Sodium regulates PLC and IP₃R‐mediated calcium signaling in invasive breast cancer cells

Abstract

Intracellular Ca²⁺ signaling and Na⁺ homeostasis are inextricably linked via ion channels and co‐transporters, with alterations in the concentration of one ion having profound effects on the other. Evidence indicates that intracellular Na⁺ concentration ([Na⁺]_i) is elevated in breast tumors, and that aberrant Ca²⁺ signaling regulates numerous key cancer hallmark processes. The present study therefore aimed to determine the effects of Na⁺ depletion on intracellular Ca²⁺ handling in metastatic breast cancer cell lines. The relationship between Na⁺ and Ca²⁺ was probed using fura‐2 and SBFI fluorescence imaging and replacement of extracellular Na⁺ with equimolar N‐methyl‐D‐glucamine (0Na⁺/NMDG) or choline chloride (0Na⁺/ChoCl). In triple‐negative MDA‐MB‐231 and MDA‐MB‐468 cells and Her2+ SKBR3 cells, but not ER+ MCF‐7 cells, 0Na⁺/NMDG and 0Na⁺/ChoCl resulted in a slow, sustained depletion in [Na⁺]_i that was accompanied by a rapid and sustained increase in intracellular Ca²⁺ concentration ([Ca²⁺]_i). Application of La³⁺ in nominal Ca²⁺‐free conditions had no effect on this response, ruling out reverse‐mode NCX activity and Ca²⁺ entry channels. Moreover, the Na⁺‐linked [Ca²⁺]_i increase was independent of membrane potential hyperpolarization (NS‐1619), but was inhibited by pharmacological blockade of IP₃ receptors (2‐APB), phospholipase C (PLC, U73122) or following depletion of endoplasmic reticulum Ca²⁺ stores (cyclopiazonic acid). Thus, Na⁺ is linked to PLC/IP₃‐mediated activation of endoplasmic reticulum Ca²⁺ release in metastatic breast cancer cells and this may have an important role in breast tumors where [Na⁺]_i is perturbed.

Intracellular Na⁺ concentration is elevated in breast cancer cells, and aberrant Ca²⁺ signaling regulates numerous key cancer hallmark processes. Here, we found that Na⁺ depletion in metastatic breast cancer cell lines led to a rapid and sustained increase in intracellular Ca²⁺ concentration via PLC/IP₃‐mediated activation of endoplasmic reticulum Ca²⁺ release. Our findings suggest that Na⁺‐dependent alteration of Ca²⁺ handling may have an important role in breast tumors where Na⁺ is perturbed.

1. INTRODUCTION

Early detection has played an important role for timely clinical intervention in breast cancer; however, development of novel therapies which selectively kill malignant cells remains a central research goal. Key hallmarks of cancer include unlimited replicative capacity, self‐sufficiency of growth, apoptosis resistance, angiogenesis, and ultimately tissue invasion (Hanahan & Weinberg, 2016). Importantly, intracellular ion signaling pathways (in particular Ca²⁺ and Na⁺ signaling) play a key role in regulating many of these hallmark processes in breast cancer (Bruce & James, 2020; Capatina et al., 2022; Leslie et al., 2019; Yang & Brackenbury, 2022), and thus ion signaling may be a novel therapeutic locus.

Intracellular [Na⁺] ([Na⁺]_i) and intracellular [Ca²⁺] ([Ca²⁺]_i) signaling is achieved via a multiplicity of ion channels and transporters located both at the plasma membrane and on intracellular organelles. These channels harness the electrochemical gradient of Na⁺ and Ca²⁺ across cell membranes to achieve their function, with [Na⁺]_i and [Ca²⁺]_i typically tightly regulated by various ion transporters (Bruce, 2018; Leslie et al., 2019). Importantly, Ca²⁺ and Na⁺ signaling are inextricably linked via mechanisms mutually regulated by both ions, such as the plasma membrane Na⁺/Ca²⁺ exchanger (NCX; which is responsible for bulk export of Ca²⁺ from the cytosol) and the mitochondrial Na⁺/Ca²⁺/Li⁺ exchanger (which regulates mitochondrial Ca²⁺ and Na⁺ and thus bioenergetics, Hernansanz‐Agustín et al., 2020). Thus, changes in how the cell handles one ion would be expected to impact upon the other. Moreover, the interplay between [Na⁺]_i and [Ca²⁺]_i handling has important implications in breast cancer since [Na⁺]_i is elevated in malignant breast cancer cells, particularly in those exhibiting aberrant Na⁺ channel expression (Cameron et al., 1980; James et al., 2022; Ouwerkerk et al., 2007; Yang et al., 2020).

