Phosphate (P_i)-regulated heterodimerization of the high-affinity sodium-dependent P_i transporters PiT1/Slc20a1 and PiT2/Slc20a2 underlies extracellular P_i sensing independently of P_i uptake

Abstract

Extracellular phosphate (P_i) can act as a signaling molecule that directly alters gene expression and cellular physiology. The ability of cells or organisms to detect changes in extracellular P_i levels implies the existence of a P_i-sensing mechanism that signals to the body or individual cell. However, unlike in prokaryotes, yeasts, and plants, the molecular players involved in P_i sensing in mammals remain unknown. In this study, we investigated the involvement of the high-affinity, sodium-dependent P_i transporters PiT1 and PiT2 in mediating P_i signaling in skeletal cells. We found that deletion of PiT1 or PiT2 blunted the P_i-dependent ERK1/2-mediated phosphorylation and subsequent gene up-regulation of the mineralization inhibitors matrix Gla protein and osteopontin. This result suggested that both PiTs are necessary for P_i signaling. Moreover, the ERK1/2 phosphorylation could be rescued by overexpressing P_i transport–deficient PiT mutants. Using cross-linking and bioluminescence resonance energy transfer approaches, we found that PiT1 and PiT2 form high-abundance homodimers and P_i-regulated low-abundance heterodimers. Interestingly, in the absence of sodium-dependent P_i transport activity, the PiT1-PiT2 heterodimerization was still regulated by extracellular P_i levels. Of note, when two putative P_i-binding residues, Ser-128 (in PiT1) and Ser-113 (in PiT2), were substituted with alanine, the PiT1-PiT2 heterodimerization was no longer regulated by extracellular P_i. These observations suggested that P_i binding rather than P_i uptake may be the key factor in mediating P_i signaling through the PiT proteins. Taken together, these results demonstrate that P_i-regulated PiT1-PiT2 heterodimerization mediates P_i sensing independently of P_i uptake.

Introduction

Phosphorus is the sixth most abundant element in the human body, constituting ∼1% of total body weight (1). About 85% of total phosphate can be found in the skeleton, where it is a major constituent of hydroxyapatite crystals deposited on the extracellular organic matrix during the mineralization process. The remaining 15% is found mainly in cells from soft tissues and in extracellular volume, where it represents <1% of total phosphate (2–4). In plasma, ∼16% of circulating phosphate is found as organic phosphate bound to proteins and lipids, whereas the major part (84%) is orthophosphate, or free inorganic phosphate (P_i),⁵ that can be filtered by the kidney (1). At physiological pH, the monovalent H₂PO₄⁻ and the divalent HPO₄²⁻ forms are present at a 1:4 molar ratio (5). Although this plasma P_i represents a small fraction of total body phosphorus, it serves as an exchange pool between the various P_i-containing and -regulating organs, and disturbances in P_i homeostasis can affect almost all organ systems.

In addition to the widespread structural and metabolic functions of P_i, it has become increasingly apparent during the past 15 years that extracellular P_i can act as a signaling molecule directly altering gene expression and cell phenotype (6–9). The abundance of P_i in the skeleton has led to early studies describing the effects of extracellular P_i in this organ. Exposing cultured chondrocytes to a high level of extracellular P_i leads to their terminal maturation and subsequent matrix mineralization (10–13). Similarly, the apoptosis of terminally differentiated hypertrophic chondrocytes was shown to be dependent upon the circulating plasma P_i levels in vivo (14). In both of these in vitro and in vivo approaches, the P_i-mediated apoptosis of chondrocytes was dependent upon the activation of the MAPK ERK1/2 pathway (15–17), but not of other mitogen-activated protein kinases, such as p38 or c-Jun N-terminal kinase. Interestingly, the P_i-dependent activation of the ERK1/2 pathway up-regulated the gene expression of the mineralization inhibitors matrix Gla protein (Mgp) and osteopontin (Opn), most likely setting off a feedback mechanism to control P_i-induced mineralization (7, 16, 18, 19). The elevated extracellular P_i level was also shown to be important in osteoblast proliferation and differentiation (16, 20–24), cementoblast formation (25), odontoblast differentiation (26, 27), and osteoclast differentiation (28–30).

The P_i-mediated signaling underlies the notion that cells must possess a P_i-sensing mechanism on the surface of or within the cell that is able to detect and respond to the variation of extracellular P_i levels. The ability of organisms to detect changes in extracellular levels of other metabolites (such as Ca²⁺, glucose, or amino acids) has already been described (31–33), and emerging evidence suggests that similar events are at work to mediate the cellular effects of P_i (8, 34–36). Although the identity of the molecules involved in these mechanisms is still unknown in mammals, P_i-sensing machineries have been characterized in prokaryotic and eukaryotic unicellular organisms (37). In Escherichia coli, the phosphate transporter PstS and other periplasmic proteins (PstC, Pst, and PstB/PhoU) detect the variation of external P_i concentrations. In case of a low extracellular P_i level, this system increases the efficiency of P_i retention in the bacteria (38). In Saccharomyces cerevisiae, a low extracellular P_i level resulted in induction of the P_i transporter Pho84, now identified as the essential component of the P_i-sensing system (38, 39). Interestingly, following P_i restriction, it was demonstrated that Pho84 could trigger the rapid activation of protein kinase A without transporting P_i (40).

