pmc.ncbi.nlm.nih.gov

Reciprocal regulation of two G protein-coupled receptors sensing extracellular concentrations of Ca2+ and H+

Significance

The composition of the extracellular fluid surrounding all cells changes in an activity-dependent manner. Cell surface receptors allow cells to respond to components of the fluid, which is vital for proper functioning of cells and tissues. Ca²⁺ and H⁺ are crucial for cell survival and functioning. Their extracellular concentrations are monitored by two receptors, extracellular calcium-sensing receptor (CaSR) and ovarian cancer gene receptor 1 (OGR1), respectively. We report that these two receptors can be regulated in a seesaw manner; conditions favoring activity of one receptor inhibit signaling through the other, and vice versa, allowing cells to detect subtle changes in the extracellular concentration of these ions. We provide evidence that dysregulated activity of CaSR and OGR1 may contribute to the formation and progression of pathologies.

Keywords: OGR1, pH sensing, extracellular acidosis, CaSR, microenvironment

Abstract

G protein-coupled receptors (GPCRs) are cell surface receptors that detect a wide range of extracellular messengers and convey this information to the inside of cells. Extracellular calcium-sensing receptor (CaSR) and ovarian cancer gene receptor 1 (OGR1) are two GPCRs that sense extracellular Ca²⁺ and H⁺, respectively. These two ions are key components of the interstitial fluid, and their concentrations change in an activity-dependent manner. Importantly, the interstitial fluid forms part of the microenvironment that influences cell function in health and disease; however, the exact mechanisms through which changes in the microenvironment influence cell function remain largely unknown. We show that CaSR and OGR1 reciprocally inhibit signaling through each other in central neurons, and that this is lost in their transformed counterparts. Furthermore, strong intracellular acidification impairs CaSR function, but potentiates OGR1 function. Thus, CaSR and OGR1 activities can be regulated in a seesaw manner, whereby conditions promoting signaling through one receptor simultaneously inhibit signaling through the other receptor, potentiating the difference in their relative signaling activity. Our results provide insight into how small but consistent changes in the ionic microenvironment of cells can significantly alter the balance between two signaling pathways, which may contribute to disease progression.

Cells are surrounded by interstitial fluid, the composition of which is influenced by neighboring cells and which constitutes a key part of the microenvironment in which cells have to operate and survive. Changes in this microenvironment influence cell physiology (1, 2) and may promote disease (3, 4). Extracellular Ca²⁺ ([Ca²⁺]_o) and H⁺ ([H⁺]_o) concentrations are important components of the microenvironment, and their extracellular concentration changes in an activity- and state-dependent manner (5, 6). [Ca²⁺]_o is required for membrane stability, serves as a reservoir to allow Ca²⁺ influx into cells, and contributes to the membrane potential. [H⁺]_o sets the local pH, thereby influencing protein function as well as contributing to the membrane potential. Furthermore, Ca²⁺ and H⁺ can cross the membrane via ion channels and transporters, meaning that both can serve as intracellular and extracellular messengers (7–10).

Levels of [Ca²⁺]_o and [H⁺]_o are communicated to cells via cell surface receptors that change their activity in a manner dependent on [Ca²⁺]_o and [H⁺]_o. These receptors include G protein-coupled receptors (GPCRs), such as the extracellular Ca²⁺-sensing receptor (CaSR), ovarian cancer gene receptor 1 (OGR1), G protein-coupled receptor 4 (GPR4), and T-cell death-associated gene 8 (TDAG8), all of which sense [H⁺]_o, as well as a range of ion channels (8).

Intriguingly, Ca²⁺ and H⁺ signaling can be intimately linked, and changes in extracellular pH (pH_o) and intracellular pH (pH_i) may affect intracellular Ca²⁺ ([Ca²⁺]_i) signaling directly and indirectly (8, 11, 12). This is exemplified by OGR1, which, like CaSR (13), can couple to G_q and hence trigger Ca²⁺ release from intracellular Ca²⁺ stores via activation of the phospholipase C pathway (14). Neither CaSR nor OGR1 desensitizes (13, 14); thus, they continually monitor [Ca²⁺]_o and [H⁺]_o levels, respectively, and faithfully report any changes in their extracellular concentration. Because of the vital importance of Ca²⁺ and H⁺ to cells, information about their extracellular presence is crucial for cells, and lack of or altered signaling through these receptors may contribute to disease pathways.

We have previously found that OGR1 activation in DAOY cells, a human cerebellar granule cancer cell line, leads to complex [Ca²⁺]_i signals and activation of the ERK signaling pathway, thereby providing a mechanistic explanation of how the acidic environment may influence transformed cell function and/or survival (15). This action is lost on differentiation, suggesting a link between OGR1 activity and proliferative behavior of the transformed neurons (16). To better understand the role played by OGR1 in central neurons, we investigated OGR1 activation in primary cerebellar granule cells, the nontransformed equivalent of DAOY cells. We found that OGR1 and CaSR reciprocally inhibit [Ca²⁺]_i signaling through each other, and that intracellular acidosis, which accompanies extracellular acidification, promotes OGR1 but inhibits CaSR activity. Finally, CaSR-dependent inhibition of OGR1activity is absent in DAOY cells.

Results

We first established that OGR1 was expressed in primary wild-type (WT) murine cerebellar granule cell cultures throughout their culturing period [days in vitro (DIV) 2–15] using RT-PCR (Fig. 1A). We next carried out fluorescence Ca²⁺ imaging experiments to see whether extracellular acidosis could trigger changes in [Ca²⁺]_i concentration. We first dropped pH_o from 8 to 6 in the absence of [Ca²⁺]_o but in the presence of 2 mM extracellular Mg²⁺ ([Mg²⁺]_o) (Ca²⁺-free conditions). Increases in [Ca²⁺]_i under these conditions reflect Ca²⁺ release from intracellular Ca²⁺ stores, suggesting functional OGR1 expression.