Emerging evidence implicates altered Na⁺ and Ca²⁺ signaling in the progression of breast cancer (Bruce & James, 2020; Leslie et al., 2019). For example, aberrantly expressed voltage‐gated Na⁺ channels (VGSCs) drive metastasis (Bon et al., 2016; Brackenbury et al., 2007; Brisson et al., 2013; Driffort et al., 2014; Fraser et al., 2005; Nelson et al., 2014; Nelson, Yang, Dowle, et al., 2015; Nelson, Yang, Millican‐Slater, & Brackenbury, 2015), elevations in intracellular Na⁺ correlate with malignancy and treatment response Ouwerkerk et al., 2007; James et al., 2022) and upregulation of numerous Ca²⁺ channels and ATPases (and thus alterations in Ca²⁺ handling) are linked with breast cancer progression and poor prognosis (Davis et al., 2014; Feng et al., 2010; Jeong et al., 2016; Middelbeek et al., 2012; So et al., 2019; VanHouten et al., 2010). The apparent elevation in [Na⁺]_i exhibited by breast cancer cells might be expected to affect Na⁺‐dependent Ca²⁺ handling mechanisms; for example, NCX, which can operate in reverse, Ca²⁺ entry mode following changes in [Na⁺]_i (Pappalardo et al., 2014; Verkhratsky et al., 2018. Nevertheless, the interplay between Na⁺ and Ca²⁺ handling in the context of cancer is poorly understood (Bruce & James, 2020; Leslie et al., 2019).

The present study aimed to determine the effects of Na⁺ depletion on intracellular Ca²⁺ handling in metastatic breast cancer cell lines in order to probe the relationship between Na⁺ and Ca²⁺ in breast cancer. A commonly recognized method to probe the role of Na⁺ transport in cell physiology is by replacement of extracellular Na⁺ with equimolar N‐methyl‐D‐glucamine (0Na⁺/NMDG) or choline chloride (0Na⁺/ChoCl). These replacement cations maintain the osmotic balance across the cell membrane, yet are not transported via Na⁺ channels or transporters. Breast cancer cells can be subdivided into three main categories based on expression of the estrogen receptor (ER), progesterone receptor and human epidermal growth factor 2 (Her2). In triple‐negative MDA‐MB‐231 and MDA‐MB‐468 cells (lacking all three receptors) and Her2+ SKBR3 cells, removal of Na⁺ from the extracellular space led to a slow, sustained depletion in [Na⁺]_i that was accompanied by a rapid and sustained increase in intracellular [Ca²⁺]_i. Interestingly, this response was absent in ER+ MCF‐7 cells. The observed [Ca²⁺]_i transient was not due to Ca²⁺ entry from the extracellular space, ruling out reverse‐mode NCX activity. Moreover, it was independent of membrane potential (V_m) and was inhibited by depletion of the endoplasmic reticulum Ca²⁺ store or by pharmacological blockade of inositol (1,4,5) trisphosphate (IP₃) receptors (IP₃Rs) or phospholipase C (PLC). These data reveal a previously unreported Na⁺‐linked activation of endoplasmic reticulum Ca²⁺ release in metastatic breast cancer cells, which may have an important role in breast tumors where [Na⁺]_i is elevated or perturbed. Moreover, the dramatic Ca²⁺ release observed upon Na⁺ depletion serves as a cautionary note for those utilizing similar Na⁺ replacement methods to study the role of Na⁺ transport in cellular processes.

2. MATERIALS AND METHODS

2.1. Cell culture

Cells were cultured at 37°C in a humidified atmosphere of air/CO₂ (95:5%) in Dulbecco's modified Eagle's medium (DMEM 219690–35, Thermo Fisher Scientific) supplemented with 4 mM L‐glutamine and 5% fetal bovine serum, as described previously (Brisson et al., 2013; Ding & Djamgoz, 2004; Pan & Djamgoz, 2008; Yang et al., 2012). Cells were routinely tested for mycoplasma using the DAPI method (Uphoff et al., 1992). MDA‐MB‐231 and MCF‐7 cells were gifts from M. Djamgoz (Imperial College London) and SKBR3 cells were a gift from J. Rae (University of Michigan). Molecular identity of MDA‐MB‐231, MCF‐7, and SKBR3 cells was verified by short tandem repeat analysis (Masters et al., 2001). MDA‐MB‐468 cells were obtained for this study directly from the American Type Culture Collection (ATCC).

