic Ca²⁺ ions bind to SERCA pump in its higher Ca²⁺ affinity state (known as E₁-ATP), it results in rotational changes within the transmembrane and cytoplasmic domains, thus bringing N-domain ATP closer to Asp³⁵¹ for transfer of γ-phosphate. From this high-energy intermediate state (Ca₂-E₁-P-ADP) the pump prepares to transition into a lower energy state, Ca₂-E₂-P-ADP by completely transferring the γ-phosphate to P-domain and coordinating Mg²⁺ ions to all the phosphate groups present. In a two-step process again, first ADP is exchanged for ATP in the ER causing appropriate conformational changes to create a channel and exposing the Ca²⁺-binding residues in its lumen (E₂P-ATP). Ca²⁺ ions are then released in exchange for protons and the pump restores its resting state. ADP, adenosine diphosphate; ATP, adenosine triphosphate; SERCA, sarcoendoplasmic reticulum Ca²⁺ ATPase.

P-type Ca²⁺-ATPases also exist within the plasma membrane and maintain cytosolic Ca²⁺ levels by transferring them into the extracellular space. The Plasma Membrane Ca²⁺ ATPases (PMCAs) were earlier known only as housekeeping proteins required for intracellular Ca²⁺ homeostasis, but some isoforms and splice variants are now known to have a more active role [33,34]. PMCAs transport one Ca²⁺ ion per ATP molecule which differs from two Ca²⁺ ions per ATP molecule stoichiometry of SERCA [35]. Four PMCA isoforms are known in mammals, each one with many splice variants [34]. PMCA1 (especially PMCA1b) has a ubiquitous expression with its presence being essential even during embryonic development. PMCA2 and PMCA3 are expressed more selectively in tissues such as the brain, pancreatic β cells, inner ear cells, mammary glands, and the heart. The pattern of PMCA4 tissue expression of overlaps with that of PMCA1, however, its absence does not cause embryonic lethality. The isoforms and variants also differ in the activation/inactivation rates which makes it possible for PMCAs to manage intracellular Ca²⁺ in the form of fast-acting spikes or slowly released puffs [34,36]. The general structure of such Ca²⁺ transporters comprises 10 transmembrane segments with large cytosolic loops TM 1–2 and TM 3–4, a cytosolic N- and C-termini tails [34,36,37] (Figure 4A). The first cytosolic loop consists of splice site A (regulates membrane-targeting) and phospholipid-binding sites, the second loop has an aspartyl-containing phosphorylation site, and the C-terminus is strewn with the calmodulin-binding site, splice site C and PDZ domain. The activity of all PMCA isoforms (and variants) is heavily regulated. Short-term regulation of catalytic activity of most of the “b” or splice site 2 variants is mainly calmodulin-dependent [37,38,39,40]. The binding of CaM reverses auto-inhibition of the pump due to conformational shifts which displace C-tail from cytosolic loops. Other means of autoinhibition reversal include phosphorylation of C-tail (Ser/Thr residues) by protein kinase A or C, proteolytic cleavage of C-tail, or dimerization via C-terminus. Change in the localization via interaction with PDZ proteins like MAGUK (membrane-associated guanylate kinase) at the C-tail or transcriptional and translational modulation influences the long-term activity of PMCAs.

PMCAs also partner with sodium-Ca²⁺ exchanger (NCX) in some cells to remove Ca²⁺ from the cytosol [41]. PMCA has high Ca²⁺ ion affinity and low capacity when compared to NCX that has low Ca²⁺ ion affinity but a high capacity for ion efflux. This means that PMCA maintains basal cytosolic levels or small bursts of Ca²⁺ ion entry while NCX is responsible for regulating large but transient increase in intracellular Ca²⁺. NCX or SLC8 belongs to a superfamily of the Ca²⁺ ion/cation antiporter gene family. Within the SLC8 family, NCX1, 2, and 3 are the identified functional members encoded by separate genes in mammals [42]. NCX1 is expressed ubiquitously, NCX2 is found in the brain and skeletal muscles, and NCX3 in neurons. Topological analysis of this family based on NCX1 predicts a structure with ten transmembrane alpha-helices. The first five helices form the N-terminus which is separated by a cytosolic loop from the remaining helices forming the C-terminus. The cytosolic loop (500 a.a.) is a regulatory site—a beta repeat region with two Ca²⁺ ion binding regions, CBD1 and CBD2. CBD1 is the primary site for detecting small changes in cytosolic Ca²⁺ ion concentration resulting in greater structural changes that activate the exchanger. CBD2 only responds to moderate change in cytosolic Ca²⁺ ion levels [42,43,44,45] (Figure 4B).

Physiological relevance: Blocking the function of SERCA isoforms can lead to disproportionate Ca²⁺ levels in the cell cytosol, thus activating apoptosis signals in select cell types. For instance, reduced SERCA2b activity in hepatocytes results in ER stress followed by cell death due to accumulation of excessive cytosolic Ca²⁺ [46]. PMCA can help maintain the local intracellular Ca²⁺ ion [Ca²⁺]i gradient required for cellular motility. Migrating endothelial cells have higher expression of PMCA at their leading edges to maintain low basal [Ca²⁺]i levels thereby, preventing continued activation of Myosin Light Chain Kinase (MLCK) and extended contraction of the cell membrane at the migration frontier [47]. On the other hand, NCX1 inhibition and thus impairment of [Ca²⁺]i extrusion allows the proliferation of pancreatic beta cells [48].

Figure 4. Structures of intracellular Ca²⁺ efflux machinery. (A) Plasma membrane Ca²⁺-ATPase (PMCA) is comprised of cytosolic N- and C-termini, two Ca²⁺-binding sites (akin to SERCA; not shown here), and 10 transmembrane (TM) segments. Between the N- and C-tails lie three loop structures named like the domains in SERCA—A domain (red circle between TM 2–3), P domain (blue circle between TM 4–5), and N domain (yellow circle between TM 4–5) [34,36,37,38,39,40]. Various binding motifs and regulatory or interaction sites exist on each terminus and cytosolic domain with variations observed in their presence between PMCA isoforms and splice variants. Ca²⁺-bound calmodulin or Ca²⁺-CaM is a major regulator for all PMCA family members. The proximity of Ca²⁺-CaM binding sites on the C-tail to their corresponding motifs in A- and N-domains determine the open or closed states of the pump; (B) Sodium–Ca²⁺ exchanger (NCX) also has a ten transmembrane topology with N- and C-tails facing the extracellular space [42,43,44,45,49]. The cytosolic side consists of an f-loop (between TM 5–6) that has two Ca²⁺ ion binding sites, an alternate splicing site, a regulatory site, a sodium ion-dependent inactivation site, and an inhibitory domain. The structure also has two conserved alpha-helical repeats (α1 and α2) that influence ion binding and transport. ATP, adenosine triphosphate; CaM, calmodulin; NCX2, sodium-Ca²⁺ exchanger 2; PKA, protein kinase A; PDZ, a structural domain; XIP, exchanger inhibitory peptide.

Initially ascribed to Ca²⁺ homeostasis in sensory neurons, Transient Receptor Potential (TRP) channels have lately been appreciated for a similar function in epithelial and immune cells [50]. Mammalian TRP channel superfamily is composed of 28 family members belonging to six subfamilies—TRPC (Canonical), TRPA (Ankyrin), TRPM (Melastatin), TRPV (Vanilloid), TRPP (Polycystin), and TRPML (Mucopilin)—that differ in their sensitivity to various sensory stimulations and affinity for cations (including Ca²⁺ ions) sequestration [51]. Commonly, TRP family members share a structure with six transmembrane domains, intracellular N- and C-termini, and a pore-forming TM 5–6 loop [52] Figure 5. Cation selectivity of a TRP channel is determined via an extracellular portion of the pore-forming TM 5–6 helices [51,53]. To have a functional channel for Ca²⁺ transport, TRP members form homo- or hetero-tetramers within and across the subfamily members [54]. The N-terminus of each tetramer subunit of a TRP channel along with corresponding transmembrane helices are associated with channel assembly and pore regulation [55]. The N-terminus within and across each TRP subfamily varies in the presence and number of ankyrin repeats; having such repeats in general at the amino end provides a site for protein-protein interaction or ligand binding [51,56,57]. The intracellular C-terminus of each subunit is a site for protein interaction and post-translational modification. It also brings structural and functional diversity between subfamilies [52,53,58]. For example, TRPC, TRPV, and TRPM subfamilies share a C-terminus TRP box motif—a short cytosolic hydrophobic stretch at the end of the transmembrane domain putatively holding the channel in a closed conformation [53,59]. Moreover, the C-tail of these channels can have PDZ protein binding domains (TRPV and C), sites for interaction with G-proteins (Gq/11)/calmodulin/PLCβ, ADP-ribose binding (NUDIX; TRPM2), or PLC-interacting kinase (PLIK; TRPM6 and 7) domains [59] (Table 2).

Physiological relevance: On a high level, TRP channels act as activators, integrators, as well as downstream effectors of Ca²⁺ signaling at the plasma membrane and in intracellular compartments [54,55,69] (Figure 6). Almost all TRP channels permeable to Ca²⁺ ions (other than TRPM4 and 5), can directly activate intracellular Ca²⁺ signaling [51]. Taking TRPC as an example, many members of this subfamily are activated by DAG (Diacylglycerol) which is produced by PLC β- or γ-mediated cleavage of phosphatidylinositol 4,5-bisphosphate (PIP₂) after the ligand binding at GPCRs or RTKs [59]. Being a family of non-selective cation channels reacting to numerous stimuli, TRP channels can also indirectly influence the activation of receptor-operated bulk entry of Ca²⁺ from the extracellular space [58]. TRPP1/2, TRPA1, TRPM8, and TRPV1-4 are all expressed on the ER membrane [54,70]. At this site, PLC-independent activation of the TRP channels (such as TRPV1) is suggested to induce ER Ca²⁺ release via inositol triphosphate receptor (IP3R) which further triggers bulk entry of extracellular Ca²⁺ into the cell (discussed further in a later section) [71,72,73,74]. On the flip side, cytosolic Ca²⁺ regulates the activity of TRP channels in response to physiological stimuli. This regulatory effect is usually through CaM binding (inhibition of TRPV5, TRPV6, and sensitization of TRPV3) and indirectly through CaM-binding kinase II (CaMKII) [69]. Among the extra-neuronal TRP channels, TRPA1, TRPC1, TRPM8, TRPV1, and TRPV4 have been recognized for their role in epithelial and immune cell Ca²⁺ homeostasis [50,75]. By regulating intracellular Ca²⁺, some of these channels like TRPV1 and TRPM8 which are expressed in human bronchial and lung epithelium, respectively, aid the release of chemoattractants and promote immune cell–epithelial cell interaction.

2.3. Cellular Organelles

2.3.1. Endoplasmic Reticulum: STIM, ORAI, IP₃Rs, and TRPC1 in SOCE and SOCIC Ca²⁺ Entry Models

The ER serves as the largest and most dynamic organelle reservoir for intracellular Ca²⁺ and is therefore central to an array of cell signaling processes for protein synthesis, folding, and post-translational modifications [79,80,81,82]. In contrast to the cytosol, ER Ca²⁺ ion levels can range from 100 uM to 1 mM based on the cell type [72,80]. ER and other intracellular organelles buffer excessive cytosolic Ca²⁺ by both housing Ca²⁺-binding proteins (example: calreticulin in ER) and via active transport (example: SERCA pumps in ER) [5,79,80]. Depletion of Ca²⁺ from the ER lumen actuates an indirect mode of Ca²⁺ entry into the organelle which is termed Store-Operated Ca²⁺ Entry (SOCE) or Ca²⁺ Release Activated Ca²⁺ (CRAC) entry; it is activated when plasma membrane receptors like PLC-coupled GPCRs (but not voltage-gated channels) trigger Ca²⁺ ion release from the organelle [83,84]. Exhaustion of the intraluminal ER Ca²⁺ ion store following such prolonged release is then sensed by STIM (Stromal Interaction Molecule) tethered to the ER membrane and subsequently relayed to the CRAC channels on the plasma membrane.

Physiological relevance: SOCE and its key players participate in multiple normal cellular processes that go awry during cancer progression. For example, siRNA mediated downregulation of STIM1 and ORAI1 in keratinocytes impairs cellular differentiation [85], whereas SOCE is found to be inactivated during mitosis [86]. By contrast, stimulation of IP₃Rs (and RyRs) promotes cell cycle progression of stem cells, pancreatic beta cells, renal cells, and more.

STIM, a type I transmembrane protein, was originally identified during a search for transmembrane and secretory proteins in stromal and pre-B lymphocytes [84,87]. The role of STIM in [Ca²⁺]_i signaling was later confirmed by independent work of two groups using high-throughput RNAi screens to identify inhibitors of thapsigargin evoked CRAC current (ICRAC) [88,89,90]. Both the isoforms, STIM1 and STIM2 have a luminal N-terminus and a cytoplasmic C-terminus [91]. Starting at the N-terminus, the basic STIM structure is comprised of EF-hand motifs (canonical and non-canonical) and a sterile α-motif (SAM), all of which help in intraluminal Ca²⁺ ion sequestration [83,88]; Figure 7A. The C-terminus has various coiled-coiled domains (CC1-3) that include CRAC activating domain (CAD; also known as STIM-ORAI activating region (SOAR)), and a polybasic site (PBS). The cytoplasm-facing C-terminus of STIM is essential for localization to ER-PM junctions and for interaction with CRAC channels. Even with greater than 74 percent sequence similarity, the STIM isoforms perform differently as ER Ca²⁺ sensors—the Ca²⁺ affinity of EF-SAM domains being greater for STIM1 (200 mM; 400 mM for STIM2) while the time to form oligomers for plasma membrane CRAC channel association is nearly 70 times longer for STIM2 [84,91,92,93]. This makes STIM2 more suitable for the regulation of basal ER Ca²⁺, although only a small contribution to SOCE was recorded in some studies [54,76]. In fact, STIM2 has been shown to inhibit STIM1-mediated SOCE [94]. With the expression of STIM1 on both the plasma membrane and the ER membrane, the difference between the cellular localization of these isoforms also highlights their functional disparity [90].

A year after STIM molecules were recognized as ER Ca²⁺ ion sensors, ORAI1 was confirmed as Ca²⁺ Release-Activated Ca²⁺ Modulator 1 (CRACM1) or simply, the CRAC channel [84,95]. Genome-wide RNAi screening, linkage analysis, and positional cloning isolated and identified ORAI1 as the mutated gene responsible for severe combined immunodeficiency in humans (with low ICRAC in T lymphocytes). Three isoforms of ORAI proteins exist in mammals—ORAI1, ORAI2, and ORAI3 [95,96]. Topologically, an ORAI channel-forming monomer has four transmembrane domains forming two intracellular (I-II, III-IV) and one extracellular peptide loop (II-III) [97,98] (Figure 7A). Both N- and C-termini are cytoplasmic with each having a STIM1-binding site or CRAC activation domain (CAD; 73–91 a.a. on amino side and 268–291 a.a. on the carboxyl side of ORAI1). A Ca²⁺-binding site is present at E106 that resides in the conserved TM1 segment. The three isoforms have high sequence homology in the transmembrane domains (92%) but only 62 percent overall due to some differences in the C-terminus (coiled-coiled domain) and III-IV loop. This results in variations between the isoforms in terms of activation time, Ca²⁺ ion-dependent inactivation kinetics, activation, or inhibition by 2-aminoethoxydiphenyl borate, affinity for STIM1, redox sensitivity, and activation by STIM2 [96,99,100].

The mechanistic model of SOCE has evolved over the last two decades. Going by the widely accepted “interaction model” or “diffusion trap model”, STIM1 exists as a dimer in resting (inactive) state where the Ca²⁺-bound canonical EF-hand, the non-canonical EF-hand, and SAM domain on each monomer together impart structural stability [95,99,100] (Figure 7B). It is complemented by the conformation of the cytosolic segment that keeps the ORAI-binding CAD domain, and the plasma membrane interacting polybasic C-tail hidden away. Depletion of unbound ER Ca²⁺ then triggers the freely diffusing STIM1 dimers on the ER surface to lose Ca²⁺ ions from cEF; this causes a conformational change on the C-termini exposing STIM CAD domains and extending the polybasic tails to bind ORAI hexamers and PIP2, respectively, at the ER-PM junction. Some studies have recognized STIM1 expressed on PM along with ORAI1 as key players for a store-independent, arachidonic acid-regulated Ca²⁺ current in association with ORAI3 [101,102]. Nonetheless, the function of plasma membrane STIM1 is debatable to date.

The release of intraluminal ER Ca²⁺ that evokes an influx of extracellular Ca²⁺ via ORAI1-STIM1complex at the ER-PM puncta is best known to be mediated by activated IP3R [103,104]. Certain types of ligand-stimulated GPCRs and RTKs generate IP3 and DAG as secondary effectors; the IP3 molecules then bind to its receptors expressed on the ER surface. Sampieri and Vaca et al. discovered IP3Rs to be spatially and functionally associated with STIM1 localized at the puncta [105]. In their study, activated IP3Rs immediately localized to STIM1 to form a complex that allowed the cEF domain of the latter to effectively sense nearby intraluminal Ca²⁺ depletion. Furthermore, overexpression of IP3Rs was shown to enhance the SOCE Ca²⁺ influx. Contrastingly, IP3Rs remain inactive during ER Ca²⁺ leak due to SERCA pump inhibition. Consequently, Ca²⁺ ion depletion around STIM1 cEF is slow and ineffectivewhich leads to smaller ORAI1 currents.

Figure 7. Major molecules in store-operated Ca²⁺ entry. (A) STIM, ORAI, and IP3Rs are central to the influx of Ca²⁺ ions from extracellular space into cytosol re-quired for replenishing the depleted intracellular stores. In humans, STIM has two isoforms. Both, STIM1 and STIM2 have a type I transmembrane structure comprised of a signal peptide at the N-terminus for localization to the ER, an EF-hand motif, and a sterile alpha motif for binding of luminal Ca²⁺ ions. SAM is also a site for protein–protein interactions), and a short transmembrane segment followed by three coiled-coiled domains (CC1, CC2, and CC3, that form ORAI-activating STIM fragment or OASF). CC2-CC3 region (or SOAR) is adequate as well for STIM–ORAI interactions [106,107,108,109]. The ORAI interacting sites on the domain are hidden in inactive STIM dimers. A Ca²⁺-dependent inhibitory domain, a proline/serine-rich region (polybasic; PBS), and a lysine-rich tail segment are found at the C-terminus. ORAI channels have three isoforms in humans. All the ORAI isoforms have an N-terminal ETON segment (cytosolic), four transmembrane domains, and a cytosolic C-terminus. The structure between TM 1–2 and TM 3–4 is exposed to the extracellular space. The yellow vertical bars depict regions of amino acid residues (positive, negative, hydrophobic, or special) that influence ORAI channel pore selectivity. ORAI channel exists as a hexamer that is formed by oligomerization of 3 dimer subunits. Inositol 1,4,5-triphosphate receptor subunits have most of their structure (5 domains) exposed to the cytosol. Each IP3R is formed of four subunits [110,111,112,113]. The transmembrane segments and the C-terminus are involved in pore-forming, gate-keeping, and tetramerization; (B) Subsequent to the activation of IP3Rs on the ER membrane by ligand IP3 and cytosolic Ca²⁺, STIM dimers are activated once the luminal Ca²⁺ concentration drops below basal levels [114,115,116,117,118,119]. The coiled-coiled domains experience a shift in their conformation, thus exposing the SOAR/OASF region for interaction with ORAI channels on the plasma membrane [120]. STIM-ORAI functional coupling for SOCE pore formation requires 3 STIM dimers interacting with one ORAI hexamer [87]. STIM dimer composition and function vary under different cellular states. Under resting or physiologically inactive state (or with low stimuli), STIM dimers are largely composed of STIM2 (S2) which has an inhibitory effect on IP3R activation. As the activation stimuli become stronger and intracellular store gets depleted of Ca²⁺, STIM dimers composed mostly of STIM1 takeover to participate in SOCE. IP3Rs are also expressed on parts of ER membrane away from the ER-PM microdomain involved in SOCE. These receptors provide intracellular Ca²⁺ ions for downstream Ca²⁺ signaling including NFAT-mediated transcription. Red semi-circle in the ER represents high luminal Ca²⁺ levels, pink semi-circle is for moderately low Ca²⁺ ion concentration, and pale semi-circle indicates extremely low Ca²⁺ concentration. CC, coiled-coil; NFAT, nuclear factor of activated T-cells; SAM, sterile alpha motif; SOAR, STIM1 Orai1-activating region; TM, transmembrane.

Mikoshiba et al. were the first to identify and characterize IP3Rs in Purkinje cells of cerebellar mutated mice as binding sites for the second messenger, IP3 [121]. Three members of this receptor have been identified in mammals all of which share a 2700 amino acid structure composed of six transmembrane segments and five distinct domains—IP3-binding suppressor/coupling domain at N-terminus, IP3-binding domain, internal modulatory/coupling domain, Ca²⁺ pore-forming transmembrane domain, gate-keeper domain at the C-terminal [122] (Figure 7A). Four IP3R subunits, each with the five domains, come together to form a functional IP3R in a homomeric or heteromeric fashion. Other than the pore-forming domain, the majority of the IP3R structure is extracellular (Figure 7A). The affinity of the IP3-binding domain for IP3 is similar between the isoforms. However, the binding affinity is modified by the N-terminal coupling domain [123]. It has been now revealed that the 225 a.a. segment in the N-terminus of IP3R1 decreases the binding of IP3 to the receptor-binding site by directly interacting with it. Different studies agree that IP3R2 is the most sensitive and IP3R3 is the least sensitive for IP3 docking [122]. The N-terminal domain also allows for receptor regulation via proteins like HOMER, CaM, Ca²⁺ Binding Protein 1 (CaBP1), Ankyrin, and IRBIT (IP3R binding protein released with inositol 1,4,5-trisphosphate) [124,125,126,127,128,129]. IP3 and Ca²⁺ ions are the main regulators of IP3R channel activity [123]. Each subunit of the functional IP3R tetramer has a binding site for IP3 and the predominant observation is that channel opening requires occupancy of more than one but not all four IP3-binding sites. N-terminal residues 1–223 and internal modulatory domain residues 651–1130 have been shown to be essential for coupling IP3 binding to channel opening [130]. Studies have found IP3 docking to also be necessary for IP3 receptor cluster formation in the ER membrane for localized Ca²⁺ release and SOCE activation [131,132]. Regulation by cytosolic Ca²⁺ ions is biphasic; a minor increase in its concentration accelerates channel activity in response to IP3 while higher levels inhibit channel opening [72,122,133]. Purportedly, intraluminal Ca²⁺ influences IP3 binding to the receptor as well with high levels sensitizing the ligand–receptor interaction and vice versa [133]. IP3R-mediated Ca²⁺ release has another positive regulator, OAG (1-oleoyl-2-acyl-sn-glycerol) that indirectly amplifies the channel activity by increasing IP3 production through the PLC pathway [134].

Physiologically, intracellular Ca²⁺ release by IP3R and store refilling by SOCE occur less in phasic format and overlap to quite an extent [88]. This means that in between the IP3-dependent Ca²⁺ ion leaks, closure of IP3Rs, and initiation of Ca²⁺ influx by SOCE, there is a time window when some IP3Rs are active and releasing Ca²⁺ while ion influx by STIM-ORAI complex and SERCA pump-mediated ER store repletion are occurring. The active IP3Rs in that duration are, however, not clustered the ER-PM junction and thereby contribute to differential activation of SOCE downstream effectors situated near puncta but away from the Ca²⁺ influx nanodomain [88,98]. Apart from IP3R, the translocon, which is a complex of proteins that help newly formed polypeptides having signaling sequence to be transported from cytosol to ER, acts as another Ca²⁺-leak channel [135]. When bound to a ribosome, a translocon complex forms a leak pore of 4–5 nm in size that is permeable to intraluminal Ca²⁺ during the resting state of ER.

Newer studies have provided evidence for more proteins besides STIM, ORAI, and IP3R to be participants in store-operated Ca²⁺ entry. Luis Vaca was the first to describe a protein complex termed Store-Operated Ca²⁺ Influx Complex or SOCIC that involves many key proteins such as TRPC1, SERCA, and microtubule end-tracking protein, EB1 [136]. The quintessential CRAC channels required for SOCE have a voltage-independent, highly Ca²⁺ ion-selective, inwardly rectifying (reversal potential near 40 mV), low amplitude (6 fA) current at physiological negative membrane potential which is blocked by low concentration of lanthanides and a high dose of 2-APB (2-aminoethoxydiphenyl borate) [137]. On the other hand, the delayed global Ca²⁺ influx mediated by SOCIC complex that simultaneously utilizes TRPC as its ion channel generates a current amplitude up to 1–2 pA [84]. TRPCs were originally speculated to be CRAC channels [138]. Typically, electrophysiological properties of TRPC homomers include an inwardly rectifying (reversal potential about 15 mV) Ca²⁺ ion-selective current that is activated by store depletion (via thapsigargin or IP3 inclusion in micropipette during whole-cell patch clamp) and inhibited by lanthanides as well as 2APB [139]. Experimental evidence has also demonstrated a decline in SOCE response corresponding to downregulation or no expression of endogenous TRPCs in vitro and in vivo, respectively. Nonetheless, certain dissimilarities in their current properties with that of ICRAC made the role of these channels in SOCE debatable. Presently, TRP channels are known as the key components of receptor-operated Ca²⁺ entry (ROC) where these channels open for Ca²⁺ entry in response to increased DAG levels from PIP2 hydrolysis upon Gq-coupled receptor activation [139]. However, with the proposed SOCIC model and some recent studies, the contribution of TRPC channels to ISOC currents is beginning to unravel [136,139].

The SOCIC model is based on the findings that TRPC-mediated Ca²⁺ influx is dependent on several interactors [138]. STIM1 is a major interactor and regulator of TRPC expressed within the ER-PM junctions. Vesicular TRPC1 is positioned near ER–PM junctions in unstimulated cells with the help of Rab4-dependent fast-recycling endosomes [140]. Store depletion and subsequent Ca²⁺ entry by ORAI1-STIM1 complexes translocate vesicular TRPC1s into the proximity of ORAI1-STIM1 clusters present on the cell surface. At this point, critical interaction between TRPC1s and caveolin-1 allows insertion of these channel proteins into cholesterol-rich lipid rafts in the junctions, thereby placing them proximal to ORAI1 for subsequent activation by Ca²⁺ influx. To be activated and participating as a store-operated Ca²⁺ channel, TRPC1 in the lipid rafts dissociates from caveolin-1 before engaging STIM1. STIM1 gates TRPC1 through the interaction of lysine residues at 684–685 a.a. in its polybasic tail with the aspartate residues at 639–640 a.a in the tetrameric channel protein. Other regulatory interactions between these proteins involve STIM1 ERM and SOAR domains, and TRPC1 coiled-coiled domains. Following a sustained and global Ca²⁺ release inside the cells, TRPC1 dissociates from STIM1 to re-associate with caveolin-1 for internalization into Rab5-linked early endosomes (via Arf6-dependent pathway). Due to the experimental proof of direct interaction between ORAI1-TRPC1-STIM1 shown by coimmunoprecipitation, Luis Vaca et al. considered TRPC1 to be the pore-forming component and the surrounding ORAI1 to be the regulator in their proposed SOCIC model [136]. Lately, studies have diverged from this model with mounting evidence supporting the formation of distinct but interacting ORA1-STIM1 and TRPC1-STIM1 Ca²⁺ ion conducting pores within the lipid rafts [139,141]. Using the new model, ORAI1-STIM1 based Ca²⁺ entry explains the “oscillatory” changes in intracellular levels of the ion whereas, the larger scale intracellular Ca²⁺ modulations are attributed to TRPC1-STIM1 associated influx. Physiologically, ICRAC activates proteins like calcineurin that promote translocation of NFAT and subsequent gene expression that is distinct from activation of NFκB based transcription triggered by SOCIC complex (ORAI-STIM1 + TRPC1-STIM1)-mediated ISOC.

Knockdown and mutation analyses reveal that the TRPC1-mediated store-operated Ca²⁺ currents are very much dependent on the expression and normal function of both STIM1 and ORAI1 [141]. Scaffolding proteins (example: Homer1), junction stabilizing proteins (example: Junctate, junctophilins (JP), and extended synaptotagmins (E-Syts)), vesicle-membrane fusion protein (example: synaptosome-associated protein (SNAP-25)), and STIM1 inhibitor protein called SARAF (store-operated Ca²⁺ entry (SOCE)-associated regulatory factor) also regulate TRPC1-mediated store-operated Ca²⁺ entry [142,143]. Some SOCIC modulators directly impact ORAI1 and STIM1 function too. For instance, Homer1 (cytoplasmic) and junctate proteins coimmunoprecipitate and apparently interact with both ORAI and STIM1, thereby promoting the SOCE complex formation [139]. Similarly, CLCA2 or human chloride channel accessory protein 2, a putative tumor suppressor expressed on the cell surface known to enhance SOCE response, colocalizes and coimmunoprecipitates with ORAI1 and STIM1 [144]. STIMATE (STIM-Activating Enhancer) or TMEM110, an ER-resident protein that colocalizes with STIM1 positively impacts SOCE in a two-prong manner. It modulates STIM1-ORAI1 mediated Ca²⁺ signaling by promoting STIM1 translocation to ER-PM junctions and via stabilizing the puncta [145,146]. By contrast, proteins such as CaM (cytoplasmic) and SARAF (ER membrane) directly interact with the ORAI1-STIM1 complex that causes suppression of SOCE response by ORAI1 inactivation (both) and deoligomerization of STIM1 (SARAF only) [141]. The revised SOCIC-based current model for SOCE is thus, visualized, to begin with, ER store depletion that results in the release of STIM1 dimers from the impact of regulators like SARAF and calsequestrin [141,147,148] (Figure 8). As the “free” STIM1 dimers oligomerize, ORAI1 molecules cluster into homo or hetero hexamers in plasma membrane domains that are low in PIP₂ and cholesterol. ORAI1 clustering triggers the recruitment of TRPC1 tetramers from proximal vesicles into the lipid raft domains. In coordination with the ER-PM junctions coming closer and being stabilized by the tethering proteins (such as junctates, junctophilins (JPH3, JPH4), and E-Syt1), the STIM1 oligomers form independent complexes with the ORAI1 and TRPC1 clusters. APC (adenomatous polyposis coli) facilitates this localization of STIM1 into ER-PM puncta by dissociating the ER Ca²⁺ sensor from microtubule end-tracking protein, EB1. Subsequently, the ORAI1-STIM1 and TRPC1-STIM1 complexes move into cholesterol and PIP2 rich domains to activate ICRAC and ISOC for replenishing Ca²⁺ stores.

2.3.2. Mitochondria and Acidic Vesicles (Mainly Lysosomes)

Mitochondria, known to be the “powerhouse of the cell”, also play a critical role in maintaining Ca²⁺ ion levels in the cytosol and endoplasmic reticulum [83,149,150]. These sphero-cylindrical organelles that are found mostly aggregated around the nucleus store similar levels of intracellular Ca²⁺ as the cytosol (0.1 μM) [140]. Electrochemical proton gradient or membrane potential (~Ψmt = −150 to −180 mV) and close apposition to ER are the two key factors responsible for Ca²⁺ uptake in mitochondria [151]. The free movement of small molecules (less than 5 kDa) from the outer mitochondrial membrane (OMM) into the inner mitochondrial space and their impermeability across the latter generates a high electrochemical proton gradient for ATP synthesis [152]. This gradient simultaneously draws Ca²⁺ ions from the cytosol.

Transfer of Ca²⁺ ions from ER to mitochondria occurs at specialized microdomains or contact sites known as Mitochondrial Associated Membranes (MAMs). These are characterized by the ER and OMM apposed at 10–25 nm from each other and are strewn with a cluster of channels, transporters, exchangers, and tethering proteins for facilitating Ca²⁺ ion transfer [151,152,153,154]. IP3Rs localized at the ER side of the MAMs release Ca²⁺ ions that gate voltage-dependent anion channels (VDACs) located on the OMM [149,153]; Figure 9A. VDACs (1, 2, and 3) are 30 kDa polypeptides having a 19-strand beta-barrel structure that regulates the flux of metabolites (polyvalent anions like ADP and ATP) across the outer mitochondria membranes [155]. These channels transport cations including Ca²⁺ more readily than anions like chloride. Due to voltage-dependent electrostatic gating, the ion selectivity and flux across VDACs change between open and closed states. For instance, the movement of alpha-helix positive charge to the channel outer walls in the closed state increases Ca²⁺ ion flux by 10 times relative to the open state. The importance of proximity between IP3Rs and VDACs in MAMs became clearer when it was realized that the channels on the inner mitochondrial membrane (IMM) transporting Ca²⁺ ions into the matrix have a low affinity (Km ~5–10 mM, KD ~10–50 mM) to these cations [151,156]. These channels, known as Mitochondrial Ca²⁺ Uniporters (MCUs), are highly selective for Ca²⁺ ions, and their opening demonstrates sigmoidal dependence on the cation concentration partly due to lowering of Ψmt that subsequently diminishes drive for cation flux. MCU (40 kDa protein) oligomers form a functional multi-protein complex with their regulators—the mitochondria Ca²⁺ uptake proteins (MICU1, 2, 3) and the essential MCU regulators (EMRE). MICU1 or CBARA1 and MICU2 form obligate heterodimers together in IMM to regulate MCU. These proteins have EF-hands that sense Ca²⁺ ions concentration in the IMS and accordingly inhibit or promote MCU activity. MICU1 is known to stimulate the rapid agonist-mediated Ca²⁺ ions uptake while MICU2 acts as a gatekeeper for MCU during low Ca²⁺ ion concentrations.

MAMs are stabilized by several chaperones and tethering proteins for optimal functioning of key Ca²⁺ ion transporting components, IP3R-VDAC-MCU/MICU1/MICU2 [157,158] (Figure 9A). Mitofusin 1 and 2 (MFN1/2) promote and regulate ER-mitochondria connectivity at MAMs by maintaining ER shape along with the interaction between adjacent mitochondria. Out of the OMM-linked GTPases, MFN2 is directly involved in MAM formation and tends to cluster more in the microdomain via either homotypic or heterotypic (with mitostatin or THCP) interactions, whereas MFN1 has a dominant role in mitochondrial fusion [159]. GRP75, a member of the heat shock protein 70 family (glucose-regulated protein 75 or HSPA9 or mortalin), is an essential cytosolic tethering protein that stabilizes the interaction between IP3R N-terminus and VDAC by acting as a bridge [160]. PML (or pro-myelocytic leukemia), a tumor-suppressor protein enriched on the ER side of MAM microdomains, modulates Ca²⁺ ion release from IP3R by forming a multi-protein complex with the receptor, AKT (or protein kinase B), and protein phosphatase A, thereby modifying the ER-mitochondria Ca²⁺ ion transfer [158]. MAMs are also enriched with ER chaperone proteins like Sigma 1 Receptor (Sig1R) and BiP (immunoglobulin heavy chain binding protein or GRP78) that interact with each other under normal cytosolic Ca²⁺ ion levels. However, with the release of ER Ca²⁺ ions, SigR1 dissociates from BiP to bind and prevent IP3R degradation, thus enhancing Ca²⁺ transfer to mitochondria.