Fig. 1. — CaSR inhibits extracellular acidification-mediated [Ca²⁺]_i signaling. (A) Murine cerebellar granule cells express OGR1 mRNA at different DIV stages. (B) Average fluorescence traces recorded in WT granule cells in response to extracellular acidification from pH 8 to pH 6 in the absence (0Ca0Mg) and presence of 2 mM Ca²⁺_o and Mg²⁺_o (2Ca2Mg), and in the additional presence of CaSR inhibitors 10 μM NPS2143 (2Ca2Mg+NPS2143) or NPS2390 (2Ca2Mg+NPS2390). n = 46–56 cells. (C) Average fluorescence traces in WT granule cells in response to extracellular acidification in the presence of extracellular divalents following knockdown of CaSR (shCaSR) and using a scrambled sh construct (shScrambled) as a negative control. n = 38–49 cells. (D) Representative raw traces showing the effects of extracellular acidification in presence of increasing [Ca²⁺]_o in WT granule cells. All experiments were performed in the absence of [Mg²⁺]_o. (E) Average peak fluorescence signals in response to acidification from pH_o8–6 for a given [Ca²⁺]_o were normalized to average peak fluorescence signal at 0 mM Ca²⁺_o (taken as 100%). Experimental conditions were as in D. Dotted lines indicate the position of 50% of the control fluorescence signal at 0 mM Ca²⁺_o. n = 85–92 cells. (F) Average time delay between extracellular acidification and peak response in presence of increasing [Ca²⁺]_o, measured in seconds. Same cells as for E. ***P < 0.0001.

Results in response to extracellular acidification under Ca²⁺-free conditions were very variable (Fig. S1); thus, we repeated the experiments in the presence of Ca²⁺_o to obtain more robust and reliable Ca²⁺ signals, as both Ca²⁺ release from stores and Ca²⁺ influx through plasma membrane channels should contribute to the overall fluorescence signal. However, extracellular acidification did not give rise to any change in [Ca²⁺]_i under these conditions (Fig. 1B), suggesting that the presence of Ca²⁺_o interfered with acidosis-mediated changes in [Ca²⁺]_i in granule cells.

We therefore considered that extracellular divalents might inhibit OGR1 signaling and repeated extracellular acidification experiments in the absence of [Ca²⁺]_o and [Mg²⁺]_o (in the additional presence of 0.1 mM EGTA and EDTA; divalent-free conditions). Under these conditions, we measured robust increases in [Ca²⁺]_i in all cells tested subsequent to extracellular acidification (Fig. 1B). Thus, [Ca²⁺]_o and [Mg²⁺]_o interfere with OGR1 signaling.

Both Ca²⁺ and Mg²⁺ are agonists of CaSR (13), and GPCRs have been documented to inhibit signaling through one another (17–19). Therefore, we considered the possibility that CaSR might inhibit OGR1 activity. To address this, we used two distinct pharmacologic inhibitors of CaSR, NPS2390 and NPS2143, and tested their impact on [Ca²⁺]_i signaling in the presence of [Ca²⁺]_o and [Mg²⁺]_o, when CaSR is active. There was a prominent rise in [Ca²⁺]_i on extracellular acidification in the presence of these antagonists that was observed in all cells and that was larger than the Ca²⁺ signal in the absence of extracellular divalents (Fig. 1B), suggesting that extracellular acidosis can evoke Ca²⁺ influx and Ca²⁺ release from intracellular Ca²⁺_i stores.

Importantly, in the absence of [Ca²⁺]_o and [Mg²⁺]_o, neither NPS2143 nor NPS2390 had any impact on the acidosis-mediated rise in [Ca²⁺]_i (Fig. S2), demonstrating selectivity of the inhibitors for CaSR.

To confirm a role for CaSR in inhibiting acidosis-mediated [Ca²⁺]_i signals, we used an shRNA internal ribosome entry site–red fluorescent protein (IRES-RFP) tag approach to knock down CaSR. In successfully transfected cells, extracellular acidosis triggered a rise in [Ca²⁺]_i in the presence of extracellular divalents (Fig. 1C) (extent of knockdown shown in Fig. S3 A and B). Thus, knockdown of CaSR led to disinhibition of acidosis-dependent [Ca²⁺]_i signaling in granule cells that was not observed in cells transfected with a control (scrambled) sh-construct (Fig. 1C). Fig. S3C lists the shScramble controls to demonstrate the specificity of the sh-CaSR approach.

Fig. S3. — Control experiments for CaSR knockdown in WT granule cells using an shRNA approach. To assess the extent of CaSR knockdown using the shRNA approach, we carried out experiments in which we compared CaSR protein expression using a CaSR antibody in cells that had been successfully transfected with shRNA against CaSR, a scrambled sh construct as a negative control, and untransfected cells (*SI Materials and Methods*). These experiments showed that shCaSR-transfection resulted in a 66.9 ± 1.7% knockdown of CaSR protein compared with untransfected cells and cells transfected with scrambled sh construct (Fig. S3B). In another set of experiments, we used an automatic cell sorter to separate nontransfected cells from shCaSR-transfected cells. Protein extraction from these two cell populations and subsequent Western blot analyses revealed that CaSR was knocked down by 66.8 ± 5.9% in shCaSR-transfected cells. (A) Bar charts depicting average relative CaSR protein expression in WT granule cells under control conditions (control; n = 54 cells) and following transfection of cells with shRNA targeted against CaSR (shCaSR; n = 31 cells) or with scrambled shRNA as a negative transfection control (sh Scrambled; n = 46 cells). Protein expression levels were normalized to control CaSR expression. Error bars represent SEM. The quantification method is described in *Materials and Methods*. (B) Bar charts depicting average relative CaSR protein expression in WT granule cells under control conditions and following transfection of cells with shRNA targeted against CaSR (shCaSR). Cells were sorted automatically on the basis of the fluorescent tag of the sh construct. Protein was isolated, and Western blot analyses were carried out with an antibody against CaSR and α-tubulin for quantification (n = 3 repeats). (*Left*) Average relative CaSR expression. (*Right*) Representative Western blot results. Molecular weights are given in kDa. Error bars represent SEM. Experimental procedures are described in *Materials and Methods*. (C) Transfection of murine WT granule cells with scrambled shRNA for CaSR (shScramble) does not lead to a disinhibition of acidosis-mediated [Ca²⁺]_i signals in these cells. Experiments serve as negative controls for experiments using shRNA against CaSR and were carried out in the absence of extracellular divalents (0Ca0Mg + shScramble; n = 30 cells), in the presence of extracellular divalents (2Ca2Mg + shScramble; n = 49 cells), and in the additional presence of 10 μM NPS2143 (2Ca2Mg + NPS2143 + shScramble; n = 48 cells). shScramble-expressing cells were identified with the help of their RFP signals.