In mammals, the Slc20a1/PiT1 and Slc20a2/PiT2 proteins are expressed at the plasma membrane and have been described as mediating the intracellular uptake of P_i with a high affinity (41–43). PiT1 and PiT2 have a wide tissue distribution, being the only P_i transporters expressed in bone (44, 45). Interestingly, their expression can be modulated by extracellular P_i (21, 43, 44, 46, 47), and previous studies have suggested that they can mediate downstream effects of extracellular P_i. In bone, the elegant study of Kimata et al. (48) suggests that the chondrocyte response to extracellular P_i is mediated by a PiT1-dependent up-regulation of cyclin D1 through ERK1/2 pathway activation. The authors hypothesize that P_i-driven conformational changes of PiT1 could be involved in the P_i-sensing mechanism. In parathyroid cells, PiT1 was suggested to act as a P_i sensor to modulate the secretion of the phosphaturic parathyroid hormone (49). On the other hand, based on its property of oligomerizing upon extracellular P_i variation, PiT2 was also proposed to serve as a P_i sensor (50). Although these data support a possible role for PiT1 or PiT2 as P_i sensors, little is known about the underlying mechanisms. Because PiT1 and PiT2 have very close P_i transport characteristics (51), they may also share P_i-sensing properties and thus have interconnected roles in P_i sensing. Moreover, because P_i-independent functions have been highlighted recently for PiT1 (52–56), the involvement of P_i transport in the P_i sensing by PiT1 or PiT2 remains to be investigated.

In this report, we investigated the role of PiT1 and PiT2 as P_i sensors in osteoblastic and chondrocytic cell lines. We show that both PiT1 and PiT2 are required for mediating P_i-dependent signaling. We demonstrate that PiT1 and PiT2 could interact together and that extracellular P_i modulates this interaction. Finally, we show that cellular P_i uptake is not required to mediate P_i signaling through the PiT proteins.

Results

Requirement of both PiT1 and PiT2 for P_i-mediated signaling

We first investigated whether PiT1 and/or PiT2 were involved in the P_i-dependent up-regulation of Mgp and Opn expression. To this aim, using RNA interference, we established stably transfected osteoblastic MC3T3-E1 clones in which PiT1 or PiT2 expression was knocked down. In MC3T3-E1 shPiT1 clones, PiT1 gene expression showed a 63% reduction, together with a significant up-regulation of PiT2 (Fig. 1A). Similarly, the MC3T3-E1 shPiT2 clones displayed a 62% decrease in PiT2 mRNA level, together with a significant up-regulation of PiT1 (Fig. 1A). Comparable results were observed when cells were incubated with 10 mm extracellular P_i concentration (Fig. S1A). Interestingly, the sodium-dependent P_i uptake was similar in MC3T3-E1 shPiT1 and shPiT2 clones and control MC3T3-E1 cells (Fig. 1B), suggesting that a depletion of either PiT may be compensated by the remaining PiT, as was previously suggested (56). Consistent with this possibility, MC3T3-E1 clones stably transfected with both shPiT1 and shPiT2 resulted in a 52% reduction of both PiTs (Fig. 1A), resulting in a similar decrease in sodium-dependent P_i uptake (Fig. 1B). In contrast to wildtype differentiated MC3T3-E1 cells in which Mgp and Opn expression was up-regulated following stimulation with 10 mm extracellular P_i for 24 h, the up-regulation of Mgp and Opn expression in PiT-depleted MC3T3-E1 clones was blunted (Fig. 1C). The defect in P_i-dependent Mgp and Opn up-regulation arose despite a normal P_i transport in the shPiT1 or shPiT2 MC3T3-E1 clones, suggesting that a variation in intracellular P_i content is unlikely to account for defects in P_i-dependent signaling in the absence of either PiTs. Because the ERK1/2 signaling pathway was shown to be required for P_i-dependent regulation of Mgp and Opn expression (16, 19), we investigated the P_i-dependent ERK1/2 activation in differentiated PiT-depleted MC3T3-E1 clones. We showed that following a 30-min (Fig. S1B) or 24-h (Fig. 1D) stimulation with 10 mm extracellular P_i, the activation of ERK1/2 pathway was blunted in shPiT1, shPiT2, or shPiT1/shPiT2 clones, as compared with untransfected and shScramble-transfected cells. Similar data were obtained in three separate PiT-depleted MC3T3-E1 clones (Fig. S1, B–D) and in transiently transfected MC3T3-E1 cells (Fig. S2). Interestingly, the effect of PiT depletion on the activation of ERK1/2 pathway following stimulation with 10% fetal bovine serum (FBS) was not as pronounced as P_i stimulation, arguing for a specificity of PiT1 and PiT2 in the P_i-dependent activation of the ERK1/2 pathway.

Figure 1.

P_i-dependent Mgp and Opn gene regulation and ERK1/2 signaling require both PiT1 and PiT2 in MC3T3-E1 cells. A, RT-qPCR analysis of PiT1 (white bars) and PiT2 (black bars) mRNA levels in untransfected (UT) or stably transfected MC3T3-E1 cells, as indicated. Data are means ± S.E. (*, versus shScramb, p < 0.05, n = 3). B, sodium-dependent P_i uptake was measured in untransfected or stably transfected MC3T3-E1 cells, as indicated. Data are means ± S.E. (n = 3). C, RT-qPCR analysis of Mgp and Opn mRNA levels in untransfected or stably transfected MC3T3-E1 cells, as indicated. Cells were incubated in low-serum (0.5%) medium for 24 h and stimulated with 1 mm (white bars) or 10 mm (black bars) extracellular P_i concentration for 24 h. Data are means ± S.E. (error bars) (#, p < 0.05; ##, p < 0.01 versus 1 mm P_i control; and *, p < 0.05; **, p < 0.01 versus shScramb; n = 3) D, Western blot analysis of ERK1/2 phosphorylation (P-ERK 1/2) in untransfected or stably transfected MC3T3-E1 cells, as indicated. Cells were incubated in low-serum (0.5%) medium for 24 h and stimulated for another 24 h with 1 mm or 10 mm extracellular P_i concentration or with 10% FBS used as a positive control for ERK1/2 phosphorylation. Total ERK1/2 proteins were used as a loading control.