2.2. Fura‐2 and SBFI fluorescence microscopy

Cells were seeded onto glass coverslips and allowed to adhere overnight. Cells were loaded with either fura‐2 AM (4 μM) or SBFI AM (4 μM) in HEPES‐buffered physiological saline solution (HEPES‐PSS: 144 mM NaCl, 5.4 mM KCl, 1 mM MgCl₂, 1.28 mM CaCl₂, 5 mM HEPES and 5.6 mM glucose, pH 7.2) with 0.08% Pluronic F‐127 at room temperature (fura‐2 AM, 40 min; SBFI AM, 2 h). Cells were then rinsed once with HEPES‐PSS and incubated in dye‐free HEPES‐PSS to allow uncleaved dye to re‐equilibrate (fura‐2 AM, 20 min; SBFI AM, 40 min). For all experiments except those shown in Supplementary Figure S1, loaded coverslips were mounted within a perfusion chamber (RC26G, Warner Instruments) fitted to an imaging system comprising an Eclipse TE‐200 inverted microscope (Nikon), a Plan Fluor ELWD 20×/0.45 Ph1 objective (Nikon Corporation) and a Rolera XR 12 Bit Fast 1394 CCD camera (QImaging, Surrey, British Columbia) controlled by SimplePCI software (Hamamatsu). Excitation (340 and 380 nm, 50 ms exposure, mercury bulb) and emission light were separated using a 400 DCLP dichroic with a D510/80 m filter. Data shown in Supplementary Figure S1 were acquired using a system comprised of an BX51WI (Olympus) microscope, a LUMPLFLN 40XW water‐dipping lens (Olympus), a pE‐340fura LED illumination system (CoolLED) and a SciCam Pro (Scientifica), controlled by μManager 2.0. Cells were perfused with HEPES‐PSS using a gravity‐fed perfusion system at a continuous rate of ~2 mL/min. Na⁺‐free HEPES PSS was prepared as for HEPES‐PSS above but with NaCl replaced with equimolar N‐methyl‐D‐glucamine (0Na⁺/NMDG) or choline chloride (0Na⁺/ChoCl). Nominal Ca²⁺‐free HEPES PSS was prepared as for HEPES‐PSS above, omitting CaCl₂.

2.3. Drug preparation

Stocks of NS‐1619, 2‐APB, dantrolene, cyclopiazonic acid (CPA), and ionomycin were prepared in DMSO. The final DMSO concentration in working solutions was 0.1%. Stocks of La³⁺ were prepared using ultrapure water. Drugs were diluted into HEPES‐PSS immediately prior to use.

2.4. Measurement of [Ca²⁺]_i responses

Changes in [Ca²⁺]_i in response to treatment were measured as either the maximum fura‐2 ratio change (ΔR_max) or the area under the curve (AUC). The fura‐2 fluorescence ratios for each trace were first normalized to the ratio at time 0 (R/R₀) to control for variability in starting ratio between cells. To obtain a pretreatment baseline, the average ratio across the six timepoints (30 seconds at an acquisition interval of 5 seconds) immediately prior to treatment application was determined. ΔR_max was defined as the maximum increase from this baseline during the treatment period. The AUC was defined as the area under the curve during the entire treatment period (i.e., 10 min) in fura‐2 ratio unit seconds (R.s). Where no [Ca²⁺]_i response was elicited in response to treatment, cells were stimulated with ionomycin (3 μM) to ensure a change in [Ca²⁺]_i could be measured. Cells in which no fura‐2 fluorescence change was observed were excluded from analysis.

2.5. Data analysis

Data analysis and statistical comparisons were performed using Microsoft Excel and GraphPad Prism 9. Statistical comparisons were performed using a nested t‐test for side‐by‐side comparisons and a nested one‐way ANOVA with posthoc Tukey test for multiple comparisons. Within each figure, nested data are presented from individual experiments (3–6 independent repeats, each containing 12–40 independent cells) and the individual experimental means ± S.E.M.

