Critical contributions of pre-S1 shoulder and distal TRP box in DAG-activated TRPC6 channel by PIP2 regulation

Mori, Masayuki X.; Okada, Ryo; Sakaguchi, Reiko; Hase, Hideharu; Imai, Yuko; Polat, Onur K.; Itoh, Satoru G.; Okumura, Hisashi; Mori, Yasuo; Okamura, Yasushi; Inoue, Ryuji

doi:10.1038/s41598-022-14766-x

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Published: 24 June 2022

Critical contributions of pre-S1 shoulder and distal TRP box in DAG-activated TRPC6 channel by PIP₂ regulation

Masayuki X. Mori^1,3,
Ryo Okada^1,2,
Reiko Sakaguchi¹,
Hideharu Hase³,
Yuko Imai⁴,
Onur K. Polat³,
Satoru G. Itoh^5,6,
Hisashi Okumura^5,6,
Yasuo Mori³,
Yasushi Okamura⁷ &
…
Ryuji Inoue⁴

Scientific Reports volume 12, Article number: 10766 (2022) Cite this article

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Abstract

Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P₂ or PIP₂) regulates the activities of numerous membrane proteins, including diacylglycerol(DAG)-activated TRPC3/6/7 channels. Although PIP₂ binding is known to support DAG-activated TRP channel activity, its binding site remains unknown. We screened for PIP₂ binding sites within TRPC6 channels through extensive mutagenesis. Using voltage-sensitive phosphatase (DrVSP), we found that Arg437 and Lys442, located in the channel’s pre-S1 domain/shoulder, are crucial for interaction with PIP₂. To gain structural insights, we conducted computer protein–ligand docking simulations with the pre-S1 domain/shoulder of TRPC6 channels. Further, the functional significance of PIP₂ binding to the pre-S1 shoulder was assessed for receptor-operated channel functions, cross-reactivity to DAG activation, and the kinetic model simulation. These results revealed that basic residues in the pre-S1 domain/shoulder play a central role in the regulation of PIP₂-dependent gating. In addition, neutralizing mutation of K771 in the distal TRP box reversed the effect of PIP₂ depletion from inhibiting to potentiating channel activity. A similar effect was seen in TRPV1 channels, which suggests that TRPC6 possesses a common but robust polarity switch mediating the PIP₂-dependent effect. Overall, these mutagenesis studies reveal functional and structural insights for how basic residues and channel segments in TRP channels are controlled through phosphoinositides recognition.

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Introduction

Phosphatidylinositol 4,5-bisphosphate, also known simply as PI(4,5)P₂ or PIP₂, a phospholipid component of cell membranes, contributes to the activity of numerous molecules, including ion channels^1,2,3. Among those are transient receptor potential (TRP) channels, which are known to sense diverse thermal, mechanical, and chemical stimuli and to be related to a variety of pathophysiological functions⁴. Nearly all TRP channels are positively or negatively regulated by PIP₂^5,6,7. For instance, reductions in membrane PIP₂ levels inhibit most mammalian TRP canonical or classical (TRPC) channels^8,9,10,11,12, while hydrolysis products of PIP₂ such as DAG and polyunsaturated fatty acids (PUFAs) activate these channels^13,14. Thus, the decrease in PIP₂ level and resultant degradation products convey opposing signals to TRPC channels. These complicated but exquisite regulatory mechanism serves to control TRPC regulation in self-limiting and autonomic manner^8,9,10,12. We previously showed that reducing PIP₂ inhibits DAG-activated TRPC channels (TRPC3/6/7) channels, even in the presence of DAG, which indicates that the channel likely possesses distinct sites to interact with PIP₂ and DAG, respectively^8,15. However, no exact PIP₂ binding/interaction site that could affect TRPC channel gating has yet been identified.

Our focus has been to gain knowledge into the molecular and the structural insight of the PIP₂-interaction site on DAG-activated channels, especially TRPC6. TRPC6 plays an important role in the development of several fibrotic diseases including FSGS^16,17,18, and does not conserve a classically established PIP₂ binding domain, the Pleckstrin homology (PH)¹⁹. It has been reported that a complementary PH-like domain fragment in the N-terminal domain and the C-terminal domain serve as PIP₂ binding sites in DAG-activated TRPC channels^20,21. In addition, several other regions conserved in non-DAG-related TRP channels including the Ankyrin(ANK)-repeat domain (ARD) and TRP box, also reportedly bind PIP₂^22,23. Given this complexity, analysis that provides a broad perspective of PIP₂ interaction sites is essential for a comprehensive understanding of the regulation of TRP channels.