Moreover, we performed similar experiments in the MC615 chondrogenic cell line. We used a transient transfection approach leading to a 50 and 56% deletion of PiT1 and PiT2 mRNA levels, respectively (Fig. 2A). A similar extent of PiT deletion was obtained at the protein level, as shown by immunofluorescence (Fig. S3). MC615 cells were also used to rescue PiT deletion by overexpressing human PiT1 and PiT2 in PiT1- and PiT2-depleted cells, respectively (Fig. 2B). Similar to what was observed in MC3T3-E1 cells, depletion of either PiT1 or PiT2 in MC615 cells blunted the activation of the ERK1/2 pathway by 10 mm extracellular P_i, despite the up-regulation of the remaining PiT (Fig. 2C). When human PiT1 was overexpressed in PiT1-depleted MC615 cells, we could rescue the P_i-dependent ERK1/2 phosphorylation (Fig. 2C). Similar results were obtained when human PiT2 was overexpressed in PiT2-depleted MC615 cells or when both human PiT1 and PiT2 were overexpressed in PiT1-PiT2–depleted MC615 cells (Fig. 2C). This further illustrated the requirement of both PiT1 and PiT2 for the P_i-dependent ERK signaling in cell lines of skeletal origin.

Figure 2.

P_i-dependent ERK1/2 signaling requires both PiT1 and PiT2 in MC615 cells. A, RT-qPCR analysis of mPiT1 (white bars) and mPiT2 (black bars) mRNA levels in transiently transfected MC615 cells, as indicated. Data are means ± S.E. (error bars) (**, p < 0.01; ***, p < 0.001 versus shScramb; n = 3). B, RT-qPCR analysis of hPiT1 (white bars) and hPiT2 (black bars) mRNA levels in transiently transfected MC615 cells, as indicated. The endogenous murine PiT1 or PiT2 genes were used as reference genes to evaluate the overexpression of the transfected human PiT genes. Data are expressed as mean ± S.E. (***, p < 0.001 versus shScramb+pcDNA, n = 3). N/A, not applicable. C, Western blot analysis of ERK1/2 phosphorylation (P-ERK 1/2) in transiently transfected MC615 cells, as indicated. Cells were incubated in low-serum (0.5%) medium for 24 h and stimulated for 30 min with 1 mm or 10 mm extracellular P_i concentration as indicated. Total ERK1/2 proteins were used as a loading control.

PiT1 and PiT2 form hetero-oligomers upon variation of extracellular P_i concentrations

The requirement of both PiT1 and PiT2 for P_i-dependent ERK1/2 signaling may indicate the existence of a functional protein complex comprising both PiTs. This possibility is also reinforced by the presence in PiT1 and PiT2 protein sequences of a conserved and highly hydrophobic 127-amino acid domain that was suggested to be important for determining the quaternary structure of the protein (57). To investigate the formation of hetero- and homo-oligomers, we used HEK293T cells that are easy to transfect, allow a robust expression of PiTs at the plasma membrane, and have been shown to have similar P_i-mediated ERK1/2-specific activation (58). In addition, because PiT proteins are often difficult to detect using total cell extracts in Western blot experiments, we used a crude cell fractionation approach to analyze an enriched plasma membrane protein fraction revealed by the specific expression of the Na/K-ATPase, as shown in Fig. 3A. When analyzing the enriched plasma membrane fraction from PiT1- and/or PiT2-transfected cells by Western blotting, we could observe protein complexes after cell surface cross-linking using BS³, a membrane-impermeable cross-linker (Fig. 3B). The apparent molecular mass of 142–165 kDa that we detected from cells transfected with hPiT2 alone recapitulated the results obtained by Salaün et al. (50) demonstrating the formation of PiT2 homodimers. Similarly, the 151–188 kDa apparent molecular mass band detected from cells transfected with hPiT1 is consistent with the association of two PiT1 molecules. When cells were co-transfected with both hPiT1 and hPiT2, no change was observed in the band profile, apart from a less intense signal due to transfection with 50% fewer hPiT1 or hPiT2 plasmids. In this condition, we could not detect a distinct band at the theoretical PiT1-PiT2 heterodimer molecular mass, most likely due to the expected similar molecular weights of PiT homo- and heterodimers.

Figure 3.

Specific interaction between PiT1 and PiT2 varies upon extracellular P_i concentration. A, Western blotting analysis of Na/K-ATPase, hPiT1, and hPiT2 expression in pmaxGFP-transfected (Ctrl) or hPiT1 and hPiT2 co-transfected HEK293T cells after crude cellular fractionation, as indicated (see “Experimental procedures” for details). Overexpression of hPiT1 and PiT2 allowed a better signal than Ctrl. B, Western blotting analysis (IB) of hPiT1 (left) and hPiT2 (right) expression in HEK293T cells untransfected (UT) or transfected with hPiT1, hPiT2, or both after cell surface cross-linking using bis(sulfosuccinimidyl)suberate and cellular fractionation. Enriched plasma membrane fraction (F1) was analyzed. C, schematic representation of chimeric hPiT1-eYFP or -Rluc and hPiT2-eYFP or -Rluc proteins used for BRET assays. The DNA region encoding for the large internal loop of PiT1 and PiT2 (iLoop1 and iLoop2, respectively) was replaced by coding sequences of eYFP or Rluc. Transmembrane domains (black rectangles) are shown. D, confocal images obtained from transfected HEK293T cells, as indicated. Rluc was detected after immunofluorescence and nuclei were stained using Hoechst (blue). Scale bar, 10 μm. E, representative BRET saturation curves of co-transfected HEK293T cells, as indicated. F, representative BRET saturation curves (left panel) and BRET 50 index (right panel) of HEK293T co-transfected with hPiT1-eYFP and hPiT2-Rluc obtained following a 10-min incubation with 1, 3, or 10 mm extracellular P_i concentration, as indicated. BRET 50 data are means ± S.E. (error bars) (**, p < 0.01 versus 1 mm P_i condition, n = 8). G, the BRET 50 index was measured from BRET saturation curves obtained from HEK293T co-transfected with hPiT1-eYFP and hPiT1-Rluc (left) or hPiT2-eYFP and hPiT2-Rluc (right) following a 10-min incubation with 1, 3, or 10 mm extracellular P_i concentration. BRET 50 data are means ± S.E., and no statistically difference was observed (n = 4–8).