CaSR is activated by [Ca²⁺]_o in the physiological range (0.1–1 mM) (13); therefore, we examined whether this was also the Ca²⁺ concentration range inhibiting acidosis-mediated Ca²⁺ signals. Under divalent-free conditions, extracellular acidification gave rise to a robust [Ca²⁺]_i signal, but already at 0.1 mM Ca²⁺, the peak Ca²⁺ signal was reduced, and full block was observed at 1 mM [Ca²⁺]_o (Fig. 1 D and E). A 50% block of acidosis-mediated [Ca²⁺]_i signals was achieved in the presence of 0.36 mM [Ca²⁺]_o (Fig. 1E). Moreover, there was a significant delay in the time to peak of the [Ca²⁺]_i signal (Fig. 1 D and F; P < 0.0001).

We next wanted to establish whether the rises in [Ca²⁺]_i observed in granule cells in response to extracellular acidosis were mediated by OGR1, or whether there might be a role for other acid-sensing proteins in this process, by establishing granule cell cultures from Ogr1 knockout (KO) mice (Ogr1^−/−). In these cells, extracellular acidification did not give rise to any changes in [Ca²⁺]_i even when CaSR activity was inhibited (Fig. 2A), demonstrating that the changes in [Ca²⁺]_i signaling observed in response to extracellular acidification in WT cells were not mediated by proteins other than OGR1.

Fig. 2. — OGR1 underlies the acidification-mediated [Ca²⁺]_i signaling. (A) Average fluorescence response to acidification from pH_o8–6 measured in *Ogr1*^−/− granule cells in the absence (0Ca0Mg), and presence of 2 mM Ca²⁺_o and Mg²⁺_o (2Ca2Mg), and in the additional presence of NPS2143 (10 μM; 2Ca2Mg+NPS2143) or NPS2390 (10 μM; 2Ca2Mg+NPS2390). n = 46–56 cells. (B) The same experiments as in A, but following transfection of murine *Ogr1* into *Ogr1*^−/− cells (*Ogr1*^−/− + mOGR1). n = 26–49 cells. (C) Representative raw traces showing the dose–response curve of OGR1 signaling to extracellular acidosis in absence of extracellular divalent cations in WT granule cells. (D) Experiments performed as in C. Average peak Ca²⁺ signals were plotted against the pH_o at which they occurred. All data were obtained in WT granule cells. n = 47–53 cells. Dotted lines indicate the pH_o at which the measured response is one-half that measured at pH 6.

To further prove this latter point, we introduce murine OGR1 (RFP-tagged) back into Ogr1^−/− cells. This resulted in acidosis-dependent Ca²⁺_i signals in successfully transfected cells (Fig. 2B) that exhibited the same CaSR-mediated suppression of OGR1 signaling observed in WT cells (Figs. 1B and 2B). Extracellular acidosis-induced changes in [Ca²⁺]_i were not observed in Ogr1^−/− cells transfected with an empty RFP vector control (Fig. S4).

Fig. S4. — Control experiments for OGR1 transfection into granule cells derived from *Ogr1*^−/− mice. Transfection of the empty RFP vector into granule cells derived from *Ogr1*^−/− mice was done to rule out that the possibility that the transfection procedure could give rise to responses resembling OGR1 responses. Experiments were carried out in the absence of extracellular divalents (0Ca0Mg + RFP; n = 29 cells), in the presence of extracellular divalents (2Ca2Mg + RFP; n = 25 cells), and in the additional presence of 10 μM NPS2390 (2Ca2Mg + NPS2390 + RFP; n = 30 cells). Only RFP-positive cells were used for analysis.

We next determined the pH dependence of OGR1 activation in WT cells (divalent-free conditions). Dropping pH_o from 8 to 7.35 did not produce any appreciable increase in [Ca²⁺]_i, but there was a significant rise in [Ca²⁺]_i at pH_o 6.8 and below (P < 0.0001; Fig. 2C). The peak [Ca²⁺]_i rises showed a clear dependence on pH_o (Fig. 2D). Assuming maximal activation of OGR1 at pH_o 6, half-maximal activation of OGR1 in WT cells was achieved when pH_o dropped to around 6.6 (Fig. 2D). Furthermore, the kinetics also strongly depended on pH_o (P < 0.0001; Fig. S5). Thus, OGR1 in cerebellar granule cells has a more acidic pH dependence than has been reported previously for some cells (14, 15), but not for others (20, 21).

Fig. S5. — Kinetics of OGR1 responses in response to various extents of extracellular acidification. Same cells and experimental procedures as in Fig. 2D. Shown is the average time to peak of the [Ca²⁺]_i signal in response to extracellular acidification to pH_o 6.8, 6.5, and 6, measured in seconds. ***P < 0.0001.

CaSR is inhibited by extracellular acidosis owing to pH effects on its agonist binding site (22); a role for OGR1 in influencing CaSR signaling was not considered in that study. Thus, we investigated whether OGR1 and CaSR could engage in reciprocal regulation by activating CaSR (increase in [Ca²⁺]_o from 0 to 2 mM in the absence of [Mg²⁺]_o) at different pH_o values in WT and Ogr1^−/− cells. There was a significant reduction in CaSR responsiveness with increasing extracellular acidification in both cell types (peak Ca²⁺ signal, B Ca²⁺ integral; P < 0.0001 for both; Fig. 3A). However, for almost every given pH_o value, the CaSR-mediated Ca²⁺ responses were smaller in WT cells than in Ogr1^−/− cells (P < 0.0002).