3. RESULTS

3.1. Na⁺ depletion drives an increase in [Ca²⁺]_i in breast cancer cells

Metastatic breast cancer cells exhibit elevated [Na⁺]_i (Leslie et al., 2019) that may regulate reverse‐mode (Ca²⁺‐entry) NCX activity (Pappalardo et al., 2014; Verkhratsky et al., 2018) in these cells. To assess the effects of lowering extracellular Na⁺ on [Ca²⁺]_i, MDA‐MB‐231 cells were loaded with fura‐2 AM (4 μM) and perfused with sequential pulses of HEPES‐PSS where Na⁺ had been replaced with either equimolar N‐methyl D glucamine (0Na⁺/NMDG) or equimolar choline chloride (0Na⁺/ChoCl) to remove extracellular Na⁺ while maintaining osmotic balance. Fura‐2 fluorescence imaging revealed that, using either maneuver, removal of extracellular Na⁺ resulted in a steep increase in [Ca²⁺]_i that returned to baseline upon reperfusion with standard Na⁺‐containing HEPES‐PSS (Figure 1ai,bi). In parallel experiments, SBFI fluorescence imaging revealed that both 0Na⁺/NMDG and 0Na⁺/ChoCl resulted in a concomitant depletion of [Na⁺]_i, suggesting a link between [Na⁺]_i depletion and subsequent [Ca²⁺]_i. A similar increase in [Ca²⁺]_i upon perfusion with 0Na⁺/NMDG was observed in a second triple‐negative breast cancer cell line, MDA‐MB‐468 (Supplementary Figure S1A). Interestingly, perfusion of 0Na⁺/NMDG had no effect on [Ca²⁺]_i in the ER+ breast cancer cell line MCF‐7 (Figure 1c), but induced a [Ca²⁺]_i transient in the Her2+ breast cancer cell line SKBR3 (Figure 1d); nevertheless, both MCF‐7 and SKBR3 cells exhibited depletion of [Na⁺]_i upon perfusion with 0Na⁺/NMDG (Supplementary Figure S1b,c). Taken together, these data suggest that the [Ca²⁺]_i rise upon removal of extracellular Na⁺ and concomitant depletion of [Na⁺]_i is present only in certain breast cancer cell lines.

FIGURE 1.

Na⁺‐free conditions deplete [Na⁺]_i and induce elevated [Ca²⁺]_i in breast cancer cells. Fura‐2 AM (4 μM) or SBFI AM (4 μM) fluorescence microscopy was used to measure [Ca²⁺]_i and [Na⁺]_i in cultured human breast cancer cells. Following perfusion with HEPES‐PSS, MDA‐MB‐231 cells were perfused with Na⁺‐free HEPES PSS; extracellular Na⁺ was replaced with equimolar N‐methyl‐D‐glucamine (0Na⁺/NMDG) or choline chloride (0Na⁺/ChoCl) to maintain osmotic balance. Representative traces show the effects of 0Na⁺/NMDG and 0Na⁺/ChoCl on [Ca²⁺]_i (ai and bi, respectively) and [Na⁺]_i (aii and bii, respectively) in MDA‐MB‐231 cells. Additional experiments were performed in MCF‐7 (c) and SKBR3 (d) cells. Ionomycin (3 μM) was used to elicit a [Ca⁺]_i increase as a positive control.

3.2. The [Ca²⁺]_i increase induced by Na⁺‐depletion is not due to Ca²⁺ influx from the extracellular space

We reasoned that the increase in [Ca²⁺]_i observed under 0Na⁺ conditions likely originated from one of two compartments: Ca²⁺ release from intracellular stores (most likely the endoplasmic reticulum) or Ca²⁺ influx from the extracellular space. To determine whether Ca²⁺ influx from the extracellular space contributed to the [Ca²⁺]_i responses observed, MDA‐MB‐231 cells were perfused with 0Na⁺/NMDG in nominal Ca²⁺‐free conditions with La³⁺ (1 mM); La³⁺ in the millimolar range is a known inhibitor of both Ca²⁺ entry from and Ca²⁺ efflux to the extracellular space (James et al., 2013). La³⁺ was applied for 5 minutes prior to application of 0Na⁺/NMDG (Figure 2b). Compared with control cells (Figure 2a), La³⁺ had no effect on the [Ca²⁺]_i transients elicited following 0Na⁺/NMDG application, as measured by either the maximum change in fura‐2 ratio (ΔR_max, Figure 2c) or the area under the curve (AUC, Figure 2d). These results indicate that Ca²⁺ influx via agonist‐induced and store‐operated Ca²⁺ entry mechanisms plays no role in the [Ca²⁺]_i transients induced following application of 0Na⁺/NMDG, and that the provenance of this Ca²⁺ response must be some compartment other than the extracellular space.

FIGURE 2.