Figure 9. Mitochondrial and lysosomal impact on intracellular Ca²⁺ signal. (A) Primary components of Ca²⁺ signaling at the mitochondrial associated membranes (MAMs) include IP₃R3 on the endoplasmic reticulum, VDAC1 on the outer mitochondrial membrane, and MCU complex on the inner mitochondrial membrane [151,154,156,161]. Transport of Ca²⁺ ions from ER to mitochondria plays a crucial role in cellular metabolism (autophagy), cell survival (during unfolded protein response and impinging cell death signals), lipid production, and distribution [162,163,164,165,166]. For these cellular processes to proceed normally, the integrity of MAMs is essential. Various tethering proteins regulate MAM structural integrity. Such secondary MAM components include GRP75, PML, GRP78 (or BiP), mitofusins (MFN1 and 2), and phosphofurin acidic cluster sorting protein (PACS2) [158,159,160,164,167,168,169]. Localized to the ER side of MAMs, growth factor-activated mTORC2 utilizes phosphorylated PACS2 (via Akt) to help avoid structural disruption of this sub-compartment [170]. It also phosphorylates IP3R3 to promote Ca²⁺ release at MAM sites. Other key MAM localized proteins maintain Ca²⁺ homeostasis in the region. These are calreticulin and calmodulin in the ER lumen, and SERCA on the ER membrane (184). Excessive Ca²⁺ ions in the mitochondrial matrix are extruded mainly by a permeability transition pore (mPTP). This Ca²⁺ efflux pore is situated on the inner mitochondrial membrane and is comprised of F1F0 ATPase with cyclophilin D (CyPD), adenine nucleotide translocase (ANT), and mitochondrial phosphate carrier (PiC) as its regulators [171,172]. Proapoptotic molecules Bax and Bak influence mPTP opening by controlling outer mitochondrial membrane permeability [173]. Ca²⁺ exchangers such as NCLX and HCX are less understood efflux mechanisms that may prevent overload of these cations in the organelle [151,174]; (B) Lysosomal Ca²⁺ promotes IP₃R3 mediated Ca²⁺ release from the endoplasmic reticulum [175,176]. Activated by NAADP, two-pore channels (TPCs) and TRPMLs are the main mode of Ca²⁺ extrusion from lysosomes. Multi-subunit V-ATPases actively transport hydrogen ions into the lysosomal lumen to maintain acidic pH that has been previously considered as the driving force for Ca²⁺ entry into lysosomes via HCX. Although any direct association between acidic lumen and Ca²⁺ store maintenance is now considered controversial, any indirect impact on lysosomal Ca²⁺ stores has not been ruled out. Parallel to this, the dependence of lysosomal Ca²⁺ uptake on ER Ca²⁺ stores is speculated [177]. Lysosomes can also play a more active role in intracellular Ca²⁺ homeostasis as indicated by their inhibition of SOCE [178]. Akt, a serine-threonine kinase; BiP, binding immunoglobulin protein; CyPD, cyclophilin D or peptidyl-prolyl cis-trans isomerase D; EMRE, essential mitochondria regulator; GRP75, glucose-regulated protein 75; HCX, hydrogen-Ca²⁺ exchanger; MCU; mitochondrial Ca²⁺ uniporter; MCUR1, mitochondrial Ca²⁺ uniporter regulator 1; MICU1/2, mitochondrial Ca²⁺ uptake 1 or 2; MFN1/2, mitofusin 1 or 2; mPTP, mitochondrial permeability transition pore; ML-SA1, mucolipin synthetic agonist 1; mTORC2, mTOR complex 2; NAADP, nicotinic acid adenine dinucleotide phosphate; NCLX, mitochondrial sodium Ca²⁺ exchanger; PML, promyeloctic leukemia protein; PACS2, phosphofurin acidic cluster sorting protein 2; SERCA, sarcoendoplasmic reticulum Ca²⁺ ATPase; Sig1R, sigma 1 receptor; TFEB, transcription factor EB; TPC, two-pore channel; TRPML, transient receptor potential mucolipin channel; V-ATPase, vacuolar ATPase; VDAC, voltage-dependent anion channel.

Physiological relevance: Mitochondrial-associated membranes are not just linkage points for ER and mitochondria. These sites finely regulate the movement of ions, metabolites (including Reactive Oxygen Species and lipids), and signaling molecules between the two organelles and thus, are central to normal ER function and mitochondrial biogenesis [164]. Note that triggers for vital processes such as autophagy and apoptosis hinge on the functioning of these dynamic protein bridges that maintain Ca²⁺ ion flow between the two organelles. For example, it is evident from certain studies that post-metabolic stress, autophagy-inducing proteins (like Beclin 1) and the ones involved in autophagosome formation (Atg 14L and Atg5) localize at MAMs. Moreover, the knockdown of MAM complex proteins such as mitofusins impairs autophagosome formation.

Ca²⁺ extrusion is as important as uptake for homeostasis in mitochondria. Two Ca²⁺ exchangers that localize to IMM, a sodium-Ca²⁺-lithium exchanger (NCLX) and a hydrogen-Ca²⁺ exchanger (mCHE) [151,174]. NCLX or SLC24A6 (solute carrier family 24 member 6) is an isoform of plasma membrane sodium-Ca²⁺ exchanger and is mainly expressed in excitable cells [179,180]. It is assumed to exchange three Na⁺ inside the matrix for one Ca²⁺ ion and its electrogenic activity (Ψmt dependent) is inhibited by a selective inhibitor (CGP-37157). Much less is known about the hydrogen-Ca²⁺ exchanger that extrudes one Ca²⁺ ion per 2-3 hydrogen ions [159]. Mitochondrial permeability transition pore (PTP; 3 nm pore diameter), a high conductance non-selective ion channel also has a considerable role in mitochondrial Ca²⁺ ion efflux [181,182]. The remarkable difference in Vmax (maximum rate of reaction) of MCU and the combined efflux rate of Ca²⁺ exchangers creates a huge kinetic imbalance that predisposes mitochondria toward Ca²⁺ ion overload [181]. Opening of PTP prevents this overload via fast Ca²⁺ ion efflux leading to depolarization and subsequent increase in permeability of the inner mitochondrial membrane. The open state of PTP is favored by the presence of Ca²⁺ ions, reactive oxygen species (ROS), mitochondrial matrix pH (around 7.4), and cyclophilin D (CypD) [183]. Inhibitors of PTP include divalent ions like Mg²⁺, Sr²⁺, Mn²⁺, cyclosporin (directly inhibits cyclophilin D), nucleotides, and matrix acidification [181]. The quest to molecularly identify PTP led to the discovery of it being formed by F₀F₁ ATPase dimers, though the exact location of the pore within the dimer remains to be identified [184]. PTP is active under normal physiological conditions as well where it functions in a transient low conductance mode to maintain cytosolic Ca²⁺ level without irreversibly changing Ψmt [171]. The positioning of these sphero-cylindrical organelles within the cell, too impacts cytosolic and mitochondrial Ca²⁺ buffering. For instance, the rise in cytosolic Ca²⁺ is limited to the apical or secretory side of pancreatic acinar cells by a “belt” of mitochondria unless the organellar Ca²⁺ buffering capacity is superseded [185].

While IP3 acts as the dominant Ca²⁺-mobilizing messenger, cADPR (cyclic ADP-ribose) and NAADP (nicotinic acid adenine dinucleotide phosphate) are also known to modulate intracellular Ca²⁺ stores [186]. cADPR evokes Ca²⁺ ion release from ER by acting on ryanodine receptors (RyR; counterpart of IP3R in myocytes and co-expressed in some other cell types). NAADP releases Ca²⁺ from acidic and/or secretory vesicles such as lysosomes and endosomes [187]. Although the exact stimulus for intracellular NAADP synthesis has not been established, some studies link activation of certain GPCRs and tyrosine kinase receptors to the formation of this Ca²⁺ messenger.

In most mammalian cells, lysosomes comprise ~5 percent of the cell volume and store similar levels of intracellular Ca²⁺ (0.5 mM) as the ER [175,188] Due to relatively smaller size than ER, lysosomes release nearly undetectable amounts of intracellular Ca²⁺ in response to NAADP trigger [175]. It is hypothesized that this weak Ca²⁺ ion signal is subsequently amplified by ER Ca²⁺ release. Such a model of anterograde Ca²⁺ signal coupling can work only when ER and lysosomes are adjacent. Two-pore channels (TPCs; TPC1 in both lysosomes and endosomes; TPC2 only in endosomes) and TRPML are the prime modes for Ca²⁺ ion release (others include P2X4, VGCCs, TRPA1, and TRPM2) while V-type H⁺ ATPase and H⁺/Ca²⁺ exchanger may concertedly transport Ca²⁺ ions into lysosomes [175,176] (Figure 9B). Some studies support that anterograde ER-lysosomal coupling happens at specialized membrane contact sites (MCS) and have experimentally demonstrated how the removal of TPCs from such sites disrupts the NAADP-based inter-organelle signaling [189,190]. Nonetheless, without a clear understanding of the MCS structure, its functional significance is difficult to verify in anterograde ER-lysosomal Ca²⁺ coupling. Absent the anterograde Ca²⁺ release, it is suggested that lysosomes can participate in refining the ER Ca²⁺ release signal by sequestering Ca²⁺ ions in the MCS microdomain and inactivating background IP3R activity [170,175]. Few cell types are predisposed to retrograde or reverse Ca²⁺ ion signal coupling between ER and lysosomes where IP3 triggers ER Ca²⁺ release that eventually stimulates NAADP synthesis and lysosomal Ca²⁺ ion release via TPCs [191] (Figure 9B). Research is still underway to determine why some cell types have this form of ER-lysosomal signaling and what its physiological relevance is.

3. Redistribution of Intracellular Ca²⁺ and Hijack of Its Regulatory Machinery in Cancer Cells

The relationship between Ca²⁺ and carcinogenesis goes back to the 1946 pioneer study by Carruthers et al. where [Ca²⁺]_i in mouse and human squamous cell carcinoma was measured to be 57 percent and 47 percent lower, respectively, than respective normal epidermises [192]. Following research further highlighted reduced cellular Ca²⁺ dependency with an increasing degree of neoplastic transformation [193,194,195,196,197,198,199]. Separately, some studies consistently showed how a reduction in extracellular Ca²⁺ beyond a certain point accelerated cellular transformation [193,200,201,202]. Moreover, reduction in cell adhesiveness, enhancement in cell motility, and increased leakage of proteolytic enzymes in cancer cells have been attributed to the loss of cellular Ca²⁺ [203,204,205]. To complement the in vitro findings, various epidemiological studies have yielded evidence for the inverse correlation between intake of dietary or supplementary Ca²⁺ and cancers of colon, rectum, breast, gastric tract, endometrium, renal system, and ovaries [206]. Although the above-mentioned evidence on the differential levels of Ca²⁺ ion between normal and tumor cells would suggest this cation to have a tumor-inhibitory effect, newer studies indicate a rather complex dynamic [207,208,209]. In this section, we will bring to light multiple ways in which cancer cells manipulate intracellular [Ca²⁺]_i levels and the associated molecular machinery during different stages of tumor progression. Each subsection will provide an overview of studies supporting distortion of signaling in a specific intracellular Ca²⁺ pool, thereby explaining the mechanisms underlying various cancer hallmarks—excessive proliferation, inhibition of growth suppressors, activation of invasion and metastasis, replicative immortality, induction of angiogenesis, resistance to cell death, dysregulated cellular metabolism, and immune surveillance evasion [210]. Additionally, the tables summarize the role of individual Ca²⁺ signaling components described earlier during each of the hallmark cellular processes involved in tumorigenesis and cancer progression. They also provide a corresponding list of potential therapeutic drugs targeting Ca²⁺ signaling proteins under preclinical conditions.

3.1. Intracellular Ca²⁺ Pool in the Endoplasmic Reticulum and at the ER-PM Junction

Because of the substantial observed association of aberrant expression or dysfunction of ion channels with tumor initiation and progression, cancer is at times termed as “oncochannelopathy” [211]. Multiple Ca²⁺ channels expressed on the plasma membrane are functionally altered in cancer cells [210,211,212,213] including voltage-gated Ca²⁺ channels (VGCCs), ligand-gated channels, TRP channels, and ORAI channels. Bioinformatic analysis has revealed abundant expression of genes encoding most VGCCs (L-, R-, N-, P/Q-, and T-type) in cancer tissues [214,215,216,217,218]. Accordingly, CACNA1G encoding the pore-forming alpha subunit of T-type VGCC, Cav 3.1, is expressed in lung adenocarcinoma (A549), colon cancer (HCT116), breast cancer (MCF-7, MDA-MB-231), ovarian cancer (A2780), and melanoma [208,215,217,218,219,220,221,222]. Similarly, Cav 1.3 (CACNA1D encoded L-type VGCC) is shown to be upregulated in breast, endometrial, prostate, neuroblastoma, and colorectal cancer biopsies [217,223]. Consequently, blocking VGCC activity either chemically (BK10040 and KYSO5090; T-type channel blockers or Verapamil; L-type blocker) or by gene silencing is reported to inhibit growth, induce death, or reduce migration of tumor cells by causing a drop in intracellular Ca²⁺ [217,223,224,225,226,227].

The somewhat controvertible, but the generally accepted, role of ligand-gated Ca²⁺ channels in cancer cells is anti-inflammatory/protumorigenic for P1 receptors and proinflammatory/anti-tumorigenic for P2 [228,229,230]. Due to higher levels of ATP in tumors than normal tissues, purinergic receptors, P2X and P2Y, are the most extensively studied ligand-gated Ca²⁺ channels in tumors [231]. ATP binding to P2X receptors results in increased Ca²⁺ influx through them while P2Y receptor (G-protein coupled) activation by the nucleotide enhances ER Ca²⁺ release or cAMP production [231,232]. The receptor expression levels and related Ca²⁺ signaling are both altered during growth factor-induced epithelial–mesenchymal transition (EMT) [232,233]. For instance, EGF-mediated EMT in MDA-MB-468 breast cancer cells is reported to reduce the sensitivity of P2Rs to ATP binding with simultaneous downregulation of P2Y13 and upregulation of P2X5 and P2Y6 transcript levels. ATP-mediated increase in [Ca²⁺]_i can exert anti-migratory and antiangiogenic effects on tumor-derived endothelial cells. Two separate studies have shown that treatment of Breast Tumor-derived Endothelial cells (BTEC) and Renal Tumor-derived Endothelial Cells (RTEC) with 100 mM ATP leads to a biphasic increase in [Ca²⁺]_i—an initial transient release from ER followed by SOCE and then a prolonged release rise due to Ca²⁺ influx via P2X7R and P2Y11R [234,235]. This raised [Ca²⁺]_i then activates adenylate cyclase10/cAMP/EPAC-1 pathway to disrupt actin cytoskeleton and thus, inhibit migration of tumor-derived endothelial cells.

The modification of Ca²⁺ ion efflux machinery at the ER-PM junction during tumor formation or progression also requires attention. PMCA and Na⁺/Ca²⁺ Exchanger (NCX) lead a concerted effort on a normal cell surface to expel excessive free cytosolic Ca²⁺ ions and thereby prevent cytotoxicity [236]. Functionally, it is logical to consider deactivation or negligible expression of the efflux machinery in cancer cells so that high intracellular Ca²⁺ is available to drive tumor proliferation and metabolism [41,237,238]. Such has been observed in colon cancer where PMCA4 mRNA levels were found to be significantly lower in high-grade colon adenocarcinoma, lymph node metastasis, and benign tumors relative to healthy tissue [36,239,240]. Using the HT29 colon cancer cell line, Aung et al. showed that PMCA4 overexpression indeed minimizes cell proliferation in colon cancer [239]. Moreover, PMCA4 expression is observed to be induced during differentiation of colon cancer (HT29), neuroblastoma (IMR-32), and breast cancer (MCF7) [36,241,242,243]. By contrast, pancreatic cancer cells that exhibit high glycolytic over mitochondrial metabolism need PMCA for cell survival [244]. PMCA4 function in these PDAC (pancreatic duct adenocarcinoma) cell lines relies on ATP derived from glycolysis. Therefore, inhibiting ATP production by disrupting glycolysis stops PMCA driven Ca²⁺ ion efflux resulting in excessive Ca²⁺ filling and cell death [245]. Expression of PMCA4 mRNA is muted in breast cancer cell lines too. However, expression of its splice variant PMCA4b is complex due to distinctive regulation by histone deacetylase inhibitors and ERα [243,245,246]. As an example, PMCA4b exhibits low expression with high sensitivity to HDAC inhibitors in MCF-7 (ER-positive cell line), but high expression and low sensitivity to 17β estradiol or HDAC inhibitors in MDA-MB-231 cells (triple-negative breast cancer) [245]. Kenealey et al. have also shown that inhibition of PMCA activity (likely PMCA1 and 4) by resveratrol induces programmed cell death in MDA-MB-231 cells [241].

The Ca²⁺ antiporter NCX1 is the most widely distributed and well-studied isoform within its family. Its mRNA and protein expression levels are dramatically reduced in renal cancer cells and nephroblastoma, conferring several advantages [247]. Knockdown of NCX1 in MDCK cells induces EMT via the Ca²⁺-dependent ERK signaling activation. However, the ability of NCX1-knockdown kidney cells to grow in an anchorage-independent manner and their increased junctional permeability is independent of the Ca²⁺ transport function of the exchanger [248]. In penile tumors, knockdown of NCX1 lifts the brakes on proliferation and reduces apoptosis [247,249]. For therapy-resistant medulloblastoma and ovarian cancer, on the other hand, knockdown of NCX1 results in sensitization of these cancers to ionizing radiation and cisplatin [250,251]. Similarly, OSW-1, a natural saponin, and potential anticancer treatment blocks NCX activity in acute leukemia cell line (HL-60) and induces cytotoxicity via accumulation of excess Ca²⁺ ion in the cytosol [252]. In cancer cells exposed to hypoxia, the reverse mode NCX functioning is coupled to carbonic anhydrase IX (CA1X) and sodium-hydrogen exchanger (NHE1) for converting the intracellular proton load (occurring due to metabolic changes during hypoxia) into interstitial acidosis [253,254]. This allows the breakdown of the extracellular matrix and consequently, promotes tumor cell migration and invasion [253,255].

SOCE is central to intracellular Ca²⁺ signaling as it forms the major Ca²⁺ ion influx route in non-excitable cells [256]. Conceivably, the expression and activity of SOCE components are remodeled during each stage of cancer progression. However, the pattern of SOCE alteration for advancing cancer hallmarks is complex because it is based on the cancer cell type, progression stage, and isoforms of participating components [257,258,259]. As discussed earlier, STIM1 and ORAI1 are the “classical SOC” channel components with STIM2, ORAI2, and ORAI3 taking the center stage in selective cases for SOCE-mediated Ca²⁺ influx [257,260]. Evidence from pharmacological and molecular inhibition studies underscores the role of SOCE in cell cycle progression [257,261,262,263,264,265]. Store-operated Ca²⁺ entry is elevated during G1/S transition but decreased during entry into the G2/M phase [266]. Diminishing SOCE via STIM1 inhibited the proliferation of cervical cancer, glioblastoma (U251), osteosarcoma (143B and U2OS), lung carcinoma (A549 and SK-MES-1), and breast cancer (MCF7) [265,266,267,268,269,270,271,272]. Knockdown of STIM1 in cervical cancer (SiHa and HeLa) caused inhibition of CDK2 phosphorylation, increase in cyclin inhibitors p21 and p27, and accumulation of Cyclin E [272]. In hepatoma cell lines (Huh-7 and HepG2), simultaneous knockdown of STIM1 and ORAI1 dropped protein levels of cyclin D1, thus causing G0/G1 cell cycle arrest [272]. Various studies have also shown that ORAI3-actuated SOCE in MCF7 upregulates c-myc/NFAT/p-ERK axis to increase in CDK2/4, cyclins D1 and E, and promote G1/S transition [266,273,274]. Interestingly, STIM2 upregulation in melanoma cells has been known to contribute to antiproliferative but invasive phenotype [275]. Modulation of SOCE components and the associated Ca²⁺ uptake by cancer cells also influences programmed cell death. Based on the cell type and stimuli, SOCE can either aid apoptosis or provide apoptotic resistance in cancer cells [276,277,278,279,280]. The pivotal proapoptotic role of STIM1/ORAI1 was first noted in pancreatic cancer where downregulation of ORAI1 or ectopic expression of its dominant-negative mutant reduced the susceptibility of tumor cells to apoptotic stimuli [280]. On the other hand, overexpression of ORAI1 in androgen-independent prostate cancer cells can reinstate the normal level of apoptosis. The proapoptotic role of STIM1/ORAI1-based SOCE is reversed in non-small cell lung cancer (A549), ovary carcinoma (A2780), pancreatic adenocarcinoma (HC67, Panc1, Capan1, ASPC1, and MiaPaca2), multiple myeloma (immortalized human cell lines and patient-derived tumor cells), and melanoma (B16BL6-8) with the general accompaniment of elevated Akt pathway activity [277,281,282,283,284,285]. ORAI1-based Ca²⁺ entry also slows down the rate of CD95 mediated apoptosis in leukemic T cells [286]. Moreover, STIM2, found abundantly in colon cancer cells, can elicit an anti-apoptotic effect on such tumor cells [287]. Likewise, ORAI3 overexpression in ER-positive T47D cell line confers apoptotic resistance to cisplatin, paclitaxel, or other chemotherapeutics [288].

Unlike apoptosis, all STIM and ORAI isoforms exert a uniform positive effect on tumor migration and invasiveness [276]. Studies on human cervical cancer have revealed that patients with STIM1 upregulated primary tumors have poorer clinical outcomes due to excessive tumors and lymph node metastasis [257,289]. Similarly, STIM1/ORAI1 overexpression in multiple myeloma or upregulation of ORAI1 in esophageal squamous cell carcinoma is associated with poorer progression-free survival [257,269,284]. One of the pioneer studies demonstrating the proinvasive role of STIM1/ORAI1 in breast cancer cells determined that SOCE enhanced turnover of Rac- and Ras-based focal adhesions to increase cancer cell migration [257,290]. A similar role for STIM1/ORAI mediated SOCE has been observed in cervical cancer, hepatocellular carcinoma, renal cell carcinoma, nasopharyngeal cancer, and glioblastoma where this Ca²⁺ influx regulates focal adhesion turnover, cytoskeletal reorganization, and actomyosin-based mechanotransduction [291,292]. Except for melanoma, STIM2 overexpression in cancer cells and their invasiveness are inversely correlated [284,293]. Understandably, therefore, a high STIM1/STIM2 ratio in breast cancer cells combined with increased SOCE correlates with poorer prognosis in patients [276,284].

Increased angiogenesis is another quintessential feature of cancer [210]. Therefore, manipulation of SOCE has been carefully studied in cancer-associated stromal cells, especially endothelial cells [294]. Silencing of STIM1 or ORAI1 in vascular endothelial cells (HUVECs) attenuates cell proliferation and VEGF-triggered Ca²⁺ influx [276,295]. On the other hand, STIM1 in vivo overexpression in cancer cells positively correlates with increased VEGF secretion, endothelial cell proliferation, and thus angiogenesis [296]. Studies have also shown ORAI1 to stimulate in vitro tubulogenesis and in vivo angiogenesis [269]. Immune cells are another integral and active part of the tumor microenvironment. Since SOCE components were first discovered and studied in T cells, it is presumable that their function in cancer-promoting or -killing immune cells has been extensively explored [297,298]. Surprisingly, distortion of the role of SOCE components and thereby, the intracellular Ca²⁺ flux in cancer-related T cells is not well investigated. One of the early studies in this area delineated how downregulation or absence of STIM1 and STIM2 in cytotoxic T lymphocytes (CTLs; a subset of anti-tumor immune cells) prevents the production of cytolytic factors (interferon-γ and TNF-α), thus releasing control on tumor cell engraftment and in vivo tumor growth [299]. Contradictorily, partial inhibition of ORAI1 in a separate study resulted in the killing of CTLs by tumor cells; more research is, therefore, warranted [300].

IP3Rs, as the principal Ca²⁺ ion release channels for endoplasmic reticulum, are found localized at various sites on the ER membrane: near the SOCE nanodomain, the ER-PM junction outside of the nanodomain, and at the mitochondrial-associated membranes (MAMs) [301,302,303,304]. The functionality of these receptors is determined by their localization and isoform type. Receptors expressed in the ER-PM junction participate in the initiation and propagation of SOCE, whereas the receptors expressed at MAMs regulate ER-mitochondrial Ca²⁺ ion movement, thereby influencing cellular bioenergetics and apoptosis [165]. As our focus here is on the ER-PM junction, the role of IP3Rs at MAMs will be discussed in the latter part of this article.

IP3Rs (and the smooth muscle counterparts, RyRs) control cellular functions like contractility, cellular motility, and migration [305]. Although the research on the role of IP3Rs in cell migration is still new, studies, in general, support a promigratory effect [305,306]. A migrating cell has polarized front and rear ends that differ in the cytoskeletal arrangement and Ca²⁺ ion gradient. The repetitive attachment-detachment of leading and trailing edges during cell migration are accompanied by Ca²⁺ ion oscillations. As a result, migrating tumor cells should have a strong dependency on Ca²⁺ channels [307,308,309]. This hypothesis is validated by Baljinnyam et al. via their study demonstrating that the activation of PLCε/IP3/IP3R in melanoma cell lines (Mel-2 and SK-Mel-24) increases cell migration; the resulting Ca²⁺ ion flux promotes interaction between S100A4 (Ca²⁺-activated invasion protein) and myosin light chain kinase II which rearranges the actin cytoskeleton via EPAC/cAMP-induced Ca²⁺ elevation [305,310]. IP3R3 as well has been found to cause peritoneal dissemination of gastric cancer [305,311]. Ano1, or TMEM16A, a Ca²⁺-activated chloride channel (CaCC) at the ER-PM junction, is highly upregulated in several cancer types—gastrointestinal tumors, head and neck squamous carcinoma, glioblastoma, pancreatic, breast, and colorectal cancer— as a pro-proliferative and pro-migratory signaling molecule [312,313,314,315,316,317]. IP3Rs are now known to tether Ano1 to ER-PM junctions, provide localized Ca²⁺ ion for their activation, and link the opening of CaCCs to SOCE augmentation along with GPCR stimulation [318,319].

By maintaining intracellular Ca²⁺ homeostasis, voltage-independent Ca²⁺ channel family known as Transient Receptor Potential or TRP channels influence various Ca²⁺ ion-sensitive downstream effectors of cellular processes like proliferation, motility, and apoptosis [320,321]. TRP channels having a pro-proliferative effect in cancer cells are mostly derived from TRPC (5, 6), TRPM (2, 4, 5, 7, 8), and TRPV (1, 2, 6) subfamilies [322,323]. TRPC5 downregulation reduces growth in adriamycin-resistant breast cancer cells due to concomitant decline in drug resistance imparting P-glycoproteins [324]. TRPV1, TRPV2, and TRPM2 have been observed to stimulate the proliferation of prostate cancer [325]. Moreover, overexpression of TRPV2 or upregulation of TRPV6 positively correlates with poor prognosis of esophageal squamous cancer and malignant transformation in leukemia or other tumors, respectively [322,326,327,328]. Interestingly, TRPV channels trigger cancer cell proliferation based on cellular subtypes. For example, TRPV1 is expressed in several breast cancer cell lines, peculiarly is found to inhibit cellular proliferation in triple-negative breast cancer [329]. Proapoptotic members of TRP channels largely belong to TRPC, TRPM, and TRPV subfamilies [322]. TRPC1 in conjunction with STIM1 has been recently shown to induce cisplatin cytotoxicity in non-small cell lung cancer cells via reactive oxygen species and DNA damage response [280,330]. In addition, TRPC1 and TRPC4 are postulated to sensitize triple-negative breast cancer to various chemotherapeutics [331]. Among the TRPM members, TRPM2 activation with subsequent Ca²⁺ ion elevation causes oxidative stress and apoptotic death in MCF7 and CaCo-2 cell lines treated with 5-fluorouracil and leucovorin [332]. Conversely, TRPM8 protects pancreatic cancer cell lines Panc-1 and BxPC3 from gemcitabine by maintaining the expression of multi-drug resistance proteins like P-glycoprotein [333]. TRPV1 and downstream Ca²⁺ ion signaling get activated in breast cancer MCF7 cells and glioblastoma treated with TRPV1 positive allosteric modulator MRS1477 or capsaicin, respectively, to trigger cell death [334,335]. Members of the TRP family are also involved in mechanotransduction that allows the cancer cell to invade and metastasize [336]. TRPM7 is the best-studied example in this context with its expression in nasopharyngeal and pancreatic cancer related to poor prognosis [337,338]. This non-selective Ca²⁺ ion-permeable channel forms a mechanosensory complex in breast cancer cells [336,339]. Thus, silencing TRPM7 in ER-positive MCF7 and triple-negative breast cancer MDA-MB-231 redistributes filamentous actin in cell cortices, raises the number of focal adhesions due to rearrangement of local Ca²⁺ ion levels, and phosphorylates myosin light chain and paxillin [339]. Amongst the TRPV subfamily, TRPV4 is significantly upregulated during breast cancer metastasis, and in the clinical samples with poor overall survival has been observed [340,341].

By dint of its central role in ER Ca²⁺ replenishment, SERCA pumps are essential for proper protein folding and maturation and other functions required for cancer cell proliferation and survival [342,343]. Therefore, various cancer types such as colon, lung, and prostate carcinomas are rife with mutations and changes in SERCA expression levels (expression datasets of human cancer cell lines) [343,344,345,346,347,348,349]. On the other hand, SERCA3 expression is induced during differentiation but negatively correlated to colon cancer progression (DLD-1, COLO-205, and Caco-2) [345,349]. Following suit, high expression of SERCA3 is observed in differentiated forms of gastric cancer and myeloid leukemias [350,351]. SERCA pumps are also recognized as regulators of Notch1 receptors that are common drivers of tumor proliferation, especially for leukemia [352]. As a result, SERCA inhibition blocks the activity of mutated Notch1 receptors (by impairing protein folding) which leads to G0/G1 arrest in leukemia cells. SERCA isoforms are required even for the survival of cancer stem cells (CSCs) against metabolic stress [353]. In a study utilizing metabolic stress-resistant breast cancer CSCs derived from MCF-7 and MDA-MB-231 cell lines, SERCA expression was induced in a calmodulin kinase 2α (CaMK2α)/NFκB-dependent manner to prevent stress elicited by 2-deoxy-glucose treatment. Treatment of such CSCs with a combination of 2-deoxy-glucose and thapsigargin, however, sensitized the cells to apoptosis via cytosolic Ca²⁺ ion overload. To prevent unnecessary cytotoxicity induced by thapsigargin in normal cells, its soluble prodrug, G-202 or mipsagargin, was designed and tested in a Phase II study against sorafenib-resistant hepatocellular carcinoma (HCC) [354]. In the study, mipsagargin was shown to stabilize tumor progression by specifically targeting PSMA (prostate-specific membrane antigen) expressing endothelial cells that form the HCC-associated vasculature. It has also been tested against glioblastoma, prostate cancer, and renal cell carcinoma; however, results of those trials have not yet been published.

The anomalous signaling, localization, and mobilization of Ca²⁺ ions seen in tumor cells are partly attributed to peculiar expression levels of calmodulin (CaM) and its target proteins [355]. It is a common observation for tumor cells to express higher levels of CaM which complements the raised levels of cytosolic Ca²⁺ ions relative to benign tissues [356,357,358,359,360]. Although it was contended that this observation was cell type-, culture condition-, or transformation agent-specific, Wang et al. (1992) confirmed a significant increase in CaM levels at the individual cell level between transformed and normal cells during G1 to S phase transition [361]. It was further validated that the surge in CaM levels was specific to cellular transformation and not due to a rise in total intracellular protein during cell cycle progression. Inhibition of CaM by the naphthalensulfonamide-derived selective CaM antagonists W-7 and W-13 caused p21cip1-dependent growth arrest and apoptosis of multiple myeloma tumors in a mouse xenograft model [362]. Some of the protumorigenic effects of calmodulin are facilitated through its binding to the Ca^2+/calmodulin-stimulated protein kinase (CaMK) family that modulate cell cycle progression by interacting with phase-specific cyclins and cyclin-dependent kinases [363]. For instance, CaMKI allows G1/S transition via phosphorylation and activation of cdk4 while CaMKII triggers metaphase to anaphase progression by stimulating cdc2 [364,365,366]. Therefore, pharmacological (STO-609, KN-62, KN-93, berbamine, etc.) and siRNA inhibition of CaMKs have demonstrated anti-proliferative effects in numerous cancer cell lines [367]. Direct interaction between CaM and nuclear hormone receptors is another exploitable mechanism for limiting tumor proliferation. This is exemplified in studies where the growth of estrogen receptor- and androgen receptor-positive cancer cell lines, MCF7 and LnCaP, respectively, is inhibited by CaM antagonists. In such cell lines, CaM antagonists disrupt stabilization and activation of estrogen receptor and androgen receptor, both of which are dependent upon direct interaction with CaM [368,369,370]. The functional interdependencies of calmodulin and numerous [Ca²⁺]_i transporters too can be leveraged for killing cancer cells. It is well known that calmodulin regulates the slow inactivation of CRAC currents by direct interaction with the STIM1 SOAR domain, thereby disrupting the STIM1-ORAI1 complex [371]. It has also been established that calmodulin-binding is required for IP3R activation [372]. On the other hand, CaMKII potentiates SOCE by promoting STIM1-ORAI1 complex formation [373]. CaM and CaMKs are not only regulators of SOCE, but also form integral components of its downstream signaling cascade. This is well evident by their activation of Akt, ERK1/2, and Raf/Pyk2 (via cytosolic Ca²⁺ elevation) for tumor survival, growth, migration, and invasion [374,375,376,377,378].

Calreticulin (CRT) and calbindin are other major EF-hand containing Ca²⁺-binding proteins that are altered by cancer cells [379,380,381,382]. Cancer cells gain many advantages over normal tissue by dysregulating calreticulin since it is a critical modulator of Ca²⁺ ion-dependent processes including proliferation, differentiation, cell adhesion, migration, intercellular interactions, immune response, and apoptosis [383,384,385,386,387,388,389,390]. Accordingly, upregulation of CRT expression is recorded in various tumors such as invasive breast carcinoma, high microsatellite instability colorectal carcinoma, squamous cell carcinoma, leukemia, and many more [391,392,393,394,395]. CRT expression also positively correlates with tumor size, grade, and development stage in breast and lung cancers [396,397]. Even more, its high expression levels are commensurate with poor survival in prostate cancer and neuroblastoma patients [398,399]. The effect of CRT on tumor proliferation is cell type-dependent with most cancer cells responding to upregulation of this protein with rapid growth [400,401]. For instance, overexpression of CRT in pancreatic and gastric cancer cells causes remarkable growth of these tumor types while stable knockdown of the same in oral squamous cell carcinoma led to significant G0/G1 arrest with a negative impact on anchorage-independent growth and colony formation. In certain cancers like gastric carcinoma, CRT overexpression is reported to promote VEGF expression as well, thus leading to enhanced angiogenesis, tumor invasiveness, and migration [401]. Under normal conditions, calreticulin is essential for cellular adhesion via integrin-associated Ca²⁺ signaling with the Wnt pathway as the potential downstream effector [402,403]. In contrast, CRT N-terminus expressed on the cancer cell surface is found to be essential for thrombospondin (TSP) mediated invasion and metastasis [404]. Studies have elucidated that the N-terminus of TSP binds to the CRT and low-density lipoprotein receptor co-complex to induce disassembly of focal adhesion kinase, thus reducing cellular adhesion. CRT as a promoter of cancer cell invasion and metastasis has been verified through overexpression (MDCK cells, gastric cancer cell line AGS) and knockdown (HL60 leukemia cell line and J82 bladder cancer cells) studies as well [405,406,407,408]. Strikingly, CRT expressed on the cancer cell surface can engage in immunomodulatory activities and pose a threat to tumor survival. Work from de Bruyn and Bremer et al. revealed that TRAIL-induced apoptosis relocates CRT from ER lumen to cell surface where the Ca²⁺-binding protein is shown to attract dendritic cells and macrophages for phagocytosis [409]. In glioma, an increase in CRT expression likewise correlates with higher radiosensitivity and, thus, a greater rate of apoptosis [410]. In pancreatic adenocarcinoma, cancer cell survival and chemoresistance decrease significantly post CRT knockdown [411]. Very limited research has been conducted on modifications of calbindin activities in cancer cells. This Ca²⁺ ion buffer which has dominant expression in neurons has so far only been found to protect osteosarcoma cells from apoptotic stimuli [412].