The foregoing finding could reflect increased CaSR expression in Ogr1^−/− cells compared with WT cells, thereby resulting in larger CaSR responses. Consequently, we compared CaSR protein expression levels in WT and Ogr1^−/− DIV2 and DIV15 granule cells and found no difference in CaSR expression levels (Fig. 3 C and D).

Furthermore, at pH_o 8, CaSR responses were identical in WT and Ogr1^−/− cells (Fig. 3 A and B). At this pH, OGR1 is not active (14), and WT cells should behave like Ogr1^−/− cells, which is what we observed. Thus, the reduced CaSR responses in WT cells compared with Ogr1^−/− cells is a likely consequence of OGR1 interfering with CaSR-mediated [Ca²⁺]_i signaling in WT cells.

Our data also show that the impact of OGR1 on CaSR-dependent signaling is not restricted to influencing peak Ca²⁺ signals. CaSR-mediated Ca²⁺ influx in WT was smaller than that in Ogr1^−/− cells; this was particularly evident in the integral Ca²⁺ response at pH_o 6.8 and below (Fig. S6).

Fig. S6. — OGR1 inhibits Ca²⁺ influx on CaSR activation. Shown are average fluorescence traces for experiments in WT and *Ogr1*^−/− cerebellar granule cells on increasing [Ca²⁺]_o from 0 to 2 mM in the absence of [Mg²⁺]_o (arrow) at pH_o 6.8, 6.5, and 6. Scale bars are identical for all three graphs; error bars represent SEM. Same cells as used for bar charts for pH 6, 6.5, and 6.8 in Fig. 3 A and B; n = 38–71 cells.

Extracellular acidification also may lead to (transient) intracellular acidification (15). To investigate whether this could contribute to the inhibition of CaSR-mediated signaling following extracellular acidification, we looked at pH_i changes in response to pH_o changes using fluorescence H⁺ imaging with BCECF [2′,7′-Bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein] as the H⁺ dye. First, cells were exposed to different pH_o conditions for 5 min, and then pH_i was measured. A clear dependence of pH_i on pH_o was seen; the more acidic the pH_o, the more acidic the pH_i, albeit to a lesser extent (Fig. S7A).

Fig. S7. — Control experiments for intracellular acidification. (A) pH_i depending on pH_o. pH_i was measured using BCECF following an pH_o change (5 min after changing pH_o) for 2 min in WT granule cells, recording fluorescence ratios every 5 s. Each data point represents the average of n = 33–59 cells; error bars represent SEM. (B) pH_i following application of 0, 12.5, 25, or 50 mM NaAc at pH_o 8; pH_i was measured using BCECF (with fluorescence ratios recorded every 5 s) in WT granule cells, and the peak decrease in pH_i was used for calculation of average pH_i in response to NaAc application. Each data point represents the average of n = 19–33 cells; error bars represent SEM. (C) Comparison of average pH_i following acidification of pH_o to pH 6 or application of 25 and 50 mM NaAc, respectively (data derived from A and B; n = 35–22 cells/condition). The decreases in pH_i are not statistically significantly different among the three conditions (P = 0.4836, ANOVA). (D) pH_i change in response to acidification of pH_o in WT and *Ogr1*^−/− granule cells. n = 33–59. Change in pH_i was assessed using the fluorescent H⁺ dye BCECF (details in *Materials and Methods*).

To study impact of intracellular acidosis on CaSR function, we induced intracellular acidification in the absence of extracellular acidification using different concentrations of extracellular sodium acetate ([NaAc]_o) (15, 23) (Fig. S7B). These experiments showed that 25 and 50 mM [NaAc]_o produced the same intracellular acidification as an extracellular acidification to pH 6 (Fig. S7C). Thus, the use of 25 or 50 mM NaAc mimics intracellular acidification in response to extracellular acidification to pH 6.

Finally, we determined whether WT and Ogr1^−/− granule cells showed the same changes in pH_i in response to extracellular acidification. OGR1 reportedly affects pH_i regulation (24), and lack of OGR1 expression in Ogr1^−/− cells might affect the cells’ ability to handle [H⁺]_i, resulting in variations in intracellular acidification. This could then affect CaSR function, thereby explaining some of the disparities in CaSR responses observed between WT and Ogr1^−/− cells.

Consequently, we compared levels of intracellular acidification to acute extracellular acidification in WT and Ogr1^−/− cells and found no differences (Fig. S7D). Thus, differences in extent of inhibition of CaSR by extracellular acidosis in WT cells vs. Ogr1^−/− cells are unlikely to result from varying degrees of intracellular acidification.

We next studied the impact of intracellular acidosis on CaSR-dependent Ca²⁺ signaling in WT cells to assess the possible contribution of this to the decreased CaSR responses measured on extracellular acidification. We studied CaSR activation in response to increasing [Ca²⁺]_o from 0 to 2 mM under control conditions (at pH_o 8) and under conditions of intracellular acidification (+ 25 mM NaAc, at pH_o 8). There was a clear change in the time course of the CaSR response under conditions of intracellular acidosis (Fig. 4A); the response was slower to develop and displayed a smaller peak and plateau phase. Thus, intracellular acidification interferes with CaSR-dependent signaling independent of extracellular acidosis. CaSR is not the only GPCR affected by intracellular acidosis, however; we also found inhibited signaling ability of metabotropic ATP receptors P2Y1 and 6 under conditions of intracellular acidosis (Fig. S8).

Fig. 4. — Intracellular acidosis inhibits CaSR and potentiates OGR1. (A) Average (± SEM) CaSR fluorescence responses at pH_o8 under control conditions (black) and following intracellular acidification with 25 mM NaAc (purple). n = 30–37 cells. Responses were evoked by increasing [Ca²⁺]_o from 0 to 2 mM (in the absence of [Mg²⁺]_o). (B) Average graph showing the impact of intracellular acidification by 50 mM NaAc at extracellular pH 7.35 in the presence of extracellular divalents and 10 μM NPS2390 in WT cells (n = 52). (C and D) Same experimental protocol as for B, but either at pH_o 8 in WT granule cells (n = 27) (C) or using *Ogr1*^−/−granule cells (n = 27) (D). (E) Black, pH_o 8 acidified to pH_o 6 after 150 s. Purple, pH_o 8 constant and pH_i acidified with 50 mM sodium acetate after 50 s. Green, pH_i acidified with 50 mM sodium acetate after 50 s (pH_o 8), and pH_o acidified to 6 after 150 s. n = 66–74 cells. (F) Same experiments as in E, but carried out in *Ogr1*^−/− granule cells. n = 31–45 cells.