To study further the PiT oligomerization, we then used a BRET approach. To this aim, we constructed hPiT1 and hPiT2 chimeric proteins expressing the eYFP acceptor or Rluc donor. Because structure-function studies have excluded a role of the large intracellular loop (iLoop) in P_i transport and retrovirus binding (59–62) and showed no overlapping between iLoop and the highly hydrophobic domain (57), we substituted the iLoop with eYFP and Rluc sequences (Fig. 3C). When expressed in HEK293T, the chimeric hPiT1-eYFP or -Rluc and hPiT2-eYFP or -Rluc proteins could be visualized at the plasma membrane, as shown by confocal microscopy (Fig. 3D), enabling us to study their role in detecting the variation of extracellular P_i levels. We performed saturation BRET experiments in living cells to investigate their potential hetero- and homo-oligomerization. As shown in Fig. 3E, we obtained typical BRET-saturable curves when using hPiT1-eYFP and hPiT2-Rluc proteins, together with a high BRET ratio. In contrast, when the BTN3A2 protein was used instead of either PiT, no saturation could be achieved, together with a weak BRET ratio (Fig. S4, A and B). These data support the notion that PiTs can form hetero- and homo-oligomers specifically. We confirmed the specificity of the hetero-oligomers by a competition assay (Fig. S4C) whereby overexpression of untagged PiT1 or PiT2 reduces the BRET ratio, whereas unrelated BTN3A2 expression does not. We next determined whether the interaction of PiTs could be modulated by the variation of extracellular P_i concentration. We therefore performed the same saturation BRET experiments after a 10-min stimulation with 1, 3, or 10 mm extracellular P_i. Results reported in Fig. 3 (F and G) and Fig. S5A showed that saturation curves were different upon extracellular P_i concentration only for hPiT1-hPiT2 hetero-oligomers. Indeed, calculating the BRET 50 values from these curves, we showed a significant decrease at 10 mm extracellular P_i, suggesting a stronger interaction between PiT1 and PiT2 in this condition, whereas no variation was observed for homo-oligomers. In all conditions, the BRET max values did not vary significantly (Fig. S5B).

PiT1-PiT2–mediated ERK1/2 activation by extracellular P_i and regulation of PiT hetero-oligomerization are P_i transport–independent

Our results illustrated that both PiT proteins are important for P_i-dependent ERK signaling and that P_i can modulate PiT hetero-oligomerization. To elucidate whether the P_i transport function of PiTs was important to mediate P_i-dependent ERK1/2 signaling, we used the previously reported hPiT1^S128A and hPiT2^S113A P_i transport–deficient mutants (56, 63, 64). When hPiT1^S128A was overexpressed in PiT1-depleted cells in which P_i-dependent ERK1/2 activation was lost, we could rescue the activation of ERK1/2 signaling (Fig. 4A). Similarly, we could rescue the P_i-dependent ERK1/2 activation in PiT2-depleted cells by overexpressing hPiT2^S113A mutant. This was also true when cells depleted from both PiTs were transfected by both hPiT1^S128A and hPiT2^S113A (Fig. 4A). These data demonstrated that the P_i transport function of PiTs was dispensable for the P_i-dependent ERK1/2 activation. To further study whether the sodium-dependent P_i transport function of PiTs was important to mediate P_i-dependent PiT hetero-oligomerization, we performed BRET experiments in the absence of Na⁺. In this condition where P_i transport was blunted, the variation of extracellular P_i concentrations was still able to modulate PiT interaction (Fig. 4 (B and C) and Fig. S5C). Because P_i was able to modulate PiT interaction without being transported, this suggests that the binding of P_i to PiT proteins rather than its actual uptake into the cell may be involved in the modulation of PiT interaction. We next generated mutated versions of the chimeric hPiT1-eYFP and hPiT2-Rluc proteins in which Ser¹²⁸ or Ser¹¹³ was replaced by an alanine residue. As expected, overexpression of chimeric hPiT1^S128A-eYFP and hPiT2^S113A-Rluc resulted in decreased sodium-dependent P_i transport compared with transporting chimeric PiT proteins (Fig. 4D). Interestingly, using a BRET approach, we showed that although hPiT1^S128A-eYFP and hPiT2^S113A-Rluc were still able to interact together, this interaction was not modulated anymore by extracellular P_i variations (Fig. 4E (left) and Fig. S5C), consistent with a role for Ser¹²⁸ or Ser¹¹³ in P_i binding. When BRET experiments were performed with a transport-deficient mutant (hPiT1^S128A-eYFP or hPiT2^S113A-Rluc) and a transporting chimeric PiT protein (hPiT1-eYFP and hPiT2-Rluc), we could recover the modulation of PiT interaction by P_i (Fig. 4E (middle and left) and Fig. S5C), further supporting a role of Ser¹²⁸ or Ser¹¹³ in the P_i-dependent interaction of PiTs.

Figure 4.