3.2. Intracellular Ca²⁺ Pool at ER-Mitochondrial Junction

Aberrant cellular metabolism, a hallmark of a wide variety of tumors, stems from the dysfunction of mitochondrial pathways for ATP production [210]. The shift from oxidative phosphorylation to glycolysis for ATP generation in a subset of cancers is induced by the cumulative effect of mutations in the participating mitochondrial enzymes and Ca²⁺ signaling [413,414,415,416]. Specifically, activities of TCA cycle enzymes, alpha-ketoglutarate, isocitrate dehydrogenase, and pyruvate dehydrogenase rely on proper Ca²⁺ ion uptake by mitochondria. Thus, plummeting Ca²⁺ ion transport from ER to mitochondria in transformed cells portends tumor progression.

ER-mitochondria junctions or Mitochondrial-Associated Membranes (MAMs) are hubs for proteins that are integral to vital processes such as phospholipid synthesis and translocation, mitochondrial Ca²⁺ ion transport, ER stress, ER Ca²⁺ release, mitochondrial morphology regulation, cellular bioenergetics, apoptosis and survival, autophagy, and ROS generation [165]. IP3R, as discussed earlier in this review, is one of the MAM proteins involved in ER to mitochondria Ca²⁺ ion release, and thus crucial for autophagy and apoptosis induction [417,418,419]. Simplistically, inhibition of IP3R (mainly IP3R3) mediated Ca²⁺ ionrelease into mitochondria creates an imbalance in ATP/ADP ratio that derepresses AMPK/mTOR/ULK-1 axis and initiates autophagy flux (based on nutrient starvation) [418]. Notably, abrogation of IP3R activity outside of MAMs perturbs autophagy induction independent of mTOR suppression—no binding of IP3R to Beclin1, a protein required for autophagosome formation, is observed [420]. Conversely, IP3Rs located away from MAM regions of the ER can promote autophagy induction by acting as Ca²⁺ ion leak channels as less Ca²⁺ storage in the ER manifests into poor ion supply for ATP production [418]. Depending on the tumor stage and type, autophagy flux can either cause a cancer cell to thrive or perish [421,422]. By and large, autophagy activation supports tumor suppression during the early stages of cancer development whereas, more advanced tumors utilize it for drug resistance and sustained growth. Autophagy induction due to inadequate ATP production in normal cells halts G1/S cell cycle progression because of an increase in p53/p21 expression and activity [423,424,425]. In cancer cells with mutant p53, however, aberrant proliferation continues post autophagy induction that results in mitotic catastrophe and eventually cell death due to deprivation of mitochondrial metabolites [423,424,425,426,427]. Removal of dysfunctional proteins and mitochondria during autophagy can also prevent tumorigenesis [428]. In highly developed tumors and related stem cells, autophagy induction contradictorily promotes survival from chemotherapeutic (such as 5-FU or bortezomib) assault; competition between a plethora of tumor suppressors and promoters or oncoproteins to interact with IP3R is consistent with the above-stated observation [429,430,431]. Within this context, Bax inhibitor-1 (BI-1) and Bcl2, both located on the ER membrane, stymie IP3R mediated Ca²⁺ ion release into the mitochondria, ergo inducing autophagy [432,433,434]. Inversely, tumor suppressors such as Beclin-1 prevent inhibition of IP3R-mediated Ca²⁺ ion release into the mitochondria partly by interacting with Bcl2 which then dissociates from IP3R, thereby restoring the ER-mitochondrial Ca²⁺ ion transfer [420]. Chemotherapeutics like Arsenic trioxide also increase this Ca²⁺ ion transport by upregulating the expression of tumor suppressor PML (promyelocytic leukemia) that blocks phosphorylation and inactivation of IP3R by p-Akt [418,435,436].

The ER-mitochondria Ca²⁺ ion transport has emerged as an integral component of the crosstalk between autophagy and apoptosis [437]. Inhibition of autophagy or excessive mitochondrial Ca²⁺ ion uptake via MAMs triggers intrinsic apoptosis pathways. Thus, tumor suppressors and proapoptotic molecules like BRCA1 or PTEN bind to IP₃R to boost Ca²⁺ ion uptake at MAMs while tumor promoters and anti-apoptotic molecules such as Bcl2 or Bcl-xL repress mitochondrial Ca²⁺ ion overload by binding to and inhibiting the channel activity [438,439]. VDAC1 and GRP75, members of a trio comprising IP3R and forming a Ca²⁺ ion transporting complex at MAMs are also exploited by cancer cells [440,441]. It is noteworthy that the impact of VDAC1 manipulation on cancer cell bioenergetics and survival stretches beyond its role in the trio [442,443]. Protumorigenic proteins, hexokinase I and II (HKI and HKII) partner with VDAC1 on the outer mitochondrial membrane to utilize its nucleotide shuttling property (ATP/ADP) for generating high energy storage forms like glucose-6-phosphate and fuel rapid cell proliferation via glycolysis [442,444]. Aside from this, Bcl2, Bcl-xL, or hexokinase bind to the N-terminus of VDAC1 and prevent the release of cytochrome C through a VDAC1 multimer pore that assembles in response to apoptotic stimuli [445,446,447]. At the inner mitochondrial membrane, the Ca²⁺ uptake channel, MCU with its regulators MICU1/2 act as the key mediators of Ca²⁺-based mitochondrial functions [448,449]. Because MICU1/2 limits Ca²⁺ ion uptake via MCU and its absence causes cell death via mitochondrial overload, tumor cells are widely observed to have elevated levels of these proteins, although expression of MCU is subtype-dependent [172,450]. For example, in invasive breast carcinoma such as Triple-Negative Breast Cancer, upregulated MCU expression is critical for xenograft size, lymph node metastasis, and lung infiltration [451,452]. On the other hand, downregulation of MCU can negatively impact breast cancer cell size and motility [452].

3.3. Intracellular Ca²⁺ Pool at ER-Lysosome Junction

Ca²⁺ signaling at lysosomal membranes and surfaces of acidic vesicles provides a functional scaffolding for endocytic traffic and autophagy, thereby directly influencing cellular health [453,454].

Alteration of such signaling in cancer contributes to cancer hallmarks such as uninhibited growth, angiogenesis, and metastasis [455,456,457]. TPC1 and TPC2 are considered the drivers of lysosomal Ca²⁺ signaling [458]. Initial research in a mouse model of B16 melanoma cells and xenografts established the role of these Ca²⁺ ion release channels in tumor invasiveness [459]. The study found that VEGF treatment caused NAADP-mediated, sustained Ca²⁺ ion signals via TPCs on lysosomal membranes, and this promoted G0/G1 cell cycle transition, tumor vascularization, focal adhesion kinase (FAK) formation, and migration. Both in vitro and in vivo silencing or pharmacological inhibition of TPC1/2 have also been shown to block invasion and migration of urinary bladder carcinoma (T24), hepatoma (HUH7), and mammary carcinoma [189,460]. Nguyen et. al used a mouse mammary carcinoma model and silenced TPC expression to demonstrate reduction in lung metastasis. The delineated mechanism involved a downstream failure in the trafficking of β1-integrin to plasma membrane that further prevented phosphorylation of FAK, Src, and vinculin, and thus lamellipodia formation [460]. This complements the previously established role of lysosomes in enhancing tumor migration via cathepsin-based disintegration of extracellular matrix [189,461].

TRPML1 is another Ca²⁺ ion release channel expressed on lysosomal membranes and is known to finely control autophagy [458,462]. Presently, there is low evidence of its involvement in cancer progression, but the expression of its downstream Ca²⁺ activated transcription factor, TFEB is noticeably correlated with cellular malignancy [189,463,464,465,466]. Nutrient starvation and ROS production can activate TRPML1 mediated Ca²⁺ ion release followed by calcineurin activation that subsequently dephosphorylates TFEB and releases it from 14-3-B-B; free TFEB is then translocated to the nucleus to evoke transcription of autophagy-related genes [467]. In short, cancer cells benefit by mai17ntaining dephosphorylated TFEB levels. Supporting this hypothesis, non-small cell lung carcinoma patients with the poor outcome often have TFEB (dephosphorylated) overexpression with simultaneous upregulation of other lysosomal markers like LAMP2a and cathepsin D [468]. TFEB upregulation is also linked to higher invasiveness in colorectal cancer cells [469]. Although a less known function of TFEB, DNA repair triggered by this protein is exploited by Triple-Negative Breast Cancer cells for chemoresistance against doxorubicin [470].

4. Conclusions and Discussion

Intracellular Ca²⁺ signaling involves a smorgasbord of proteins that chelate or transport Ca²⁺ ions across various cellular compartments and thereby assist in signal induction, relay, or integration. With Ca²⁺ signaling being pivotal for vital cellular processes, distortion of this machinery can be highly advantageous to cancer cells. Although there has been a steady upward trend in the number of published studies that delineate the functional exploitation of various intracellular Ca²⁺ signaling components in tumors, limited headway has occurred in translating those research outcomes into clinical applications. To demonstrate this, we first utilized the PubMed database to get a list of all the publications within the realm of intracellular Ca²⁺ in cancer. Using the advanced search feature, we selected “intracellular Ca²⁺” and “cancer” as the keywords to look for in all fields (author, title, abstract, MeSH words, journal name, etc.) of PubMed articles. We got more than 11,000 studies (not shown). As the published articles between 1960 and 1990 only accounted for less than 0.1 percent of the total, we set our year range for subsequent searches as 1990–2020. To minimize the occurrence of non-relevant articles in our dataset (that is, studies only focused on one of the two keywords and not both), we used stricter filtration criteria [Figure 10A]. Based on the distribution pattern of articles by the year and the overall categorization per major intracellular Ca²⁺ molecules, Revised Filter 2 was determined to be a sub-set of Revised Filter 1 [Figure 10B–D]. A notable difference between the two filters in terms of sorting of articles by the intracellular Ca²⁺ signaling molecules in focus was that Revised Filter 2 had a lower percentage of articles on “CaSR or Vitamin D” amongst all the ones with main Ca²⁺ signaling molecules from the endoplasmic reticulum or its interorganellar junctions [Figure 10C,D; from “Calmodulin” up to “Other ER” categories] in Revised Filter 2 (6% (30/454)) than Revised Filter 1 (19% (394/2061)). Therefore, it seemed that the search criteria used in Revised Filter 1 captured most studies relevant to our search without being too stringent; we chose to use the list of articles generated from Revised Filter 1 for subsequent analysis.

Clinical studies that were extracted from the list of articles in Revised Filter 1 were labeled in PubMed with one or more of these terms— Clinical Study, Clinical Trial, Clinical Trial Phase I, Clinical Trial Phase II, Clinical Trial Phase III, Clinical Trial Phase IV, Comparative Study, Controlled Clinical Trial, Meta-Analysis, Multi-Center Study, Randomized Controlled Trial, Systematic Review. Two-hundred-and-seventy-seven clinical studies were identified with most of them concentrated between year ranges 1991 to 1996 and 2010 to 2017 (with approximately 10 or more studies published each year) [Figure 11A]. With only 0.1% (277 out of 2941) of the clinical studies for over 30 years, it is interesting to observe that most of such research has explored the role of dietary Ca²⁺ and Vitamin D on cancer (31%) [Figure 11B]. Translational research on any other key Ca²⁺ signaling player in the endoplasmic reticulum or its interorganellar junctions accounted for less than 10 percent of all the published clinical articles. We further evaluated from the main set of published articles the number of studies that used antineoplastic agents (mainly chemotherapeutics) by searching for appropriate keywords. There are 819 articles in the main set that had utilized antineoplastic agent(s) to address a hypothesis related to the role of a major Ca²⁺ signaling molecule in cancer [Figure 11C]. Out of those studies, there are 215 ER-related and 247 that have mitochondrial Ca²⁺ signaling molecules as major components of research. Studies with bisphosphonates and not chemotherapeutics were predominant in the antineoplastic group of articles followed by the ones that utilized dietary Ca²⁺ and/or Vitamin D [Figure 11D]. Among the remaining articles, hormonal, phytogenic or natural, and organometallic agents were the most popular anticancer chemotherapeutics to be used. The above conclusions have been derived with the caveat that PubMed is not an exhaustive database of biomedical research. It is nevertheless surprising to note the lack of clinical development of anticancer therapeutics targeting Ca²⁺ signaling.

Research using small molecule drugs targeting numerous Ca²⁺ signaling proteins is currently in the early stages. Some of the examples include RP4010 (ORAI1 inhibitor; Phase I/IB clinical study terminated), Synta66 (SOCE inhibitor; preclinical), CAD204520 (SERCA inhibitor; preclinical), and SKF96365 (TRP channel and SOCE inhibition; preclinical) [471,472,473,474,475,476]. The time-intensive process (median time 7.3 years) of drug development that simultaneously demands heavy intellectual and monetary investments (roughly $648 million per drug in R&D) could be a possible reason for this gap [477]. Toxicity associated with targeting ubiquitously expressed proteins, the heterogeneity within populations of cancer cells, and the development of multidrug resistance are other critical barriers to be overcome [478,479,480,481]. The effectiveness of small molecule inhibitors of Ca²⁺ signaling could be increased by leveraging the advancements made in the nanocarrier-based targeted drug delivery systems. For example, taking from the design of Antp-LP4 related peptides (VDAC1 inhibitors), Venetoclax (a Bcl2 inhibitor) can be encapsulated in inorganic (such as gold nanoparticle or quantum dots) or organic nanocarriers (such as liposomes) coated with transferrin to specifically target cancer cells that demonstrate significant surface expression of transferrin receptors [445,482,483]. However, some studies have also highlighted the independent effect of nanoparticles themselves on the regulation of Ca²⁺ signaling [479]. Drug repurposing is another way to find effective therapeutic modalities targeting Ca²⁺ signaling in cancer. Several FDA-approved anticancer agents (such as 5-fluorouracil, cisplatin, tamoxifen, paclitaxel, and doxorubicin) and drugs against other disease conditions (such as leflunomide, tolvaptan, and teriflunomide) can impact Ca²⁺ signaling machinery [484,485]. Some of these drugs, formulated as prodrugs or nanocarrier loads, are under investigation as antagonists of cancer-promoting Ca²⁺ signaling See Figure 12 [46,226,227,252,440,441,486,487,488,489,490,491,492,493,494,495,496,497,498,499,500,501,502,503,504,505,506,507,508,509,510,511,512,513,514,515,516,517,518,519,520,521,522,523,524,525,526,527,528,529,530,531,532,533,534,535,536,537,538,539,540,541,542,543,544,545,546,547,548,549,550,551,552,553,554,555,556,557,558,559,560,561,562,563,564,565,566,567,568,569,570,571,572,573,574,575,576,577,578,579,580,581,582,583,584,585,586,587,588,589,590,591,592,593,594,595,596,597,598,599,600,601,602,603,604,605,606,607,608,609,610,611,612,613,614,615,616,617,618,619,620,621,622,623,624,625,626,627,628,629,630,631,632,633,634,635,636,637,638,639,640,641,642,643,644,645,646,647,648,649,650,651,652,653,654,655,656,657,658,659,660,661,662,663,664,665,666,667,668,669,670,671,672,673,674,675,676,677,678,679,680,681,682,683,684,685,686,687,688,689,690,691,692,693,694,695,696,697,698,699,700,701,702,703,704,705,706,707,708,709,710,711,712,713].

Even after decades of research on intracellular Ca²⁺ in tumor cells, the vast clinical potential of targeting key players in this area remains underappreciated. Whether it is through redesigning existing therapeutics or developing novel treatments for directly modulating the intracellular Ca²⁺ levels, greater efforts are needed to find ways to reverse the hijack of intracellular Ca²⁺ signaling in cancer.

Author Contributions

Conceptualization, R.C.E. and A.S.; analysis, A.S.; writing-review and editing, A.S., G.T.R. and R.C.E.; visualization, A.S.; supervision, R.C.E. All authors have read and agreed to the published version of the manuscript.

Funding

RCE was funded by an Excellence in Academic Medicine award from the State of Illinois.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