[Ca²⁺]_i transients induced by removal of extracellular Na⁺ in breast cancer cells are not due to Ca²⁺ entry. MDA‐MB‐231 cells were loaded with fura‐2 AM (4 μM) and perfused with HEPES‐PSS. Extracellular Na⁺ was replaced with equimolar 0Na⁺/NMDG to maintain osmotic balance (a). To block all Ca²⁺ entry from the extracellular space, cells were pretreated with 5 min of perfusion with Ca²⁺‐free HEPES containing La³⁺ (1 mM, b). Ionomycin (3 μM) was applied at the end of an experiment to elicit a [Ca⁺]_i increase as a positive control. The maximum change in fluorescence ratio (ΔR_max, c) and area under the curve (AUC, d) were compared using a nested t‐test for side‐by‐side comparisons; n = 4 independent experiments for each condition, ns, not significant compared with control. Data presented are nested values for individual cells grouped by experiment (gray dots) and experimental means ± SEM (black line and bars).

3.3. Elevations in [Ca²⁺]_i induced by Na⁺‐depletion are not due to changes in membrane potential

Maneuvers that deplete extracellular Na⁺ such as 0Na⁺/NMDG are known to cause V_m hyperpolarization (Yang et al., 2020), thereby altering ion flux across the plasma membrane. Moreover, V_m hyperpolarization would increase the driving force for Ca²⁺ entry. To determine whether such changes in V_m led to the [Ca²⁺]_i transient observed under 0Na⁺/NMDG conditions, we applied the large‐conductance Ca²⁺‐activated K⁺ channel (K_Ca1.1) activator NS‐1619 at a concentration (10 μM) that hyperpolarizes V_m in MDA‐MB‐231 cells to a similar degree to 0Na⁺/NMDG (Yang et al., 2020). Similar to control cells (Figure 3a), MDA‐MB‐231 cells treated with NS‐1619 (10 μM, Figure 3b) showed no change in [Ca²⁺]_i. However, an increased concentration of NS‐1619 (40 μM, Figure 3c) elicited an increase in [Ca²⁺]_i that was significantly greater than any change observed in either the control cells (p < 0.01 for both ΔR_max and AUC, Figure 3e,f) or those treated with 10 μM NS‐1619 (p < 0.01 for both ΔR_max and AUC, Figure 3e,f), indicating that dramatic changes in V_m can indeed alter [Ca²⁺]_i. Importantly, the effects of 40 μM NS‐1619 on [Ca²⁺]_i were abolished following pretreatment with La³⁺ (1 mM, p < 0.05 for both ΔR_max and AUC, Figure 3c–f), indicating that the NS‐1619 induced rise was due to Ca²⁺ influx from the extracellular space (Figure 3d). Given that Ca²⁺ influx plays no role in the [Ca²⁺]_i transients observed following application of 0Na⁺/NMDG (Figure 2), these results suggest that neither Ca²⁺ influx nor hyperpolarization of V_m play a role in changes in [Ca²⁺]_i induced by application 0Na⁺/NMDG.

FIGURE 3.

Hyperpolarization of the membrane potential (V_m) plays no role in [Ca²⁺]_i transients in breast cancer cells induced by removal of extracellular Na⁺. MDA‐MB‐231 cells were loaded with fura‐2 AM (4 μM) and perfused with HEPES‐PSS in the presence of the large‐conductance Ca²⁺‐activated K⁺ channel (K_Ca1.1) activator NS‐1619 to hyperpolarize V_m. Representative traces are shown for cells perfused without NS‐1619 (a) and with 10 μM NS‐1619 (b) or 40 μM NS‐1619 (c). Application of La³⁺ (d) determined that [Ca²⁺]_i changes observed following treatment with 40 μM NS‐1619 were due to Ca²⁺ entry. Ionomycin (3 μM) was applied at the end of an experiment to elicit a [Ca²⁺]_i increase as a positive control. The maximum change in fluorescence ratio (ΔR_max, e) and area under the curve (AUC, f) were compared using a nested one‐way ANOVA with post‐hoc Tukey test for multiple comparisons; n = 3–4 independent experiments for each condition. *p < 0.05; **p < 0.01 compared with control. Data presented are nested values for individual cells grouped by experiment (gray dots) and experimental means ± SEM (black line and bars).