P_i-dependent ERK1/2 signaling and PiT1-PiT2 hetero-oligomerization are independent of P_i transport. A, Western blot analysis of ERK1/2 phosphorylation (P-ERK 1/2) in transiently transfected MC615 cells, as indicated. Overexpression of P_i transport–deficient hPiT1 (PiT1^S128A) and/or hPiT2 (PiT2^S113A) was performed in PiT1–, PiT2–, or PiT1-PiT2–depleted cells, respectively. Cells were incubated in low-serum (0.5%) medium for 24 h and stimulated for 30 min with 1 or 10 mm extracellular P_i concentration. Total ERK1/2 proteins were used as a loading control. B, sodium-dependent and -independent P_i uptake was measured in HEK293T cells transfected as indicated. Data are means ± S.E. (error bars) (*, p < 0.05 versus sodium-dependent, n = 4). C, BRET 50 index was measured from BRET saturation curves obtained after a 10-min stimulation with 1, 3, or 10 mm extracellular P_i concentration from HEK293T co-transfected with hPiT1-eYFP and hPiT2-Rluc in an Na⁺-free medium. Data are means ± S.E. (*, p < 0.05 versus 1 mm P_i condition, n = 4). D, sodium-dependent P_i uptake was measured in HEK293T cells transfected with pcDNA6A plasmid (Ctrl) or with plasmids containing normal (hPiT1 and hPiT2) or P_i transport–deficient mutants (hPiT1^S128A and hPiT2^S113A) with or without BRET chimeric acceptor or donor, as indicated. Data are means ± S.E. (*, p < 0.05 versus P_i-transporting PiTs, n = 4). E, BRET 50 index was measured from BRET saturation curves obtained after 1, 3, or 10 mm extracellular P_i concentration stimulation for 10 min from HEK293T co-transfected with the indicated plasmids. Data are means ± S.E. (*, p < 0.05 versus 1 mm P_i condition, n = 4–5).

Discussion

The ability of a cell to detect changes in extracellular P_i levels is paramount for its adequate response to environmental fluctuations and critical for the appropriate modulation of P_i homeostasis and skeletal mineralization. In this work, we provided mechanistic insights into the molecular events leading to the detection of changes in extracellular P_i concentrations using skeletal cell lines as a model.

Skeletal cells are constantly exposed to high local extracellular P_i levels, mainly due to the continual resorption and formation of the mineralized extracellular matrix of bone and the need for tremendous quantities of P_i for mineralization purposes (65). This has made bone a model of choice to study P_i signaling, where it has been shown in early studies to regulate the programmed cell death of hypertrophic chondrocytes (10, 11, 13). In subsequent studies, we and others have shown that P_i could regulate skeletal mineralization by controlling the expression of the mineralization inhibitors Mgp and Opn through the activation of the ERK1/2 pathway (7, 15, 16, 18, 19). Our present study brings evidence for a role of PiT1 and PiT2 in transmitting the P_i signal to the cell by showing that the P_i-dependent up-regulation of Mgp and Opn and ERK1/2 phosphorylation were blunted in PiT1- or PiT2-depleted cells. A role for PiT1 in mediating ERK1/2 signaling has been reported earlier (48, 55); however, a similar role for PiT2 has never been illustrated before.

A critical aspect in studying the molecular events involved in P_i signaling is whether or not P_i needs to be transported within the cell to fulfill its role. This question is particularly relevant in view of the role of PiT1 and PiT2 as mediators of P_i signaling because these two proteins have well-described P_i transport functions (41–43). By using P_i transport–deficient mutants of PiT proteins, we could rescue the ERK1/2 signaling, demonstrating that PiT1 and PiT2 can mediate a P_i signal without transporting the ion. Furthermore, the loss of ERK1/2 signaling in the absence of PiT1 or PiT2 was not associated with a change in cellular P_i uptake, further indicating that the transport of P_i into the cell is not necessary for P_i signaling.

These data have major implications for the understanding of the P_i-signaling mechanism. The absence of functional compensation by PiT1 in PiT2-depleted cells, and vice versa, together with the requirement of both PiTs for P_i signaling, suggested that they could interact functionally and/or physically to mediate the P_i signal. Consistent with this hypothesis, using a BRET approach, we showed that PiT proteins could form homo- and heterodimers. More strikingly, we could illustrate that the formation of PiT1-PiT2 heterodimers only was affected by the variation of extracellular P_i levels. We could not quantify the relative importance of hetero- versus homodimers, but the cross-linking data supported the idea that the P_i-sensitive PiT1-PiT2 heterodimers were present at much lower quantities. It is possible that a low-abundance P_i-sensitive PiT1-PiT2 heterodimer may be more effectively tunable than a highly abundant P_i sensor. This abundance may also be consistent with the on/off P_i effect on ERK1/2 signaling that we have observed when deleting the PiT proteins.

Although a P_i-sensitive PiT1-PiT2 heterodimer is likely to represent an important component of the P_i-sensing machinery, deciphering the detailed functioning of such a sensor requires additional work. Nevertheless, our data provide several important mechanistic insights that may give clues to the understanding of the P_i-signaling cascade. We showed that in the absence of Na⁺, which blunts the P_i transport activity of PiT proteins, the formation of PiT1-PiT2 heterodimers was still sensitive to extracellular P_i variations. This is consistent with the idea that binding of P_i, rather than transport itself, is important for PiT1-PiT2 heterodimerization. This also suggests that Ser¹²⁸ and Ser¹¹³ residues that were demonstrated to be critical for P_i transport by PiT1 and PiT2 (56, 63) may indeed be involved in P_i binding, a prerequisite step for P_i transport (51). In line with this possibility, when BRET experiments were performed using PiT1 and PiT2 chimeric proteins in which Ser¹²⁸ and Ser¹¹³ were substituted with alanine residues, the PiT1-PiT2 heterodimer was not responsive anymore at extracellular P_i variations. The substitution of only one serine, however, rescued P_i sensitivity of the PiT1-PiT2 heteroduplex, illustrating the complex relationship between the structural arrangement of the P_i binding site and P_i sensitivity. A determination of the crystal structure of PiT proteins is therefore necessary to help determine the identity of the P_i binding site and its role in P_i transport and sensing.