References

Campbell, A.K. Intracellular Calcium; John Wiley & Sons: Hoboken, NJ, USA, 2014. [Google Scholar] [CrossRef]
Carafoli, E.; Krebs, J. Why Calcium? How Calcium Became the Best Communicator. J. Biol. Chem. 2016, 291, 20849–20857. [Google Scholar] [CrossRef] [Green Version]
Bagur, R.; Hajnóczky, G. Intracellular Ca²⁺ Sensing: Its Role in Calcium Homeostasis and Signaling. Mol. Cell 2017, 66, 780–788. [Google Scholar] [CrossRef] [Green Version]
Schwaller, B. Cytosolic Ca²⁺ Buffers. Cold Spring Harb. Perspect. Biol. 2010, 2, a004051. [Google Scholar] [CrossRef]
Prins, D.; Michalak, M. Organellar Calcium Buffers. Cold Spring Harb. Perspect. Biol. 2011, 3, a004069. [Google Scholar] [CrossRef] [Green Version]
Chin, D.; Means, A.R. Calmodulin: A prototypical calcium sensor. Trends Cell Biol. 2000, 10, 322–328. [Google Scholar] [CrossRef]
Nelson, M.R.; Chazin, W.J. An interaction-based analysis of calcium-induced conformational changes in Ca²⁺ sensor proteins. Protein Sci. 1998, 7, 270–282. [Google Scholar] [CrossRef] [Green Version]
O’Day, D.H.; Eshak, K.; Myre, M.A. Calmodulin Binding Proteins and Alzheimer’s Disease. J. Alzheimer’s Dis. 2015, 46, 553–569. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Faas, C.; Raghavchri, S.; Lisman, J.E.; Mody, I. Calmodulin as a Direct Detector of Ca²⁺ Signals. Nat. Neurosci. 2011, 14, 301–304. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Villalobo, A.; Hiriaki, I.; Vogel, H.J.; Berchtold, M.W. Calmodulin as a protein linker and a regulator of adaptor/scaffold proteins. Biochim. Biophys. Acta (BBA)—Mol. Cell Res. 2018, 1865, 507–521. [Google Scholar] [CrossRef]
Schmidt, H. Three functional facets of calbindin D-28k. Front. Mol. Neurosci. 2012, 5, 25. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Kawasaki, H.; Kretsinger, R.H. Structural and functional diversity of EF-hand proteins: Evolutionary perspectives. Protein Sci. 2017, 26, 1898–1920. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Rogers, J.H. Calretinin: A gene for a novel calcium-binding protein expressed principally in neurons. J. Cell Biol. 1987, 105, 1343–1353, Erratum in 1990, 110, 1845. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Lugli, A.; Forster, Y.; Haas, P.; Nocito, A.; Bucher, C.; Bissig, H.; Mirlacher, M.; Storz, M.; Mihatsch, M.J.; Sauter, G. Calretinin expression in human normal and neoplastic tissues: A tissue microarray analysis on 5233 tissue samples. Hum. Pathol. 2003, 34, 994–1000. [Google Scholar] [CrossRef]
Marenholz, I.; Heizmann, C.W.; Fritz, G. S100 proteins in mouse and man: From evolution to function and pathology (Including an update of the nomenclature). Biochem. Biophys. Res. Commun. 2004, 322, 1111–1122. [Google Scholar] [CrossRef] [PubMed]
Dudek, E.; Michalak, M. Calnexin and calreticulin. In Encyclopedia of Metalloproteins; Kretsinger, R.H., Uversky, V.N., Permyakov, E.A., Eds.; Springer: Berlin/Heidelberg, Germany, 2013; pp. 555–562. [Google Scholar] [CrossRef]
Ben Johny, M.; Yang, P.S.; Bazzazi, H.; Yue, D.T. Dynamic switching of calmodulin interactions underlies Ca²⁺ regulation of CaV1.3 channels. Nat. Commun. 2013, 4, 1717. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Lambers, T.T.; Weidema, A.F.; Nilius, B.; Hoenderop, J.G.J.; Bindels, R.J.M. Regulation of the mouse epithelial Ca²⁺ channel trpv6 by the Ca²⁺-sensor calmodulin. J. Biol. Chem. 2004, 279, 28855–28861. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Nitsche, J.; Josts, I.; Heidemann, J.; Mertens, H.D.; Maric, S.; Moulin, M.; Haertlein, M.; Busch, S.; Forsyth, V.T.; Svergun, D.I.; et al. Structural basis for activation of plasma-membrane Ca²⁺-ATPase by calmodulin. Commun. Biol. 2018, 1, 206. [Google Scholar] [CrossRef]
Kahl, C.R.; Means, A.R. Regulation of cell cycle progression by calcium/calmodulin-dependent pathways. Endocr. Rev. 2003, 24, 719–736. [Google Scholar] [CrossRef] [Green Version]
Lambers, T.T.; Mahieu, F.; Oancea, E.; Hoofd, L.; de Lange, F.; Mensenkamp, A.R.; Voets, T.; Nilius, B.; Clapham, D.E.; Hoenderop, J.G.; et al. Calbindin-D28K dynamically controls TRPV5-mediated Ca²⁺ transport. EMBO J. 2006, 25, 2978–2988. [Google Scholar] [CrossRef] [Green Version]
Berlin, J.R.; Bassani, J.W.; Bers, D.M. Intrinsic cytosolic calcium buffering properties of single rat cardiac myocytes. Biophys. J. 1994, 67, 1775–1787. [Google Scholar] [CrossRef] [Green Version]
Chandrashekera, P.C.; Kargacin, M.E.; Deans, J.P.; Lytton, J. Determination of apparent calcium affinity for endogenously expressed human sarco(endo)plasmic reticulum calcium-ATPase isoform SERCA3. Am. J. Physiol. Cell Physiol. 2009, 296, C1105–C1114. [Google Scholar] [CrossRef] [Green Version]
Gorski, P.A.; Ceholski, D.K.; Young, H.S. Structure-function relationship of the serca pump and its regulation by phospholamban and sarcolipin. In Membrane Dynamics and Calcium Signaling; Krebs, J., Ed.; Springer: Berlin/Heidelberg, Germany, 2017; pp. 77–119. [Google Scholar] [CrossRef]
Vandecaetsbeek, I.; Trekels, M.; De Maeyer, M.; Ceulemans, H.; Lescrinier, E.; Raeymaekers, L.; Wuytack, F.; Vangheluwe, P. Structural basis for the high affinity of the ubiquitous SERCA2b Ca²⁺ pump. Proc. Natl. Acad. Sci. USA 2009, 106, 18533–18538. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Toyoshima, C.; Nakasako, M.; Nomura, H.; Ogawa, H. Crystal structure of the calcium pump of sarcoplasmic reticulum at 2.6 A resolution. Nature 2000, 405, 647–655. [Google Scholar] [CrossRef] [PubMed]
Zhang, Z.; Lewis, D.; Strock, C.; Inesi, G.; Nakasako, M.; Nomura, H.; Toyoshima, C. Detailed Characterization of the Cooperative Mechanism of Ca²⁺ Binding and Catalytic Activation in the Ca²⁺ Transport (SERCA) ATPase. Biochemistry 2000, 39, 8758–8767. [Google Scholar] [CrossRef] [PubMed]
Inoue, M.; Skuta, N.; Watanabe, S.; Zhang, Y.; Yoshikaie, K.; Tanaka, Y.; Ushioda, R.; Kato, Y.; Takagi, J.; Tsukazaki, T.; et al. Structural Basis of Sarco/Endoplasmic Reticulum Ca²⁺-ATPase 2b Regulation via Transmembrane Helix Interplay. Cell Rep. 2019, 27, 1221–1230. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Anderson, D.M.; Makarewich, C.A.; Anderson, K.M.; Shelton, J.M.; Bezprozvannaya, S.; Bassel-Duby, R.; Olson, E.N. Widespread control of calcium signaling by a family of SERCA-inhibiting micropeptides. Sci. Signal. 2016, 9, ra119. [Google Scholar] [CrossRef] [Green Version]
Zhang, Y.; Inoue, M.; Tsutsumi, A.; Watanabe, S.; Nishizawa, T.; Nagata, K.; Kikkawa, M.; Inaba, K. Cryo-EM structures of SERCA2b reveal the mechanism of regulation by the luminal extension tail. Sci. Adv. 2020, 6, eabb0147. [Google Scholar] [CrossRef]
Dencher, N.A.; Choli, T.; Dresselhaus, D.; Fimmel, F.; Grzesiek, S.; Papadopoulos, G.; Wittmann-Liebold, B.; Büldt, G. Structure-function relationship of the light-driven proton pump bacteriorhodopsin. J. Protein Chem. 1989, 8, 340–343. [Google Scholar] [CrossRef] [PubMed]
A buffering serca pump in models of calcium dynamics. Biophys. J. 2006, 91, 151–163. [CrossRef] [PubMed] [Green Version]
Espinoza-Fonseca, L.M. Probing the effects of nonannular lipid binding on the stability of the calcium pump SERCA. Sci. Rep. 2019, 9, 3349. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Strehler, E.E.; Caride, A.J.; Filoteo, A.G.; Xiong, Y.; Penniston, J.T.; Enyedi, A. Plasma membrane Ca²⁺-ATPases as dynamic regulators of cellular calcium handling. Ann. N. Y. Acad. Sci. 2007, 1099. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Stafford, N.; Wilson, C.; Oceandy, D.; Neyses, L.; Cartwright, E.J. The plasma membrane calcium atp ases and their role as major new players in human disease. Physiol. Rev. 2017, 97, 1089–1125. [Google Scholar] [CrossRef] [Green Version]
Padányi, R.; Pászty, K.; Hegedűs, L.; Varga, K.; Papp, B.; Penniston, J.T.; Enyedi, A. Multifaceted plasma membrane Ca²⁺ pumps: From structure to intracellular Ca²⁺ handling and cancer. Biochim. Biophys. Acta (BBA) Mol. Cell Res. 2016, 1863, 1351–1363. [Google Scholar] [CrossRef] [Green Version]
Lopreiato, R.; Giacomello, M.; Carafoli, E. The plasma membrane calcium pump: New ways to look at an old enzyme. J. Biol. Chem. 2014, 289, 10261–10268. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Pérez-Gordones, M.C.; Ramírez-Iglesias, J.R.; Cervino, V.; Uzcanga, G.L.; Benaim, G.; Mendoza, M. Evidence of the presence of a calmodulin-sensitive plasma membrane Ca²⁺-ATPase in Trypanosoma equiperdum. Mol. Biochem. Parasitol. 2017, 213, 1–11. [Google Scholar] [CrossRef]
Bruce, J.I.E. Metabolic regulation of the PMCA: Role in cell death and survival. Cell Calcium 2018, 69, 28–36. [Google Scholar] [CrossRef]
Strehler, E.E. Plasma membrane calcium atpases as novel candidates for therapeutic agent development. J. Pharm. Pharm. Sci. 2013, 16, 190. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Brini, M.; Carafoli, E. The Plasma Membrane Ca²⁺ ATPase and the Plasma Membrane Sodium Calcium Exchanger Cooperate in the Regulation of Cell Calcium. Cold Spring Harb Perspect Biol. 2011, 3, a004168. [Google Scholar] [CrossRef] [PubMed]
Hilge, M. Ca²⁺ regulation of ion transport in the Na⁺/Ca²⁺ exchanger. J. Biol. Chem. 2012, 287, 31641–31649. [Google Scholar] [CrossRef] [Green Version]
Giladi, M.; Lee, S.Y.; Ariely, Y.; Teldan, Y.; Granit, R.; Strulovich, R.; Haitin, Y.; Chung, K.Y.; Khananshvili, D. Structure-based dynamic arrays in regulatory domains of sodium-calcium exchanger (Ncx) isoforms. Sci. Rep. 2017, 7, 993. [Google Scholar] [CrossRef] [Green Version]
Emery, L.; Whelan, S.; Hirschi, K.D.; Pittman, J.K. Protein phylogenetic analysis of Ca²⁺/cation antiporters and insights into their evolution in plants. Front. Plant. Sci. 2012, 3, 1. [Google Scholar] [CrossRef] [Green Version]
Annunziato, L.; Pignataro, G.; Di Renzo, G.F. Pharmacology of brain Na⁺/Ca²⁺ exchanger: From molecular biology to therapeutic perspectives. Pharmacol. Rev. 2004, 56, 633–654. [Google Scholar] [CrossRef] [Green Version]
Chemaly, E.R.; Troncone, L.; Lebeche, D. SERCA control of cell death and survival. Cell Calcium 2018, 69, 46–61. [Google Scholar] [CrossRef]
Tsai, F.-C.; Seki, A.; Yang, H.W.; Hayer, A.; Carrasco, S.; Malmersjö, S.; Meyer, T. A polarized Ca²⁺, diacylglycerol and STIM1 signalling system regulates directed cell migration. Nat. Cell Biol. 2014, 16, 133–144. [Google Scholar] [CrossRef] [Green Version]
Papin, J.; Zummo, F.P.; Pachera, N.; Guay, C.; Regazzi, R.; Cardozo, A.K.; Herchuelz, A. Na⁺/Ca²⁺ Exchanger a Druggable Target to Promote β-Cell Proliferation and Function. J. Endocr. Soc. 2018, 2, 631–645. [Google Scholar] [CrossRef] [PubMed]
Molinaro, P.; Pannaccione, A.; Sisalli, M.J.; Secondo, A.; Cuomo, O.; Sirabella, R.; Cantile, M.; Ciccone, R.; Scorziello, A.; Di Renzo, G.; et al. A New Cell-penetrating Peptide That Blocks the Autoinhibitory XIP Domain of NCX1 and Enhances Antiporter Activity. Mol. Ther. 2015, 23, 465–476. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Khalil, M.; Alliger, K.; Weidinger, C.; Yerinde, C.; Wirtz, S.; Becker, C.; Engel, M.A. Functional Role of Transient Receptor Potential Channels in Immune Cells and Epithelia. Front. Immunol. 2018, 9, 174. [Google Scholar] [CrossRef] [PubMed]
Schnitzler, M.M.; Wäring, J.; Gudermann, T.; Chubanov, V. Evolutionary determinants of divergent calcium selectivity of TRPM channels. FASEB J. 2008, 22, 1540–1551. [Google Scholar] [CrossRef]
Voolstra, O.; Huber, A. Post-translational modifications of trp channels. Cells 2014, 3, 258–287. [Google Scholar] [CrossRef] [Green Version]
van Goor, M.K.C.; Hoenderop, J.G.J.; van der Wijst, J. TRP channels in calcium homeostasis: From hormonal control to structure-function relationship of TRPV5 and TRPV6. Biochim. Biophys. Acta (BBA) Mol. Cell Res. 2017, 1864, 883–893. [Google Scholar] [CrossRef]
Gees, M.; Colsoul, B.; Nilius, B. The Role of Transient Receptor Potential Cation Channels in Ca²⁺ Signaling. Cold Spring Harb. Perspect. Biol. 2010, 2, a003962. [Google Scholar] [CrossRef] [Green Version]
Gaudet, R. Structural Insights into the Function of TRP Channels (Chapter 25). In TRP Ion Channel Function in Sensory Transduction and Cellular Signaling Cascades; CRC Press: Boca Raton, FL, USA; Taylor & Francis: Abingdon, UK, 2007. [Google Scholar]
Cohen, M.; Moiseenkova-Bell, V.Y. Structure of Thermally Activated TRP Channels. Curr. Top. Membr. 2014, 74, 181–211. [Google Scholar]
Gaudet, R. A primer on ankyrin repeat function in TRP channels and beyond. Mol. Biosyst. 2008, 4, 372–379. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Earley, S.; Brayden, J.E. Transient receptor potential channels in the vasculature. Physiol. Rev. 2015, 95, 645–690. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Clapham, D.E. TRP channels as cellular sensors. Nature 2003, 426, 517–524. [Google Scholar] [CrossRef]
Muller, C.; Morales, P.; Reggio, P.H. Cannabinoid ligands targeting trp channels. Front. Mol. Neurosci. 2019, 11, 487. [Google Scholar] [CrossRef] [PubMed]
Chen, J.; Hackos, D.H. TRPA1 as a drug target—Promise and challenges. Naunyn-Schmiedeberg’s Arch. Pharmacol. 2015, 388, 451–463. [Google Scholar] [CrossRef] [Green Version]
Li, M.; Yu, Y.; Yang, J. Structural biology of trp channels. Adv. Exp. Med. Biol. 2011, 704, 1–23. [Google Scholar] [CrossRef] [Green Version]
Zimova, L.; Barvikova, K.; Macikova, L.; Vyklicka, L.; Sinica, V.; Barvik, I.; Vlachova, V. Proximal c-terminus serves as a signaling hub for trpa1 channel regulation via its interacting molecules and supramolecular complexes. Front. Physiol. 2020, 11, 189. [Google Scholar] [CrossRef]
Eder, P.; Schindl, R.; Romanin, C.; Groschner, K. Protein–protein interactions in trpc channel complexes. In TRP Ion Channel Function in Sensory Transduction and Cellular Signaling Cascades; Liedtke, W.B., Heller, S., Eds.; CRC Press: Boca Raton, FL, USA; Taylor & Francis: Abingdon, UK, 2007. [Google Scholar]
Noyer, L.; Lemonnier, L.; Mariot, P.; Gkika, D. Partners in crime: Towards new ways of targeting calcium channels. Int. J. Mol. Sci. 2019, 20, 6344. [Google Scholar] [CrossRef] [Green Version]
Shin, Y.-C.; Shin, S.-Y.; Chun, J.N.; Cho, H.S.; Lim, J.M.; Kim, H.-G.; So, I.; Kwon, D.; Jeon, J.-H. Trip database 2.0: A manually curated information hub for accessing trp channel interaction network. PLoS ONE 2012, 7. [Google Scholar] [CrossRef]
Shin, Y.-C.; Shin, S.-Y.; So, I.; Kwon, D.; Jeon, J.-H. TRIP Database: A manually curated database of protein–protein interactions for mammalian TRP channels. Nucleic Acids Res. 2011, 39 (Suppl. 1), D356–D361. [Google Scholar] [CrossRef] [Green Version]
Chun, J.N.; Lim, J.M.; Kang, Y.; Kim, E.H.; Shin, Y.-C.; Kim, H.-G.; Jang, D.; Kwon, D.; Shin, S.-Y.; So, I.; et al. A network perspective on unraveling the role of TRP channels in biology and disease. Pflügers Arch. Eur. J. Physiol. 2014, 466, 173–182. [Google Scholar] [CrossRef]
Vangeel, L.; Voets, T. Transient receptor potential channels and calcium signaling. Cold Spring Harb. Perspect. Biol. 2019, 11. [Google Scholar] [CrossRef]
Dong, X.; Wang, X.; Xu, H. Trp channels of intracellular membranes. J. Neurochem. 2010, 113, 313–328. [Google Scholar] [CrossRef] [Green Version]
Pecze, L.; Blum, W.; Henzi, T.; Schwaller, B. Endogenous TRPV1 stimulation leads to the activation of the inositol phospholipid pathway necessary for sustained Ca²⁺ oscillations. Biochim. Biophys. Acta (BBA) Mol. Cell Res. 2016, 1863, 2905–2915. [Google Scholar] [CrossRef] [PubMed]
Thillaiappan, N.B.; Chakraborty, P.; Hasan, G.; Taylor, C.W. IP3 receptors and Ca²⁺ entry. Biochim. Biophys. Acta (BBA) Mol. Cell Res. 2019, 1866, 1092–1100. [Google Scholar] [CrossRef] [PubMed]
Cullen, P.J.; Steinberg, F. To degrade or not to degrade: Mechanisms and significance of endocytic recycling. Nat. Rev. Mol. Cell Biol. 2018, 19, 679–696. [Google Scholar] [CrossRef] [PubMed]
Peinelt, C.; Beck, A.; Monteilh-Zoller, M.K.; Penner, R.; Fleig, A. IP3 receptor subtype-dependent activation of store-operated calcium entry through ICRAC. Cell Calcium 2009, 45, 326–330. [Google Scholar] [CrossRef] [PubMed]
Rao, J.N.; Platoshyn, O.; Golovina, V.A.; Liu, L.; Zou, T.; Marasa, B.S.; Turner, D.J.; Yuan, J.X.-J.; Wang, J.-Y. TRPC1 functions as a store-operated Ca²⁺ channel in intestinal epithelial cells and regulates early mucosal restitution after wounding. Am. J. Physiol. Liver Physiol. 2006, 290, G782–G792. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Haustrate, A.; Prevarskaya, N.; Lehen’kyi, V. Role of the trpv channels in the endoplasmic reticulum calcium homeostasis. Cells 2020, 9, 317. [Google Scholar] [CrossRef] [Green Version]
Firth, A.L.; Yuan, J.X.-J. Ion channels and transporters in the pulmonary vasculature: A focus on smooth muscle. In Textbook of Pulmonary Vascular Disease; Yuan, J.X.-J., Garcia, J.G.N., West, J.B., Hales, C.A., Rich, S., Archer, S.L., Eds.; Springer: Jersey, NJ, USA, 2011; pp. 223–244. [Google Scholar] [CrossRef]
Sukumaran, P.; Schaar, A.; Sun, Y.; Singh, B.B. Functional role of TRP channels in modulating ER stress and Autophagy. Cell Calcium 2016, 60, 123–132. [Google Scholar] [CrossRef] [Green Version]
Lam, A.K.; Galione, A. The endoplasmic reticulum and junctional membrane communication during calcium signaling. Biochim. Biophys. Acta (BBA)—Bioenerg. 2013, 1833, 2542–2559. [Google Scholar] [CrossRef] [Green Version]
Michalak, M.; Robert Parker, J.M.; Opas, M. Ca²⁺ signaling and calcium binding chaperones of the endoplasmic reticulum. Cell Calcium 2002, 32, 269–278. [Google Scholar] [CrossRef] [PubMed]
Berridge, M.J. The endoplasmic reticulum: A multifunctional signaling organelle. Cell Calcium 2002, 32, 235–249. [Google Scholar] [CrossRef] [PubMed]
Burdakova, D.; Petersenb, O.H.; Verkhratskya, A. Intraluminal calcium as a primary regulator of endoplasmic reticulum function. Cell Calcium. 2005, 38, 303–310. [Google Scholar] [CrossRef]
Raffaello, A.; Mammucari, C.; Gherardi, G.; Rizzuto, R. Calcium at the Center of Cell Signaling: Interplay between Endoplasmic Reticulum, Mitochondria, and Lysosomes. Trends Biochem. Sci. 2016, 41, 1035–1049. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Prakriya, M.; Lewis, R.S. Store-Operated Calcium Channels. Physiol. Rev. 2015, 85, 757–810. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Numaga-Tomita, T.; Putney, J.W. Role of STIM1- and Orai1-mediated Ca²⁺ entry in Ca²⁺-induced epidermal keratinocyte differentiation. J. Cell Sci. 2013, 126, 605–612. [Google Scholar] [CrossRef] [Green Version]
Hodeify, R.; Yu, F.; Courjaret, R.; Nader, N.; Dib, M.; Sun, L.; Adap, E.; Hubrack, S.; Machaca, K. Regulation and role of store-operated Ca²⁺ entry in cellular proliferation. In Calcium Entry Channels in Non-Excitable Cells; Kozak, J.A., Putney, J.W., Eds.; CRC Press: Boca Raton, FL, USA; Taylor & Francis: Abingdon, UK, 2018. [Google Scholar] [CrossRef]
Oritani, K.; Kincade, P.W. Identification of stromal cell products that interact with pre-B cells. J. Cell Biol. 1996, 134, 771–782. [Google Scholar] [CrossRef] [PubMed]
Roos, J.; DiGregorio, P.J.; Yeromin, A.Y.; Ohlsen, K.; Lioudyno, M.; Zhang, S.; Safrina, O.; Kozak, J.A.; Wagner, S.L.; Cahalan, M.D.; et al. STIM1, an essential and conserved component of store-operated Ca²⁺ channel function. J. Cell Biol. 2005, 169, 435–445. [Google Scholar] [CrossRef] [Green Version]
Liou, J.; Kim, M.L.; Heo, W.D.; Jones, J.T.; Myers, J.W.; Ferrell, J.E., Jr.; Meyer, T. STIM is a Ca²⁺ sensor essential for Ca²⁺-store-depletion-triggered Ca²⁺ influx. Curr. Biol. 2005, 15, 1235–2141. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Hewavitharana, T.; Deng, X.; Soboloff, J.; Gill, D.L. Role of Stim and Orai proteins in calcium signaling pathways. Cell Calcium 2007, 42, 173–182. [Google Scholar] [CrossRef]
Zheng, L.; Stathopulos, P.B.; Schindl, R.; Li, G.-Y.; Romanin, C.; Ikura, M. Auto-inhibitory role of the EF-SAM domain of STIM proteins in store-operated calcium entry. Proc. Natl. Acad. Sci. USA 2011, 108, 1337–1342. [Google Scholar] [CrossRef] [Green Version]
Stathopulos, P.B.; Zheng, L.; Ikura, M. Stromal Interaction Molecule (STIM) 1 and STIM2 Calcium Sensing Regions Exhibit Distinct Unfolding and Oligomerization Kinetics. J. Biol. Chem. 2009, 284, 728–732. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Brandman, O.; Liou, J.; Park, W.S.; Meyer, T. STIM2 is a feedback regulator that stabilizes basal cytosolic and endoplasmic reticulum Ca²⁺ levels. Cell 2007, 131, 1327–1339. [Google Scholar] [CrossRef] [Green Version]
Soboloff, J.; Spassova, M.A.; Hewavitharana, T.; He, L.-P.; Xu, W.; Johnstone, L.S.; Dziadek, M.A.; Gill, D.L. STIM2 is an inhibitor of STIM1-mediated store-operated Ca²⁺ Entry. Curr. Biol. 2006, 16, 1465–1470. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Vig, M.; Peinelt, C.; Beck, A.; Koomoa, D.L.; Rabah, D.; Koblan-Huberson, M.; Kraft, S.; Turner, H.; Fleig, A.; Penner, R.; et al. Cracm1 is a plasma membrane protein essential for store-operated Ca²⁺ entry. Science 2006, 312, 1220–1223. [Google Scholar] [CrossRef] [Green Version]
Lis, A.; Peinelt, C.; Beck, A.; Parvez, S.; Monteilh-Zoller, M.; Fleig, A.; Penner, R. CRACM1, CRACM2, and CRACM3 Are Store-Operated Ca²⁺ Channels with Distinct Functional Properties. Curr. Biol. 2007, 27, 794–800. [Google Scholar] [CrossRef] [Green Version]
Prakriya, M. Store-Operated Orai Channels: Structure and Function. Curr. Top. Membr. 2013, 71, 1–32. [Google Scholar] [PubMed] [Green Version]
Hogan, P.G.; Rao, A. Store-operated calcium entry: Mechanisms and modulation. Biochem. Biophys. Res. Commun. 2015, 460, 40–49. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Hoover, P.J.; Lewis, R.S. Stoichiometric requirements for trapping and gating of Ca²⁺ release-activated Ca²⁺ (Crac) channels by stromal interaction molecule 1 (Stim1). Proc. Natl. Acad. Sci. USA 2011, 108, 13299–13304. [Google Scholar] [CrossRef] [Green Version]
Wu, M.M.; Covington, E.D.; Lewis, R.S. Single-molecule analysis of diffusion and trapping of STIM1 and Orai1 at endoplasmic reticulum—Plasma membrane junctions. Mol. Biol. Cell 2014, 25, 3672–3685. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Thompson, J.L.; Shuttleworth, T.J. Molecular basis of activation of the arachidonate-regulated Ca²⁺ (Arc) channel, a store-independent Orai channel, by plasma membrane STIM1. J. Physiol. 2013, 591, 3507–3523. [Google Scholar] [CrossRef]
Facilitation of Orai3 targeting and store-operated function by Orai1. Biochim. Biophys. Acta (BBA) Mol. Cell Res. 2015, 1853, 1541–1550. [CrossRef] [Green Version]
Putney, J.W.; McKay, R.R. Capacitative calcium entry channels. BioEssays 1999, 21, 38–46. [Google Scholar] [CrossRef]
Mikoshiba, K.; Hattori, M. Ip3 receptor-operated calcium entry. Sci. Signal. 2000, 2000, pe1. [Google Scholar] [CrossRef]
Sampieri, A.; Santoyo, K.; Asanov, A.; Vaca, L. Association of the IP3R to STIM1 provides a reduced intraluminal calcium microenvironment, resulting in enhanced store-operated calcium entry. Sci. Rep. 2018, 8, 13252. [Google Scholar] [CrossRef]
Derler, I.; Jardin, I.; Romanin, C. Molecular mechanisms of STIM/Orai communication. Am. J. Physiol. Physiol. 2016, 310, C643–C662. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Tiffner, A.; Derler, I. Molecular Choreography and Structure of Ca²⁺ Release-Activated Ca²⁺ (CRAC) and K_Ca²⁺ Channels and Their Relevance in Disease with Special Focus on Cancer. Membranes 2020, 10, 425. [Google Scholar] [CrossRef] [PubMed]
Derler, I.; Madl, J.; Schütz, G.; Romanin, C. Structure, regulation and biophysics of icrac, stim/orai1. In Calcium Signaling; Md. Islam, S., Ed.; Springer: Dordrecht, The Netherlands, 2012; pp. 383–410. [Google Scholar] [CrossRef]
Chung, S.; Zhang, M.; Stathopulos, P. The 2β Splice Variation Alters the Structure and Function of the Stromal Interaction Molecule Coiled-Coil Domains. Int. J. Mol. Sci. 2018, 19, 3316. [Google Scholar] [CrossRef] [Green Version]
Hamada, K.; Miyatake, H.; Terauchi, A.; Mikoshiba, K. IP3-mediated gating mechanism of the IP3 receptor revealed by mutagenesis and X-ray crystallography. Proc. Natl. Acad. Sci. USA 2017, 114, 4661–4666. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Narayanan, D.; Adebiyi, A.; Jaggar, J.H. Inositol trisphosphate receptors in smooth muscle cells. Am. J. Physiol. Circ. Physiol. 2012, 302, H2190–H2210. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Taylor, C.W.; da Fonseca, P.; Morris, E. IP3 receptors: The search for structure. Trends Biochem. Sci. 2004, 29, 210–219. [Google Scholar] [CrossRef] [PubMed]
Serysheva, I.I. Toward a high-resolution structure of IP3R channel. Cell Calcium 2014, 56, 125–132. [Google Scholar] [CrossRef] [Green Version]
Zhou, Y.; Nwokonko, R.M.; Baraniak, J.H., Jr.; Trebak, M.; Lee, K.P.K.; Gill, D.L. The remote allosteric control of Orai channel gating. PLoS Biol. 2019, 17, e3000413. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Cai, X.; Zhou, Y.; Nwokonko, R.; Loktionova, N.A.; Wang, X.; Xin, P.; Trebak, M.; Wang, Y.; Gill, D.L. The Orai1 Store-operated Calcium Channel Functions as a Hexamer. J. Biol. Chem. 2016, 291, 25764–25775. [Google Scholar] [CrossRef] [Green Version]
Pan, Z.; Ma, J.; Zui, P.; Jianjie, M. Open Sesame: Treasure in store-operated calcium entry pathway for cancer therapy. Sci. China Life Sci. 2014, 58, 48–53. [Google Scholar] [CrossRef] [Green Version]
Zhai, X.; Sterea, A.M.; El Hiani, Y. Lessons from the Endoplasmic Reticulum Ca²⁺ Transporters—A Cancer Connection. Cells 2020, 9, 1536. [Google Scholar] [CrossRef] [PubMed]
Emrich, S.M.; Yoast, R.E.; Xin, P.; Arige, V.; Wagner, L.E.; Hempel, N.; Gill, D.L.; Sneyd, J.; Yule, D.I.; Trebak, M. Omnitem-poral choreographies of IP₃R and all five STIM/Orai underlie the complexity of mammalian Ca²⁺ signaling. BioRxiv 2020. [Google Scholar] [CrossRef]
Korzeniowski, M.K.; Baird, B.; Holowka, D. STIM1 activation is regulated by a 14 amino acid sequence adjacent to the CRAC activation domain. AIMS Biophys. 2016, 3, 99–118. [Google Scholar] [CrossRef] [PubMed]
Lopez, J.J.; Jardin, I.; Sanchez-Collado, J.; Salido, G.M.; Smani, T.; Rosado, J.A. TRPC Channels in the SOCE Scenario. Cells 2020, 9, 126. [Google Scholar] [CrossRef] [Green Version]
Furuichi, T.; Yoshikawa, S.; Miyawaki, A.; Wada, K.; Maeda, N.; Mikoshiba, K. Primary structure and functional expression of the inositol 1,4,5-trisphosphate-binding protein P400. Nat. Cell Biol. 1989, 342, 32–38. [Google Scholar] [CrossRef] [PubMed]
Taylor, C.W.; Tovey, S.C. IP3 Receptors: Toward Understanding Their Activation. Cold Spring Harb. Perspect. Biol. 2010, 2, a004010. [Google Scholar] [CrossRef]
Hisatsune, C.; Mikoshiba, K. IP3receptor mutations and brain diseases in human and rodents. J. Neurochem. 2017, 141, 790–807. [Google Scholar] [CrossRef] [Green Version]
Prole, D.L.; Taylor, C. Inositol 1,4,5-trisphosphate receptors and their protein partners as signalling hubs. J. Physiol. 2016, 594, 2849–2866. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Tu, J.C.; Xiao, B.; Yuan, J.P.; A Lanahan, A.; Leoffert, K.; Li, M.; Linden, D.J.; Worley, P.F. Homer Binds a Novel Proline-Rich Motif and Links Group 1 Metabotropic Glutamate Receptors with IP3 Receptors. Neuron 1998, 21, 717–726. [Google Scholar] [CrossRef] [Green Version]
Li, C.; Enomoto, M.; Rossi, A.M.; Seo, M.-D.; Rahman, T.; Stathopulos, P.; Taylor, C.; Ikura, M.; Ames, J.B. CaBP1, a neuronal Ca²⁺ sensor protein, inhibits inositol trisphosphate receptors by clamping intersubunit interactions. Proc. Natl. Acad. Sci. 2013, 110, 8507–8512. [Google Scholar] [CrossRef] [Green Version]
Hirota, J.; Michikawa, T.; Natsume, T.; Furuichi, T.; Mikoshiba, K. Calmodulin inhibits inositol 1,4,5-trisphosphate-induced calcium release through the purified and reconstituted inositol 1,4,5-trisphosphate receptor type 1. FEBS Lett. 1999, 456, 322–326. [Google Scholar] [CrossRef] [Green Version]
Bourguignon, L.Y.; Jin, H.; Iida, N.; Brandt, N.R.; Zhang, S.H. The involvement of ankyrin in the regulation of inositol 1,4,5-trisphosphate receptor-mediated internal Ca²⁺ release from Ca²⁺ storage vesicles in mouse T-lymphoma cells. J. Biol. Chem. 1993, 268, 7290–7297. [Google Scholar] [CrossRef]
Ando, H.; Mizutani, A.; Matsu-Ura, T.; Mikoshiba, K. IRBIT, a Novel Inositol 1,4,5-Trisphosphate (IP3) Receptor-binding Protein, Is Released from the IP3 Receptor upon IP3 Binding to the Receptor. J. Biol. Chem. 2003, 278, 10602–10612. [Google Scholar] [CrossRef] [Green Version]
Uchida, K.; Miyauchi, H.; Furuichi, T.; Michikawa, T.; Mikoshiba, K. Critical Regions for Activation Gating of the Inositol 1,4,5-Trisphosphate Receptor. J. Biol. Chem. 2003, 278, 16551–16560. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Rahman, T. Dynamic clustering of IP3 receptors by IP3. Biochem. Soc. Trans. 2012, 40, 325–330. [Google Scholar] [CrossRef] [PubMed]
Taylor, C.W.; Konieczny, V. IP3receptors: Take four IP3to open. Sci. Signal. 2016, 9, pe1. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Bezprozvanny, L.; Watras, J.; Ehrlich, B. Bell-shaped calcium-response curves of lns(l,4,5)P3- and calcium-gated channels from endoplasmic reticulum of cerebellum. Nat. Cell Biol. 1991, 351, 751–754. [Google Scholar] [CrossRef] [PubMed]
Hisatsune, C.; Nakamura, K.; Kuroda, Y.; Nakamura, T.; Mikoshiba, K. Amplification of Ca²⁺ Signaling by Diacylglycerol-mediated Inositol 1,4,5-Trisphosphate Production. J. Biol. Chem. 2005, 280, 11723–11730. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Van Coppenolle, F.; Vanden Abeele, F.; Slomianny, C.; Flourakis, M.; Hesketh, J.; Dewailly, E.; Prevarskaya, N. Ribosome-translocon complex mediates calcium leakage from endoplasmic reticulum stores. J. Cell Sci. 2004, 117, 4135–4142. [Google Scholar] [CrossRef] [Green Version]
Vaca, L. SOCIC: The store-operated calcium influx complex. Cell Calcium 2010, 47, 199–209. [Google Scholar] [CrossRef] [PubMed]
Gudlur, A.; Hogan, P.G. The STIM-Orai Pathway: Orai, the Pore-Forming Subunit of the CRAC Channel. Adv. Exp. Med. Biol. 2017, 993, 39–57. [Google Scholar] [CrossRef] [Green Version]
DeHaven, W.I.; Jones, B.F.; Petranka, J.G.; Smyth, J.T.; Tomita, T.; Bird, G.S.; Putney, J.W. TRPC channels function independently of STIM1 and Orai1. J. Physiol. 2009, 587, 2275–2298. [Google Scholar] [CrossRef]
Cheng, K.T.; Ong, H.L.; Liu, X.; Ambudkar, I.S. Contribution and Regulation of TRPC Channels in Store-Operated Ca²⁺ Entry. Curr. Top. Membr. 2013, 71, 149–179. [Google Scholar] [CrossRef] [Green Version]
De Souza, L.B.; Ong, H.L.; Liu, X.; Ambudkar, I.S. Fast endocytic recycling determines TRPC1–STIM1 clustering in ER–PM junctions and plasma membrane function of the channel. Biochim. Biophys. Acta (BBA)—Bioenerg. 2015, 1853, 2709–2721. [Google Scholar] [CrossRef] [Green Version]
Ambudkar, I.S.; de Souza, L.B.; Ong, H.L. TRPC1, Orai1, and STIM1 in SOCE: Friends in tight spaces. Cell Calcium 2017, 63, 33–39. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Pacheco, J.; Vaca, L. Stim-trp pathways and microdomain organization: Auxiliary proteins of the stim/orai complex. In Store-Operated Ca²⁺ Entry (SOCE) Pathways: Emerging Signaling Concepts in Human (Patho)physiology; Groschner, K., Graier, W.F., Romanin, C., Eds.; Springer: Berlin/Heidelberg, Germany, 2017; pp. 189–210. [Google Scholar] [CrossRef]
Redondo, P.C.; Harper, A.G.S.; Salido, G.M.; Pariente, J.A.; Sage, S.O.; Rosado, J.A. A role for SNAP-25 but not VAMPs in store-mediated Ca²⁺entry in human platelets. J. Physiol. 2004, 558, 99–109. [Google Scholar] [CrossRef]
Sharma, A.; Ramena, G.; Yin, Y.; Premkumar, L.; Elble, R.C. CLCA2 is a positive regulator of store-operated calcium entry and TMEM16A. PLoS ONE 2018, 13, e0196512. [Google Scholar] [CrossRef]
Lopez, J.J.; Albarran, L.; Gómez, L.J.; Smani, T.; Salido, G.M.; Rosado, J.A. Molecular modulators of store-operated calcium entry. Biochim. Biophys. Acta (BBA)—Bioenerg. 2016, 1863, 2037–2043. [Google Scholar] [CrossRef] [PubMed]
Quintana, A.; Rajanikanth, V.; Farber-Katz, S.; Gudlur, A.; Zhang, C.; Jing, J.; Zhou, Y.; Rao, A.; Hogan, P.G. TMEM110 regulates the maintenance and remodeling of mammalian ER–plasma membrane junctions competent for STIM–ORAI signaling. Proc. Natl. Acad. Sci. USA 2015, 112, E7083–E7092. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Zhang, L.; Wang, L.; Li, S.; Xue, J.; Luo, D. Calsequestrin-1 Regulates Store-Operated Ca²⁺ Entry by Inhibiting STIM1 Aggregation. Cell. Physiol. Biochem. 2016, 38, 2183–2193. [Google Scholar] [CrossRef]
Albarran, L.; Lopez, J.J.; Ben Amor, N.; Cano, F.E.M.; Erro, A.B.; Smani, T.; Salido, G.M.; Rosado, J.A. Dynamic interaction of SARAF with STIM1 and Orai1 to modulate store-operated calcium entry. Sci. Rep. 2016, 6, 24452. [Google Scholar] [CrossRef]
Malli, R.; Frieden, M.; Trenker, M.; Graier, W.F. The Role of Mitochondria for Ca²⁺ Refilling of the Endoplasmic Reticulum. J. Biol. Chem. 2005, 280, 12114–12122. [Google Scholar] [CrossRef] [PubMed] [Green Version]
DeLuca, H.F.; Engstrom, G.W. Calcium uptake by rat kidney mitochondria. Proc. Natl. Acad. Sci. USA 1961, 47, 1744–1750. [Google Scholar] [CrossRef] [Green Version]
Finkel, T.; Menazza, S.; Holmström, K.; Parks, R.J.; Liu, J.; Sun, J.; Liu, J.; Pan, X.; Murphy, E. The Ins and Outs of Mitochondrial Calcium. Circ. Res. 2015, 116, 1810–1819. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Patergnani, S.; Suski, J.M.; Agnoletto, C.; Bononi, A.; Bonora, M.; De Marchi, E.; Giorgi, C.; Marchi, S.; Missiroli, S.; Poletti, F.; et al. Calcium signaling around Mitochondria Associated Membranes (MAMs). Cell Commun. Signal. 2011, 9, 19. [Google Scholar] [CrossRef] [Green Version]
Alberts, B. (Ed.) Molecular Biology of the Cell, 4th ed.; Garland Science: New York, NY, USA, 2002. [Google Scholar]
Hajnóczky, G.; Csordás, G.; Yi, M. Old players in a new role: Mitochondria-associated membranes, VDAC, and ryanodine receptors as contributors to calcium signal propagation from endoplasmic reticulum to the mitochondria. Cell Calcium 2002, 32, 363–377. [Google Scholar] [CrossRef] [PubMed]
Bononi, A.; Missiroli, S.; Poletti, F.; Suski, J.M.; Agnoletto, C.; Bonora, M.; De Marchi, E.; Giorgi, C.; Marchi, S.; Patergnani, S.; et al. Mitochondria-associated membranes (Mams) as hotspot Ca²⁺ signaling units. In Calcium Signaling; Islam, M.S., Ed.; Springer: Dordrecht, The Netherlands, 2012; Volume 740, pp. 411–437. [Google Scholar] [CrossRef]
Colombini, M. VDAC structure, selectivity, and dynamics. Biochim. Biophys. Acta (BBA) Biomembr. 2012, 1818, 1457–1465. [Google Scholar] [CrossRef] [Green Version]
Mishra, J.; Jhun, B.S.; Hurst, S.; O-Uchi, J.; Csordás, G.; Sheu, S.-S. The mitochondrial Ca²⁺ uniporter: Structure, function, and pharmacology. In Pharmacology of Mitochondria; Singh, H., Sheu, S.-S., Eds.; Springer: Berlin/Heidelberg, Germany, 2017; Volume 240, pp. 129–156. [Google Scholar] [CrossRef] [Green Version]
Paillard, M.; Csordás, G.; Szanda, G.; Golenár, T.; Debattisti, V.; Bartok, A.; Wang, N.; Moffat, C.; Seifert, E.L.; Spät, A.; et al. Tissue-specific mitochondrial decoding of cytoplasmic Ca²⁺ signals is controlled by the stoichiometry of micu1/2 and mcu. Cell Rep. 2017, 18, 2291–2300. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Sassano, M.L.; van Vliet, A.R.; Agostinis, P. Mitochondria-associated membranes as networking platforms and regulators of cancer cell fate. Front. Oncol. 2017, 7, 174. [Google Scholar] [CrossRef]
Schrepfer, E.; Scorrano, L. Mitofusins, from Mitochondria to Metabolism. Mol. Cell. 2016, 61, 683–694. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Vallese, F.; Barazzuol, L.; Maso, L.; Brini, M.; Calì, T. Er-mitochondria calcium transfer, organelle contacts and neurodegen-erative diseases. In Calcium Signaling; Islam, M.S., Ed.; Springer: Berlin/Heidelberg, Germany, 2020; pp. 719–746. [Google Scholar] [CrossRef]
Morgan, A.J.; Davis, L.C.; Wagner, S.K.T.Y.; Lewis, A.M.; Parrington, J.; Churchill, G.C.; Galione, A. Bidirectional Ca²⁺ signaling occurs between the endoplasmic reticulum and acidic organelles. J. Cell Biol. 2013, 200, 789–805. [Google Scholar] [CrossRef] [Green Version]
Gao, P.; Yan, Z.; Zhu, Z. Mitochondria-associated endoplasmic reticulum membranes in cardiovascular diseases. Front. Cell Dev. Biol. 2020, 8, 1309. [Google Scholar] [CrossRef]
van Vliet, A.R.; Agostinis, P. Mitochondria-associated membranes and er stress. In Coordinating Organismal Physiology through the Unfolded Protein Response; Wiseman, R.L., Haynes, C.M., Eds.; Springer: Berlin/Heidelberg, Germany, 2018; pp. 73–102. [Google Scholar] [CrossRef]
Perrone, M.; Caroccia, N.; Genovese, I.; Missiroli, S.; Modesti, L.; Pedriali, G.; Vezzani, B.; Vitto, V.A.M.; Antenori, M.; Lebiedzinska-Arciszewska, M.; et al. The role of mitochondria-associated membranes in cellular homeostasis and diseases. Int. Rev. Cell Mol. Biol. 2020, 350, 119–196. [Google Scholar] [CrossRef]
Betz, C.; Stracka, D.; Prescianotto-Baschong, C.; Frieden, M.; Demaurex, N.; Hall, M.N. Mtor complex 2-akt signaling at mitochondria-associated endoplasmic reticulum membranes (Mam) regulates mitochondrial physiology. Proc. Natl. Acad. Sci. USA 2013, 110, 12526–12534. [Google Scholar] [CrossRef] [Green Version]
Ouyang, Y.-B.; Giffard, R.G. Er-mitochondria crosstalk during cerebral ischemia: Molecular chaperones and er-mitochondrial calcium transfer. Int. J. Cell Biol. 2012. [Google Scholar] [CrossRef] [Green Version]
Matsuzaki, H.; Fujimoto, T.; Tanaka, M.; Shirasawa, S. Tespa1 is a novel component of mitochondria-associated endoplasmic reticulum membranes and affects mitochondrial calcium flux. Biochem. Biophys. Res. Commun. 2013, 433, 322–326. [Google Scholar] [CrossRef]
Lee, S.; Min, K.-T. The interface between er and mitochondria: Molecular compositions and functions. Mol. Cells 2018, 41, 1000–1007. [Google Scholar] [CrossRef] [PubMed]
Kilpatrick, B.S.; Eden, E.R.; Hockey, L.N.; Yates, E.; Futter, C.E.; Patel, S. An endosomal naadp-sensitive two-pore Ca²⁺ channel regulates er-endosome membrane contact sites to control growth factor signaling. Cell Rep. 2017, 18, 1636–1645. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Bernardi, P.; Di Lisa, F.; Fogolari, F.; Lippe, G. From ATP to PTP and back: A dual function for the mitochondrial ATP synthase. Circ. Res. 2015, 116, 1850–1862. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Vultur, A.; Gibhardt, C.S.; Stanisz, H.; Bogeski, I. The role of the mitochondrial calcium uniporter (Mcu) complex in cancer. Pflügers Arch. Eur. J. Physiol. 2018, 470, 1149–1163. [Google Scholar] [CrossRef]