3.4. Depletion of ER Ca²⁺ stores inhibits the increase in [Ca²⁺]_i induced by 0Na ⁺/NMDG

Having ruled out Ca²⁺ influx as being responsible for the [Ca²⁺]_i transients induced following application of 0Na⁺/NMDG, we tested whether depletion of the intracellular Ca²⁺ stores within the endoplasmic reticulum affected the observed rises in [Ca²⁺]_i. To deplete endoplasmic reticulum Ca²⁺ stores, cells were treated with the sarcoplasmic/endoplasmic reticulum Ca²⁺ ATPase (SERCA) inhibitor cyclopiazonic acid (CPA, 30 μM) in nominal Ca²⁺‐free conditions (to prevent store‐operated Ca²⁺ entry), resulting in a transient leak of Ca²⁺ from the endoplasmic reticulum that was subsequently cleared from the cell (Figure 4b), presumably by the plasma membrane Ca²⁺ ATPase (James et al., 2015). These experiments were performed without EGTA since it was observed that inclusion of EGTA (1 mM) within the Ca²⁺‐free buffer inhibited the [Ca²⁺]_i increases elicited by 0Na⁺/NMDG (Supplementary Figure S2). Compared with control cells, (Figure 4a), pretreatment with CPA completely abolished the increase in [Ca²⁺]_i elicited by 0Na⁺/NMDG (Figure 4b–d; p < 0.01 for both ΔR_max and AUC). These results indicate that the 0Na⁺/NMDG‐induced [Ca²⁺]_i transient is likely due to release of Ca²⁺ from the intracellular endoplasmic reticulum Ca²⁺ stores.

FIGURE 4.

[Ca²⁺]_i transients induced by removal of extracellular Na⁺ in breast cancer cells require ER Ca²⁺ stores. MDA‐MB‐231 cells were loaded with fura‐2 AM (4 μM) and Ca²⁺ imaging performed during replacement of extracellular Na⁺ with equimolar (0Na⁺/NMDG) in the presence or absence of the SERCA inhibitor cyclopiazonic acid (CPA, 30 μM). Representative traces are shown for cells perfused without CPA (a) or where cells were pretreated for 20 min CPA prior to application of 0Na⁺/NMDG (b). The maximum change in fluorescence ratio (ΔR_max, c) and area under the curve (AUC, d) were compared using a nested t‐test for side‐by‐side comparisons; n = 4–6 independent experiments for each condition; **p < 0.01 compared with control. Data presented are nested values for individual cells grouped by experiment (gray circles) and experimental means ± SEM (black line and bars).

3.5. Na⁺ depletion drives increases in [Ca²⁺]_i via a G‐protein coupled receptor/IP₃ receptor‐mediated mechanism

Given the evidence implicating endoplasmic reticulum Ca²⁺ release in the changes in [Ca²⁺]_i observed upon application of 0Na⁺/NMDG, we next sought to determine the key molecular players responsible. Ca²⁺ is classically released from the endoplasmic reticulum via inositol (1,4,5) triphosphate receptors (IP₃Rs) and ryanodine receptors (RyRs; Berridge et al., 2000). To determine which of these release mechanisms was responsible for [Ca²⁺]_i transients upon application of 0Na⁺/NMDG, we perfused MDA‐MB‐231 cells with 0Na⁺/NMDG following pretreatment (10 min) with either the IP₃R inhibitor 2‐APB (50 μM) or the RyR inhibitor dantrolene (10 μM). Compared to control cells (Figure 5ai,bi), 2‐APB significantly reduced the [Ca²⁺]_i transient induced by 0Na^+/NMDG (Figure 5aii–aiv; ΔR_max, p < 0.05; AUC, p < 0.001), whereas dantrolene (10 μM, Figure 5bii) had no effect (Figure 5biii,biv). These data suggest that Na⁺ depletion induces Ca²⁺ release from the endoplasmic reticulum via IP₃Rs.

FIGURE 5.

[Ca²⁺]_i transients induced by removal of extracellular Na⁺ require activation of IP₃ receptor and phospholipase C, but are independent of ryanodine receptor activation. MDA‐MB‐231 cells were loaded with fura‐2 AM (4 μM) and Ca²⁺ imaging performed during replacement of extracellular Na⁺ with equimolar (0Na⁺/NMDG) in the absence or presence of the IP₃ receptor inhibitor 2‐APB (50 μM, ai and aii, respectively), the ryanodine receptor inhibitor dantrolene (10 μM, bi and bii, respectively), and the phospholipase C inhibitor U73122 (2 μM, ci and cii, respectively). For each condition, cells were pretreated with drug for 10 min prior to application of 0Na⁺/NMDG. Ionomycin (3 μM) was applied at the end of an experiment to elicit a [Ca⁺]_i increase as a positive control. The maximum change in fluorescence ratio (ΔR_max; aiii, 2‐APB; biii, dantrolene; ciii, U73122) and area under the curve (AUC; aiv, 2‐APB; biv, dantrolene; civ, U73122) were compared using a nested t‐test for side‐by‐side comparisons; n = 3–5 independent experiments for each condition; *p < 0.05; ***p < 0.001 compared with control. Data presented are nested values for individual cells grouped by experiment (gray dots) and experimental means ± SEM (black line and bars).