An interesting consequence of our work is that PiT proteins should not be considered anymore as P_i transporters solely but also as P_i receptors able to mediate P_i signaling by activating specific downstream pathways. Such a hybrid transporter-receptor behavior may indicate that the PiT1-PiT2 heteroduplex could behave as a P_i transceptor, whereby conformational changes during the transport cycle (including the P_i-binding step) affect a signal transduction component that triggers a downstream signaling pathway (66). The difference between a transceptor and a pure receptor is that the transceptor can also transport the ligand into the cell. In recent years, evidence for transporters functioning as transceptors has been obtained in several eukaryotic systems (67). Interestingly, in yeast, the Pho84 phosphate carrier that is considered as an essential component of the P_i-sensing system was characterized as a P_i transceptor (40). In prokaryotic or lower eukaryotic organisms, true transceptors could be functionally characterized by specific mutants that either lack the signaling capacity and retain normal transport or have lost transport but retain signaling. In the case of PiT proteins, the signaling capacity is lost when either PiT1 or PiT2 is deleted, indicating that the possible transceptor function may only be revealed by heterodimerization and may not be associated with structural changes of either PiT. The gain of function provided through PiT heterodimerization implies that a unique protein complex mediates P_i signaling. This underlies the idea that specific PiT1 and PiT2 protein partners may be involved in this process. If this is the case, the identification of PiT-specific partners in the future may provide unique targets to modulate P_i signaling in specific organs or specific physiological conditions.

In summary, this study provided mechanistic insights into the P_i-signaling cascade in skeletal cell line models by unraveling PiT1-PiT2 heterodimers as essential components of a P_i-sensing machinery. Although in vivo studies are required to strengthen the physiological relevance of these findings, our work may help in deciphering the mechanisms underlying the ability of the organism to respond to the serum P_i level variations, which is the necessary first step in the P_i homeostasis-regulating cascade.

Experimental procedures

Cells and culture conditions

Murine preosteoblastic MC3T3-E1 cells were seeded at 10⁴ cells/cm² and cultured for 10 days in α-minimum essential medium GlutaMAX^TM (catalogue no. 32751, Thermo Fisher Scientific, Saint-Aubin, France) supplemented with 10% FBS and 1% penicillin/streptomycin. Murine chondrogenic MC615 cells were seeded at 10⁴ cells/cm² and maintained in a medium consisting of Dulbecco's modified Eagle's medium (DMEM) high glucose GlutaMAX^TM/Ham's F-12 (1:1) (catalogue no. 31966 and 31765, respectively, Thermo Fisher Scientific) supplemented with 10% FBS and 1% penicillin/streptomycin. HEK293T cells were maintained in DMEM high glucose GlutaMAX^TM supplemented with 10% FBS, 10 mm HEPES, and 50 μg/ml gentamicin. Cells were cultured at 37 °C in a humidified atmosphere of 5% CO₂ in air, and media were renewed every 2–3 days. When indicated, cells were incubated in low-serum (0.5%) medium for 24 h and stimulated with various concentrations of P_i for 30 min or 24 h. P_i was added as a mixture of NaH₂PO₄ and Na₂HPO₄ (pH 7.4), as described previously (10).

RNA interference

Inactivation of PiT1 or PiT2 were performed by cloning an shPiT1 (56) or shPiT2 (see sequences in Table 1) into pSUPER vector (68) expressing a puromycin resistance gene. A scramble sequence cloned into pSUPER was used as a negative control (56). Stable knockdown of PiT1 or PiT2 was obtained by transfecting MC3T3-E1 cells with 5 μg of the pSUPER-shRNAs using the T-20 program of the Amaxa nucleofector system (Cell Line Nucleofector^TM Kit V VCA-1003, Lonza, Bâle, Switzerland). Cells were plated at limiting density, and puromycin-resistant clones were picked, expanded, and tested for PiT expression. Experiments were performed with three independent stable transfectants, and the data presented illustrate a representative clone. Transient inactivation of PiT was also performed in MC3T3-E1 and MC615 cells using transfection as described above. These transient experiments were stopped 72 h after transfection.

Table 1.

Primer sequences used for RNA interference, site-directed mutagenesis and RT-qPCR

m, mouse; h, human; F, Forward primer; R, Reverse primer. N/A, not applicable.

*, siRNA sequences are underlined.

#, desired mutation is underlined.

Gene expression analysis

Total RNA was isolated from cells using the NucleoSpin RNA II kit (Macherey-Nagel, Hoerdt, France) according to the manufacturer's instructions. RNA was reverse transcribed using Affinity Script (Agilent Genomics, Santa Clara, CA) as per the manufacturer's recommendations. Real-time PCR was performed on a Mx3000P system (Stratagene, San Diego, CA) using Brilliant III Ultra-Fast SYBR QPCR Master Mix (Agilent Genomics). The following temperature profile was used: denaturation at 95 °C for 3 min, amplification during 40 cycles of 5 s at 95 °C and 20 s at 60 °C, followed by a step at 95 °C for 1 min and 65 °C for 30 s. Expression of target genes were normalized to GAPDH expression levels and were calculated as described previously (69). Primer sequences are listed in Table 1.

Immunofluorescence and confocal microscopy

One day after transient transfection, MC615 cells were fixed/permeabilized in methanol at −20 °C for 5 min and blocked in 1% bovine serum albumin for 1 h at room temperature. Immunodetection of PiT1 and PiT2 was performed using rabbit anti-mouse PiT1 or PiT2 antibody, respectively (generously provided by Dr. G. Friedlander), at a 1:200 dilution overnight at 4 °C, and goat anti-rabbit Alexa488 secondary antibody (catalogue no. A11008, Thermo Fisher Scientific) at a 1:1,000 dilution for 1 h at room temperature. The nuclei were counterstained using 1 μg/ml TO-PRO 3 iodide solution (Thermo Fisher Scientific). The stained cells were mounted in Prolong Antifade mounting medium (Thermo Fisher Scientific). Images were acquired with a Nikon Eclipse TE2000E confocal microscope (Nikon, Badhoevedorp, The Netherlands) equipped with a ×60 oil immersion objective. The averaged intensity for PiT1 and PiT2 staining was determined using Metamorph version 7.5 software.