Fig. S8. — Metabotropic ATP receptors P2Y1R and P2Y6R are affected by intracellular acidification in their signaling ability. Bar charts show the effect of intracellular acidification on average peak P2Y1R- and P2Y6R-activated Ca²⁺ release from intracellular Ca²⁺ stores. Error bars represent SEM. Black hatched bars represent control conditions (pH_o 8, in the absence of [Ca²⁺]_o); purple hatched bars are in the additional presence of 25 mM sodium acetate (+NaAc). P2Y1 receptors were activated using 1 μM MRS 2354, and P2Y6 receptors were activated using 1 μM MRS 2693; n = 151–161 cells. The observed differences were not quite statistically significant (P = 0.05979 for P2Y1; P = 0.0882 for P2Y6).

Taken together, our findings indicate that neither OGR1-dependent inhibition of CaSR, nor intracellular acidification can fully explain the large extent of inhibition of CaSR observed at pH_o 6. This suggests that extracellular acidosis inhibits CaSR by changing its agonist responsiveness (22), via activation of OGR1, and by causing intracellular acidosis.

We next wanted to establish whether or not OGR1 is also subject to modulation of its signaling ability by intracellular acidosis. The idea is that OGR1 should not be inhibited by intracellular acidification, given that this can accompany extracellular acidosis, which activates OGR1. Therefore, we considered that intracellular acidification might in fact promote OGR1 signaling.

As shown in Fig. 2C, a drop in pH_o from 8 to 7.35 did not result in any consistent Ca²⁺ responses in these cells, despite the fact that OGR1 is partially active at pH_o 7.35 (14). Thus, to investigate the impact of intracellular acidification on OGR1, we carried out experiments (under divalent-free conditions) in WT cells at pH_o 7.35, where OGR1 is partially active, in which we acidified pH_i using NaAc and monitored [Ca²⁺]_i. The idea was that if OGR1 were potentiated at this permissive pH_o, then we should see changes in [Ca²⁺]_i in response to intracellular acidification owing to increased OGR1 activity.

In these experiments, we indeed saw a distinct rise in [Ca²⁺]_i in WT cells on intracellular acidification at pH_o 7.35 (Fig. 4B) that was not observed at pH_o 8, where OGR1 is not active (Fig. 4C). Furthermore, there was no rise in [Ca²⁺]_i in Ogr1^−/− cells in response to intracellular acidification at pH_o 7.35 (Fig. 4D). This demonstrates that the [Ca²⁺]_i rise observed in WT cells at pH_o 7.35 upon intracellular acidification was related to increased OGR1 activity.

Crucially, the foregoing results confirm that intracellular acidosis inhibits CaSR signaling by acting on CaSR directly. The smaller Ca²⁺ rises in response to CaSR activation under conditions of intracellular acidification could have been the result of pH_i effects on the signaling cascades downstream of G_q activation; however, if this were the case, then OGR1 signaling should be equally impaired by intracellular acidosis. Our results show that the opposite is the case, suggesting that intracellular acidosis affects CaSR and OGR1 directly.

Given that intracellular acidosis can inhibit CaSR and promote OGR1 function, and CaSR inhibits signaling through OGR1, we wondered whether intracellular acidic preconditioning might alleviate CaSR-mediated inhibition of OGR1 function. Reduced CaSR activity and/or increased OGR1 signaling ability might permit OGR1 to signal even when CaSR is active. To address this question, we exposed WT cells (Fig. 4E) and Ogr1^−/− cells (Fig. 4F) to (i) a pH_o change from 8 to 6 only, (ii) intracellular acidification at constant pH_o 8 (using NaAc) only, and (iii) intracellular acidification followed by extracellular acidification. All experiments were carried out in the presence of extracellular divalents, to activate CaSR.

On acidification of pH_o only, there was no obvious change in [Ca²⁺]_i in WT or Ogr1^−/− cells (Fig. 4 E and F, black). Following intracellular acidification only, there was a small rise in [Ca²⁺]_i (Fig. 4 E and F, purple), which was also present in previous experiments (Fig. 4 C and D) and was independent of OGR1, given that it was also seen at pH_o 8 and in Ogr1^−/− cells. However, when intracellular acidification preceded extracellular acidification, we observed a further clear increase in [Ca²⁺]_i in WT cells, but not in Ogr1^−/− cells (Fig. 4 E and F, green). This demonstrates that intracellular acidosis can increase OGR1 function by reducing CaSR and/or promoting OGR1 signaling ability.

Our original research into OGR1 was carried out in DAOY cells, in which H⁺-induced currents in response to OGR1 activation were recorded in the presence of extracellular divalents (15), suggesting that OGR1 is not subject to inhibition by CaSR in these cells. To confirm this, we carried out fluorescence Ca²⁺ imaging experiments in DAOY cells in the presence of [Ca²⁺]_o and [Mg²⁺]_o and found that under these conditions, a drop in pH_o to 6 did indeed trigger a rise in [Ca²⁺ ]_i in virtually all cells tested (Fig. 5A), indicating OGR1 activation. These results suggest that CaSR does not interfere with OGR1 in DAOY cells, whether through lack of functional expression or through any other (additional) mechanism.

Fig. 5. — Transformed granule cells lack CaSR-mediated inhibition of OGR1-dependent [Ca²⁺]_i signals and reduced CaSR signaling activity. (A) Average data (±SEM) for OGR1-mediated intracellular fluorescence change in DAOY cells (n = 30 cells) in response to extracellular acidification from pH 7.35–6; experiments in the presence of divalents. (B) Average graphs (± SEM) showing CaSR responses in DAOY (blue; n = 46) and WT granule cells (black; n = 67). All experiments were done in the absence of [Mg²⁺]_o.