IP₃Rs are classically activated by upstream G‐protein coupled receptor signaling, with GPCRs of the G_αq/₁₁ family activating phospholipase C (PLC), which then cleaves membrane‐bound phosphatidylinositol 4,5‐bisphosphate (PIP2) to form the second messengers IP₃ and diacylglycerol (DAG; Patterson et al., 2005). To determine whether application of 0Na⁺/NMDG influenced IP₃Rs via GPCR‐mediated activation of PLC, we applied 0Na⁺/NMDG following pretreatment with the aminosteroid PLC inhibitor U73122 (2 μM, Yule & Williams, 1992). Compared with control cells (Figure 5ci), U73122 completely abolished the increase in [Ca²⁺]_i observed following application of 0Na⁺/NMDG (Figure 5cii–civ; p < 0.001 for both ΔRmax and AUC), indicating that PLC activation, potentially via GPCRs, is critical to this Na⁺‐controlled ER Ca²⁺ release mechanism.

4. DISCUSSION

The present study is the first to describe a relationship in metastatic breast cancer cells between Na⁺ depletion and a subsequent dramatic rise in [Ca²⁺]_i that is mediated by a PLC‐ and IP₃R‐dependent mechanism. These results provide further evidence of an interplay between Ca²⁺ and Na⁺ signaling in breast cancer cells that may have important implications for understanding cancer cell physiology. Through pharmacological maneuvers we rule out any role for reverse‐mode NCX activity, V_m and Ca²⁺ entry in this response. We also found that CPA abolished the [Ca²⁺]_i transients elicited by 0Na⁺/NMDG application; CPA is a well characterized SERCA blocker, providing further indirect evidence that this phenomenon was mediated by an IP₃R‐mediated endoplasmic reticulum Ca²⁺ release mechanism. Furthermore, while [Ca²⁺]_i transients were observed upon Na⁺ depletion in MDA‐MB‐231, MDA‐MB‐468, and SKBR3 cells, no such response was observed in MCF‐7 cells. This finding indicates that the Na⁺‐dependent Ca²⁺ increase is not a universal feature across all breast cancer cell lines. Further work is required to determine the underlying reason for the difference in Na⁺ and Ca²⁺ signaling between different cancer cell lines.

The present study establishes that Na⁺ regulates a PLC/IP₃‐dependent Ca²⁺ release in metastatic breast cancer cells; however, the molecular identity of the Na⁺ “sensor” that initiates this event remains unknown. Interestingly, Na⁺ has been described as an endogenous regulator of Class A GPCRs (White et al., 2018) via allosteric regulation of agonist binding (Agasid et al., 2021; Katritch et al., 2014; Zarzycka et al., 2019). While we cannot presently rule out whether it is the depletion of cytosolic [Na⁺] or removal of extracellular Na⁺ that induces this [Ca²⁺]_i transient in the current study, these previous studies provide a strong potential candidate for the underlying trigger mechanism. Indeed, in these studies, Na⁺ depletion appeared to increase basal GPCR activity in the absence of agonist, suggesting that Na⁺ may act as a negative modulator of GPCR activation (Agasid et al., 2021; Katritch et al., 2014; Zarzycka et al., 2019). Interestingly, recently obtained X‐ray structures of Class A GPCRs have identified a Na⁺ binding site in the vicinity of the agonist binding site that is exposed to the extracellular space (Yuan et al., 2013; Katritch et al., 2014). Moreover, computer simulations have suggested Class A GPCRs exhibit a transient water‐filled channel connecting this Na⁺ binding site to the cytoplasm (Hu et al., 2019; Vickery et al., 2018; Yuan et al., 2014, 2015). Na⁺ translocation via such a channel would be expected to be electrogenic, and thus impact upon V_m as well as Na⁺ handling (Katritch et al., 2014; Shalaeva et al., 2019. This has important implications for cancer cells exhibiting altered Na⁺ dynamics, since V_m is an important regulator of cancer cell behavior (Yang & Brackenbury, 2022); moreover, the altered [Na⁺]_i observed in breast tumors relative to healthy tissue (James et al., 2022) might be expected to disrupt GPCR regulation by Na⁺.