In the present study, we investigated PIP₂ interaction sites based on kinetic analysis upon the activation of DrVSP, a voltage-controllable PIP 5-phosphatase that is able to induce transient depletion of PIP₂^24,25,26. To do so, we applied this method along with extensive neutralizing mutation of the basic amino acid residues of TRPC6 channel. This enabled us to identify several residues critical for PIP₂ binding that are broadly situated within the distal N-terminal region, ARD domain, S4-S5 linker, TRP box, distal TRP box, distal C-terminal regions, and pre-S1 domain/shoulder. Among these, the neutralization of basic residues in the pre-S1 domain/shoulder severely altered the deactivation and reactivation kinetics upon the PIP₂ interaction. More specifically, channels mutated within the pre-S1 shoulder exhibited significant reductions in GPCR-activated TRPC6 current amplitudes. We also found that a mutation in the distal TRP box altered the effect of PIP₂ depletion. Intriguingly, a basic residue mutation in the distal TRP box has been shown to switch PIP₂ selectivity in the heat/capsaicin-activated TRPV1 channels²⁷. These results imply that a common switching mechanism may underlay the actions of PIP₂ recognition on TRP channels. Overall, our results provide valuable and fundamental insights into the mechanisms by which PIP₂ regulates TRP channels and how PIPs and ion channels functionally interact with one another.

Results

Evaluation of PIP₂ binding sites on the TRPC6 channel

To investigate the relationship between PIP₂ binding and TRPC6 channel activity, we evaluated three features of OAG-induced TRPC6 currents in HEK293 cells. First one was the kinetics of TRPC6 channel deactivation (decay, t_1/2) induced by DrVSP-mediated PIP₂ depletion. DrVSP was activated by membrane depolarization to + 100 mV for 700 ms. Second was the kinetics of reactivation (recovery, τ) mediated by replenishment of PIP₂ after repolarization to − 50 mV. These deactivation and reactivation kinetics can be used to estimate the dissociation and binding processes in reversible first order reactions. For wild-type TRPC6 (TRPC6_WT, C6_WT), decay (t_1/2) was 229 ± 15 ms, while recovery (τ) was 2.01 ± 0.13 s (Figs. 1B–D and 2). The third feature examined was the functional impact of PIP₂ dissociation (i.e., TRPC6 channel inhibition) elicited by DrVSP activation. This inhibition was expressed as the ratio of the current amplitudes before and after DrVSP activation (I_post/I_pre). This ratio was ~ 0.5 for TRPC6_WT (Figs. 1D and 2 bottom left). Co-expression of an inactive DrVSP mutant has no effect on TRPC6 currents (Fig. 1A,C,D). This indicates that TRPC6 channel activity led to no clear voltage-dependence, which was reported previously for other TRP channels^28,29,30.

However, the kinetics of both the deactivation and the reactivation should reflect several distinct processes including the kinetics of VSP activation/inactivation, the time course of PIP₂ dephosphorylation, and the altered channel gating upon PIP₂ dissociation and bindings. Among those processes, we measured the dynamics of PIP₂ alternation upon activation of DrVSP. The time courses of the depletion (t_1/2) and replenishment (τ) of PIP₂ were 135 ± 24 ms and 5.66 ± 0.48 s, respectively (Fig. 1E). This indicates a close correlation between PIP₂ level and TRPC6 channel activity, and the kinetics of OAG-induced currents should reflect PIP₂-dependent processes that affect TRPC6 channel functionality.

Screening of PIP₂ binding domains

TRPC6 channels possess nearly 90 positively charged (basic) residues in each subunit, among which we selected over 30 residues for the first screening based on following criteria: (1) residues were probably located on the intracellular side of the channel and (2) residues were clustered in the primary sequence with other positive residues. Mutant TRPC6 channels were co-expressed with DrVSP in HEK293 cells, and the membrane currents were measured by the whole-cell recordings. We found that following neutralization of basic residues in the distal N-terminal region (K75Q, R78Q, and R73Q/K75Q/R77Q), OAG no longer elicited any TRPC6-mediated currents (Fig. 2). This suggests that the distal N-terminal portion of the channel, where there is no available structure so far, may contribute to its translocation to the membrane, as suggested in the previous report²⁰. The other 24 constructs tested carried OAG-induced currents, and 8 out of them exhibited a significantly faster decay upon PIP₂ depletion than TRPC6_WT (Fig. 2, upper, blue bars). Among those, R437Q and R746Q mutants were the fastest (92 ± 10 ms (n = 6) and 118 ± 13 ms (n = 9), respectively).