We then examined functional CaSR expression in DAOY cells by increasing [Ca²⁺]_o from 0 to 2 mM (in the absence of [Mg²⁺]_o) at pH_o 8 (to prevent a potential impact of OGR1 on CaSR). Following this protocol, we observed a small and slow rise in [Ca²⁺]_i in response to CaSR activation that was much smaller than the Ca²⁺ rise seen in primary WT granule cells subjected to the same experimental procedure (Fig. 5B). Thus, DAOY cells do express functional CaSRs, but their level of activity is lower than that in their nontransformed counterparts.

Discussion

We show that in central neurons, two GPCRs (CaSR and OGR1) that sense extracellular concentrations of physiologically relevant ions (Ca²⁺ and H⁺, respectively) can control each other’s signaling activity. Thus, the activity of one receptor may profoundly influence the signaling ability of the other receptor. We also show that intracellular acidification, which may accompany extracellular acidosis, inhibits CaSR responses but potentiates OGR1 responses. The impact of intracellular acidosis on signaling ability is not limited to CaSR and OGR1; other receptors are affected by it as well.

Furthermore, our experimental conditions do not allow for cells to regulate their pH_i via the bicarbonate/CO₂ buffering system. The presence of this physiologically relevant buffering system would limit the extent of intracellular acidification in terms of spread, duration, and severity. Therefore, the observed opposite regulation of CaSR and OGR1 in response to intracellular acidification will be less prominent in the presence of the bicarbonate/CO₂ buffering system than in its absence.

The seesaw manner of CaSR and OGR1 regulation, whereby conditions promoting the activity of one receptor directly and indirectly inhibit that of the other receptor, may allow cells to monitor and respond to changes in environmental ion composition with exquisite sensitivity. The delicate reciprocity between CaSR and OGR1 signaling could explain why subtle changes in microdomain ion concentrations can disrupt intracellular signaling sufficiently to promote the development and amplification of pathological signaling pathways. Notably, CaSR activity is determined not only by the availability of its ligands (with Ca²⁺ the main physiological agonist), but also by its cell surface expression. Changes in this will lead to a reduction in CaSR-mediated signaling even in the absence of changes in [Ca²⁺]_o (25). Therefore, increased OGR1 activity may occur even when [Ca²⁺]_o is constant and/or at physiological levels.

Global [Ca²⁺]_o and [H⁺]_o are thought to be relatively stable, but their levels fluctuate in response to ion channel, transporter, metabolic, and functional activity in the cellular microenvironment (5, 6, 13). This is particularly true for the brain: Opening of Ca²⁺-permeable ion channels during synaptic transmission leads to a temporary decease in local [Ca²⁺]_o (13, 26), whereas release of the neurotransmitter vesicular content into the synaptic cleft causes a decrease in pH_o (27). Furthermore, opening of GABA_A channels, which are permeable to both Cl⁻ and HCO₃⁻, may contribute to fluctuations in pH (28, 29). Because neither receptor desensitizes, they continuously communicate changes in [Ca²⁺] and [H⁺] to the cells in which they are expressed. Thus, the relative activity of these two receptors could be used by neurons and other cell types as a dynamic readout of the precise composition of the extracellular ionic milieu.

DAOY cells provide a model of transformed, malignant granule cells. We demonstrate that OGR1- and CaSR-dependent [Ca²⁺]_i signaling is altered in these cells. In fact, [Ca²⁺]_i signaling through OGR1 and CaSR appears to be opposite in the two cell types; in normal granule cells, OGR1-dependent [Ca²⁺]_i signaling is slow and small, whereas CaSR-dependent [Ca²⁺]_i signaling is fast and large, whereas the opposite is seen for DAOY cells. The lack of inhibition of OGR1 signaling by CaSR in DAOY cells is likely due, at least in part, to low functional expression levels of CaSR in these cells, given that knockdown of CaSR in granule cells also leads to disinhibition of OGR1 signaling.

Expression of OGR1 in the brain has been reported (30), but the spatiotemporal expression profile has not been well characterized. Much more is known about CaSR expression and function in the brain. CaSR is expressed in a number of distinct cell types throughout the brain, including neurons (31), and is thought to play a key role in development of the brain, synaptic transmission, and plasticity (32). Moreover, it has been implicated in brain pathologies, such as ischemia, neurodegenerative disease, and brain tumors (32). Intriguingly, these conditions are accompanied by extracellular acidosis (8, 33–36). OGR1 is functionally expressed in brain tumor cells (15), and changes in its activity levels may be relevant in other acidosis-accompanied pathological states. Thus, both OGR1 and CaSR have been implicated in brain disorders that are exacerbated by changes in [H⁺]_o and [Ca²⁺]_o (37). Dysregulation of the balance between OGR1 and CaSR signaling may contribute to the development and progression of wide-ranging pathological states.

CaSR and OGR1 also are coexpressed in a number of other tissues, including kidney (24, 30, 38), bone (13, 14, 20, 39, 40), and lung (30, 41–44). These tissues also experience extracellular acidification under physiological conditions. Intriguingly, both receptors have been implicated in diseases arising in these tissues (39–46), suggesting that altered signaling through CaSR and OGR1 may have a significant impact on disease progression in tissues other than the brain.

Materials and Methods

Cell Cultures.

The cerebellar granule cell experiments were carried out using granule cell cultures derived from C57BL/6 WT mice (Charles River Laboratories) and Ogr1 (C57BL/6 background) KO mice and cultured for up to 15 d (47). The Ogr1 KO mice were a generous gift from K. Seuwen and T. Suply (Novartis). DAOY cells (American Type Culture Collection) were grown and cultured as described previously (15).

Fluorescence Imaging Experiments.

The fluorescence Ca²⁺ imaging experiments and solutions have been described previously (15). All solutions were made using HPLC-grade water. Fura 2-AM and BCECF-AM were purchased from Molecular Probes-Invitrogen. Fluorescence ratios were recorded every 1 s for Fura 2-AM (340 nm/380 nm) and every 5 s for BCECF-AM (490 nm/439 nm); emission was measured at 535 nm. Preincubation was done for 30 min at room temperature with BCECF-AM (10 μM), and for 45 min at room temperature with Fura 2-AM in standard extracellular buffer at pH 7.35. The 490 nm/439 nm ratio was converted to a pH value using a calibration curve, obtained by measuring the fluorescence ratio in cells incubated in a high-K⁺ solution at a pH range of 6.0–8.0 (four values) supplemented with nigericin (2 μM; Sigma-Aldrich) (48). Experimental conditions are described in detail in SI Materials and Methods. General chemicals for making solutions were obtained from Sigma-Aldrich.