It is worthy to note that, in the present study, the PLC inhibitor U73122 had a more potent inhibitory effect on the [Ca²⁺]_i rise than the IP₃R inhibitor 2‐APB; the reasons for this are not clear, and suggest that U73122 is a more effective inhibitor of this pathway than 2‐APB. However, 2‐APB has been shown to target other channels, including certain TRP channel subtypes (Singh et al., 2018), which may have contributed to altered cation concentrations during these experiments. These findings also present a cautionary tale for cell physiologists using the methods employed in this study for exploring the relationship between Na⁺ conductance, Ca²⁺ signaling and wider cell behavior. Application of 0Na⁺/NMDG or 0Na⁺/ChoCl is widely utilized for isolating cell responses from the influence of extracellular Na⁺; however, these maneuvers are not inert (Thuma & Hooper, 2018). In addition to assessing for reverse mode NCX activity (Chovancova et al., 2020), Na⁺ regulation of GPCRs (and resultant effects on Ca²⁺ dynamics) should be taken into careful consideration and controlled for when using 0Na⁺/NMDG, 0Na⁺/ChoCl or similar maneuvers to probe Na⁺ conductance mechanisms.

Interestingly, inclusion of EGTA in the Ca²⁺‐free extracellular buffer abolished the [Ca²⁺]_i transients elicited by 0Na⁺/NMDG application, suggesting that EGTA exerts an inhibitory effect on the PLC and IP₃‐mediated Ca²⁺ release upon Na⁺ depletion. It has previously been shown that EGTA can inhibit mobilization of IP₃‐sensitive Ca²⁺ stores (Combettes & Champeil, 1994; Finch & Goldin, 1994) in cerebellar microsomes. However, EGTA is considered membrane impermeable, and thus would not be expected to exert a direct effect on IP₃Rs in the present study. This suggests that EGTA instead exerts its effects at the extracellular side of the plasma membrane via an at present undefined mechanism. Such a mechanism could be either by a direct effect on the putative GPCR involved or via chelation of extracellular divalent cations required for proper transmembrane protein function. Alternatively, it has been argued that extracellularly‐applied EGTA may be taken up into cells via highly Ca²⁺‐permeable cation channels (e.g., TRPA1, TRPV1, P2X7) and/or via fluid‐phase endocytosis and subsequent release into the cytosol (Liu et al., 2020). Further work is required to resolve these possibilities.

Beyond the present study, very little is known about the link between Ca²⁺ signaling and Na⁺ homeostasis in the context of breast cancer. Evidence indicates that Ca²⁺ oscillations in MDA‐MB‐231 cells are regulated by aberrantly expressed VGSCs (Rizaner et al., 2016). Moreover, elevations in extracellular Na⁺ cause p‐glycoprotein‐induced resistance to paclitaxel in breast cancer cells via Ca²⁺ signaling (Babaer et al., 2018). Of particular relevance to the current study (where Na⁺ depletion activated IP₃R‐dependent Ca²⁺ signaling), IP₃Rs are upregulated in breast cancer (Foulon et al., 2022), and estradiol‐induced IP₃R signaling regulates breast cancer cell proliferation (Szatkowski et al., 2010). Either pharmacological inhibition or knockdown of IP₃Rs in breast cancer cells led to dysregulated bioenergetics, and autophagy, cell cycle arrest, and cell death (Mound et al., 2013; Singh et al., 2017). Thus, the emerging evidence for altered Na⁺ and Ca²⁺ handling and their inextricable link via common channels and transporters hint at a hitherto relatively underexplored avenue in the context of cancer. The present study provides further evidence of such a link between Ca²⁺ and Na⁺ signaling that may have important implications for regulating cancer cell behavior and future therapy.

AUTHOR CONTRIBUTIONS

The project was designed by WJB and ADJ; experiments were carried out by ADJ, NS, KU, IJ, and RS with the assistance of GJOE; data analysis was performed by ADJ, NS, KU, IJ, and RS; the manuscript was prepared by ADJ and WJB; ADJ, WJB, GJOE, and SC contributed to study design, interpretation of the data, and critical revision of the paper for important intellectual content.

CONFLICT OF INTEREST

The authors declare that they have no competing interests.

ETHICS STATEMENT

No animals or human subjects were used in this study.

Supporting information

Supplementary Figure S1.

Supplementary Figure S2.

James, A. D. , Unthank, K. P. , Jones, I. , Sajjaboontawee, N. , Sizer, R. E. , Chawla, S. , Evans, G. J. O. , & Brackenbury, W. J. (2023). Sodium regulates PLC and IP₃R‐mediated calcium signaling in invasive breast cancer cells. Physiological Reports, 11, e15663. 10.14814/phy2.15663