HEK293T cells were seeded at 3 × 10⁴ cells/cm² in μ-slide 8-well ibiTreat (catalogue no. 80826, Ibidi, Madison, WI) precoated with poly-l-lysine (Sigma-Aldrich). Cells were transfected with 0.125 μg/well of plasmid using JetPrime (Polyplus transfection, Illkirch, France) according to the manufacturer's instructions. Twenty-four hours post-transfection, HEK293T cells were fixed in 4% paraformaldehyde, permeabilized in 0.2% Triton X-100, and blocked in 1% bovine serum albumin for 1 h at room temperature. Immunodetection of hPiT1-Rluc, hPiT2-Rluc, and BTN3A2-Rluc was performed using anti-Rluc antibody (catalogue no. GTX47953, GeneTex, Irvine, CA) at a 1:200 dilution overnight at 4 °C and goat anti-rabbit Alexa488 secondary antibody (catalogue no. A11008, Thermo Fisher Scientific) at a 1:1,000 dilution for 1 h at room temperature. Nucleus staining was performed using 2 μg/ml Hoescht solution (catalogue no. H3569, Thermo Fisher Scientific), and immunofluorescence was preserved in Citifluor AF1 (Biovalley, Nanterre, France). Images were acquired with a Nikon A1Rsi confocal microscope (Nikon, Minato-ku, Tokyo, Japan) equipped with a ×60 oil immersion objective.

Phosphate uptake measurements

HEK293T cells were seeded at 10⁵ cells/cm² in poly-l-lysine (Sigma-Aldrich)–precoated 24-well plates. Cells were transfected with 0.5 μg/well of plasmid using JetPrime (Polyplus transfection) according to the manufacturer's instructions. P_i uptake in HEK293T cells was measured 24 h after transfection. To perform uptake experiments in MC3T3-E1 cells, PiT-depleted MC3T3-E1 clones were seeded at 1.5 × 10⁵ cells/cm², and uptake was performed 24 h later. P_i uptake was performed as described previously (70). Briefly, after three washing steps, cells were incubated in an uptake medium containing 100 μm [³²P]KH₂PO₄ (0.5 μCi/ml) in the presence of 137 mm NaCl (total P_i transport) or N-methyl-d-glucamine (sodium-independent transport) at 37 °C for 10 min. sodium-dependent P_i transport was calculated as the difference between total and sodium-independent P_i transports. Cells were washed three times and lysed with 0.1 m NaOH solution, and aliquots of cell lysates were taken for the determination of protein content (Pierce BCA protein assay kit, Thermo Fisher Scientific) and the radioactivity by liquid scintillation counting (Ultima Gold LLT, PerkinElmer Life Sciences) in a Hidex 300 SL β counter.

Immunoblot analysis

Cells were lysed for 30 min in ice-cold lysis buffer (20 mm Tris-HCl, pH 7.5, 100 mm potassium chloride, 1 mm EDTA, 1 mm EGTA, 1 mm dithiothreitol, 20 mm β-glycerophosphate, 2 mm Na₃VO₄, 1 mm phenylmethylsulfonyl fluoride, and 1 mm NaF). After centrifugation at 12,000 × g for 10 min at 4 °C, the protein extracts (supernatants) were boiled in Laemmli loading buffer before SDS-PAGE. For cross-linking experiments, proteins were not boiled, and electrophoresis was done on NuPAGE^TM 3–8% Tris acetate gel (catalogue no. EA0378BOX, Thermo Fisher Scientific). Proteins were transferred to PVDF membrane, and blocking was performed with 5% nonfat dry milk/TBST (10 mm Tris, 154 mm NaCl, 0.15% Tween 20) for 1 h at room temperature. Blots were probed with primary antibodies in 5% nonfat dry milk/TBST overnight at 4 °C followed by secondary antibodies for 1 h at room temperature. Anti-ERK1/2 (catalogue no. 9102) and anti-phospho-ERK1/2 (catalogue no. 4370) were from Cell Signaling (Danvers, MA) and used at a 1:2,000 dilution. Anti-PiT1 (catalogue no. 12423-1-AP) and anti-PiT2 (catalogue no. 12820-1-AP) were from ProteinTech (Rosemont, IL) and used at a 1:1,000 dilution. Anti-Na/K-ATPase (catalogue no. ab7671) was from Abcam (Cambridge, UK) and used at a 1:5,000 dilution. Monoclonal anti-β-actin clone AC-74 (Sigma-Aldrich) was used at a 1:5,000 dilution as a loading control. Anti-rabbit (catalogue no. 7074, Cell Signaling) and anti-mouse (catalogue no. A9917, Sigma-Aldrich) secondary antibodies were used at 1:2,000 and 1:80,000 dilutions, respectively. Signal detection was performed using ECL Western blotting detection reagent and ECL hyperfilm (GE Healthcare) or ChemiDoc Imaging System^TM (Bio-Rad).

Crude cellular fractionation and cross-linking

HEK293T cells were seeded at 10⁵ cells/cm² in 6-well plates and transfected with 1 μg/well each of hPiT1 and hPiT2 plasmids using JetPrime (Polyplus transfection) according to the manufacturer's instructions. Twenty-four hours post-transfection, cells were scraped and lysed in 1 ml of buffer A/well (10 mm Hepes, pH 7, 10% sucrose, 5 mm EDTA, Phosphatase Inhibitor Mixture 3 (catalogue no. P0044), and Protease Inhibitor Mixture (catalogue no. P8340) from Sigma-Aldrich. Lysates were passed 10 times through a 22-gauge 1-ml syringe and centrifuged at 800 × g for 5 min at 4 °C. Pellet (fraction 1; F1) was resuspended in 20 μl of radioimmune precipitation assay buffer (50 mm Tris-HCl, pH 8, 150 mm NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 20 mm EDTA, Phosphatase Inhibitor Mixture 3, and Protease Inhibitor Mixture). The supernatant was centrifuged at 20,000 × g for 45 min at 4 °C, and the resulting supernatant (Fraction 2; F2) was reserved. The pellet (Fraction 3; F3) was resuspended in 20 μl of radioimmune precipitation assay buffer and submitted to two freeze/thaw cycles, a 10-min incubation at 100 °C, and a 30-s sonication. All fractions were incubated 1 h at 4 °C with gentle mix. F3 was centrifuged at 12,000 × g for 10 min at 4 °C, and supernatant was used for analysis.