RNA Extraction and RT-PCR.

Total RNA from cultured mouse cerebellar granule cells and DAOY cells were extracted using a Qiagen RNeasy MiniKit according to the manufacturer’s protocol. For RT-PCR, first-strand cDNA was synthesized from 1 μg of total RNA with an oligo-dT primer and the Moloney murine leukemia virus reverse transcriptase (Promega) according to the manufacturer’s protocol. PCR reactions were optimized to 95 °C for 5 min, 30 amplification cycles for OGR1, and 20 amplification cycles for GAPDH at 95 °C for 30 s, 56 °C for 30 s, 72 °C for 30 s, and a final extension of 5 min at 72 °C. Primer sequences are presented in SI Materials and Methods.

Plasmids and Transfection.

Plasmids containing the RFP-tagged full-length cDNA of murine OGR1, CaSR shRNA, or control scrambled shRNA (Origene) were transfected into cerebellar granule cells using the Amaxa Nucleofector 2b electroporation system (Lonza) according to the manufacturer’s instructions. Cells were used at 48 h after transfection. Only transfected cells, selected on the basis of their RFP-dependent fluorescence properties, were used for experiments.

Analysis and Data Presentation.

Fluorescence traces were analyzed offline using Igor Wavemetrics 3.14. Data are shown as average ± SEM. InStat 2.03 for Mackintosh was used for statistical analysis. ANOVA was performed for comparison of more than two averages, and the unpaired Student t test was used for comparison of two averages. For Figs. 1E and 2D, data were fitted with a sigmoidal dose–response equation using GraphPad Prism. All experiments were carried out on at least two separate preparations.

SI Materials and Methods

Fluorescence Imaging Experiments.

Standard extracellular buffer contained 145 mM NaCl, 10 mM Hepes, 2.8 mM KCl, 2 mM CaCl₂, and 2 mM MgCl₂ (pH 7.35 or pH 8 with NaOH), supplemented with 10 glucose on the day of the experiments. For divalent-free conditions, MgCl₂ and CaCl₂ were omitted, and 0.1 mM EDTA and 0.1 mM EGTA were added. For Ca²⁺-free conditions, CaCl₂ was omitted, and 0.1 mM EGTA was added to the buffer (Fig. S1). To acidify pH_o during an imaging experiment, an extracellular buffer was produced with identical ionic composition to the external buffer used in the experiments with pH 3 (using HCl). Imaging experiments started in a given volume of extracellular buffer at pH 8, and pH was changed by adding a predetermined volume of pH 3 extracellular buffer to the the imaging chamber. The ratio of pH 8 to pH 3 buffer needed to change pH_o to the desired value (from pH 8 to pH 7.35, 7.1, 6.8, 6.5, or 6) was determined each time that new buffers were made and confirmed at least three times. Random checks were performed to rule out time-dependent changes in ratios. Buffers were freshly made at least twice a month, and the pH of the different buffers (pH 7.35, 8, and 3) remained stable throughout their time of use.

For OGR1 activation, cells at DIV10–15 were used for experiments. Extracellular pH was changed from pH 8 to test pH (pH 7.35/7.1/6.8/6.5/6) after 100 s, and recordings were continued for another 500 s. Experiments in the absence of [Ca²⁺]_o and/or [Mg²⁺]_o were carried out in the additional presence of 0.1 mM EGTA and/or EDTA. For CaSR activation in WT and Ogr1 KO-derived granule cells, cells were at DIV2–6. The [Ca²⁺]_o concentration was increased from Ca²⁺-free to 2 mM Ca²⁺ (always in the absence of [Mg²⁺]_o and with 0.1 mM EDTA added) after 100 s, and recordings were continued for another 500 s.

For CaSR activation, [Ca²⁺]_o was changed from Ca²⁺-free (+ 0.1 mM EGTA) to 2 mM Ca²⁺; all of these experiments were also performed in the absence of [Mg²⁺]_o (+ 0.1 mM EDTA). Because changes in [Ca²⁺]_o also may affect background Ca²⁺ flux across the membrane, experiments were repeated in the presence of 10 μM NPS 2390 (Figs. 3 and 4) or NPS2143 (Fig. 5) to identify the CaSR-independent proportion of the fluorescence Ca²⁺ signal under our experimental conditions. The averaged fluorescence response in the presence of CaSR inhibitor was then subtracted from the average fluorescence response in the absence of CaSR inhibitor, resulting in a wave that reflects CaSR response only. Data obtained after correcting for background Ca²⁺ influx were used for figures. NPS2143 and NPS2390 were obtained from R&D Systems. They were dissolved in DMSO (10 mM stock solution; aliquots frozen at −20 °C) and added to extracellular solution at a 1,000-fold dilution, yielding a final DMSO concentration of 0.1%.

To determine CaSR activity under conditions of extracellular acidosis, cells were exposed to Fura 2-AM and recovered in the presence of extracellular divalents, as described above. Cells were then transferred into extracellular solutions containing no divalents (but instead 0.1 mM EGTA and 0.1 mM EDTA) that was buffered at the desired pH value (pH 8, 7.35, 7.1, 6.8, 6.5, or 6). Cells were then immediately transferred to the imaging rig, and experiments were carried out as described above. The maximum time lapse between exposing cells to the desired pH_o (in the absence of divalents) and activation of CaSR by the addition of [Ca²⁺]_o was 5 min. To assess the impact of pH_o on pH_i under these conditions, we repeated the experiments but this time using BCECF as a fluorescent dye to monitor pH_i. Following a 5-min wait after exposing cells to the divalent-free solution at the different pH_o conditions, pH_i was measured for a period of 2 min. The average BCECF signal of those measurements was then used to calculate the average pH_i in dependence of pH_o (Fig. S3A).