Halestrap, A.P. The c ring of the f1fo atp synthase forms the mitochondrial permeability transition pore: A critical appraisal. Front. Oncol. 2014, 4, 234. [Google Scholar] [CrossRef] [Green Version]
Gomez-Suaga, P.; Paillusson, S.; Stoica, R.; Noble, W.; Hanger, D.P.; Miller, C.C.J. The ER-Mitochondria Tethering Complex VAPB-PTPIP51 Regulates Autophagy. Curr. Biol. 2017, 27, 371–385. [Google Scholar] [CrossRef] [Green Version]
Lively Lysosomes. Available online: https://www.asbmb.org/asbmb-today/science/050116/lively-lysosomes (accessed on 20 September 2020).
Yang, J.; Zhao, Z.; Gu, M.; Feng, X.; Xu, H. Release and uptake mechanisms of vesicular Ca²⁺ stores. Protein Cell 2019, 10, 8–19. [Google Scholar] [CrossRef] [Green Version]
Karch, J.; Molkentin, J.D. Identifying the components of the elusive mitochondrial permeability transition pore. Proc. Natl. Acad. Sci. USA 2014, 111, 10396–10397. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Garrity, A.G.; Wang, W.; Collier, C.M.; Levey, S.A.; Gao, Q.; Xu, H. The endoplasmic reticulum, not the pH gradient, drives calcium refilling of lysosomes. eLife 2016, 5, e15887. [Google Scholar] [CrossRef]
Natarajan, G.K.; Glait, L.; Mishra, J.; Stowe, D.F.; Camara, A.K.S.; Kwok, W.-M. Total Matrix Ca²⁺ Modulates Ca²⁺ Efflux via the Ca²⁺/H+ Exchanger in Cardiac Mitochondria. Front. Physiol. 2020, 11. [Google Scholar] [CrossRef]
Pathak, T.; Trebak, M. Mitochondrial Ca²⁺ signaling. Pharmacol. Ther. 2018, 192, 112–123. [Google Scholar] [CrossRef]
Palty, R.; Silverman, W.F.; Hershfinkel, M.; Caporale, T.; Sensi, S.L.; Parnis, J.; Nolte, C.; Fishman, D.; Shoshan-Barmatz, V.; Herrmann, S.; et al. NCLX is an essential component of mitochondrial Na⁺/Ca²⁺ exchange. Proc. Natl. Acad. Sci. USA 2010, 107, 436–441. [Google Scholar] [CrossRef] [Green Version]
Bernardi, P.; Von Stockum, S. The permeability transition pore as a Ca²⁺ release channel: New answers to an old question. Cell Calcium 2012, 52, 22–27. [Google Scholar] [CrossRef] [Green Version]
Massari, S.; Azzone, G.F. The equivalent pore radius of intact and damaged mitochondria and the mechanism of active shrinkage. Biochim. Biophys. Acta (BBA)—Bioenerg. 1972, 283, 23–29. [Google Scholar] [CrossRef]
Di Lisa, F.; Carpi, A.; Giorgio, V.; Bernardi, P. The mitochondrial permeability transition pore and cyclophilin D in cardioprotection. Biochim. Biophys. Acta (BBA)—Bioenerg. 2011, 1813, 1316–1322. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Ichas, F.; Mazat, J.-P. From calcium signaling to cell death: Two conformations for the mitochondrial permeability transition pore. Switching from low- to high-conductance state. Biochim. Biophys. Acta (BBA)—Bioenerg. 1998, 1366, 33–50. [Google Scholar] [CrossRef] [Green Version]
Tinel, H.; Cancela, J.M.; Mogami, H.; Gerasimenko, J.; Gerasimenko, O.; Tepikin, A.; Petersen, O. Active mitochondria surrounding the pancreatic acinar granule region prevent spreading of inositol trisphosphate-evoked local cytosolic Ca²⁺ signals. EMBO J. 1999, 18, 4999–5008. [Google Scholar] [CrossRef]
Gerasimenko, J.; Sherwood, M.; Tepikin, A.; Petersen, O.; Gerasimenko, O. NAADP, cADPR and IP3 all release Ca²⁺ from the endoplasmic reticulum and an acidic store in the secretory granule area. J. Cell Sci. 2006, 119, 226–238. [Google Scholar] [CrossRef] [Green Version]
Patel, S.; Ramakrishnan, L.; Rahman, T.; Hamdoun, A.; Marchant, J.; Taylor, C.; Brailoiu, E. The endo-lysosomal system as an NAADP-sensitive acidic Ca²⁺ store: Role for the two-pore channels. Cell Calcium 2011, 50, 157–167. [Google Scholar] [CrossRef] [Green Version]
Faris, P.; Shekha, M.; Montagna, D.; Guerra, G.; Moccia, F. Endolysosomal Ca²⁺ signalling and cancer hallmarks: Two-pore channels on the move, trpml1 lags behind! Cancers 2019, 11, 27. [Google Scholar] [CrossRef] [Green Version]
Phillips, M.J.; Voeltz, G.K. Structure and function of ER membrane contact sites with other organelles. Nat. Rev. Mol. Cell Biol. 2016, 17, 69–82. [Google Scholar] [CrossRef] [Green Version]
López-Sanjurjo, C.I.; Tovey, S.C.; Prole, D.L.; Taylor, C.W. Lysosomes shape Ins(1,4,5) P 3 -evoked Ca²⁺ signals by selectively sequestering Ca²⁺ released from the endoplasmic reticulum. J. Cell Sci. 2013, 126, 289–300. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Sbano, L.; Bonora, M.; Marchi, S.; Baldassari, F.; Medina, D.L.; Ballabio, A.; Giorgi, C.; Pinton, P. TFEB-mediated increase in peripheral lysosomes regulates store-operated calcium entry. Sci. Rep. 2017, 7, 40797. [Google Scholar] [CrossRef]
Carruthers, C.; Suntzeff, V. Calcium, Copper, and Zinc in the Epidermal Carcinogenesis of Mouse and Man. Cancer Res. 1946, 6, 296. [Google Scholar] [PubMed]
Borowiec, A.S.; Bidaux, G.; Pigat, N.; Goffin, V.; Bernichtein, S.; Capiod, T. Calcium Channels, External Calcium Concentration and Cell Proliferation. Eur. J. Pharmacol. 2014, 739, 19–25. [Google Scholar] [CrossRef] [PubMed]
Whitfield, J.F. Calcium signals and cancer. Crit. Rev. Oncog. 1992, 3, 55–90. [Google Scholar]
Cook, S.J.; Lockyer, P.J. Recent advances in Ca²⁺-dependent Ras regulation and cell proliferation. Cell Calcium 2006, 39, 101–112. [Google Scholar] [CrossRef]
Boynton, A.L.; Whitfield, J.F.; Isaacs, R.J. The different roles of serum and calcium in the control of proliferation of BALB/c 3T3 mouse cells. In Vitro-Plant 1976, 12, 120–123. [Google Scholar] [CrossRef]
Hazelton, B.; Mitchell, B.; Tupper, J. Calcium, magnesium, and growth control in the WI-38 human fibroblast cell. J. Cell Biol. 1979, 83, 487–498. [Google Scholar] [CrossRef]
Prevarskaya, N.; Ouadid-Ahidouch, H.; Skryma, R.; Shuba, Y. Remodelling of Ca²⁺ transport in cancer: How it contributes to cancer hallmarks? Philos. Trans. R. Soc. Lond. B Biol. Sci. 2014, 369, 20130097. [Google Scholar] [CrossRef] [Green Version]
Humeau, J.; Bravo-San Pedro, J.M.; Vitale, I.; Nuñez, L.; Villalobos, C.; Kroemer, G.; Senovill, L. Calcium signaling and cell cycle: Progression or death. Cell Calcium. 2018, 70, 3–15. [Google Scholar] [CrossRef]
Capiod, T.; Shuba, Y.; Skryma, R.; Prevarskaya, N. Calcium signalling and cancer cell growth. Subcell Biochem. 2007, 45, 405–427. [Google Scholar] [CrossRef]
Clowes, G.; Frisbie, W. No. 32. On the relationship between the rate of growth, age, and potassium and calcium content of mouse tumors (adeno-carcinoma, jensen). Am. J. Physiol. Leg. Content 1905, 14, 173–192. [Google Scholar] [CrossRef]
Carruthers, C.; Suntzeff, V. The role of calcium in carcinogenesis summary. Science 1944, 99, 245–247. [Google Scholar] [CrossRef] [PubMed]
Miller, K. Calcium and cancer. Med. Hypotheses 1977, 3, 263–264. [Google Scholar] [CrossRef]
Kadio, B.; Yaya, S.; Basak, A.; Djè, K.; Gomes, J.; Mesenge, C. Calcium role in human carcinogenesis: A comprehensive analysis and critical review of literature. Cancer Metastasis Rev. 2016, 35, 391–411. [Google Scholar] [CrossRef] [PubMed]
Peterlik, M.; Grant, W.B.; Cross, H.S. Calcium, Vitamin D and Cancer. Anticancer Res. 2009, 29, 3687–3698. [Google Scholar]
Pottle, J.; Sun, C.; Gray, L.; Li, M. Exploiting MCF-7 Cells’ calcium dependence with interlaced therapy. J. Cancer Ther. 2013, 4, 32–40. [Google Scholar] [CrossRef] [Green Version]
Taylor, J.M.; Simpson, R.U. Inhibition of Cancer Cell Growth by Calcium Channel Antagonists in the Athymic Mouse. Cancer Res. 1992, 52, 2413–2418. [Google Scholar]
Xu, M.M.; Seas, A.; Kiyani, M.; Ji, K.S.Y.; Bell, H.N. A temporal examination of calcium signaling in cancer- from tumorigenesis, to immune evasion, and metastasis. Cell Biosci. 2018, 8, 25. [Google Scholar] [CrossRef] [Green Version]
Hanahan, D.; Weinberg, R.A. Hallmarks of cancer: The next generation. Cell 2011, 144, 646–674. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Prevarskaya, N.; Skryma, R.; Shuba, Y. Ion Channels in cancer: Are cancer hallmarks oncochannelopathies? Physiol. Rev. 2018, 98, 559–621. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Monteith, G.R.; Prevarskaya, N.; Roberts-Thomson, S.J. The calcium–cancer signalling nexus. Nat. Rev. Cancer 2017, 17, 367–380. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Roberts-Thomson, S.J.; Chalmers, S.B.; Monteith, G.R. The Calcium-Signaling Toolkit in Cancer: Remodeling and Targeting. Cold Spring Harb. Perspect. Biol. 2019, 11, a035204. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Roderick, H.W.; Cook, S.J. Ca²⁺ signalling checkpoints in cancer: Remodelling Ca²⁺ for cancer cell proliferation and survival. Nat. Rev. Cancer 2008, 8, 361–375. [Google Scholar] [CrossRef] [Green Version]
Phan, N.N.; Wang, C.Y.; Chen, C.F.; Sun, Z.; Lai, M.D.; Lin, Y.L. Voltage-gated calcium channels: Novel targets for cancer therapy. Oncol. Lett. 2017, 14, 2059–2074. [Google Scholar] [CrossRef] [Green Version]
Wang, C.Y.; Lai, M.D.; Phan, N.N.; Sun, Z.; Lin, Y.C. Meta-analysis of public microarray datasets reveals voltage-gated calcium gene signatures in clinical cancer patients. PLoS ONE 2015, 10, e0125766. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Wang, X.T.; Nagaba, Y.; Cross, H.S.; Wrba, F.; Zhang, L.; Guggino, S.E. The mRNA of L-type calcium channel elevated in colon cancer: Protein distribution in normal and cancerous colon. Am. J. Pathol. 2000, 157, 1549–1562. [Google Scholar] [CrossRef]
Buchanan, P.J.; McCloskey, K.D. CaV channels and cancer: Canonical functions indicate benefits of repurposed drugs as cancer therapeutics. Eur. Biophys. J. 2016, 45, 621–633. [Google Scholar] [CrossRef] [Green Version]
Taylor, J.T.; Huang, L.; Pottle, J.E.; Liu, K.; Yang, Y.; Zeng, X.; Keyser, B.M.; Agrawal, K.C.; Hansen, J.B.; Li, M. Selective blockade of T-type Ca²⁺ channels suppresses human breast cancer cell proliferation. Cancer Lett. 2008, 267, 116–124. [Google Scholar] [CrossRef]
Azimi, I.; Roberts-Thomson, S.J.; Monteith, G.R. Calcium influx pathways in breast cancer: Opportunities for pharmacological intervention. Br. J. Pharmacol. 2014, 171, 945–960. [Google Scholar] [CrossRef] [Green Version]
Antal, L.; Martin-Caraballo, M. T-Type Calcium Channels in Cancer. Cancers 2019, 11, 134. [Google Scholar] [CrossRef] [Green Version]
Barceló, C.; Sisó, P.; Maiques, O.; de la Rosa, I.; Martí, R.M.; Macià, A. T-Type Calcium Channels in Cnacer: A Potential Target in Melanoma. Cancers 2020, 12, 391. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Choi, D.L.; Jang, S.J.; Cho, S.; Choi, H.E.; Rim, H.K.; Lee, K.T.; Lee, J.Y. Inhibition of Cellular Proliferation and Induction of Apoptosis in Human Lung Adenocarcinoma A549 Cells by T-type Calcium Channel Antagonist. Bioorg. Med. Chem. Lett. 2014, 24, 1565–1570. [Google Scholar] [CrossRef] [PubMed]
Rim, H.K.; Cho, S.; Shin, D.H.; Chung, K.S.; Cho, Y.W.; Choi, J.H.; Lee, J.Y.; Lee, K.T. T-type Ca²⁺ channel blocker, KYS05090 induces autophagy and apoptosis in A549 cells through inhibiting glucose uptake. Molecules 2014, 19, 9864–9875. [Google Scholar] [CrossRef]
Rim, H.K.; Lee, H.W.; Choi, I.S.; Park, J.Y.; Choi, H.W.; Choi, J.H.; Cho, Y.W.; Lee, J.Y.; Lee, K.T. T-type Ca²⁺ channel blocker, KYS05047 induces G1 phase cell cycle arrest by decreasing intracellular Ca²⁺ levels in human lung adenocarcinoma A549 cells. Bioorg. Med. Chem. Lett. 2012, 22, 7123–7126. [Google Scholar] [CrossRef]
Arif, T.; Amsalem, Z.; Shoshan-Barmatz, V. Metabolic Reprograming Via Silencing of Mitochondrial VDAC1 Expression Encourages Differentiation of Cancer Cells. Mol. Ther.—Nucleic Acids 2019, 17, 24–37. [Google Scholar] [CrossRef] [Green Version]
Thinnes, F.P. Neuroendocrine differentiation of LNCaP cells suggests: VDAC in the cell membrane is involved in the extrinsic apoptotic pathway. Mol. Genet. Metab. 2009, 97, 241–243. [Google Scholar] [CrossRef]
Fourbon, Y.; Guéguinou, M.; Félix, R.; Constantin, B.; Uguen, A.; Fromont, G.; Lajoie, L.; Magaud, C.; LeComte, T.; Chamorey, E.; et al. Ca²⁺ protein alpha 1D of CaV1.3 regulates intracellular calcium concentration and migration of colon cancer cells through a non-canonical activity. Sci. Rep. 2017, 7, 14199. [Google Scholar] [CrossRef] [Green Version]
Scarpellino, G.; Genova, T.; Avanzato, D.; Bernardini, M.; Bianco, S.; Petrillo, S.; Tolosano, E.; de Almeida Vieira, J.R.; Bussolati, B.; Fiorio Pla, A.; et al. Purinergic Calcium Signals in Tumor-Derived Endothelium. Cancers 2019, 11, 766. [Google Scholar] [CrossRef] [Green Version]
Chong, J.-H.; Zheng, G.-G.; Zhu, X.-F.; Guo, Y.; Wang, L.; Ma, C.-H.; Liu, S.-Y.; Xu, L.-L.; Lin, Y.-M.; Wu, K.-F. Abnormal expression of P2X family receptors in Chinese pediatric acute leukemias. Biochem. Biophys. Res. Commun. 2010, 391, 498–504. [Google Scholar] [CrossRef] [PubMed]
Vijayan, D.; Smyth, M.J.; Teng, M.W.L. Purinergic Receptors: Novel Targets for Cancer Immunotherapy. In Oncoimmunology; Zitvogel, L., Kroemer, G., Eds.; Springer: Cham, Switzerland, 2018; ISBN 978-3-319-62431-0. [Google Scholar] [CrossRef]
Maehara, Y.; Kusumoto, H.; Anai, H.; Kusumoto, T.; Sugimachi, K. Human tumor tissues have higher ATP contents than normal tissues. Clin. Chim. Acta 1987, 169, 341–343. [Google Scholar] [CrossRef]
Azimi, I.; Beilby, H.; Davis, F.M.; Marcial, D.L.; Kenny, P.A.; Thompson, E.W.; Roberts-Thomson, S.; Monteith, G.R. Altered purinergic receptor-Ca²⁺ signaling associated with hypoxia-induced epithelial-mesenchymal transition in breast cancer cells. Mol. Oncol. 2016, 10, 166–178. [Google Scholar] [CrossRef] [Green Version]
Davis, F.M.; Kenny, P.A.; Soo, E.T.L.; van Denderen, B.J.W.; Thompson, E.W.; Cabot, P.J.; Parat, M.O.; Roberts-Thomson, S.J.; Monteith, G.R. Remodeling of Purinergic Receptor-Mediated Ca²⁺ Signaling as a Consequence of EGF-induced Epithelial-Mesenchymal Transition in Breast Cancer Cells. PLoS ONE 2011, 6, e23464. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Avanzato, D.; Genova, T.; Fiorio, A.; Bernardini, M.; Bianco, S.; Bussolati, B.; Mancardi, D.; Giraudo, E.; Maione, F.; Cassoni, P.; et al. Activation of P2X7 and P2Y11 purinergic receptors inhibits migration and normalizes tumor-derived endothelial cells via cAMP signaling. Sci. Rep. 2016, 6, 32602. [Google Scholar] [CrossRef]
Feng, W.; Yang, X.; Wang, L.; Wang, R.; Yang, F.; Wang, H.; Liu, X.; Ren, Q.; Zhang, Y.; Zhu, X.; et al. P2X7 promotes the progression of MLL-AF9 induced acute myeloid leukemia by upregulation of Pbx3. Haematol. 2020, 106, 1278–1289. [Google Scholar] [CrossRef] [Green Version]
Reisner, P.D.; Brandt, P.C.; Vanaman, T.C. Analysis of plasma membrane Ca²⁺-ATPase expression in control and SV40-transformed human fibroblasts. Cell Calcium 1997, 21, 53–62. [Google Scholar] [CrossRef]
Usachev, Y.M.; Toutenhoofd, S.L.; Goellner, G.M.; Strehler, E.E.; Thayer, S.A. Differentiation induces up-regulation of plasma membrane Ca²⁺-ATPase and concomitant increase in Ca²⁺ efflux in human neuroblastoma cell line IMR-32. J. Neurochem. 2001, 76, 1756–1765. [Google Scholar] [CrossRef] [Green Version]
Roberts-Thomson, S.J.; Curry, M.C.; Monteith, G.R. Plasma membrane calcium pumps and their emerging roles in cancer. World J. Biol. Chem. 2010, 1, 248–253. [Google Scholar] [CrossRef]
Aung, C.S.; Ye, W.; Plowman, G.; Peters, A.A.; Monteith, G.; Roberts-Thomson, S.J. Plasma membrane calcium ATPase 4 and the remodeling of calcium homeostasis in human colon cancer cells. Carcinogenesis 2009, 30, 1962–1969. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Peterson, J.A.; Oblad, R.V.; Mecham, J.C.; Kenealey, J.D. Resveratrol inhibits plasma membrane Ca²⁺-ATPase inducing an increase in cytoplasmic calcium. Biochem. Biophys. Rep. 2016, 7, 253–258. [Google Scholar] [CrossRef] [Green Version]
Rüschoff, J.H.; Brandenburger, T.; Strehler, E.E.; Filoteo, A.G.; Heinmöller, E.; Aumüller, G.; Wilhelm, B. Plasma Membrane Calcium ATPase Expression in Human Colon Multistep Carcinogenesis. Cancer Investig. 2012, 30, 251–257. [Google Scholar] [CrossRef]
Ribiczey, P.; Papp, B.; Homolya, L.; Enyedi, Á.; Kovács, T. Selective upregulation of the expression of plasma membrane calcium ATPase isoforms upon differentiation and 1,25(OH)2D3-vitamin treatment of colon cancer cells. Biochem. Biophys. Res. Commun. 2015, 464, 189–194. [Google Scholar] [CrossRef]
Varga, K.; Pászty, K.; Padányi, R.; Hegedűs, L.; Brouland, J.P.; Papp, B.; Enyedi, A. Histone deacetylase inhibitor- and PMA-induced upregulation of PMCA4b enhances Ca²⁺ clearance from MCF-7 breast cancer cells. Cell Calcium 2014, 55, 78–92. [Google Scholar] [CrossRef]
James, A.D.; Patel, W.; Butt, Z.; Adiamah, M.; Dakhel, R.; Latif, A.; Uggenti, C.; Swanton, E.; Imamura, H.; Siriwardena, A.K.; et al. The Plasma Membrane Calcium Pump in Pancreatic Cancer Cells Exhibiting the Warburg Effect Relies on Glycolytic ATP. J. Biol. Chem. 2015, 290, 24760–24771. [Google Scholar] [CrossRef] [Green Version]
Varga, K.; Hollósi, A.; Pászty, K.; Hegedűs, L.; Szakács, G.; Tímár, J.; Papp, B.; Enyedi, A.; Padányi, A. Expression of Calcium Pumps Is Differentially Regulated by Histone Deacetylase Inhibitors and Estrogen Receptor Alpha in Breast Cancer Cells. BMC Cancer 2018, 18, 1029. [Google Scholar] [CrossRef] [Green Version]
Peters, A.A.; Milevskiy, M.J.G.; Lee, W.C.; Curry, M.C.; Smart, C.E.; Saunus, J.M.; Reid, L.; da Silva, L.; Marcial, D.L.; Dray, E.; et al. The calcium pump plasma membrane Ca²⁺-ATPase 2 (PMCA2) regulates breast cancer cell proliferation and sensitivity to doxorubicin. Sci. Rep. 2016, 6, 25505. [Google Scholar] [CrossRef] [PubMed]
Balasubramaniam, S.L.; Gopalakrishnapillai, A.; Petrelli, N.J.; Barwe, S.P. Knockdown of Sodium-Calcium Exchanger 1 Induces Epithelial to Mesenchymal Transition in Kidney Epithelial Cells. J. Biol. Chem. 2017, 292, 11388–11399. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Mahdi, S.H.; Cheng, H.; Li, J.; Feng, R. The effect of TGF-beta induced epithelial-mesenchymal transition on the expression of intracellular calcium-handling proteins in T47D and MCF-7 human breast cancer cells. Arch. Biochem. Biophys. 2015, 42, 1240–1251. [Google Scholar] [CrossRef] [PubMed]
Munoz, J.J.; Drigo, S.A.; Barros-Filho, M.C.; Marchi, F.A.; Scapulatempo-Neto, C.; Pessoa, G.S.; Guimaraes, G.C.; Trindade Filho, J.C.; Lopes, A.; Arruda, M.A.; et al. Down-Regulation of SLC8A1 as a Putative Apoptosis Evasion Mechanism by Modulation of Calcium Levels in Penile Carcinoma. J. Urol. 2015, 194, 245–251. [Google Scholar] [CrossRef]
Pelzl, L.; Hosseinzadeh, Z.; al-Maghout, T.; Singh, Y.; Sahu, I.; Bissinger, R.; Schmidt, S.; Alkahtani, S.; Stournaras, C.; Toulany, M.; et al. Role of Na⁺/Ca²⁺ Exchangers in Therapy Resistance of Medulloblastoma Cells. Cell Physiol. Biochem. 2017, 42, 1240–1251. [Google Scholar] [CrossRef]
Zheng, X.; Lu, S.; He, Z.; Huang, H.; Yao, Z.; Miao, Y.; Cai, C.; Zou, F. MCU-dependent negative sorting of miR-4488 to extracellular vesicles enhances angiogenesis and promotes breast cancer metastatic colonization. Oncogene 2020, 39, 6975–6989. [Google Scholar] [CrossRef]
Kucukkaya, B.; Basoglu, H.; Erdag, D.; Akbas, F.; Susgun, S.; Yalcintepe, L. Calcium homeostasis in cisplatin resistant epithelial ovarian cancer. Gen. Physiol. Biophys. 2019, 38, 353–363. [Google Scholar] [CrossRef]
Liskova, V.; Hudecova, S.; Lencesova, L.; Iuliano, F.; Sirova, M.; Ondrias, K.; Pastorekova, S.; Krizanova, O. Type 1 Sodium Calcium Exchanger Forms a Complex with Carbonic Anhydrase IX and Via Reverse Mode Activity Contributes to pH Control in Hypoxic Tumors. Cancers 2019, 11, 1139. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Svastova, E.; Witarski, W.; Csaderova, L.; Kosik, I.; Skvarkova, L.; Hulikova, A.; Zatovicova, M.; Barathova, M.; Kopacek, J.; Pastorek, J.; et al. Carbonic Anhydrase IX Interacts with Bicarbonate Transporters in Lamellipodia and Increases Cell Migration via Its Catalytic Domain. J. Biol. Chem. 2012, 287, 3392–3402. [Google Scholar] [CrossRef] [Green Version]
Sennoune, S.R.; Santos, J.M.; Hussain, F.; Martínez-Zaguilán, R. Sodium calcium ex-changer operates in the reverse mode in metastatic human melanoma cells. Cell. Mol. Biol. 2015, 61, 40–49. [Google Scholar]
Tojyo, Y.; Morita, T.; Nezu, A.; Tanimura, A. Key Components of Store-Operated Ca²⁺ Entry in Non-Excitable Cells. J. Pharmacol. Sci. 2014, 125, 340–346. [Google Scholar] [CrossRef] [Green Version]
Chen, Y.F.; Lin, P.C.; Yeh, Y.M.; Chen, L.H.; Shen, M.R. Store-Operated Ca²⁺ Entry in Tumor Progression: From Molecular Mechanisms to Clinical Implications. Cancers 2019, 11, 899. [Google Scholar] [CrossRef] [Green Version]
Stewart, T.A.; Yapa, K.T.D.S.; Monteith, G.R. Altered calcium signaling in cancer cells. Biochim. Biophys. Acta 2015, 1848, 2502–2511. [Google Scholar] [CrossRef] [Green Version]
Pierro, C.; Sneyers, F.; Bultynck, G.; Roderick, H.W. ER Ca²⁺ Release and Store-Operated Ca²⁺ Entry—Partners in Crime or Independent Actors in Oncogenic Transformation? Cell Calcium 2019, 82, 102061. [Google Scholar] [CrossRef]
Hoth, M.; Niemeyer, B.A. The neglected CRAC proteins: Orai2, Orai3, and STIM2. Curr. Top. Membr. 2013, 71, 237–271. [Google Scholar] [CrossRef] [PubMed]
Azimi, I.; Bong, A.H.; Poo, G.X.H.; Armitage, K.; Lok, D.; Roberts-Thomson, S.J.; Monteith, G.R. Pharmacological Inhibition of Store-Operated Calcium Entry in MDA-MB-468 Basal A Breast Cancer Cells: Consequences on Calcium Signalling, Cell Migration and Proliferation. Cell Mol. Life Sci. 2018, 75, 4525–4537. [Google Scholar] [CrossRef] [PubMed]
Chen, Y.W.; Chen, Y.F.; Chen, Y.T.; Chiu, W.T.; Shen, M.R. The STIM1-Orai1 pathway of store-operated Ca²⁺ entry controls the checkpoint in cell cycle G1/S transition. Sci. Rep. 2016, 6, 22142. [Google Scholar] [CrossRef] [Green Version]
Jardin, I.; Lopez, J.J.; Salido, G.M.; Rosado, J.M. Store-Operated Ca²⁺ Entry in Breast Cancer Cells: Remodeling and Functional Role. Int. J. Mol. Sci. 2018, 19, 4053. [Google Scholar] [CrossRef] [Green Version]
Yang, S.; Zhang, J.J.; Huang, X.Y. Orai1 and STIM1 are critical for breast tumor cell migration and metastasis. Cancer Cell 2009, 15, 124–134. [Google Scholar] [CrossRef] [Green Version]
El Boustany, C.; Bidaux, G.; Enfissi, A.; Delcourt, P.; Prevarskaya, N.; Capiod, T. Capacitative Calcium Entry and Transient Receptor Potential Canonical 6 Expression Control Human Hepatoma Cell Proliferation. Hepatology 2008, 47, 2068–2077. [Google Scholar] [CrossRef]
Feng, M.; Grice, D.M.; Faddy, H.M.; Nguyen, N.; Leitch, S.; Wang, Y.; Muend, S.; Kenny, P.A.; Sukumar, S.; Roberts-Thomson, S.J.; et al. Store-independent activation of Orai1 by SPCA2 in mammary tumors. Cell 2010, 143, 84–98. [Google Scholar] [CrossRef] [Green Version]
Li, G.; Zhang, Z.; Wang, R.; Ma, W.; Yang, Y.; Wei, J.; Wei, Y. Suppression of STIM1 inhibits human glioblastoma cell proliferation and induces G0/G1 phase arrest. J. Exp. Clin. Cancer Res. 2013, 32, 20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Chen, Y.F.; Chiu, W.T.; Chen, Y.T.; Lin, P.Y.; Huang, H.J.; Chou, C.Y.; Chang, H.C.; Tang, M.J.; Shen, M.R. Calcium store sensor stromal-interaction molecule 1-dependent signaling plays an important role in cervical cancer growth, migration, and angiogenesis. Proc. Natl. Acad. Sci. USA 2011, 31, 203–211. [Google Scholar] [CrossRef] [Green Version]
Zang, J.; Zuo, D.; Shogren, K.L.; Gustafson, C.T.; Zhou, Z.; Thompson, M.A.; Guo, R.; Prakash, Y.S.; Lu, L.; Guo, W.; et al. STIM1 expression is associated with osteosarcoma cell survival. Chin J Cancer Res. 2019, 31, 203–211. [Google Scholar] [CrossRef]
Cheng, H.; Wang, S.-Q.; Feng, R. STIM1 plays an important role in TGF-β-induced suppression of breast cancer cell proliferation. Oncotarget 2016, 7, 16866–16878. [Google Scholar] [CrossRef] [Green Version]
Ge, C.; Zeng, B.; Li, R.; Li, Z.; Fu, Q.; Wang, W.; Wang, Z.; Dong, S.; Lai, Z.; Wang, Y.; et al. Knockdown of STIM1 expression inhibits non-small-cell lung cancer cell proliferation in vitro and in nude mouse xenografts. Bioengineered 2019, 10, 425–436. [Google Scholar] [CrossRef] [Green Version]
Motiani, R.K.; Zhang, X.X.; Harmon, K.E.; Keller, R.S.; Matrougui, K.; Bennett, J.A.; Trebak, M. Orai3 is an estrogen receptor α-regulated Ca 2+ channel that promotes tumorigenesis. FASEB J. 2013, 27, 63–75. [Google Scholar] [CrossRef] [Green Version]
Benzerdjeb, N.; Sevestre, H.; Ahidouch, A.; Ouadid-Ahidouch, H. Orai3 is a predictive marker of metastasis and survival in resectable lung adenocarcinoma. Oncotarget 2016, 7, 81588–81597. [Google Scholar] [CrossRef] [Green Version]
Dubois, C.; Abeele, F.V.; Lehen’Kyi, V.; Gkika, D.; Guarmit, B.; Lepage, G.; Slomianny, C.; Borowiec, A.S.; Bidaux, G.; Benahmed, M.; et al. Remodeling of Channel-Forming ORAI Proteins Determines an Oncogenic Switch in Prostate Cancer. Cancer Cell 2014, 26, 19–32. [Google Scholar] [CrossRef] [Green Version]
Stanisz, H.; Saul, S.; Müller, C.S.L.; Kappl, R.; Niemeyer, B.A.; Vogt, T.; Hoth, M.; Roesch, A.; Bogeski, I. Inverse regulation of melanoma growth and migration by Orai1/STIM2-dependent calcium entry. Pigment. Cell Melanoma Res. 2014, 27, 442–453. [Google Scholar] [CrossRef]
Fiorio Pla, A.; Kondratska, K.; Prevarskaya, N. STIM and ORAI proteins: Crucial roles in hallmarks of cancer. Am. J. Physiol. Cell Physiol. 2016, 310, C509–C519. [Google Scholar] [CrossRef]
Kondratska, K.; Kondratskyi, A.; Yassine, M.; Lemonnier, L.; Lepage, G.; Morabito, A.; Skryma, R.; Prevarskaya, N. Orai1 and STIM1 mediate SOCE and contribute to apoptotic resistance of pancreatic adenocarcinoma. Biochim. Biophys. Acta (BBA)—Bioenerg. 2014, 1843, 2263–2269. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Schmidt, S.; Liu, G.; Liu, G.; Yang, W.; Honisch, S.; Pantelakos, S.; Stournaras, C.; Hönig, A.; Lang, F. Enhanced Orai1 and STIM1 expression as well as store operated Ca²⁺ entry in therapy resistant ovary carcinoma cells. Oncotarget 2014, 5, 4799–4810. [Google Scholar] [CrossRef] [Green Version]
Gualdani, R.; De Clippele, M.; Ratbi, I.; Gailly, P.; Tajeddine, N. Store-Operated Calcium Entry Contributes to Cisplatin-Induced Cell Death in Non-Small Cell Lung Carcinoma. Cancers 2019, 11, 430. [Google Scholar] [CrossRef] [Green Version]
Flourakis, M.; Lehen’Kyi, V.; Beck, B.; Raphael, M.; Vandenberghe, M.; Van Denabeele, F.; Roudbaraki, M.; Lepage, G.; Mauroy, B.; Romanin, C.; et al. Orai1 contributes to the establishment of an apoptosis-resistant phenotype in prostate cancer cells. Cell Death Dis. 2010, 1, e75. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Li, Q.; Wang, J.; He, Y.-Q.; Feng, C.; Zhang, X.-J.; Sheng, J.-Q.; Li, P.-F. MicroRNA-185 regulates chemotherapeutic sensitivity in gastric cancer by targeting apoptosis repressor with caspase recruitment domain. Cell Death Dis. 2014, 5, e1197. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Fedida-Metula, S.; Feldman, B.; Koshelev, V.; Levin-Gromiko, U.; Voronov, E.; Fishman, D. Lipid rafts couple store-operated Ca²⁺ entry to constitutive activation of PKB/Akt in a Ca²⁺/calmodulin-, Src- and PP2A-mediated pathway and promote melanoma tumor growth. Carcinogenesis 2012, 33, 740–750. [Google Scholar] [CrossRef] [Green Version]
Mimura, N.; Hideshima, T.; Shimomura, T.; Suzuki, R.; Ohguchi, H.; Rizq, O.; Kikuchi, S.; Yoshida, Y.; Cottini, F.; Jakubikova, J.; et al. Selective and potent Akt inhibition triggers anti-myeloma activities and enhances fatal endoplasmic reticulum stress induced by proteasome inhibition. Oncotarget 2014, 74, 4458–4469. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Wang, W.; Ren, Y.; Wang, L.; Zhao, W.; Dong, X.; Pan, J.; Gao, X.; Tian, Y. Orai1 and Stim1 Mediate the Majority of Store-Operated Calcium Entry in Multiple Myeloma and Have Strong Implications for Adverse Prognosis. Cell. Physiol. Biochem. 2018, 48, 2273–2285. [Google Scholar] [CrossRef]
Zhan, Z.Y.; Zhong, L.X.; Feng, M.; Wang, J.F.; Liu, D.B.; Xiong, J.P. Over-expression of Orai1 mediates cell proliferation and associates with poor prognosis in human non-small cell lung carcinoma. Int. J. Clin. Exp. Pathol. 2015, 8, 5080–5088. [Google Scholar] [PubMed]
Khadra, N.; Bresson-Bepoldin, L.; Penna, A.; Chaigne-Delalande, B.; Ségui, B.; Levade, T.; Vacher, A.M.; Reiffers, J.; Ducret, T.; Moreau, J.F.; et al. CD95 triggers Orai1-mediated localized Ca²⁺ entry, regulates recruitment of protein kinase C (PKC) β2, and prevents death-inducing signaling complex formation. Proc. Natl. Acad. Sci. USA 2011, 108, 19072–19077. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Sobradillo, D.; Hernández-Morales, M.; Ubierna, D.; Moyer, M.P.; Núñez, L.; Villalobos, C. A Reciprocal Shift in Transient Receptor Potential Channel 1 (TRPC1) and Stromal Interaction Molecule 2 (STIM2) Contributes to Ca²⁺ Remodeling and Cancer Hallmarks in Colorectal Carcinoma Cells. J. Biol. Chem. 2014, 289, 28765–28782. [Google Scholar] [CrossRef] [Green Version]
Hasna, J.; Hague, F.; Rodat-Despoix, L.; Geerts, D.; Leroy, C.; Tulasne, D.; Ouadid-Ahidouch, H.; Kischel, P. Orai3 calcium channel and resistance to chemotherapy in breast cancer cells: The p53 connection. Cell Death Differ. 2018, 25, 691–705. [Google Scholar] [CrossRef]
Zhu, H.; Zhang, H.; Jin, F.; Fang, M.; Huang, M.; Yang, C.S.; Chen, T.; Fu, L.; Pan, Z. Elevated Orai1 Expression Mediates Tumor-Promoting Intracellular Ca²⁺ Oscillations in Human Esophageal Squamous Cell Carcinoma. Oncotarget 2014, 5, 3455–3471. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Didiasova, M.; Zakrzewicz, D.; Magdolen, V.; Nagaraj, C.; Bálint, Z.; Rohde, M.; Preissner, K.T.; Wygrecka, M. STIM1/ORAI1-mediated Ca²⁺ Influx Regulates Enolase-1 Exteriorization. J. Biol. Chem. 2015, 290, 11983–11999. [Google Scholar] [CrossRef] [Green Version]
Chen, Y.F.; Hsu, K.F.; Shen, M.R. The Store-Operated Ca²⁺ Entry-Mediated Signaling Is Important for Cancer Spread. Biochim. Biophys. Acta 2016, 1863, 1427–1435. [Google Scholar] [CrossRef] [PubMed]
Mo, P.; Yang, S. The Store-Operated Calcium Channels in Cancer Metastasis: From Cell Migration, Invasion to Metastatic Colonization. Front. Biosci. 2018, 23, 1241–1256. [Google Scholar] [CrossRef] [Green Version]
McAndrew, D.; Grice, D.M.; Peters, A.A.; Davis, F.M.; Stewart, T.; Rice, M.; Smart, C.E.; Brown, M.A.; Kenny, P.A.; Roberts-Thomson, S.J.; et al. ORAI1-mediated Calcium Influx in Lactation and in Breast Cancer. Mol. Cancer Ther. 2011, 10, 448–460. [Google Scholar] [CrossRef] [Green Version]
Moccia, F. Endothelial Ca²⁺ Signaling and the Resistance to Anticancer Treatments: Partners in Crime. Int. J. Mol. Sci. 2018, 19, 217. [Google Scholar] [CrossRef] [Green Version]
Li, J.; Cubbon, R.M.; Wilson, L.A.; Amer, M.S.; McKeown, L.; Hou, B.; Majeed, Y.; Tumova, S.; Seymour, V.A.L.; Taylor, H.; et al. Orai1 and CRAC channel dependence of VEGF-activated Ca2entry and endothelial tube formation. Circ. Res. 2011, 108, 1190–1198. [Google Scholar] [CrossRef] [Green Version]
Li, J.; Bruns, A.F.; Hou, B.; Rode, B.; Webster, P.J.; Bailey, M.A.; Appleby, H.L.; Moss, N.K.; Ritchie, J.E.; Yuldasheva, N.Y.; et al. Orai3 Surface Accumulation and Calcium Entry Evoked by Vascular Endothelial Growth Factor. Arterioscler. Thromb. Vasc. Biol. 2015, 35, 1987–1994. [Google Scholar] [CrossRef] [Green Version]
Hoth, M.; Penner, R. Depletion of intracellular calcium stores activates a calcium current in mast cells. Nature 1992, 355, 353–356. [Google Scholar] [CrossRef] [PubMed]
Zweifach, A.; Lewis, R.S. Mitogen-regulated Ca²⁺ current of T lymphocytes is activated by depletion of intracellular Ca²⁺ stores. Proc. Natl. Acad. Sci. USA 1993, 90, 6295–6299. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Weidinger, C.; Shaw, P.J.; Feske, S. STIM1 and STIM2-mediated Ca²⁺ influx regulates antitumour immunity by CD8+ T cells. EMBO Mol. Med. 2013, 5, 1311–1321. [Google Scholar] [CrossRef]
Zhou, X.; Friedmann, K.S.; Lyrmann, H.; Zhou, Y.; Schoppmeyer, R.; Knörck, A.; Mang, S.; Hoxha, C.; Angenendt, A.; Backes, C.S.; et al. A calcium optimum for cytotoxic T lymphocyte and natural killer cell cytotoxicity. J. Physiol. 2018, 596, 2681–2698. [Google Scholar] [CrossRef] [Green Version]
Endo, M. Calcium Release from the Sarcoplasmic Reticulum. Physiol. Rev. 1977, 57, 71–108. [Google Scholar] [CrossRef]
Courjaret, R.; Dib, M.; Machaca, K. Spatially restricted subcellular Ca²⁺ signaling downstream of store-operated calcium entry encoded by a cortical tunneling mechanism. Sci. Rep. 2018, 8, 11214. [Google Scholar] [CrossRef]
Thillaiappan, N.B.; Chavda, A.P.; Tovey, S.C.; Prole, D.L.; Taylor, C.W. Ca²⁺ signals initiate at immobile IP3 receptors adjacent to ER-plasma membrane junctions. Nat. Commun. 2017, 8, 1505. [Google Scholar] [CrossRef]
Rizzuto, R.; Pinton, P.; Carrington, W.; Fay, F.S.; Fogarty, K.E.; Lifshitz, L.M.; Tuft, R.A.; Pozzan, T. Close Contacts with the Endoplasmic Reticulum as Determinants of Mitochondrial Ca²⁺ Responses. Science 1998, 280, 1763–1766. [Google Scholar] [CrossRef]
Mandeville, J.T.H.; Ghosh, R.N.; Maxfield, F.R. Intracellular calcium levels correlate with speed and persistent forward motion in migrating neutrophils. Biophys. J. 1995, 68, 1207–1217. [Google Scholar] [CrossRef] [Green Version]
Ritaine, A.; Shapovalov, G.; Prevarskaya, N. Metabolic Disorders and Cancer: Store-Operated Ca²⁺ Entry in Cancer: Focus on IP 3 R-Mediated Ca²⁺ Release from Intracellular Stores and Its Role in Migration and Invasion. Adv. Exp. Med. Biol. 2017, 993, 623–637. [Google Scholar] [CrossRef] [PubMed]
Wei, C.; Wang, X.; Chen, M.; Ouyang, K.; Song, L.S.; Cheng, H. Calcium flickers steer cell migration. Nature 2009, 457, 901–905. [Google Scholar] [CrossRef] [Green Version]
Brundage, R.A.; Fogarty, K.E.; Tuft, R.A.; Fay, F.A. Calcium gradients underlying polarization and chemotaxis of eosinophils. Science 1991, 254, 703–706. [Google Scholar] [CrossRef]
Okeke, E.; Parker, T.; Dingsdale, H.; Concannon, M.; Awais, M.; Voronina, S.; Molgó, J.; Begg, M.; Metcalf, D.; Knight, A.E.; et al. Epithelial–mesenchymal transition, IP3 receptors and ER–PM junctions: Translocation of Ca²⁺ signalling complexes and regulation of migration. Biochem. J. 2016, 473, 757–767. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Baljinnyam, E.; De Lorenzo, M.S.; Xie, L.H.; Iwatsubo, M.; Chen, S.; Goydos, J.S.; Nowycky, M.C.; Iwatsubo, K. Exchange Protein Directly Activated by Cyclic AMP Increases Melanoma Cell Migration by a Ca²⁺-dependent Mechanism. Cancer Res. 2010, 70, 5607–5617. [Google Scholar] [CrossRef] [Green Version]
Jin, X.; Shah, S.; Liu, Y.; Zhang, H.; Lees, M.; Fu, Z.; Lippiat, J.D.; Beech, D.J.; Sivaprasadarao, A.; Baldwin, S.A.; et al. Activation of the Cl- Channel ANO1 by Localized Calcium Signals in Nociceptive Sensory Neurons Requires Coupling With the IP3 Receptor. Sci. Signal. 2013, 6, ra73. [Google Scholar] [CrossRef] [Green Version]
Ruiz, C.; Martins, J.R.; Rudin, F.; Schneider, S.; Dietsche, T.; Fischer, C.A.; Tornillo, L.; Terracciano, L.M.; Schreiber, R.; Bubendorf, L.; et al. Enhanced Expression of ANO1 in Head and Neck Squamous Cell Carcinoma Causes Cell Migration and Correlates with Poor Prognosis. PLoS ONE. 2012, 7, e43265. [Google Scholar] [CrossRef] [Green Version]
Zeng, X.; Pan, D.; Wu, H.; Chen, H.; Yuan, W.; Zhou, J.; Shen, Z.; Chen, S. Transcriptional Activation of ANO1 Promotes Gastric Cancer Progression. Biochem. Biophys. Res. Commun. 2019, 512, 131–136. [Google Scholar] [CrossRef]
Lee, Y.S.; Lee, J.K.; Bae, Y.; Lee, B.S.; Kim, E.; Cho, C.H.; Ryoo, K.; Yoo, J.; Kim, C.H.; Yi, G.S.; et al. Suppression of 14-3-3γ-mediated surface expression of ANO1 inhibits cancer progression of glioblastoma cells. Sci. Rep. 2016, 6, 26413. [Google Scholar] [CrossRef] [Green Version]
Crottès, D.; Lin, Y.H.T.; Peters, C.J.; Gilchrist, J.M.; Wiita, A.P.; Jan, Y.N.; Jan, L.Y. TMEM16A Controls EGF-induced Calcium Signaling Implicated in Pancreatic Cancer Prognosis. Proc. Natl. Acad. Sci. USA 2019, 116, 13026–13035. [Google Scholar] [CrossRef] [Green Version]
Britschgi, A.; Bill, A.; Brinkhaus, H.; Rothwell, C.; Clay, I.; Duss, S.; Rebhan, M.; Raman, P.; Guy, C.T.; Wetzel, K.; et al. Calcium-activated chloride channel ANO1 promotes breast cancer progression by activating EGFR and CAMK signaling. Proc. Natl. Acad. Sci. USA 2013, 110, E1026–E1034. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Sakakura, C.; Hagiwara, A.; Fukuda, K.; Shimomura, K.; Takagi, T.; Kin, S.; Nakase, Y.; Fujiyama, J.; Mikoshiba, K.; Okazaki, Y.; et al. Possible involvement of inositol 1,4,5-trisphosphate receptor type 3 (IP3R3) in the peritoneal dissemination of gastric cancers. Anticancer Res. 2003, 23, 3691–3697. [Google Scholar]
Courjaret, R.; Machaca, K. Mid-range Ca²⁺ Signalling Mediated by Functional Coupling Between Store-Operated Ca²⁺ Entry and IP3-dependent Ca²⁺ Release. Nat. Commun. 2014, 5, 3916. [Google Scholar] [CrossRef] [Green Version]
Sui, Y.; Sun, M.; Wu, F.; Yang, L.; Di, W.; Zhang, G.; Zhong, L.; Ma, Z.; Zheng, J.; Fang, X.; et al. Inhibition of TMEM16A Expression Suppresses Growth and Invasion in Human Colorectal Cancer Cells. PLoS ONE 2014, 9, e115443. [Google Scholar] [CrossRef]
Prevarskaya Na Zhang, L.; Barritt, G. TRP channels in cancer. Biochim. Biophys. Acta 2007, 1772, 937–946. [Google Scholar] [CrossRef] [Green Version]
Pla, A.F.; Gkika, D. Emerging role of TRP channels in cell migration: From tumor vascularization to metastasis. Front. Physiol. 2013, 4, 311. [Google Scholar] [CrossRef] [Green Version]
Shapovalov, G.; Ritaine, A.; Skryma, R.; Prevarskaya, N. Role of TRP Ion Channels in Cancer and Tumorigenesis. Semin. Immunopathol. 2016, 38, 357–369. [Google Scholar] [CrossRef] [PubMed]
Ma, X.; Cai, Y.; He, D.; Zou, C.; Zhang, P.; Lo, C.Y.; Xu, Z.; Chan, F.L.; Yu, S.; Chen, Y.; et al. Transient receptor potential channel TRPC5 is essential for P-glycoprotein induction in drug-resistant cancer cells. Proc. Natl. Acad. Sci. USA 2012, 109, 16282–16287. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Yang, D. Emerging role of transient receptor potential (TRP) channels in cancer progression. BMB Rep. 2020, 53, 125–132. [Google Scholar] [CrossRef] [Green Version]
Zhou, K.; Zhang, S.-S.; Yan, Y.; Zhao, S. Overexpression of transient receptor potential vanilloid 2 is associated with poor prognosis in patients with esophageal squamous cell carcinoma. Med. Oncol. 2014, 31, 17. [Google Scholar] [CrossRef]
Semenova, S.B.; Vassilieva, I.; Fomina, A.F.; Runov, A.L.; Negulyaev, Y. Endogenous expression of TRPV5 and TRPV6 calcium channels in human leukemia K562 cells. Am. J. Physiol. Physiol. 2009, 296, C1098–C1104. [Google Scholar] [CrossRef]