For cross-linking experiments, transfected HEK293T cells were washed three times in cold PBS and cross-linked 30 min at 4 °C with 1 mm bis(sulfosuccinimidyl)suberate (catalogue no. 21580, Thermo Fisher Scientific). This cell surface cross-linker was blunted with ice-cold 50 mm Tris-HCl, pH 8, for 15 min and washed twice with cold PBS. Cells were then lysed in Buffer A and processed for cell fractionation as described above.

Construction of chimeric and transport-deficient PiT plasmids

The hPiT1 and hPiT2 sequences were previously cloned into pcDNA6A plasmid (56). The V5-His₆ fusion tag sequence present in pcDNA6A was excluded by introducing a stop codon at the end of the hPiT coding sequence using site-directed mutagenesis (QuikChange, Agilent Genomics) (see primers in Table 1). The iLoop of hPiT1 and hPiT2 was then replaced by eYFP or RLuc proteins using FastCloning (71). Briefly, using Phusion® high-fidelity DNA polymerase (New England Biolabs) and overlapping specific primers (Table 2), we PCR-amplified the pcDNA6A-hPiT1 and pcDNA6A-hPiT2 vectors with the exception of the iLoop regions and the coding regions of eYFP and Rluc from pEYFP-N1 (Clontech, Mountain View, CA) and phRluc-C1 (PerkinElmer Life Sciences) plasmids, respectively. After digestion of the DNA templates by DpnI, PCR-amplified overlapping sequences were reassembled by E. coli-mediated recombination-ligation following transformation in high-efficiency NEB® 10-β competent cells (New England Biolabs, Ipswich, MA). In the final hPiT1-eYFP or -Rluc chimeric constructs, the eYFP or Rluc coding sequences were inserted in place of iLoop1 (amino acids 268–492). Similarly, the eYFP or Rluc coding sequence was inserted in place of iLoop2 (amino acids 256–450) in the hPiT2-eYFP or -Rluc constructs. The integrity of the constructs was verified by sequencing. To serve as a control for BRET experiments, the BTN3A2 DNA sequence (generously provided by Dr. Scotet, INSERM UMR1232, Centre de Recherche en Cancérologie et Immunologie, Nantes-Angers, France), encoding a small cell surface–expressed protein, was fused to the N terminus of eYFP or Rluc coding sequences using the same strategy (primers used are reported in Table 2).

Table 2.

Primer sequences used for FastCloning

*, overlapping sequences underlined.

To generate transport-deficient mutant versions of hPiT1-eYFP and hPiT2-Rluc chimeric constructs, we replaced the Ser¹²⁸ of hPiT1 or Ser¹¹³ of hPiT2 by an alanine using site-directed mutagenesis (QuikChange, Agilent Genomics), as reported previously (56, 63). The sequences of the primers used are listed in Table 1.

BRET saturation assays

HEK293T cells were seeded at 5 × 10⁴ cells/cm² in 12-well plates. The next day, cells were co-transfected using JetPrime (Polyplus transfection) with a fixed amount of hPiT1-Rluc (50 ng/well), hPiT2-Rluc (100 ng/well), or BTN3A2-Rluc (10 ng/well) plasmids (encoding BRET donors) and variable amounts of hPiT1-eYFP or hPiT2-eYFP (from 12.5 to 1,500 ng/well) or BTN3A2-eYFP (from 1.56 to 50 ng/well) plasmids (encoding BRET acceptors). The pcDNA6A empty vector was used to compensate for the variable amounts of transfected DNA and to ensure equivalent transfection conditions in each well. Twenty-four hours later, transfected cells were detached using 0.5 mm EDTA solution and seeded at 1.5 × 10⁵ cells/cm² in white flat bottom 96-well plates in duplicate. BRET experiments were performed 48 h post-transfection. Cells were washed once with 0.9% NaCl solution and stimulated with various concentrations of P_i for 10 min. P_i was added as a mixture of NaH₂PO₄ and Na₂HPO₄ (pH 7.4). When indicated, cells were previously starved of P_i by incubating cells overnight with DMEM high glucose no phosphates (catalogue no. 11971, Thermo Fisher Scientific) supplemented with 10% FBS, 10 mm HEPES, and 50 μg/ml gentamicin before P_i stimulation. The coelenterazine h substrate (UPR3078, Interchim Uptima, Montluçon, France) was added at a final concentration of 5 μm by automated injection in the Mithras LB940 plate reader (Berthold Technologies, Versailles, France), and 485- and 530-nm light emissions were measured consecutively several times. The BRET ratio was calculated as the ratio of light emitted by the acceptor fusion protein at 530 nm over the light emitted by the donor fusion protein at 485 nm. Values were corrected with the background signal calculated from a well without donor fusion protein. The BRET 50 was calculated as the eYFP/Rluc value at which the BRET ratio is half of the maximum BRET ratio achieved at saturating substrate concentration.

Statistical analysis

Data are expressed as mean ± S.E. GraphPad version 5.0 software was used to perform Mann–Whitney tests. A p value of <0.05 was considered statistically significant. Unless otherwise stated, experiments were repeated at least three times.

Author contributions

N.B., G.C., and A.B. conducted most of the experiments, from conception and design to acquisition of data or analysis and interpretation of data. S.S. provided technical assistance. L.B., J.G., and S.B.C. conceived the idea and supported the coordination of the project. N.B. and L.B. wrote the paper. N.B., G.C., A.B., S.B.C., J.G., and L.B. made adjustments to the final paper version. All authors reviewed the results and approved the final version of the manuscript.