To evaluate the effects of intracellular acidification on CaSR and OGR1 function, we carried out experiments in which pH_i was acidified by replacing extracellular NaCl with sodium acetate (NaAc; 12 mM, 25 mM, or 50 mM) and the time course and extent of intracellular acidification with BCECF-AM were measured. This was compared with the extent of intracellular acidification observed on extracellular acidification. The results showed that replacement of 25 or 50 mM extracellular NaCl yielded the same extent of intracellular acidification. The difference in pH_i was 0.37 ± 0.08 (n = 34) for 25 mM NaAc and 0.3 ± 0.17 (n = 22) for 50 mM NaAc (Fig. S3B). This decrease in pH_i was not different from that observed when pH_o was dropped to pH 6.5 or 6.0. The difference in pH_i was 0.44 ± 0.1 (n = 59) for pH_o 6.5 and 0.49 ± 0.1 (n = 36) for pH_o 6.0. These experiments also established the time interval between adding NaAc and pH_i acidification to the same extent as following extracellular acidification, which informed the time point in the CaSR and OGR1 experiments at which [Ca²⁺]_o was increased or pH was dropped to ensure that intracellular acidification had reached the required extent.

Primer Sequences.

For OGR1, forward (5′-CACCCTGAAGGCAGCCGTGG-3′) and reverse (5′-TGCGGCTCTTCTGTGTGCCG-3′) primers were used to amplify a 269-bp fragment from mouse OGR1. For GAPDH, forward (5′-GTGCAGTGCCAGCCTCGTCC-3′) and reverse (5′-TTCAAGTGGGCCCCGGCCTT-3′) primers were used to amplify a 362-bp fragment from mouse GAPDH.

Quantification of shCaSR Knockdown Efficiency Using a Cellular Approach.

Plasmids containing the RFP-tagged CaSR shRNA or control scrambled were transfected into cerebellar granule cells, and at 48 h after transfection, cells were fixed with 4% (wt/vol) paraformaldehyde for 10 min and then permeabilized with 0.5% Triton X-100 for 10 min. After washing with PBS, the cells were incubated with blocking solution (Thermo Scientific) for 1 h. The cells were then incubated with specific primary antibody against CaSR (Abcam) overnight at 4 °C. After washing with PBS, the cells were stained with Alexa Fluor 488-conjugated secondary antibody against mouse IgG (Invitrogen) at room temperature for 1 h. Nuclei were stained with DAPI (Invitrogen).

Images were obtained using an inverted confocal fluorescence microscope (Fluoview FV1000; Olympus), and fluorescence intensities were compared for untransfected cells (control cells) successfully transfected with shCaSR and cells successfully transfected with scrambled sh construct. Fluorescence intensity quantifications were performed with ImageJ software.

Quantification of shCaSR Knockdown Efficiency Using a Cell Population Approach.

Plasmids containing the RFP-tagged CaSR shRNA or control scrambled shRNA (Origene) were transfected into cerebellar granule cells using the Amaxa Nucleofector electroporation system (Lonza) according to the manufacturer’s instructions. At 48 h after transfection, the cells were trypsinized and resuspended in culture medium. Cells positive for RFP were sorted using a MoFlo XDP high-speed cell sorter (Beckman Coulter). Nontransfected cells were used to determine control CaSR protein levels.

For protein extraction, cells were rinsed three times with ice-cold PBS. Whole-cell extracts were harvested in RIPA lysis buffer (Sigma-Aldrich) supplemented with 100 mM sodium orthovanadate (Na₃VO₄), 100 mM phenylmethanesulfonyl fluoride (PMSF), and protease inhibitor mixture (Roche). Protein concentrations were determined using a DC protein assay (Bio-Rad), and 30 mg of protein lysate was fractionated by SDS/PAGE and transferred onto nitrocellulose membranes (Amersham Pharmacia Biotech) using a Trans-Blot SD semidry electrophoretic transfer cell (Bio-Rad). After incubation with 5% nonfat milk in PBST for 1 h at room temperature, membranes were washed once with PBST and then incubated with antibodies against CaSR (Abcam) or α-tubulin (Santa Cruz Biotechnology) at 4 °C for 20 h. Membranes were then washed with PBST three times for 10 min, incubated with HRP-conjugated anti-mouse IgG antibodies (Jackson ImmunoResearch) for 1 h at room temperature, and then washed three more times with PBST for 10 min. Proteins of interest were visualized using an enhanced chemiluminescence system (Amersham Pharmacia Biotech) according to the manufacturer’s protocol. Fluorescence intensity quantifications were performed using ImageJ software. Data were normalized to α-tubulin levels. The same procedure was adopted for quantification of CaSR expression in WT and Ogr1^−/−-derived granule cells at DIV2 and DIV15 (Fig. 3C).

Acknowledgments

We thank Drs. Klaus Seuwen and Thomas Suply for the Ogr1 KO mouse and Professors Gero Miesenböck, Anant Parekh, and Robert Wilkins for helpful comments on this manuscript. This work was supported by Biotechnology and Biological Sciences Research Council Grant BB/1008748/1. E.B. is the recipient of a Research Fellowship from the Royal Society.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

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Reciprocal regulation of two G protein-coupled receptors sensing extracellular concentrations of Ca2+ and H+

Significance

Abstract

Results

Fig. 1.

Fig. S1.

Fig. S2.

Fig. S3.

Fig. 2.

Fig. S4.

Fig. S5.

Fig. 3.

Fig. S6.

Fig. S7.

Fig. 4.

Fig. S8.

Fig. 5.

Discussion

Materials and Methods

Cell Cultures.

Fluorescence Imaging Experiments.

RNA Extraction and RT-PCR.

Plasmids and Transfection.

Analysis and Data Presentation.

SI Materials and Methods

Fluorescence Imaging Experiments.

Primer Sequences.

Quantification of shCaSR Knockdown Efficiency Using a Cellular Approach.

Quantification of shCaSR Knockdown Efficiency Using a Cell Population Approach.

Acknowledgments

Footnotes

References