Bödding, M.; Wissenbach, U.; Flockerzi, V. The Recombinant Human TRPV6 Channel Functions as Ca²⁺Sensor in Human Embryonic Kidney and Rat Basophilic Leukemia Cells. J. Biol. Chem. 2002, 277, 36656–36664. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Weber, L.V.; Al-Refae, K.; Wölk, G.; Bonatz, G.; Altmüller, J.; Becker, C.; Gisselmann, G.; Hatt, H. Expression and functionality of TRPV1 in breast cancer cells. Breast Cancer 2016, 8, 243–252. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Elzamzamy, O.M.; Penner, R.; Hazlehurst, L.A. The Role of TRPC1 in Modulating Cancer Progression. Cells 2020, 9, 388. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Grant, C.V.; Carver, C.M.; Hastings, S.D.; Ramachandran, K.; Muniswamy, M.; Risinger, A.L.; Beutler, J.A.; Mooberry, S.L. Triple-negative breast cancer cell line sensitivity to englerin A identifies a new, targetable subtype. Breast Cancer Res. Treat. 2019, 177, 345–355. [Google Scholar] [CrossRef]
Santoni, G.; Maggi, F.; Morelli, M.B.; Santoni, M.; Marinelli, O. Transient Receptor Potential Cation Channels in Cancer Therapy. Med. Sci. 2019, 7, 108. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Liu, J.; Hu, G.; Gong, Y.; Yu, Q.; He, B.; Li, W.; He, Z.; Hao, W.; He, Z.; Liu, Y. Silencing of TRPM8 inhibits aggressive tumor phenotypes and enhances gemcitabine sensitivity in pancreatic cancer. Pancreatology 2018, 18, 935–944. [Google Scholar] [CrossRef]
Nazıroğlu, M.; Çiğ, B.; Blum, W.; Vizler, C.; Buhala, A.; Marton, A.; Katona, R.; Jósvay, K.; Schwaller, B.; Oláh, Z.; et al. Targeting breast cancer cells by MRS1477, a positive allosteric modulator of TRPV1 channels. PLoS ONE 2017, 12, e0179950. [Google Scholar] [CrossRef] [Green Version]
Amantini, C.; Mosca, M.; Nabissi, M.; Lucciarini, R.; Caprodossi, S.; Arcella, A.; Giangaspero, F.; Santoni, G. Capsaicin-induced apoptosis of glioma cells is mediated by TRPV1 vanilloid receptor and requires p38 MAPK activation. J. Neurochem. 2007, 102, 977–990. [Google Scholar] [CrossRef] [Green Version]
Canales, J.; Morales, D.; Blanco, C.; Rivas, J.; Díaz, N.; Angelopoulos, I.; Cerda, O. A TR(i)P to Cell Migration: New Roles of TRP Channels in Mechanotransduction and Cancer. Front. Physiol. 2019, 10, 757. [Google Scholar] [CrossRef]
Chen, J.-P.; Wang, J.; Luan, Y.; Wang, C.-X.; Li, W.-H.; Zhang, J.-B.; Sha, D.; Shen, R.; Cui, Y.-G.; Zhang, Z.; et al. TRPM7 promotes the metastatic process in human nasopharyngeal carcinoma. Cancer Lett. 2015, 356, 483–490. [Google Scholar] [CrossRef] [PubMed]
Rybarczyk, P.; Gautier, M.; Hague, F.; Dhennin-Duthille, I.; Chatelain, D.; Kerr-Conte, J.; Pattou, F.; Regimbeau, J.-M.; Sevestre, H.; Ouadid-Ahidouch, H. Transient receptor potential melastatin-related 7 channel is overexpressed in human pancreatic ductal adenocarcinomas and regulates human pancreatic cancer cell migration. Int. J. Cancer 2012, 131, E851–E861. [Google Scholar] [CrossRef]
Middelbeek, J.; Kuipers, A.J.; Henneman, L.; Visser, D.; Eidhof, I.; Van Horssen, R.; Wieringa, B.; Canisius, S.V.; Zwart, W.; Wessels, L.F.; et al. TRPM7 Is Required for Breast Tumor Cell Metastasis. Cancer Res. 2012, 72, 4250–4261. [Google Scholar] [CrossRef] [Green Version]
Lee, W.H.; Choong, L.Y.; Jin, T.H.; Mon, N.N.; Chong, S.; Liew, C.S.; Putti, T.; Lu, S.Y.; Harteneck, C.; Lim, Y.P. TRPV4 plays a role in breast cancer cell migration via Ca²⁺-dependent activation of AKT and downregulation of E-cadherin cell cortex protein. Oncog. 2017, 6, e338. [Google Scholar] [CrossRef] [PubMed]
Lee, W.H.; Choong, L.Y.; Mon, N.N.; Lu, S.; Lin, Q.; Pang, B.; Yan, B.; Krishna, V.S.R.; Singh, H.; Tan, T.Z.; et al. TRPV4 Regulates Breast Cancer Cell Extravasation, Stiffness and Actin Cortex. Sci. Rep. 2016, 6, 27903. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Lodish, H.; Kong, N.; Wikström, L. Calcium is required for folding of newly made subunits of the asialoglycoprotein receptor within the endoplasmic reticulum. J. Biol. Chem. 1992, 267, 12753–12760. [Google Scholar] [CrossRef]
Dang, D.; Rao, R. Calcium-ATPases: Gene disorders and dysregulation in cancer. Biochim. Biophys. Acta (BBA)—Bioenerg. 2016, 1863, 1344–1350. [Google Scholar] [CrossRef] [PubMed]
Chung, F.-Y.; Lin, S.-R.; Lu, C.-Y.; Yeh, C.-S.; Chen, F.-M.; Hsieh, J.-S.; Huang, T.-J.; Wang, J.-Y. Sarco/Endoplasmic Reticulum Calcium-ATPase 2 Expression as a Tumor Marker in Colorectal Cancer. Am. J. Surg. Pathol. 2006, 30, 969–974. [Google Scholar] [CrossRef]
Gou, W.-F.; Niu, Z.-F.; Zhao, S.; Takano, Y.; Zheng, H.-C. Aberrant SERCA3 expression during the colorectal adenoma-adenocarcinoma sequence. Oncol. Rep. 2014, 31, 232–240. [Google Scholar] [CrossRef] [Green Version]
Bergner, A.; Kellner, J.; Tufman, A.; Huber, R.M. Endoplasmic reticulum Ca²⁺-homeostasis is altered in small and non-small cell lung cancer cell lines. J. Exp. Clin. Cancer Res. 2009, 28, 25. [Google Scholar] [CrossRef] [Green Version]
Arbabian, A.; Brouland, J.P.; Apáti, Á.; Pászty, K.; Hegedűs, L.; Enyedi, Á.; Chomienne, C.; Papp, B. Modulation of endoplasmic reticulum calcium pump expression during lung cancer cell differentiation. FEBS J. 2013, 280, 5408–5418. [Google Scholar] [CrossRef] [Green Version]
O’Neill, J.P.; Velalar, C.N.; Lee, D.I.; Zhang, B.; Nakanishi, T.; Tang, Y.; Selaru, F.; Ross, D.; Meltzer, S.J.; Hussain, A. Thapsigargin resistance in human prostate cancer cells. Cancer 2006, 107, 649–659. [Google Scholar] [CrossRef]
Brouland, J.P.; Gélébart, P.; Kovàcs, T.; Enouf, J.; Grossmann, J.; Papp, B. The Loss of Sarco/Endoplasmic Reticulum Calcium Transport ATPase 3 Expression Is an Early Event during the Multistep Process of Colon Carcinogenesis. Am. J. Pathol. 2005, 167, 233–242. [Google Scholar] [CrossRef] [Green Version]
Launay, S.; Giannì, M.; Kovàcs, T.; Bredoux, R.; Bruel, A.; Gélébart, P.; Zassadowski, F.; Chomienne, C.; Enouf, J.; Papp, B. Lineage-specific Modulation of Calcium Pump Expression During Myeloid Differentiation. Blood 1999, 93, 4395–4405. [Google Scholar] [CrossRef]
Gélébart, P.; Kovács, T.; Brouland, J.P.; van Gorp, R.; Grossmann, J.; Rivard, N.; Panis, Y.; Martin, V.; Bredoux, R.; Enouf, J.; et al. Expression of Endomembrane Calcium Pumps in Colon and Gastric Cancer Cells. J. Biol. Chem. 2002, 277, 26310–26320. [Google Scholar] [CrossRef] [Green Version]
Roti, G.; Carlton, A.; Ross, K.N.; Markstein, M.; Pajcini, K.; Su, A.H.; Perrimon, N.; Pear, W.S.; Kung, A.; Blacklow, S.C.; et al. Complementary Genomic Screens Identify SERCA as a Therapeutic Target in NOTCH1 Mutated Cancer. Cancer Cell 2013, 23, 390–405. [Google Scholar] [CrossRef] [Green Version]
Park, K.C.; Kim, S.W.; Jeon, J.Y.; Jo, A.R.; Choi, H.J.; Kim, J.; Lee, H.G.; Kim, Y.; Mills, G.; Noh, S.H.; et al. Survival of Cancer Stem-Like Cells Under Metabolic Stress via CaMK2α-mediated Upregulation of Sarco/Endoplasmic Reticulum Calcium ATPase Expression. Clin. Cancer Res. 2017, 24, 1677–1690. [Google Scholar] [CrossRef] [Green Version]
Mahalingam, D.; Peguero, J.; Cen, P.; Arora, S.P.; Sarantopoulos, J.; Rowe, J.; Allgood, V.; Tubb, B.; Campos, L. A Phase II, Multicenter, Single-Arm Study of Mipsagargin (G-202) as a Second-Line Therapy Following Sorafenib for Adult Patients with Progressive Advanced Hepatocellular Carcinoma. Cancers 2019, 11, 833. [Google Scholar] [CrossRef] [Green Version]
Berchtold, M.W.; Villalobo, A. The many faces of calmodulin in cell proliferation, programmed cell death, autophagy, and cancer. Biochim. Biophys. Acta (BBA)—Bioenerg. 2014, 1843, 398–435. [Google Scholar] [CrossRef]
Colomer, J.; Agell, N.; Engel, P.; Bachs, O. Expression of calmodulin and calmodulin binding proteins in lymphoblastoid cells. J. Cell. Physiol. 1994, 159, 542–550. [Google Scholar] [CrossRef]
Ye, Q.; Wei, Y.; Fischer, R.; Borner, C.; Berchtold, M.W. Expression of calmodulin and calmodulin binding proteins in rat fibroblasts stably transfected with protein kinase C and oncogenes. Biochim. Biophys. Acta (BBA)—Bioenerg. 1997, 1359, 89–96. [Google Scholar] [CrossRef] [Green Version]
Krishnaraju, K.; Murugesan, K.; Vij, U.; Kapur, B.; Farooq, A. Calmodulin levels in oestrogen receptor positive and negative human breast tumours. Br. J. Cancer 1991, 63, 346–347. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Wei, J.W.; Morris, H.P.; A Hickie, R. Positive correlation between calmodulin content and hepatoma growth rates. Cancer Res. 1982, 42, 2571–2574. [Google Scholar] [PubMed]
Veigl, M.L.; Vanaman, T.C.; E Branch, M.; Sedwick, W.D. Differences in calmodulin levels of normal and transformed cells as determined by culture conditions. Cancer Res. 1984, 44, 3184–3189. [Google Scholar] [PubMed]
Wang, R.; Zhang, H.; Li, S.; Xue, S. Intracellular levels of calmodulin are increased in transformed cells. Cell Res. 1992, 2, 119–127. [Google Scholar] [CrossRef]
Yokokura, S.; Yurimoto, S.; Matsuoka, A.; Imataki, O.; Dobashi, H.; Bandoh, S.; Matsunaga, T. Calmodulin antagonists induce cell cycle arrest and apoptosis in vitro and inhibit tumor growth in vivo in human multiple myeloma. BMC Cancer 2014, 14, 1–11. [Google Scholar] [CrossRef] [Green Version]
Skelding, K.A.; Rostas, J.A.P.; Verrills, N.M. Controlling the cell cycle: The role of calcium/calmodulin-stimulated protein kinases I and II. Cell Cycle 2011, 10, 631–639. [Google Scholar] [CrossRef] [Green Version]
Kahl, C.R.; Means, A.R. Regulation of Cyclin D1/Cdk4 Complexes by Calcium/Calmodulin-dependent Protein Kinase I. J. Biol. Chem. 2004, 279, 15411–15419. [Google Scholar] [CrossRef] [Green Version]
Patel, R.; Holt, M.; Philipova, R.; Moss, S.; Schulman, H.; Hidaka, H.; Whitaker, M. Calcium/Calmodulin-dependent Phosphorylation and Activation of Human Cdc25-C at the G2/M Phase Transition in HeLa Cells. J. Biol. Chem. 1999, 274, 7958–7968. [Google Scholar] [CrossRef] [Green Version]
Wang, Y.; Zhao, R.; Zhe, H. The emerging role of CaMKII in cancer. Oncotarget 2015, 6, 11725–11734. [Google Scholar] [CrossRef] [Green Version]
Brzozowski, J.S.; Skelding, K.A. The Multi-Functional Calcium/Calmodulin Stimulated Protein Kinase (CaMK) Family: Emerging Targets for Anti-Cancer Therapeutic Intervention. Pharmaceuticals 2019, 12, 8. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Li, Z.; Joyal, J.L.; Sacks, D. Calmodulin Enhances the Stability of the Estrogen Receptor. J. Biol. Chem. 2001, 276, 17354–17360. [Google Scholar] [CrossRef] [Green Version]
Li, Z.; Zhang, Y.; Hedman, A.C.; Ames, J.B.; Sacks, D.B. Calmodulin Lobes Facilitate Dimerization and Activation of Estrogen Receptor-α. J. Biol. Chem. 2017, 292, 4614–4622. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Cifuentes, E.; Mataraza, J.M.; Yoshida, B.A.; Menon, M.; Sacks, D.; Barrack, E.R.; Reddy, G.P.-V. Physical and functional interaction of androgen receptor with calmodulin in prostate cancer cells. Proc. Natl. Acad. Sci. USA 2004, 101, 464–469. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Li, X.; Wu, G.; Yang, Y.; Fu, S.; Liu, X.; Kang, H.; Yang, X.; Su, X.-C.; Shen, Y. Calmodulin dissociates the STIM1-Orai1 complex and STIM1 oligomers. Nat. Commun. 2017, 8, 1–14. [Google Scholar] [CrossRef] [PubMed]
Sun, Y.; Rossi, A.M.; Rahman, T.; Taylor, C. Activation of IP3 receptors requires an endogenous 1-8-14 calmodulin-binding motif. Biochem. J. 2012, 449, 39–49. [Google Scholar] [CrossRef] [Green Version]
Li, S.; Xue, J.; Sun, Z.; Liu, T.; Zhang, L.; Wang, L.; You, H.; Fan, Z.; Zheng, Y.; Luo, D. CaMKII Potentiates Store-Operated Ca²⁺ Entry Through Enhancing STIM1 Aggregation and Interaction with Orai1. Cell. Physiol. Biochem. 2018, 46, 1042–1054. [Google Scholar] [CrossRef] [PubMed]
Nussinov, R.; Wang, G.; Tsai, C.-J.; Jang, H.; Lu, S.; Banerjee, A.; Zhang, J.; Gaponenko, V. Calmodulin and PI3K Signaling in KRAS Cancers. Trends Cancer 2017, 3, 214–224. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Umemura, M.; Baljinnyam, E.; Feske, S.; De Lorenzo, M.S.; Xie, L.-H.; Feng, X.; Oda, K.; Makino, A.; Fujita, T.; Yokoyama, U.; et al. Store-Operated Ca²⁺ Entry (SOCE) Regulates Melanoma Proliferation and Cell Migration. PLoS ONE 2014, 9, e89292. [Google Scholar] [CrossRef] [PubMed]
Villalobo, A.; Berchtold, M.W. The Role of Calmodulin in Tumor Cell Migration, Invasiveness, and Metastasis. Int. J. Mol. Sci. 2020, 21, 765. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Li, T.; Yi, L.; Hai, L.; Ma, H.; Tao, Z.; Zhang, C.; Abeysekera, I.; Zhao, K.; Yang, Y.; Wang, W.; et al. The interactome and spatial redistribution feature of Ca²⁺ receptor protein calmodulin reveals a novel role in invadopodia-mediated invasion. Cell Death Dis. 2018, 9, 1–17. [Google Scholar] [CrossRef] [PubMed]
Choi, H.S.; Kim, D.-A.; Chung, H.; Park, I.H.; Kim, B.H.; Oh, E.-S.; Kang, D.-H. Screening of breast cancer stem cell inhibitors using a protein kinase inhibitor library. Cancer Cell Int. 2017, 17, 1–12. [Google Scholar] [CrossRef] [Green Version]
Kaur, A.; Raghavan, M. A Calreticulin Tail: C-terminal Mutants of Calreticulin Allow Cancer Cells to Evade Phagocytosis. Mol. Cell 2020, 77, 683–685. [Google Scholar] [CrossRef] [PubMed]
Venkateswaran, K.; Verma, A.; Bhatt, A.N.; Shrivastava, A.; Manda, K.; Raj, H.G.; Prasad, A.; Len, C.; Parmar, V.S.; Dwarakanath, B.S. Emerging Roles of Calreticulin in Cancer: Implications for Therapy. Curr. Protein Pept. Sci. 2018, 19, 344–357. [Google Scholar] [CrossRef] [PubMed]
Han, A.; Li, C.; Zahed, T.; Wong, M.; Smith, I.; Hoedel, K.; Green, D.; Boiko, A.D. Calreticulin is a Critical Cell Survival Factor in Malignant Neoplasms. PLoS Biol. 2019, 17, e3000402. [Google Scholar] [CrossRef]
Jin, C.; Lin, T.; Shan, L. Downregulation of Calbindin 1 by miR-454-3p Suppresses Cell Proliferation in Nonsmall Cell Lung Cancer In Vitro. Cancer Biother. Radiopharm. 2019, 34, 119–127. [Google Scholar] [CrossRef]
Pfyffer, G.E.; Humbel, B.; Sträuli, P.; Mohrmann, I.; Murer, H.; Heizmann, C.W. Calcium-binding proteins in carcinoma, neuroblastoma and glioma cell lines. Virchows Archiv 1987, 412, 135–144. [Google Scholar] [CrossRef]
Lu, Y.-C.; Weng, W.-C.; Lee, H. Functional Roles of Calreticulin in Cancer Biology. BioMed Res. Int. 2015, 2015, 1–9. [Google Scholar] [CrossRef]
Sun, J.; Mu, H.; Dai, K.; Yi, L. Calreticulin: A potential anti-cancer therapeutic target. Die Pharm. 2017, 72, 503–510. [Google Scholar]
Villagomez, M.; Szabo, E.; Podcheko, A.; Feng, T.; Papp, S.; Opas, M. Calreticulin and focal-contact-dependent adhesion. Biochem. Cell Biol. 2009, 87, 545–556. [Google Scholar] [CrossRef]
Huang, G.; Sun, Z.; Wu, J.; Shui, S.; Han, X.; Guo, D.; Li, T. Calreticulin Promotes Proliferation and Migration but Inhibits Apoptosis in Schwann Cells. Med. Sci. Monit. 2016, 22, 4516–4522. [Google Scholar] [CrossRef] [Green Version]
Clark, R.A.; Li, S.-L.; Pearson, D.W.; Leidal, K.G.; Clark, J.R.; Denning, G.M.; Reddick, R.; Krause, K.-H.; Valente, A.J. Regulation of Calreticulin Expression during Induction of Differentiation in Human Myeloid Cells. J. Biol. Chem. 2002, 277, 32369–32378. [Google Scholar] [CrossRef] [Green Version]
Martins, I.; Kepp, O.; Galluzzi, L.; Senovilla, L.; Schlemmer, F.; Adjemian, S.; Menger, L.; Michaud, M.; Zitvogel, L.; Kroemer, G. Surface-exposed calreticulin in the interaction between dying cells and phagocytes. Ann. N. Y. Acad. Sci. 2010, 1209, 77–82. [Google Scholar] [CrossRef]
Raghavan, M.; Wijeyesakere, S.J.; Peters, L.R.; Del Cid, N. Calreticulin in the immune system: Ins and outs. Trends Immunol. 2013, 34, 13–21. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Zamanian, M.; Hamadneh, L.A.Q.; Veerakumarasivam, A.; Rahman, S.A.; Shohaimi, S.; Rosli, R. Calreticulin mediates an invasive breast cancer phenotype through the transcriptional dysregulation of p53 and MAPK pathways. Cancer Cell Int. 2016, 16, 56. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Banerjea, A.; Ahmed, S.; E Hands, R.; Huang, F.; Han, X.; Shaw, P.M.; Feakins, R.; A Bustin, S.; Dorudi, S. Colorectal cancers with microsatellite instability display mRNA expression signatures characteristic of increased immunogenicity. Mol. Cancer 2004, 3, 21. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Harada, K.; Takenawa, T.; Ferdous, T.; Kuramitsu, Y.; Ueyama, Y. Calreticulin is a novel independent prognostic factor for oral squamous cell carcinoma. Oncol. Lett. 2017, 13, 4857–4862. [Google Scholar] [CrossRef] [Green Version]
Chiang, W.-F.; Hwang, T.-Z.; Hour, T.-C.; Wang, L.-H.; Chiu, C.-C.; Chen, H.-R.; Wu, Y.-J.; Wang, C.-C.; Wang, L.-F.; Chien, C.-Y.; et al. Calreticulin, an endoplasmic reticulum-resident protein, is highly expressed and essential for cell proliferation and migration in oral squamous cell carcinoma. Oral Oncol. 2013, 49, 534–541. [Google Scholar] [CrossRef] [PubMed]
Hacken, E.; Gounari, M.; Back, J.W.; Shimanovskaya, E.; Scarfò, L.; Kim, E.; Burks, J.; Ponzoni, M.; Ramirez, G.A.; Wierda, W.G.; et al. Calreticulin as a novel B-cell receptor antigen in chronic lymphocytic leukemia. Haematologica 2017, 102, e394–e396. [Google Scholar] [CrossRef] [Green Version]
Lwin, Z.-M.; Guo, C.; Salim, A.; Yip, G.W.-C.; Chew, F.-T.; Nan, J.; Thike, A.A.; Tan, P.-H.; Bay, B.-H. Clinicopathological significance of calreticulin in breast invasive ductal carcinoma. Mod. Pathol. 2010, 23, 1559–1566. [Google Scholar] [CrossRef] [Green Version]
Liu, R.; Gong, J.; Chen, J.; Li, Q.; Song, C.; Zhang, J.; Li, Y.; Liu, Z.; Dong, Y.; Chen, L.; et al. Calreticulin as a potential diagnostic biomarker for lung cancer. Cancer Immunol. Immunother. 2011, 61, 855–864. [Google Scholar] [CrossRef]
Alur, M.; Nguyen, M.M.; Eggener, S.E.; Jiang, F.; Dadras, S.S.; Stern, J.; Kimm, S.; Roehl, K.; Kozlowski, J.; Pins, M.; et al. Suppressive Roles of Calreticulin in Prostate Cancer Growth and Metastasis. Am. J. Pathol. 2009, 175, 882–890. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Hsu, W.-M.; Hsieh, F.J.; Jeng, Y.-M.; Kuo, M.L.; Chen, C.-N.; Lai, D.M.; Wang, B.T.; Tsao, P.-N.; Lee, H.; Lin, M.T.; et al. Calreticulin expression in neuroblastoma—A novel independent prognostic factor. Ann. Oncol. 2005, 16, 314–321. [Google Scholar] [CrossRef] [PubMed]
Sheng, W.; Chen, C.; Dong, M.; Zhou, J.; Liu, Q.; Dong, Q.; Li, F. Overexpression of Calreticulin Contributes to the Development and Progression of Pancreatic Cancer. J. Cell. Physiol. 2014, 229, 887–897. [Google Scholar] [CrossRef]
Lee, P.-C.; Chiang, J.-C.; Chen, C.-Y.; Chien, Y.-C.; Chen, W.-M.; Huang, C.-W.; Weng, W.-C.; Chen, C.-I.; Lee, P.-H.; Chen, C.-N.; et al. Calreticulin regulates vascular endothelial growth factor-A mRNA stability in gastric cancer cells. PLoS ONE 2019, 14, e0225107. [Google Scholar] [CrossRef] [PubMed]
Fadel, M.P.; Szewczenko-Pawlikowski, M.; Leclerc, P.; Dziak, E.; Symonds, J.M.; Blaschuk, O.; Michalak, M.; Opas, M. Calreticulin Affects β-Catenin-associated Pathways. J. Biol. Chem. 2001, 276, 27083–27089. [Google Scholar] [CrossRef] [Green Version]
Coppolino, M.G.; Woodside, M.J.; Demaurex, N.; Grinstein, S.; St-Arnaud, R.; Dedhar, S. Calreticulin is essential for integrin-mediated calcium signalling and cell adhesion. Nat. Cell Biol. 1997, 386, 843–847. [Google Scholar] [CrossRef]
Goicoechea, S.; Pallero, M.A.; Eggleton, P.; Michalak, M.; Murphy-Ullrich, J.E. The Anti-adhesive Activity of Thrombospondin Is Mediated by the N-terminal Domain of Cell Surface Calreticulin. J. Biol. Chem. 2002, 277, 37219–37228. [Google Scholar] [CrossRef] [Green Version]
Hayashida, Y.; Urata, Y.; Muroi, E.; Kono, T.; Miyata, Y.; Nomata, K.; Kanetake, H.; Kondo, T.; Ihara, Y. Calreticulin Represses E-cadherin Gene Expression in Madin-Darby Canine Kidney Cells via Slug. J. Biol. Chem. 2006, 281, 32469–32484. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Chen, C.; Su, T.; Lu, Y.; Lee, H. Calreticulin regulates cell proliferation and migration in gastric cancer cell line AGS. FASEB J. 2007, 21, A1318. [Google Scholar] [CrossRef]
Yi, L.; Shan, J.; Chen, X.; Li, G.; Li, L.; Tan, H.; Su, Q. Involvement of calreticulin in cell proliferation, invasion and differentiation in diallyl disulfide-treated HL-60 cells. Oncol. Lett. 2016, 12, 1861–1867. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Lu, Y.-C.; Chen, C.-N.; Wang, B.; Hsu, W.-M.; Chen, S.-T.; Chang, K.-J.; Chang, C.-C.; Lee, H. Changes in Tumor Growth and Metastatic Capacities of J82 Human Bladder Cancer Cells Suppressed by Down-Regulation of Calreticulin Expression. Am. J. Pathol. 2011, 179, 1425–1433. [Google Scholar] [CrossRef]
de Bruyn, M.; Wiersma, V.R.; Helfrich, W.; Eggleton, P.; Bremer, E. The Ever-Expanding Immunomodulatory Role of Calreticulin in Cancer Immunity. Front. Oncol. 2015, 5, 35. [Google Scholar] [CrossRef] [PubMed]
Okunaga, T.; Urata, Y.; Goto, S.; Matsuo, T.; Mizota, S.; Tsutsumi, K.; Nagata, I.; Kondo, T.; Ihara, Y.; Kendall, H.E.; et al. Calreticulin, a Molecular Chaperone in the Endoplasmic Reticulum, Modulates Radiosensitivity of Human Glioblastoma U251MG Cells. Cancer Res. 2006, 66, 8662–8671. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Matsukuma, S.; Yoshimura, K.; Ueno, T.; Oga, A.; Inoue, M.; Watanabe, Y.; Kuramasu, A.; Fuse, M.; Tsunedomi, R.; Nagaoka, S.; et al. Calreticulin is highly expressed in pancreatic cancer stem-like cells. Cancer Sci. 2016, 107, 1599–1609. [Google Scholar] [CrossRef] [PubMed]
Huang, Z.; Fan, G.; Wang, D. Downregulation of calbindin 1, a calcium-binding protein, reduces the proliferation of osteosarcoma cells. Oncol. Lett. 2017, 13, 3727–3733. [Google Scholar] [CrossRef] [Green Version]
Favier, J.; Brière, J.-J.; Burnichon, N.; Rivière, J.; Vescovo, L.; Benit, P.; Giscos-Douriez, I.; De Reyniès, A.; Bertherat, J.; Badoual, C.; et al. The Warburg Effect Is Genetically Determined in Inherited Pheochromocytomas. PLoS ONE 2009, 4, e7094. [Google Scholar] [CrossRef] [Green Version]
Seyfried, T.N.; Shelton, L.M. Cancer as a metabolic disease. Nutr. Metab. 2010, 7, 1–9. [Google Scholar] [CrossRef] [Green Version]
Hirschey, M.D.; DeBerardinis, R.J.; Diehl, A.M.E.; Drew, J.E.; Frezza, C.; Green, M.; Jones, L.W.; Ko, Y.H.; Le, A.; Lea, M.A.; et al. Dysregulated metabolism contributes to oncogenesis. Semin. Cancer Biol. 2015, 35, S129–S150. [Google Scholar] [CrossRef] [Green Version]
Amuthan, G.; Biswas, G.; Ananadatheerthavarada, H.K.; Vijayasarathy, C.; Shephard, H.M.; Avadhani, N.G. Mitochondrial stress-induced calcium signaling, phenotypic changes and invasive behavior in human lung carcinoma A549 cells. Oncogene 2002, 21, 7839–7849. [Google Scholar] [CrossRef] [Green Version]
Rizzuto, R.; Brini, M.; Murgia, M.; Pozzan, T. Microdomains with high Ca²⁺ close to IP3-sensitive channels that are sensed by neighboring mitochondria. Science 1993, 262, 744–747. [Google Scholar] [CrossRef]
Kania, E.; Roest, G.; Vervliet, T.; Parys, J.B.; Bultynck, G. IP3 Receptor-Mediated Calcium Signaling and Its Role in Autophagy in Cancer. Front. Oncol. 2017, 7, 140. [Google Scholar] [CrossRef] [Green Version]
Parys, J.B.; Decuypere, J.-P.; Bultynck, G. Role of the inositol 1,4,5-trisphosphate receptor/Ca²⁺-release channel in autophagy. Cell Commun. Signal. 2012, 10, 17. [Google Scholar] [CrossRef] [Green Version]
Vicencio, J.M.; Ortiz, C.; Criollo, A.; E Jones, A.W.; Kepp, O.; Galluzzi, L.; Joza, N.; Vitale, I.; Morselli, E.; Tailler, M.; et al. The inositol 1,4,5-trisphosphate receptor regulates autophagy through its interaction with Beclin 1. Cell Death Differ. 2009, 16, 1006–1017. [Google Scholar] [CrossRef] [Green Version]
Choi, K.S. Autophagy and cancer. Exp. Mol. Med. 2012, 44, 109–120. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Degenhardt, K.; Mathew, R.; Beaudoin, B.; Bray, K.; Anderson, D.; Chen, G.; Mukherjee, C.; Shi, Y.; Gélinas, C.; Fan, Y.; et al. Autophagy promotes tumor cell survival and restricts necrosis, inflammation, and tumorigenesis. Cancer Cell 2006, 10, 51–64. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Zheng, K.; He, Z.; Kitazato, K.; Wang, Y. Selective Autophagy Regulates Cell Cycle in Cancer Therapy. Theranostics 2019, 9, 104–125. [Google Scholar] [CrossRef]
Jones, R.G.; Plas, D.R.; Kubek, S.; Buzzai, M.; Mu, J.; Xu, Y.; Birnbaum, M.; Thompson, C.B. AMP-Activated Protein Kinase Induces a p53-Dependent Metabolic Checkpoint. Mol. Cell 2005, 18, 283–293. [Google Scholar] [CrossRef] [PubMed]
Mathiassen, S.G.; De Zio, D.; Cecconi, F. Autophagy and the Cell Cycle: A Complex Landscape. Front. Oncol. 2017, 7, 51. [Google Scholar] [CrossRef] [Green Version]
Sorokina, I.V.; Denisenko, T.V.; Imreh, G.; Tyurin-Kuzmin, P.A.; Kaminskyy, V.; Gogvadze, V.; Zhivotovsky, B. Involvement of autophagy in the outcome of mitotic catastrophe. Sci. Rep. 2017, 7, 1–12. [Google Scholar] [CrossRef]
Blandino, G.; Di Agostino, S. New therapeutic strategies to treat human cancers expressing mutant p53 proteins. J. Exp. Clin. Cancer Res. 2018, 37, 1–13. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Chen, H.-Y.; White, E. Role of Autophagy in Cancer Prevention. Cancer Prev. Res. 2011, 4, 973–983. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Li, X.; Zhou, Y.; Li, Y.; Yang, L.; Ma, Y.; Peng, X.; Yang, S.; Liu, J.; Li, H. Autophagy: A novel mechanism of chemoresistance in cancers. Biomed. Pharmacother. 2019, 119, 109415. [Google Scholar] [CrossRef] [PubMed]
Smith, A.; MacLeod, K.F. Autophagy, cancer stem cells and drug resistance. J. Pathol. 2019, 247, 708–718. [Google Scholar] [CrossRef] [Green Version]
Sui, X.; Chen, R.; Wang, Z.; Huang, Z.; Kong, N.; Zhang, M.; Han, W.; Lou, F.; Yang, J.; Zhang, Q.; et al. Autophagy and chemotherapy resistance: A promising therapeutic target for cancer treatment. Cell Death Dis. 2013, 4, e838. [Google Scholar] [CrossRef]
Liu, E.Y.; Ryan, K.M. Autophagy and cancer—Issues we need to digest. J. Cell Sci. 2012, 125, 2349–2358. [Google Scholar] [CrossRef] [Green Version]
Patergnani, S.; Missiroli, S.; Marchi, S.; Giorgi, C. Mitochondria-associated endoplasmic reticulum membranes microenvironment: Targeting autophagic and apoptotic pathways in cancer therapy. Front. Oncol. 2015, 5, 173. [Google Scholar] [CrossRef] [Green Version]
Pihán, P.; Carreras-Sureda, A.; Hetz, C. BCL-2 family: Integrating stress responses at the ER to control cell demise. Cell Death Differ. 2017, 24, 1478–1487. [Google Scholar] [CrossRef] [Green Version]
Lallemand, V.; Zhu, J.; Puvion, F.; Koken, M.; Honoré, N.; Doubeikovsky, A.; Duprez, E.; Pandolfi, P.P.; Puvion, E.; Freemont, P.; et al. Role of Promyelocytic Leukemia (Pml) Sumolation in Nuclear Body Formation, 11s Proteasome Recruitment, and as2O3-Induced Pml or Pml/Retinoic Acid Receptor α Degradation. J. Exp. Med. 2001, 193, 1361–1372. [Google Scholar] [CrossRef] [PubMed]
Missiroli, S.; Bonora, M.; Patergnani, S.; Poletti, F.; Perrone, M.; Gafà, R.; Magri, E.; Raimondi, A.; Lanza, G.; Tacchetti, C.; et al. PML at Mitochondria-Associated Membranes Is Critical for the Repression of Autophagy and Cancer Development. Cell Rep. 2016, 16, 2415–2427. [Google Scholar] [CrossRef] [Green Version]
Endoplasmic Reticulum-Mitochondria Ca ²⁺ Crosstalk in the Control of the Tumor Cell Fate. Biochim. Biophys. Acta Mol. Cell Res. 2017, 1864, 858–864. [CrossRef]
Bultynck, G.; Campanella, M. Tumor suppressive Ca²⁺ signaling is driven by IP3 receptor fitness. Cell Stress 2017, 1, 73–78. [Google Scholar] [CrossRef]
Hedgepeth, S.C.; Garcia, M.I.; Wagner, L.E., II; Rodriguez, A.M.; Chintapalli, S.V.; Snyder, R.R.; Hankins, G.D.V.; Henderson, B.R.; Brodie, K.M.; Yule, D.I.; et al. The BRCA1 Tumor Suppressor Binds to Inositol 1,4,5-Trisphosphate Receptors to Stimulate Apoptotic Calcium Release. J. Biol. Chem. 2015, 290, 7304–7313. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Nussinov, R.; Muratcioglu, S.; Tsai, C.-J.; Jang, H.; Gursoy, A.; Keskin, O. K-Ras4B/calmodulin/PI3Kα: A promising new adenocarcinoma-specific drug target? Expert Opin. Ther. Targets 2016, 20, 831–842. [Google Scholar] [CrossRef] [PubMed]
Chen, W.; An, P.; Quan, X.-J.; Zhang, J.; Zhou, Z.-Y.; Zou, L.-P.; Luo, H.-S. Ca²⁺/calmodulin-dependent protein kinase II regulates colon cancer proliferation and migration via ERK1/2 and p38 pathways. World J. Gastroenterol. 2017, 23, 6111–6118. [Google Scholar] [CrossRef] [PubMed]
Szabadkai, G.; Bianchi, K.; Várnai, P.; De Stefani, D.; Wieckowski, M.; Cavagna, D.; Nagy, A.I.; Balla, T.; Rizzuto, R. Chaperone-mediated coupling of endoplasmic reticulum and mitochondrial Ca²⁺ channels. J. Cell Biol. 2006, 175, 901–911. [Google Scholar] [CrossRef] [Green Version]
Rizzuto, R.; De Stefani, D.; Raffaello, A. Cristina Mammucari Mitochondria as sensors and regulators of calcium signaling. Nat. Rev. Mol. Cell Biol. 2012, 13, 566–578. [Google Scholar] [CrossRef] [PubMed]
Shoshan-Barmatz, V.; Mizrachi, D.; Mizrachi, D. VDAC1: From structure to cancer therapy. Front. Oncol. 2012, 2, 164. [Google Scholar] [CrossRef] [Green Version]
Shoshan-Barmatz, V.; Krelin, Y.; Shteinfer-Kuzmine, A.; Arif, T. Voltage-Dependent Anion Channel 1 As an Emerging Drug Target for Novel Anti-Cancer Therapeutics. Front. Oncol. 2017, 7, 154. [Google Scholar] [CrossRef]
Mathupala, S.P.; Ko, Y.H.; Pedersen, P.L. Hexokinase II: Cancer’s double-edged sword acting as both facilitator and gatekeeper of malignancy when bound to mitochondria. Oncogene 2006, 25, 4777–4786. [Google Scholar] [CrossRef] [Green Version]
Abu-Hamad, S.; Arbel, N.; Calo, D.; Arzoine, L.; Israelson, A.; Keinan, N.; Ben-Romano, R.; Friedman, O.; Shoshan-Barmatz, V. The VDAC1 N-terminus is essential both for apoptosis and the protective effect of anti-apoptotic proteins. J. Cell Sci. 2009, 122, 1906–1916. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Mallilankaraman, K.; Doonan, P.; Cárdenas, C.; Chandramoorthy, H.C.; Müller, M.; Miller, R.; Hoffman, N.E.; Gandhirajan, R.K.; Molgó, J.; Birnbaum, M.J.; et al. MICU1 is an essential gatekeeper for MCU-mediated mitochondrial Ca(²⁺) uptake that regulates cell survival. Cell 2012, 151, 630–644. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Plovanich, M.; Bogorad, R.L.; Sancak, Y.; Kamer, K.J.; Strittmatter, L.; Li, A.A.; Girgis, H.S.; Kuchimanchi, S.; De Groot, J.; Speciner, L.; et al. MICU2, a paralog of MICU1, resides within the mitochondrial uniporter complex to regulate calcium handling. PLoS ONE 2013, 8, e55785. [Google Scholar] [CrossRef] [Green Version]
Graier, W.F.; Malli, R. Mitochondrial calcium: A crucial hub for cancer cell metabolism? Transl. Cancer Res. 2017, 6, S1124–S1127. [Google Scholar] [CrossRef]
Curry, M.C.; Peters, A.A.; Kenny, P.A.; Roberts-Thomson, S.J.; Monteith, G.R. Mitochondrial calcium uniporter silencing potentiates caspase-independent cell death in MDA-MB-231 breast cancer cells. Biochem. Biophys. Res. Commun. 2013, 434, 695–700. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Tosatto, A.; Sommaggio, R.; Kummerow, C.; Bentham, R.B.; Blacker, T.; Berecz, T.; Duchen, M.; Rosato, A.; Bogeski, I.; Szabadkai, G.; et al. The mitochondrial calcium uniporter regulates breast cancer progression via HIF -1α. EMBO Mol. Med. 2016, 8, 569–585. [Google Scholar] [CrossRef] [PubMed]
Lawrence, R.E.; Zoncu, R. The lysosome as a cellular centre for signalling, metabolism and quality control. Nat. Cell Biol. 2019, 21, 133–142. [Google Scholar] [CrossRef] [PubMed]
Galione, A. A primer of NAADP-mediated Ca²⁺ signalling: From sea urchin eggs to mammalian cells. Cell Calcium 2015, 58, 27–47. [Google Scholar] [CrossRef] [PubMed]
Piao, S.; Amaravadi, R.K. Targeting the lysosome in cancer. Ann. N. Y. Acad. Sci. 2016, 1371, 45–54. [Google Scholar] [CrossRef] [Green Version]
Yang, S.; Wang, X.; Contino, G.; Liesa, M.; Sahin, E.; Ying, H.; Bause, A.; Li, Y.; Stommel, J.M.; Dell’Antonio, G.; et al. Pancreatic cancers require autophagy for tumor growth. Genes Dev. 2011, 25, 717–729. [Google Scholar] [CrossRef] [Green Version]
Hämälistö, S.; Jäättelä, M. Lysosomes in cancer—living on the edge (of the cell). Curr. Opin. Cell Biol. 2016, 39, 69–76. [Google Scholar] [CrossRef] [PubMed]
Yamaguchi, S.; Jha, A.; Li, Q.; Soyombo, A.; Dickinson, G.; Churamani, D.; Brailoiu, E.; Patel, S.; Muallem, S. Transient Receptor Potential Mucolipin 1 (TRPML1) and Two-pore Channels Are Functionally Independent Organellar Ion Channels. J. Biol. Chem. 2011, 286, 22934–22942. [Google Scholar] [CrossRef] [Green Version]
Favia, A.; Pafumi, I.; Desideri, M.; Padula, F.; Montesano, C.; Passeri, D.; Nicoletti, C.; Orlandi, A.; Del Bufalo, D.; Sergi, M.; et al. NAADP-Dependent Ca²⁺ Signaling Controls Melanoma Progression, Metastatic Dissemination and Neoangiogenesis. Sci. Rep. 2016, 6, 18925. [Google Scholar] [CrossRef] [Green Version]
Nguyen, O.N.P.; Grimm, C.; Schneider, L.S.; Chao, Y.-K.; Atzberger, C.; Bartel, K.; Watermann, A.; Ulrich, M.; Mayr, D.; Wahl-Schott, C.; et al. Two-Pore Channel Function Is Crucial for the Migration of Invasive Cancer Cells. Cancer Res. 2017, 77, 1427–1438. [Google Scholar] [CrossRef] [Green Version]
Kern, U.; Wischnewski, V.; Biniossek, M.L.; Schilling, O.; Reinheckel, T. Lysosomal protein turnover contributes to the acquisition of TGFβ-1 induced invasive properties of mammary cancer cells. Mol. Cancer 2015, 14, 39. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Zhang, X.; Cheng, X.; Yu, L.; Yang, J.; Calvo, R.; Patnaik, S.; Hu, X.; Gao, Q.; Yang, M.; Lawas, M.; et al. MCOLN1 is a ROS sensor in lysosomes that regulates autophagy. Nat. Commun. 2016, 7, 12109. [Google Scholar] [CrossRef] [Green Version]
Settembre, C.; Di Malta, C.; Polito, V.A.; Arencibia, M.G.; Vetrini, F.; Serkan, E.; Erdin, S.U.; Huynh, T.; Medina, D.; Colella, P.; et al. TFEB Links Autophagy to Lysosomal Biogenesis. Science 2011, 332, 1429–1433. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Durinck, S.; Stawiski, E.W.; Pavía-Jiménez, A.; Modrusan, Z.; Kapur, P.; Jaiswal, B.S.; Zhang, N.; Toffessi-Tcheuyap, V.; Nguyen, T.; Pahuja, K.B.; et al. Spectrum of diverse genomic alterations define non–clear cell renal carcinoma subtypes. Nat. Genet. 2015, 47, 13–21. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Calcagni, A.; Kors, L.; Verschuren, E.; De Cegli, R.; Zampelli, N.; Nusco, E.; Confalonieri, S.; Bertalot, G.; Pece, S.; Settembre, C.; et al. Modelling TFE renal cell carcinoma in mice re-veals a critical role of WNT signaling. eLife 2016, 5, e17047. [Google Scholar] [CrossRef] [Green Version]
Perera, R.M.; Stoykova, S.; Nicolay, B.N.; Ross, K.N.; Fitamant, J.; Boukhali, M.; Lengrand, J.; Deshpande, V.; Selig, M.K.; Ferrone, C.R.; et al. Transcriptional control of autophagy–lysosome function drives pancreatic cancer metabolism. Nat. Cell Biol. 2015, 524, 361–365. [Google Scholar] [CrossRef]
Medina, D.L.; Di Paola, S.; Peluso, I.; Armani, A.; De Stefani, D.; Venditti, R.; Montefusco, S.; Rosato, A.S.; Prezioso, C.; Forrester, A.; et al. Lysosomal calcium signalling regulates autophagy through calcineurin and TFEB. Nat. Cell Biol. 2015, 17, 288–299. [Google Scholar] [CrossRef] [Green Version]
Giatromanolaki, A.; Kalamida, D.; Sivridis, E.; Karagounis, I.V.; Gatter, K.C.; Harris, A.L.; Koukourakis, M.I. Increased expression of transcription factor EB (TFEB) is associated with autophagy, migratory phenotype and poor prognosis in non-small cell lung cancer. Lung Cancer 2015, 90, 98–105. [Google Scholar] [CrossRef]
Liang, J.; Jia, X.; Wang, K.; Zhao, N. High expression of TFEB is associated with aggressive clinical features in colorectal cancer. OncoTargets Ther. 2018, 11, 8089–8098. [Google Scholar] [CrossRef] [Green Version]
Slade, L.; Biswas, D.; Ihionu, F.; El Hiani, Y.; Kienesberger, P.C.; Pulinilkunnil, T. A Lysosome Independent Role for TFEB in Activating DNA Repair and Inhibiting Apoptosis in Breast Cancer Cells. Biochem. J. 2020, 477, 137–160. [Google Scholar] [CrossRef] [PubMed]
Cui, C.; Chang, Y.; Zhang, X.; Choi, S.; Tran, H.; Penmetsa, K.V.; Viswanadha, S.; Fu, L.; Pan, Z. Targeting Orai1-mediated store-operated calcium entry by RP4010 for anti-tumor activity in esophagus squamous cell carcinoma. Cancer Lett. 2018, 432, 169–179. [Google Scholar] [CrossRef]
Serafini, M.; Cordero-Sanchez, C.; Di Paola, R.; Bhela, I.P.; Aprile, S.; Purghè, B.; Fusco, R.; Cuzzocrea, S.; Genazzani, A.A.; Riva, B.; et al. Store-Operated Calcium Entry as a Therapeutic Target in Acute Pancreatitis: Discovery and Development of Drug-Like SOCE Inhibitors. J. Med. Chem. 2020, 63, 14761–14779. [Google Scholar] [CrossRef] [PubMed]
Waldherr, L.; Tiffner, A.; Mishra, D.; Sallinger, M.; Schober, R.; Frischauf, I.; Schmidt, T.; Handl, V.; Sagmeister, P.; Köckinger, M.; et al. Blockage of Store-Operated Ca²⁺ Influx by Synta66 is Mediated by Direct Inhibition of the Ca²⁺ Selective Orai1 Pore. Cancers 2020, 12, 2876. [Google Scholar] [CrossRef] [PubMed]
Marchesini, M.; Gherli, A.; Montanaro, A.; Patrizi, L.; Sorrentino, C.; Pagliaro, L.; Rompietti, C.; Kitara, S.; Heit, S.; Olesen, C.E.; et al. Blockade of Oncogenic NOTCH1 with the SERCA Inhibitor CAD204520 in T Cell Acute Lymphoblastic Leukemia. Cell Chem. Biol. 2020, 27, 678–697.e13. [Google Scholar] [CrossRef]
Pagliaro, L.; Marchesini, M.; Roti, G. Targeting oncogenic Notch signaling with SERCA inhibitors. J. Hematol. Oncol. 2021, 14, 1–17. [Google Scholar] [CrossRef]
Patergnani, S.; Danese, A.; Bouhamida, E.; Aguiari, G.; Previati, M.; Pinton, P.; Giorgi, C. Various Aspects of Calcium Signaling in the Regulation of Apoptosis, Autophagy, Cell Proliferation, and Cancer. Int. J. Mol. Sci. 2020, 21, 8323. [Google Scholar] [CrossRef]
Prasad, V.; Mailankody, S. Research and development spending to bring a single cancer drug to market and revenues after approval. JAMA Internal Med. 2017, 177, 1569–1575. [Google Scholar] [CrossRef]
Senapati, S.; Mahanta, A.K.; Kumar, S.; Maiti, P. Controlled drug delivery vehicles for cancer treatment and their performance. Signal Transduct. Target. Ther. 2018, 3, 7. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Lee, D.; Hong, J.H. Physiological application of nanoparticles in calcium-related proteins and channels. Nanomedicine 2019, 14, 2479–2486. [Google Scholar] [CrossRef] [PubMed]
Colone, M.; Calcabrini, A.; Stringaro, A. Drug Delivery Systems of Natural Products in Oncology. Molecules 2020, 25, 4560. [Google Scholar] [CrossRef]
Bong, A.H.; Monteith, G.R. Calcium signaling and the therapeutic targeting of cancer cells. Biochim. Biophys. Acta (BBA)—Bioenerg. 2018, 1865, 1786–1794. [Google Scholar] [CrossRef]
Shen, Y.; Li, X.; Dong, D.; Zhang, B.; Xue, Y.; Shang, P. Transferrin receptor 1 in cancer: A new sight for cancer therapy. Am. J. Cancer Res. 2018, 8, 916–931. [Google Scholar] [PubMed]
Choudhary, D.; Goykar, H.; Karanwad, T.; Kannaujia, S.; Gadekar, V.; Misra, M. An understanding of mitochondria and its role in targeting nanocarriers for diagnosis and treatment of cancer. Asian J. Pharm. Sci. 2020. [Google Scholar] [CrossRef]
Rahman, S.; Rahman, T. Unveiling some FDA-approved drugs as inhibitors of the store-operated Ca²⁺ entry pathway. Sci. Rep. 2017, 7, 12881. [Google Scholar] [CrossRef] [Green Version]
Frandsen, S.K.; Vissing, M.; Gehl, J. A Comprehensive Review of Calcium Electroporation—A Novel Cancer Treatment Modality. Cancers 2020, 12, 290. [Google Scholar] [CrossRef] [Green Version]
Munk, M.; Alcalde, J.; Lorentzen, L.; Villalobo, A.; Berchtold, M.W.; Panina, S. The impact of calmodulin on the cell cycle analyzed in a novel human cellular genetic system. Cell Calcium 2020, 88, 102207. [Google Scholar] [CrossRef]
Chai, S.; Xu, X.; Wang, Y.; Zhou, Y.; Zhang, C.; Yang, Y.; Xu, H.; Xu, R.; Wang, K.; Yang, Y. Ca²⁺/calmodulin-dependent protein kinase IIγ enhances stem-like traits and tumorigenicity of lung cancer cells. Oncotarget 2015, 6, 16069–16083. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Chi, M.; Evans, H.; Gilchrist, J.; Mayhew, J.; Hoffman, A.; Pearsall, E.A.; Jankowski, H.; Brzozowski, J.; Skelding, K.A. Phosphorylation of calcium/calmodulin-stimulated protein kinase II at T286 enhances invasion and migration of human breast cancer cells. Sci. Rep. 2016, 6, 33132. [Google Scholar] [CrossRef]
Daft, P.G.; Yang, Y.; Napierala, D.; Zayzafoon, M. The Growth and Aggressive Behavior of Human Osteosarcoma Is Regulated by a CaMKII-Controlled Autocrine VEGF Signaling Mechanism. PLoS ONE 2015, 10, e0121568. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Jung, H.J.; Kim, J.H.; Shim, J.S.; Kwon, H.J. A Novel Ca²⁺/Calmodulin Antagonist HBC Inhibits Angiogenesis and Down-regulates Hypoxia-inducible Factor. J. Biol. Chem. 2010, 285, 25867–25874. [Google Scholar] [CrossRef] [Green Version]
Can, G.; Akpinar, B.; Baran, Y.; Zhivotovsky, B.; Olsson, M. 5-Fluorouracil signaling through a calcium–calmodulin-dependent pathway is required for p53 activation and apoptosis in colon carcinoma cells. Oncogene 2013, 32, 4529–4538. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Ren, T.; Tang, Y.; Wang, M.; Wang, H.; Liu, Y.; Qian, X.; Chang, C.; Chen, M. Triptolide induces apoptosis through the calcium/calmodulin-dependent protein kinase kinaseβ/AMP-activated protein kinase signaling pathway in non-small cell lung cancer cells. Oncol. Rep. 2020, 44, 2288–2296. [Google Scholar] [CrossRef] [PubMed]
Karacosta, L.G.; Foster, B.A.; Azabdaftari, G.; Feliciano, D.; Edelman, A.M. A Regulatory Feedback Loop Between Ca²⁺/Calmodulin-dependent Protein Kinase Kinase 2 (CaMKK2) and the Androgen Receptor in Prostate Cancer Progression. J. Biol. Chem. 2012, 287, 24832–24843. [Google Scholar] [CrossRef] [Green Version]
Chimote, A.A.; Gawali, V.; Newton, H.S.; Wise-Draper, T.M.; Conforti, L. A Compartmentalized Reduction in Membrane-Proximal Calmodulin Reduces the Immune Surveillance Capabilities of CD8+ T Cells in Head and Neck Cancer. Front. Pharmacol. 2020, 11, 143. [Google Scholar] [CrossRef] [Green Version]
Racioppi, L.; Nelson, E.R.; Huang, W.; Mukherjee, D.; Lawrence, S.A.; Lento, W.; Masci, A.M.; Jiao, Y.; Park, S.; York, B.; et al. CaMKK2 in myeloid cells is a key regulator of the immune-suppressive microenvironment in breast cancer. Nat. Commun. 2019, 10, 1–16. [Google Scholar] [CrossRef] [Green Version]
Blum, W.; Schwaller, B. Calretinin is essential for mesothelioma cell growth/survival in vitro: A potential new target for malignant mesothelioma therapy? Int. J. Cancer 2013, 133, 2077–2088. [Google Scholar] [CrossRef] [Green Version]
Winn, B.; Tavares, R.; Fanion, J.; Noble, L.; Gao, J.; Sabo, E.; Resnick, M.B. Differentiating the undifferentiated: Immunohistochemical profile of medullary carcinoma of the colon with an emphasis on intestinal differentiation. Hum. Pathol. 2009, 40, 398–404. [Google Scholar] [CrossRef] [Green Version]
Wörthmüller, J.; Oberson, A.; Salicio, V.; Blum, W.; Schwaller, B. Calretinin Functions in Malignant Mesothelioma Cells Cannot Be Replaced by the Closely Related Ca²⁺-Binding Proteins Calbindin-D28k and Parvalbumin. Int. J. Mol. Sci. 2018, 19, 4015. [Google Scholar] [CrossRef] [Green Version]
Dodla, P.; Bhoopalan, V.; Khoo, S.K.; Miranti, C.; Sridhar, S. Gene expression analysis of human prostate cell lines with and without tumor metastasis suppressor CD82. BMC Cancer 2020, 20, 1–17. [Google Scholar] [CrossRef] [PubMed]
Bignotti, E.; Tassi, R.A.; Calza, S.; Ravaggi, A.; Bandiera, E.; Rossi, E.; Donzelli, C.; Pasinetti, B.; Pecorelli, S.; Santin, A.D. Gene expression profile of ovarian serous papillary carcinomas: Identification of metastasis-associated genes. Am. J. Obstet. Gynecol. 2007, 196, 245.e1–245.e11. [Google Scholar] [CrossRef] [PubMed]
Wörthmüller, J.; Blum, W.; Pecze, L.; Salicio, V.; Schwaller, B. Calretinin promotes invasiveness and EMT in malignant mesothelioma cells involving the activation of the FAK signaling pathway. Oncotarget 2018, 9, 36256–36272. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Gordillo, C.H.; Sandoval, P.; Muñoz-Hernández, P.; Pascual-Antón, L.; López-Cabrera, M.; Jiménez-Heffernan, J.A. Mesothelial-to-Mesenchymal Transition Contributes to the Generation of Carcinoma-Associated Fibroblasts in Locally Advanced Primary Colorectal Carcinomas. Cancers 2020, 12, 499. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Jung, E.M.; Choi, K.C.; Jeung, E.B. Expression of calbindin-D28k is inversely correlated with proapototic gene expression in hydrogen peroxide-induced cell death in endometrial cancer cells. Int. J. Oncol. 2011, 38, 1059–1066. [Google Scholar] [CrossRef] [PubMed]
Stevenson, L.; Allen, W.L.; Proutski, I.; Stewart, G.; Johnston, L.; McCloskey, K.; Wilson, P.M.; Longley, D.B.; Johnston, P.G. Calbindin 2 (CALB2) Regulates 5-Fluorouracil Sensitivity in Colorectal Cancer by Modulating the Intrinsic Apoptotic Pathway. PLoS ONE 2011, 6, e20276. [Google Scholar] [CrossRef] [Green Version]
Sergeev, I.N. Vitamin D and cellular Ca²⁺ signaling in breast cancer. Anticancer. Res. 2012, 32, 299–302. [Google Scholar]
Kim, J.H.; sil Hong, B.; Heo, W.; Han, J.M.; Han, W.; Noh, D.-Y.; Moon, H.-G. Abstract 33: Calsequestrin 2 regulates prolif-eration, migration, and invasion in triple-negative breast cancer cells. Cancer Res. 2018, 78 (Suppl. 13), 33. [Google Scholar] [CrossRef]
Toquet, C.; Jarry, A.; Bou-Hanna, C.; Bach, K.; Denis, M.G.; Mosnier, J.F.; Laboisse, C.L. Altered Calreticulin expression in human colon cancer: Maintenance of Calreticulin expression is associated with mucinous differentiation. Oncol. Rep. 2007, 17, 1101–1107. [Google Scholar] [CrossRef]
Vanoverberghe, K.; Abeele, F.V.; Mariot, P.; Lepage, G.; Roudbaraki, M.; Bonnal, J.L.; Mauroy, B.; Shuba, Y.; Skryma, R.; Prevarskaya, N. Ca²⁺ homeostasis and apoptotic resistance of neuroendocrine-differentiated prostate cancer cells. Cell Death Differ. 2003, 11, 321–330. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Han, Y.; Liao, Q.; Wang, H.; Rao, S.; Yi, P.; Tang, L.; Tian, Y.; Oyang, L.; Wang, H.; Shi, Y.; et al. High expression of calreticulin indicates poor prognosis and modulates cell migration and invasion via activating Stat3 in nasopharyngeal carcinoma. J. Cancer 2019, 10, 5460–5468. [Google Scholar] [CrossRef] [PubMed]
Pouliquen, D.L.; Boissard, A.; Coqueret, O.; Guette, C. Biomarkers of tumor invasiveness in proteomics (Review). Int. J. Oncol. 2020, 57, 409–432. [Google Scholar] [CrossRef]
Amuthan, G.; Biswas, G.; Zhang, S.; Klein-Szanto, A.; Vijayasarathy, C.; Avadhani, N.G. Mitochondria-to-nucleus stress signaling induces phenotypic changes, tumor progression and cell invasion. EMBO J. 2001, 20, 1910–1920. [Google Scholar] [CrossRef] [Green Version]
Pike, S.E.; Yao, L.; Setsuda, J.; Jones, K.D.; Cherney, B.; Appella, E.; Sakaguchi, K.; Nakhasi, H.; Atreya, C.D.; Teruya-Feldstein, J.; et al. Calreticulin and calreticulin fragments are endothelial cell inhibitors that suppress tumor growth. Blood 1999, 94, 2461–2468. [Google Scholar] [CrossRef] [PubMed]
Delom, F.; Emadali, A.; Cocolakis, E.; Lebrun, J.-J.; Nantel, A.; Chevet, E. Calnexin-dependent regulation of tunicamycin-induced apoptosis in breast carcinoma MCF-7 cells. Cell Death Differ. 2006, 14, 586–596. [Google Scholar] [CrossRef] [PubMed]
Li, D.-D.; Xie, B.; Wu, X.-J.; Li, J.-J.; Ding, Y.; Wen, X.-Z.; Zhang, X.; Zhu, S.-G.; Liu, W.; Zhang, X.-S.; et al. Late-stage inhibition of autophagy enhances calreticulin surface exposure. Oncotarget 2016, 7, 80842–80854. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Obeid, M.; Tesniere, A.; Ghiringhelli, F.; Fimia, G.M.; Apetoh, L.; Perfettini, J.-L.; Castedo, M.; Mignot, G.; Panaretakis, T.; Casares, N.; et al. Calreticulin exposure dictates the immunogenicity of cancer cell death. Nat. Med. 2007, 13, 54–61. [Google Scholar] [CrossRef]
Chen, Y.; Ma, D.; Wang, X.; Fang, J.; Liu, X.; Song, J.; Li, X.; Ren, X.; Li, Q.; Li, Q.; et al. Calnexin Impairs the Antitumor Immunity of CD4+ and CD8+ T Cells. Cancer Immunol. Res. 2018, 7, 123–135. [Google Scholar] [CrossRef] [Green Version]
Loulousis, M.; Krager, S.L.; Darcy, Y.L.; Tischkau, S.A.; Copello, J.A. Drugs that inhibit the sarcoplasmic reticulum Ca²⁺ atpase (Serca) and prevention of breast cancer cell proliferation. FASEB J. 2016, 30, 768.4. [Google Scholar] [CrossRef]
Lee, W.J.; Robinson, J.A.; Holman, N.A.; McCall, M.N.; Roberts-Thomson, S.; Monteith, G. Antisense-mediated Inhibition of the Plasma Membrane Calcium-ATPase Suppresses Proliferation of MCF-7 Cells. J. Biol. Chem. 2005, 280, 27076–27084. [Google Scholar] [CrossRef] [Green Version]
Chovancova, B.; Liskova, V.; Babula, P.; Krizanova, O. Role of Sodium/Calcium Exchangers in Tumors. Biomolecules 2020, 10, 1257. [Google Scholar] [CrossRef]
Sritangos, P.; Alarcon, E.P.; James, A.; Sultan, A.; Richardson, D.A.; Bruce, J.I.E. Plasma Membrane Ca²⁺ ATPase Isoform 4 (PMCA4) Has an Important Role in Numerous Hallmarks of Pancreatic Cancer. Cancers 2020, 12, 218. [Google Scholar] [CrossRef] [Green Version]
Fondello, C.; Agnetti, L.; Glikin, G.C.; Finocchiaro, L.M. Mechanisms Enhancing the Cytotoxic Effects of Bleomycin plus Suicide or Interferon-β Gene Lipofection in Metastatic Human Melanoma Cells. Anti-Cancer Agents Med. Chem. 2019, 18, 1338–1348. [Google Scholar] [CrossRef]
Szadvari, I.; Hudecova, S.; Chovancova, B.; Matuskova, M.; Cholujova, D.; Lencesova, L.; Valerian, D.; Ondrias, K.; Babula, P.; Krizanova, O. Sodium/calcium exchanger is involved in apoptosis induced by H2S in tumor cells through decreased levels of intracellular pH. Nitric Oxide 2019, 87, 1–9. [Google Scholar] [CrossRef]
Ho, P.-C.; Bihuniak, J.D.; Macintyre, A.; Staron, M.; Liu, X.; Amezquita, R.; Tsui, Y.-C.; Cui, G.; Micevic, G.; Perales, J.C.; et al. Phosphoenolpyruvate Is a Metabolic Checkpoint of Anti-tumor T Cell Responses. Cell 2015, 162, 1217–1228. [Google Scholar] [CrossRef] [PubMed] [Green Version]
James, A.; Richardson, D.A.; Oh, I.-W.; Sritangos, P.; Attard, T.; Barrett, L.; Bruce, J.I.E. Cutting off the fuel supply to calcium pumps in pancreatic cancer cells: Role of pyruvate kinase-M2 (PKM2). Br. J. Cancer 2020, 122, 266–278. [Google Scholar] [CrossRef] [PubMed]
Márián, T.; Szabó-Péli, J.; Németh, E.; Trón, L.; Friedländer, E.; Szabó, A.; Balkay, L.; Veress, G.; Krasznai, Z. Na⁺/Ca²⁺ exchanger inhibitors modify the accumulation of tumor-diagnostic PET tracers in cancer cells. Eur. J. Pharm. Sci. 2007, 30, 56–63. [Google Scholar] [CrossRef] [PubMed]
Ghosh, S.; Adhikary, A.; Chakraborty, S.; Nandi, P.; Mohanty, S.; Chakraborty, S.; Bhattacharjee, P.; Mukherjee, S.; Putatunda, S.; Chakraborty, S.; et al. Nifetepimine, a Dihydropyrimidone, Ensures CD4+ T Cell Survival in a Tumor Microenvironment by Maneuvering Sarco(endo)plasmic Reticulum Ca²⁺ ATPase (SERCA). J. Biol. Chem. 2012, 287, 32881–32896. [Google Scholar] [CrossRef] [Green Version]
Bhargava, A.; Saha, S. T-Type voltage gated calcium channels: A target in breast cancer? Breast Cancer Res. Treat. 2019, 173, 11–21. [Google Scholar] [CrossRef] [PubMed]
Xie, R.; Xu, J.; Wen, G.; Jin, H.; Liu, X.; Yang, Y.; Ji, B.; Jiang, Y.; Song, P.; Dong, H.; et al. The P2Y2 Nucleotide Receptor Mediates the Proliferation and Migration of Human Hepatocellular Carcinoma Cells Induced by ATP. J. Biol. Chem. 2014, 289, 19137–19149. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Adinolfi, E.; Melchiorri, L.; Falzoni, S.; Chiozzi, P.; Morelli, A.; Tieghi, A.; Cuneo, A.; Castoldi, G.; Di Virgilio, F.; Baricordi, O.R. P2X7 receptor expression in evolutive and indolent forms of chronic B lymphocytic leukemia. Blood 2002, 99, 706–708. [Google Scholar] [CrossRef]
Ledur, P.F.; Villodre, E.S.; Paulus, R.; Cruz, L.A.; Flores, D.G.; Lenz, G. Extracellular ATP reduces tumor sphere growth and cancer stem cell population in glioblastoma cells. Purinergic Signal. 2011, 8, 39–48. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Conigrave, A.D.; van der Weyden, L.; Holt, L.; Jiang, L.; Wilson, P.; I Christopherson, R.; Morris, M.B. Extracellular ATP-dependent suppression of proliferation and induction of differentiation of human HL-60 leukemia cells by distinct mechanisms. Biochem. Pharmacol. 2000, 60, 1585–1591. [Google Scholar] [CrossRef]
Maiques, O.; Macià, A.; Moreno, S.; Barceló, C.; Santacana, M.; Vea, A.; Herreros, J.; Gatius, S.; Ortega, E.; Valls, J.; et al. Immunohistochemical analysis of T-type calcium channels in acquired melanocytic naevi and melanoma. Br. J. Dermatol. 2016, 176, 1247–1258. [Google Scholar] [CrossRef] [Green Version]
Zhou, X.; Wang, W.; Zhang, S.; Wang, X.; Tang, Z.; Gu, J.; Li, J.; Huang, J. CACNA1B (Cav2.2) Overexpression and Its Association with Clinicopathologic Characteristics and Unfavorable Prognosis in Non-Small Cell Lung Cancer. Dis. Markers 2017, 2017, 1–8. [Google Scholar] [CrossRef] [PubMed]
Schumacher, D.; Strilic, B.; Sivaraj, K.K.; Wettschureck, N.; Offermanns, S. Platelet-Derived Nucleotides Promote Tumor-Cell Transendothelial Migration and Metastasis via P2Y2 Receptor. Cancer Cell 2013, 24, 130–137. [Google Scholar] [CrossRef] [Green Version]
Takai, E.; Tsukimoto, M.; Harada, H.; Kojima, S. Autocrine signaling via release of ATP and activation of P2X7 receptor influences motile activity of human lung cancer cells. Purinergic Signal. 2014, 10, 487–497. [Google Scholar] [CrossRef] [Green Version]
Rumjahn, S.M.; A Javed, M.; Wong, N.; E Law, W.; O Buxton, I.L. Purinergic regulation of angiogenesis by human breast carcinoma-secreted nucleoside diphosphate kinase. Br. J. Cancer 2007, 97, 1372–1380. [Google Scholar] [CrossRef]
Dziegielewska, B.; Gray, L.S.; Dziegielewski, J. T-type calcium channels blockers as new tools in cancer therapies. Pflügers Archiv—Eur. J. Physiol. 2014, 466, 801–810. [Google Scholar] [CrossRef]
Gilbert, S.M.; Oliphant, C.J.; Hassan, S.; Peille, A.L.; Bronsert, P.; Falzoni, S.; Di Virgilio, F.; McNulty, S.; Lara, R. ATP in the tumour microenvironment drives expression of nfP2X7, a key mediator of cancer cell survival. Oncogene 2019, 38, 194–208. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Höpfner, M.; Maaser, K.; Barthel, B.; Von Lampe, B.; Hanski, C.; Riecken, E.-O.; Zeitz, M.; Scherübl, H. Growth inhibition and apoptosis induced by P2Y2 receptors in human colorectal carcinoma cells: Involvement of intracellular calcium and cyclic adenosine monophosphate. Int. J. Color. Dis. 2001, 16, 154–166. [Google Scholar] [CrossRef]
Sallán, M.C.; Visa, A.; Shaikh, S.; Nàger, M.; Herreros, J.; Cantí, C. T-type Ca²⁺ Channels: T for Targetable. Cancer Res. 2018, 78, 603–609. [Google Scholar] [CrossRef] [Green Version]
Amoroso, F.S.; Capece, M.; Rotondo, A.; Cangelosi, D.; Ferracin, M.; Franceschini, A.; Raffaghello, L.; Pistoia, V.; Varesio, L.; Adinolfi, E. The P2X7 receptor is a key modulator of the PI3K/GSK3β/VEGF signaling network: Evidence in experimental neuroblastoma. Oncogene 2015, 34, 5240–5251. [Google Scholar] [CrossRef]
Pfaffenzeller, M.S.; Franciosi, M.L.M.; Cardoso, A.M. Purinergic signaling and tumor microenvironment in cervical Cancer. Purinergic Signal. 2020, 16, 123–135. [Google Scholar] [CrossRef]
Wu, L.; Lin, W.; Liao, Q.; Wang, H.; Lin, C.; Tang, L.; Lian, W.; Chen, Z.; Li, K.; Xu, L.; et al. Calcium Channel Blocker Nifedipine Suppresses Colorectal Cancer Progression and Immune Escape by Preventing NFAT2 Nuclear Translocation. Cell Rep. 2020, 33, 108327. [Google Scholar] [CrossRef] [PubMed]
Karacicek, B.; Erac, Y.; Tosun, M. Functional consequences of enhanced expression of STIM1 and Orai1 in Huh-7 hepatocellular carcinoma tumor-initiating cells. BMC Cancer 2019, 19, 1–10. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Faouzi, M.; Kischel, P.; Hague, F.; Ahidouch, A.; Benzerdjeb, N.; Sevestre, H.; Penner, R.; Ouadid-Ahidouch, H. ORAI3 silencing alters cell proliferation and cell cycle progression via c-myc pathway in breast cancer cells. Biochim. Biophys. Acta 2013, 1833, 752–760. [Google Scholar] [CrossRef] [PubMed] [Green Version]
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.; Chang, W.-C. Orai1/CRACM1 overexpression suppresses cell proliferation via attenuation of the store-operated calcium influx-mediated signalling pathway in A549 lung cancer cells. Biochim. Biophys. Acta (BBA)—Gen. Subj. 2011, 1810, 1278–1284. [Google Scholar] [CrossRef]
Diez-Bello, R.; Jardin, I.; Salido, G.M.; Rosado, J. Orai1 and Orai2 mediate store-operated calcium entry that regulates HL60 cell migration and FAK phosphorylation. Biochim. Biophys. Acta (BBA)—Bioenerg. 2017, 1864, 1064–1070. [Google Scholar] [CrossRef]
Zhang, Z.; Liu, X.; Feng, B.; Liu, N.; Wu, Q.; Han, Y.; Nie, Y.; Wu, K.; Shi, Y.; Fan, D. STIM1, a direct target of microRNA-185, promotes tumor metastasis and is associated with poor prognosis in colorectal cancer. Oncogene 2015, 34, 4808–4820. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Lee, S.H.; Rigas, N.K.; Lee, C.-R.; Bang, A.; Srikanth, S.; Gwack, Y.; Kang, M.K.; Kim, R.H.; Park, N.-H.; Shin, K.-H. Orai1 promotes tumor progression by enhancing cancer stemness via NFAT signaling in oral/oropharyngeal squamous cell carcinoma. Oncotarget 2016, 7, 43239–43255. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Wu, S.; Chen, M.; Huang, J.; Zhang, F.; Lv, Z.; Jia, Y.; Cui, Y.-Z.; Sun, L.-Z.; Wang, Y.; Tang, Y.; et al. ORAI2 Promotes Gastric Cancer Tumorigenicity and Metastasis through PI3K/Akt Signaling and MAPK-Dependent Focal Adhesion Disassembly. Cancer Res. 2021, 81, 986–1000. [Google Scholar] [CrossRef] [PubMed]
Miao, Y.; Shen, Q.; Zhang, S.; Huang, H.; Meng, X.; Zheng, X.; Yao, Z.; He, Z.; Lu, S.; Cai, C.; et al. Calcium-sensing stromal interaction molecule 2 upregulates nuclear factor of activated T cells 1 and transforming growth factor-β signaling to promote breast cancer metastasis. Breast Cancer Res. 2019, 21, 1–12. [Google Scholar] [CrossRef]
Sun, J.; Lu, F.; He, H.; Shen, J.; Messina, J.; Mathew, R.; Wang, D.; Sarnaik, A.A.; Chang, W.-C.; Kim, M.; et al. STIM1- and Orai1-mediated Ca²⁺ oscillation orchestrates invadopodium formation and melanoma invasion. J. Cell Biol. 2014, 207, 535–548. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Pan, S.; Zhao, X.; Shao, C.; Fu, B.; Huang, Y.; Zhang, N.; Dou, X.; Zhang, Z.; Qiu, Y.; Wang, R.; et al. STIM1 promotes angiogenesis by reducing exosomal miR-145 in breast cancer MDA-MB-231 cells. Cell Death Dis. 2021, 12, 1–15. [Google Scholar] [CrossRef]
Liu, X.; Wan, X.; Kan, H.; Wang, Y.; Yu, F.; Feng, L.; Jin, J.; Zhang, P.; Ma, X. Hypoxia-induced upregulation of Orai1 drives colon cancer invasiveness and angiogenesis. Eur. J. Pharmacol. 2018, 832, 1–10. [Google Scholar] [CrossRef]
Sun, X.; Wei, Q.; Cheng, J.; Bian, Y.; Tian, C.; Hu, Y.; Li, H. Enhanced Stim1 expression is associated with acquired chemo-resistance of cisplatin in osteosarcoma cells. Hum. Cell 2017, 30, 216–225. [Google Scholar] [CrossRef] [Green Version]
Faouzi, M.; Hague, F.; Potier, M.; Ahidouch, A.; Sevestre, H.; Ouadid-Ahidouch, H. Down-regulation of Orai3 arrests cell-cycle progression and induces apoptosis in breast cancer cells but not in normal breast epithelial cells. J. Cell. Physiol. 2011, 226, 542–551. [Google Scholar] [CrossRef]
Liu, H.; Hughes, J.D.; Rollins, S.; Chen, B.; Perkins, E. Calcium entry via ORAI1 regulates glioblastoma cell proliferation and apoptosis. Exp. Mol. Pathol. 2011, 91, 753–760. [Google Scholar] [CrossRef]
Zhao, H.; Yan, G.; Zheng, L.; Zhou, Y.; Sheng, H.; Wu, L.; Zhang, Q.; Lei, J.; Zhang, J.; Xin, R.; et al. STIM1 is a metabolic checkpoint regulating the invasion and metastasis of hepatocellular carcinoma. Theranostics 2020, 10, 6483–6499. [Google Scholar] [CrossRef] [PubMed]
Frisch, J.; Angenendt, A.; Hoth, M.; Prates Roma, L.; Lis, A. STIM-Orai Channels and Reactive Oxygen Species in the Tumor Microenvironment. Cancers 2019, 11, 457. [Google Scholar] [CrossRef] [Green Version]
Tang, B.-D.; Xia, X.; Lv, X.-F.; Yu, B.-X.; Yuan, J.-N.; Mai, X.-Y.; Shang, J.-Y.; Zhou, J.-G.; Liang, S.-J.; Pang, R.-P. Inhibition of Orai1-mediated Ca²⁺entry enhances chemosensitivity of HepG2 hepatocarcinoma cells to 5-fluorouracil. J. Cell. Mol. Med. 2016, 21, 904–915. [Google Scholar] [CrossRef] [PubMed]
Fleur-Lominy, S.S.; Maus, M.; Vaeth, M.; Lange, I.; Zee, I.; Suh, D.; Liu, C.; Wu, X.; Tikhonova, A.; Aifantis, I.; et al. STIM1 and STIM2 Mediate Cancer-Induced Inflammation in T Cell Acute Lymphoblastic Leukemia. Cell Rep. 2018, 24, 3045–3060.e5. [Google Scholar] [CrossRef] [Green Version]
Xu, Y.; Zhang, S.; Niu, H.; Ye, Y.; Hu, F.; Chen, S.; Li, X.; Luo, X.; Jiang, S.; Liu, Y.; et al. STIM1 accelerates cell senescence in a remodeled microenvironment but enhances the epithelial-to-mesenchymal transition in prostate cancer. Sci. Rep. 2015, 5, 11754. [Google Scholar] [CrossRef] [Green Version]
Szatkowski, C.; Parys, J.B.; Ouadid-Ahidouch, H.; Matifat, F. Inositol 1,4,5-trisphosphate-induced Ca²⁺ signalling is involved in estradiol-induced breast cancer epithelial cell growth. Mol. Cancer 2010, 9, 156. [Google Scholar] [CrossRef] [Green Version]
Nougarède, A.; Popgeorgiev, N.; Kassem, L.; Omarjee, S.; Borel, S.; Mikaelian, I.; Lopez, J.; Gadet, R.; Marcillat, O.; Treilleux, I.; et al. Breast Cancer Targeting through Inhibition of the Endoplasmic Reticulum-Based Apoptosis Regulator Nrh/BCL2L10. Cancer Res. 2018, 78, 1404–1417. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Thoppil, R.J.; Adapala, R.K.; Cappelli, H.C.; Kondeti, V.; Dudley, A.C.; Gary Meszaros, J.; Paruchuri, S.; Thodeti, C.K. TRPV4 channel activation selectively inhibits tumor endothelial cell proliferation. Sci. Rep. 2015, 5, 14257. [Google Scholar] [CrossRef] [Green Version]
Bomben, V.C.; Sontheimer, H. Disruption of transient receptor potential canonical channel 1 causes incomplete cytokinesis and slows the growth of human malignant gliomas. Glia 2010, 58, 1145–1156. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Jiang, J.; Li, M.-H.; Inoue, K.; Chu, X.-P.; Seeds, J.; Xiong, Z.-G. Transient Receptor Potential Melastatin 7–like Current in Human Head and Neck Carcinoma Cells: Role in Cell Proliferation. Cancer Res. 2007, 67, 10929–10938. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Sun, C.; Shui, B.; Zhao, W.; Liu, H.; Li, W.; Lee, J.C.; Doran, R.; Lee, F.K.; Sun, T.; Shen, Q.S.; et al. Central role of IP3R2-mediated Ca²⁺ oscillation in self-renewal of liver cancer stem cells elucidated by high-signal ER sensor. Cell Death Dis. 2019, 10, 1–13. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Bidaux, G.; Flourakis, M.; Thébault, S.; Zholos, A.; Beck, B.; Gkika, D.; Roudbaraki, M.; Bonnal, J.-L.; Mauroy, B.; Shuba, Y.; et al. Prostate cell differentiation status determines transient receptor potential melastatin member 8 channel subcellular localization and function. J. Clin. Investig. 2007, 117, 1647–1657. [Google Scholar] [CrossRef] [Green Version]
Jiang, H.-N.; Zeng, B.; Zhang, Y.; Daskoulidou, N.; Fan, H.; Qu, J.-M.; Xu, S.-Z. Involvement of TRPC Channels in Lung Cancer Cell Differentiation and the Correlation Analysis in Human Non-Small Cell Lung Cancer. PLoS ONE 2013, 8, e67637. [Google Scholar] [CrossRef] [Green Version]
Middelbeek, J.; Visser, D.; Henneman, L.; Kamermans, A.; Kuipers, A.J.; Hoogerbrugge, P.M.; Jalink, K.; Van Leeuwen, F.N. TRPM7 maintains progenitor-like features of neuroblastoma cells: Implications for metastasis formation. Oncotarget 2015, 6, 8760–8776. [Google Scholar] [CrossRef]
Kang, S.S.; Han, K.-S.; Ku, B.M.; Lee, Y.K.; Hong, J.; Shin, H.Y.; Almonte, A.; Woo, D.H.; Brat, D.J.; Hwang, E.M.; et al. Caffeine-Mediated Inhibition of Calcium Release Channel Inositol 1,4,5-Trisphosphate Receptor Subtype 3 Blocks Glioblastoma Invasion and Extends Survival. Cancer Res. 2010, 70, 1173–1183. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Huang, X.; Jin, M.; Chen, Y.-X.; Wang, J.; Chang, Y.; Yuan, Q.; Yao, K.-T.; Ji, G. ERP44 inhibits human lung cancer cell migration mainly via IP3R2. Aging 2016, 8, 1276–1286. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Xu, N.; Zhang, D.; Chen, J.; He, G.; Gao, L. Low expression of ryanodine receptor 2 is associated with poor prognosis in thyroid carcinoma. Oncol. Lett. 2019, 18, 3605–3612. [Google Scholar] [CrossRef] [PubMed] [Green Version]
De Quirós, S.B.; Merlo, A.; Secades, P.; Zambrano, I.; de Santa María, I.S.; Ugidos, N.; Jantus-Lewintre, E.; Sirera, R.; Suarez, C.; Chiara, M.-D. Identification of TRPC6 as a possible candidate target gene within an amplicon at 11q21-q22.2 for migratory capacity in head and neck squamous cell carcinomas. BMC Cancer 2013, 13, 116. [Google Scholar] [CrossRef] [Green Version]
Ramer, R.; Merkord, J.; Rohde, H.; Hinz, B. Cannabidiol inhibits cancer cell invasion via upregulation of tissue inhibitor of matrix metalloproteinases-1. Biochem. Pharmacol. 2010, 79, 955–966. [Google Scholar] [CrossRef] [Green Version]
Gaunt, H.J.; Vasudev, N.S.; Beech, D.J. Transient receptor potential canonical 4 and 5 proteins as targets in cancer therapeutics. Eur. Biophys. J. 2016, 45, 611–620. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Kanugula, A.K.; Adapala, R.K.; Midha, P.; Cappelli, H.C.; Meszaros, J.G.; Paruchuri, S.; Chilian, W.M.; Thodeti, C.K. Novel noncanonical regulation of soluble VEGF/VEGFR2 signaling by mechanosensitive ion channel TRPV4. FASEB J. 2019, 33, 195–203. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Bernardini, M.; Brossa, A.; Chinigo, G.; Grolez, G.P.; Trimaglio, G.; Allart, L.; Hulot, A.; Marot, G.; Genova, T.; Joshi, A.; et al. Transient Receptor Potential Channel Expression Signatures in Tumor-Derived Endothelial Cells: Functional Roles in Prostate Cancer Angiogenesis. Cancers 2019, 11, 956. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Santoni, G.; Morelli, M.; Santoni, M.; Nabissi, M. New deals on the transcriptional and post-transcriptional regulation of TRP channel target genes during the angiogenesis of glioma. J. Exp. Integr. Med. 2011, 1, 221–234. [Google Scholar] [CrossRef]
Mariot, P.; Prevarskaya, N.; Roudbaraki, M.M.; Le Bourhis, X.; Van Coppenolle, F.; Vanoverberghe, K.; Skryma, R. Evidence of functional ryanodine receptor involved in apoptosis of prostate cancer (Lncap) cells. Prostate 2000, 43, 205–214. [Google Scholar] [CrossRef]
Shin, D.-H.; Leem, D.-G.; Shin, J.-S.; Kim, J.-I.; Kim, K.-T.; Choi, S.Y.; Lee, M.-H.; Choi, J.-H.; Lee, K.-T. Compound K induced apoptosis via endoplasmic reticulum Ca²⁺ release through ryanodine receptor in human lung cancer cells. J. Ginseng Res. 2018, 42, 165–174. [Google Scholar] [CrossRef]
Rezuchova, I.; Hudecova, S.; Soltysova, A.; Matuskova, M.; Durinikova, E.; Chovancova, B.; Zuzcak, M.; Cihova, M.; Burikova, M.; Penesova, A.; et al. Type 3 inositol 1,4,5-trisphosphate receptor has antiapoptotic and proliferative role in cancer cells. Cell Death Dis. 2019, 10, 186. [Google Scholar] [CrossRef] [Green Version]
Akl, H.; Monaco, G.; La Rovere, R.; Welkenhuyzen, K.; Kiviluoto, S.; Vervliet, T.; Molgó, J.; Distelhorst, C.W.; Missiaen, L.; Mikoshiba, K.; et al. IP3R2 levels dictate the apoptotic sensitivity of diffuse large B-cell lymphoma cells to an IP3R-derived peptide targeting the BH4 domain of Bcl-2. Cell Death Dis. 2013, 4, e632. [Google Scholar] [CrossRef] [PubMed]
Boutin, B.; Tajeddine, N.; Monaco, G.; Molgo, J.; Vertommen, D.; Rider, M.; Parys, J.; Bultynck, G.; Gailly, P. Endoplasmic reticulum Ca²⁺ content decrease by PKA-dependent hyperphosphorylation of type 1 IP3 receptor contributes to prostate cancer cell resistance to androgen deprivation. Cell Calcium 2015, 57, 312–320. [Google Scholar] [CrossRef]
Kiss, F.; Pohóczky, K.; Szállási, A.; Helyes, Z. Transient Receptor Potential (TRP) Channels in Head-and-Neck Squamous Cell Carcinomas: Diagnostic, Prognostic, and Therapeutic Potentials. Int. J. Mol. Sci. 2020, 21, 6374. [Google Scholar] [CrossRef]
Monet, M.; Gkika, D.; Lehen’Kyi, V.; Pourtier, A.; Abeele, F.V.; Bidaux, G.; Juvin, V.; Rassendren, F.; Humez, S.; Prevarsakaya, N. Lysophospholipids stimulate prostate cancer cell migration via TRPV2 channel activation. Biochim. Biophys. Acta (BBA)—Bioenerg. 2009, 1793, 528–539. [Google Scholar] [CrossRef] [Green Version]
Liu, X.; Zou, J.; Su, J.; Lu, Y.; Zhang, J.; Li, L.; Yin, F. Downregulation of transient receptor potential cation channel, subfamily C, member 1 contributes to drug resistance and high histological grade in ovarian cancer. Int. J. Oncol. 2015, 48, 243–252. [Google Scholar] [CrossRef] [Green Version]
Cardenas, C.; Lovy, A.; Silva-Pavez, E.; Urra, F.; Mizzoni, C.; Ahumada-Castro, U.; Bustos, G.; Jaňa, F.; Cruz, P.; Farias, P.; et al. Cancer cells with defective oxidative phosphorylation require endoplasmic reticulum–to–mitochondria Ca²⁺transfer for survival. Sci. Signal. 2020, 13, eaay1212. [Google Scholar] [CrossRef]
Law, B.Y.K.; Michelangeli, F.; Qu, Y.Q.; Xu, S.-W.; Han, Y.; Mok, S.W.F.; de Seabra Rodrigues Dias, I.R.; Javed, M.-H.; Chan, W.-K.; Xue, W.-W.; et al. Neferine induces autophagy-dependent cell death in apoptosis-resistant cancers via ryanodine receptor and Ca²⁺-dependent mechanism. Sci. Rep. 2019, 9, 20034. [Google Scholar] [CrossRef] [PubMed]
Fels, B.; Bulk, E.; Pethő, Z.; Schwab, A. The Role of TRP Channels in the Metastatic Cascade. Pharmaceuticals 2018, 11, 48. [Google Scholar] [CrossRef] [Green Version]
Liu, X.; Zhang, P.; Xie, C.; Sham, K.W.-Y.; Ng, S.S.M.; Chen, Y.; Cheng, C.H.K. Activation of PTEN by inhibition of TRPV4 suppresses colon cancer development. Cell Death Dis. 2019, 10, 1–14. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Huang, Y.; Li, S.; Jia, Z.; Zhao, W.; Zhou, C.; Zhang, R.; Ali, D.W.; Michalak, M.; Chen, X.-Z.; Tang, J. Transient Receptor Potential Melastatin 8 (TRPM8) Channel Regulates Proliferation and Migration of Breast Cancer Cells by Activating the AMPK-ULK1 Pathway to Enhance Basal Autophagy. Front. Oncol. 2020, 10, 2645. [Google Scholar] [CrossRef]
Lan, X.; Zhao, J.; Song, C.; Yuan, Q.; Liu, X. TRPM8 facilitates proliferation and immune evasion of esophageal cancer cells. Biosci. Rep. 2019, 39, BSR20191878. [Google Scholar] [CrossRef] [Green Version]
Yu, S.; Huang, S.; Ding, Y.; Wang, W.; Wang, A.; Lu, Y. Transient receptor potential ion-channel subfamily V member 4: A potential target for cancer treatment. Cell Death Dis. 2019, 10, 1–17. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Yamamoto, M.; Horie, I.; Isohama, Y.; Tsukimoto, M. Activation of TRPV4 Channel Regulates Differentiation to and Function of Myeloid-Derived Suppressor Cells. BPB Rep. 2020, 3, 70–75. [Google Scholar] [CrossRef]
Eliaa, S.G.; Al-Karmalawy, A.A.; Saleh, R.M.; ElShal, M.F. Empagliflozin and Doxorubicin Synergistically Inhibit the Survival of Triple-Negative Breast Cancer Cells via Interfering with the mTOR Pathway and Inhibition of Calmodulin: In Vitro and Molecular Docking Studies. ACS Pharmacol. Transl. Sci. 2020, 3, 1330–1338. [Google Scholar] [CrossRef]
Pawar, P.; Ma, L.; Byon, C.H.; Liu, H.; Ahn, E.-Y.; Jhala, N.; Arnoletti, J.P.; McDonald, J.M.; Chen, Y. Molecular Mechanisms of Tamoxifen Therapy for Cholangiocarcinoma: Role of Calmodulin. Clin. Cancer Res. 2009, 15, 1288–1296. [Google Scholar] [CrossRef] [Green Version]
Mine, N.; Yamamoto, S.; Saito, N.; Yamazaki, S.; Suda, C.; Ishigaki, M.; Kufe, D.W.; Von Hoff, D.D.; Kawabe, T. CBP501-Calmodulin Binding Contributes to Sensitizing Tumor Cells to Cisplatin and Bleomycin. Mol. Cancer Ther. 2011, 10, 1929–1938. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Chen, P.; Wu, Q.; Feng, J.; Yan, L.; Sun, Y.; Liu, S.; Xiang, Y.; Zhang, M.; Pan, T.; Chen, X.; et al. Erianin, a novel dibenzyl compound in Dendrobium extract, inhibits lung cancer cell growth and migration via calcium/calmodulin-dependent ferroptosis. Signal Transduct. Target. Ther. 2020, 5, 1–11. [Google Scholar] [CrossRef] [PubMed]
Merlot, A.M.; Shafie, N.H.; Yu, Y.; Richardson, V.; Jansson, P.J.; Sahni, S.; Lane, D.; Kovacevic, Z.; Kalinowski, D.S.; Richardson, D. Mechanism of the induction of endoplasmic reticulum stress by the anti-cancer agent, di-2-pyridylketone 4,4-dimethyl-3-thiosemicarbazone (Dp44mT): Activation of PERK/eIF2α, IRE1α, ATF6 and calmodulin kinase. Biochem. Pharmacol. 2016, 109, 27–47. [Google Scholar] [CrossRef] [PubMed]
Rewcastle, G.W.; Baguley, B.C.; Atwell, G.J.; Denny, W.A. Potential antitumor agents. 52. Carbamate analogs of amsacrine with in vivo activity against multidrug-resistant P388 leukemia. J. Med. Chem. 1987, 30, 1576–1581. [Google Scholar] [CrossRef]
Mayur, Y.C.; Jagadeesh, S.; Thimmaiah, K.N. Targeting calmodulin in reversing multi drug resistance in cancer cells. Mini-Reviews Med. Chem. 2006, 6, 1383–1389. [Google Scholar] [CrossRef]
Kim, C.; Kim, B. Anti-Cancer Natural Products and Their Bioactive Compounds Inducing ER Stress-Mediated Apoptosis: A Review. Nutrients 2018, 10, 1021. [Google Scholar] [CrossRef] [Green Version]
Koido, S.; Kan, S.; Yoshida, K.; Yoshizaki, S.; Takakura, K.; Namiki, Y.; Tsukinaga, S.; Odahara, S.; Kajihara, M.; Okamoto, M.; et al. Immunogenic modulation of cholangiocarcinoma cells by chemoimmunotherapy. Anticancer. Res. 2014, 34, 6353–6361. [Google Scholar] [PubMed]
Humeau, J.; Sauvat, A.; Cerrato, G.; Xie, W.; Loos, F.; Iannantuoni, F.; Bezu, L.; Lévesque, S.; Paillet, J.; Pol, J.; et al. Inhibition of transcription by dactinomycin reveals a new characteristic of immunogenic cell stress. EMBO Mol. Med. 2020, 12, e11622. [Google Scholar] [CrossRef] [PubMed]
Charlier, H.A., Jr.; Olson, R.D.; Thornock, C.M.; Mercer, W.K.; Olson, D.R.; Broyles, T.S.; Muhlestein, D.J.; Larson, C.L.; Cusack, B.J.; Shadle, S.E. Investigations of Calsequestrin as a Target for Anthracyclines: Comparison of Functional Effects of Daunorubicin, Daunorubicinol, and Trifluoperazine. Mol. Pharmacol. 2005, 67, 1505–1512. [Google Scholar] [CrossRef]
Gorini, S.; De Angelis, A.; Berrino, L.; Malara, N.; Rosano, G.; Ferraro, E. Chemotherapeutic Drugs and Mitochondrial Dysfunction: Focus on Doxorubicin, Trastuzumab, and Sunitinib. Oxidative Med. Cell. Longev. 2018, 2018, 1–15. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Tadini-Buoninsegni, F.; Smeazzetto, S.; Gualdani, R.; Moncelli, M.R. Drug Interactions With the Ca²⁺-ATPase From Sarco(Endo)Plasmic Reticulum (SERCA). Front. Mol. Biosci. 2018, 5. [Google Scholar] [CrossRef]
Yiallouris, A.; Patrikios, I.; Johnson, E.O.; Sereti, E.; Dimas, K.; De Ford, C.; Fedosova, N.U.; Graier, W.F.; Sokratous, K.; Kyriakou, K.; et al. Annonacin promotes selective cancer cell death via NKA-dependent and SERCA-dependent pathways. Cell Death Dis. 2018, 9, 1–13. [Google Scholar] [CrossRef]
Varghese, E.; Samuel, S.M.; Sadiq, Z.; Kubatka, P.; Liskova, A.; Benacka, J.; Pazinka, P.; Kruzliak, P.; Büsselberg, D. Anti-Cancer Agents in Proliferation and Cell Death: The Calcium Connection. Int. J. Mol. Sci. 2019, 20, 3017. [Google Scholar] [CrossRef] [Green Version]
Stenius, U.; Miraglia, E.; Högberg, J. Statins exhibit anticancer effects through modifications of the pAkt signaling pathway. Int. J. Oncol. 2011, 40, 867–875. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Stokes, L.; Bidula, S.; Bibič, L.; Allum, E. To Inhibit or Enhance? Is There a Benefit to Positive Allosteric Modulation of P2X Receptors? Front. Pharmacol. 2020, 11, 627. [Google Scholar] [CrossRef] [PubMed]
Mitrugno, A.; Sylman, J.L.; Rigg, R.A.; Yunga, S.T.; Shatzel, J.J.; Williams, C.D.; Mccarty, O.J. Carpe low-dose aspirin: The new anti-cancer face of an old anti-platelet drug. Platelets 2017, 29, 773–778. [Google Scholar] [CrossRef] [PubMed]
Ballerini, P.; Dovizio, M.; Bruno, A.; Tacconelli, S.; Patrignani, P. P2Y12 Receptors in Tumorigenesis and Metastasis. Front. Pharmacol. 2018, 9, 66. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Bentham Science Publisher Bentham Science Publisher Pharmacological Inhibition of Voltage-gated Ca²⁺ Channels for Chronic Pain Relief. Curr. Neuropharmacol. 2013, 11, 606–620. [CrossRef] [Green Version]
Vashisht, A.; Trebak, M.; Motiani, R.K. STIM and Orai proteins as novel targets for cancer therapy. A Review in the Theme: Cell and Molecular Processes in Cancer Metastasis. Am. J. Physiol. Physiol. 2015, 309, C457–C469. [Google Scholar] [CrossRef] [Green Version]
Echakraborty, S.; Eghosh, S.; Ebanerjee, B.; Santra, A.; Eadhikary, A.; Misra, A.K.; Sen, P.C. Phemindole, a Synthetic Di-indole Derivative Maneuvers the Store Operated Calcium Entry (SOCE) to Induce Potent Anti-Carcinogenic Activity in Human Triple Negative Breast Cancer Cells. Front. Pharmacol. 2016, 7, 114. [Google Scholar] [CrossRef] [Green Version]
Ali, S.; Doan, D.; Ojong, T.; Solomon, H.; Corpuz, E.; Huynh, T.; Haziq, M.; Shahid, M.; Siddiqui, M.R.; Newaz, M.; et al. Metformin Attenuates High Glucose-Induced Coronary Vascular Endothelial Hyper Permeability Via Inhibition of Orai-1 Mediated Store-Operated Calcium Entry. FASEB J. 2020, 34, 1. [Google Scholar] [CrossRef]
Song, M.; Chen, D.; Yu, S.P. The TRPC channel blocker SKF 96365 inhibits glioblastoma cell growth by enhancing reverse mode of the Na⁺/Ca²⁺exchanger and increasing intracellular Ca²⁺. Br. J. Pharmacol. 2014, 171, 3432–3447. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Rodrigues, T.; Sieglitz, F.; Bernardes, G. Natural product modulators of transient receptor potential (TRP) channels as potential anti-cancer agents. Chem. Soc. Rev. 2016, 45, 6130–6137. [Google Scholar] [

Article Menu

Advances in Intracellular Calcium Signaling Reveal Untapped Targets for Cancer Therapy

Abstract

1. Introduction

2. Intracellular Ca2+ Buffers in Normal Cells

2.1. Soluble and Unbound Intracellular Proteins: Calmodulin, Calbindin, and Calretinin

2.2. Intramembranous Molecular Buffers: SERCA, PMCA, NCX, and TRP

2.3. Cellular Organelles

2.3.1. Endoplasmic Reticulum: STIM, ORAI, IP3Rs, and TRPC1 in SOCE and SOCIC Ca2+ Entry Models

2.3.2. Mitochondria and Acidic Vesicles (Mainly Lysosomes)

3. Redistribution of Intracellular Ca2+ and Hijack of Its Regulatory Machinery in Cancer Cells

3.1. Intracellular Ca2+ Pool in the Endoplasmic Reticulum and at the ER-PM Junction

3.2. Intracellular Ca2+ Pool at ER-Mitochondrial Junction

3.3. Intracellular Ca2+ Pool at ER-Lysosome Junction

4. Conclusions and Discussion

Author Contributions

Funding

Data Availability Statement

Conflicts of Interest

References

2. Intracellular Ca²⁺ Buffers in Normal Cells

2.3.1. Endoplasmic Reticulum: STIM, ORAI, IP₃Rs, and TRPC1 in SOCE and SOCIC Ca²⁺ Entry Models

3. Redistribution of Intracellular Ca²⁺ and Hijack of Its Regulatory Machinery in Cancer Cells

3.1. Intracellular Ca²⁺ Pool in the Endoplasmic Reticulum and at the ER-PM Junction

3.2. Intracellular Ca²⁺ Pool at ER-Mitochondrial Junction

3.3. Intracellular Ca²⁺ Pool at ER-Lysosome Junction