On the other hand, the recovery of TRPC6 current from DrVSP-mediated inhibition, with restoration of PIP₂ was significantly accelerated or delayed in 9 out of 26 constructs (Fig. 2, middle, green bars). The current recovery was markedly delayed in R437Q and R865Q mutants, 4.78 ± 0.53 s and 3.70 ± 0.51 s, respectively. These mutations are respectively located in the pre-S1 domain/shoulder and calmodulin/inositol-1,4,5-trisphosphate receptor binding domain (CIRB)³¹, the latter is also known as PIP₂/calmodulin binding domain^21,32. Only the single mutation R758Q and double mutation K781Q/K782Q reduced the effect of PIP₂ depletion, as indicated by the weak inhibition ratio (I_post/I_pre), (Fig. 2, bottom, yellow bars). However, none of the mutations led to more than 50% current inhibition upon PIP₂ depletion. This suggests that there is a 50% PIP₂-independent component in the OAG-induced currents or the maximal efficacy of DrVSP to deplete endogenous PIP₂ may not be 100%, possibly due to fast replenishment of PIP₂ in some compartmentalized area.

Pre-S1 is a PIP₂ binding domain

In contrast to the aforementioned constructs, R629Q (in S4–S5 linker) and K748Q (within TRP box) mutants exhibited accelerated recovery from the inhibition (1.17 ± 0.05 s and 1.47 ± 0.11 s, respectively), suggesting that these basic residues may have less contribution to rebinding of PIP₂. Thus, among the first screened mutants, the pre-S1 domain/shoulder mutation R437Q exerted the most critical effect on both the decay and the recovery of TRPC6 channel currents (Fig. 2). This finding was further confirmed using the depolarizing step-pulse protocols over a wide range of membrane potentials (Fig. 3A–F). The results revealed that throughout the membrane potential tested, the R437Q mutation decays faster and recovers more slowly than TRPC6_WT.

Because the pre-S1 shoulder is enriched in basic residues (Fig. 3G), this prompted us to conduct a second mutational screening focused on the pre-S1 shoulder, which revealed the importance of K442, in addition to K431 and K434. The time constants for the decay and recovery of K442Q were 54 ± 11 ms and 3.71 ± 0.53 s, respectively (Fig. 3H). Amino acid sequence alignment of the pre-S1 domain/shoulder of mammalian TRPC channels shows identical positively charged residues at sites equivalent to R437 and K442 (Fig. 3G). This indicates that R437 and K442 in the pre-S1 shoulder play an essential role in PIP₂ binding to TRPC channels, including TRPC6. Moreover, the contributions of basic residues within the TRP box were also undoubtful in the kinetic analysis, which showed R746Q to produce the fastest current decay among the tested mutations (Fig. 2). To understand whether the pre-S1 shoulder and the TRP box exert a cooperative effect on PIP₂ binding, we examined the effect of R437Q/R746Q double mutations. The decay and recovery kinetics of this double mutant were indistinguishable to R437Q mutant (Fig. 4C,D), indicating that the contribution of R437 is dominant to R746 for PIP₂ binding, rather than showing an additive effect.

To compare PIP₂-affinity to respective mutants, we calculated the ratio of the recovery (τ) and decay (t_1/2) as a value related to the dissociation constant (Fig. 3I). Based on this estimation, the R437Q and K442Q channels exhibited nearly five to eight-fold less affinity for PIP₂ than the TRPC6_WT channel (Fig. 3I). Given that the dissociation constant for PIP₂ binding to TRPC6_WT was previously reported to be 2 µM¹⁵, we propose that those for PIP₂ binding to the pre-S1 domain/shoulder mutants (R437Q and K442Q) are greater than 10 µM.

PIP₂ binding to the pre-S1 domain/shoulder, linker domain, and TRP box

Recent cryo-EM structures of TRPC3/6 channels showed that the pre-S1 domain is exposed to the outside of the channel complex^33,34. Within the pre-S1 domain, positively charged residues are situated every three or five amino acids (K431, K434, R437, K442) to form an amphipathic helix termed the “pre-S1 shoulder” (Fig. 4A). Intriguingly, the pre-S1 shoulder is positioned at the inner surface of the membrane, where it encounters the residues H358 and R360. Neutralizing mutations of those residues slows the recovery, implying weaker PIP₂ affinities for TRPC6 channel (Fig. 2). This suggests the basic residues in the pre-S1 shoulder as well as the H358/R360 residues are crucial for recognition of PIP₂.

To confirm this idea, we performed a docking simulation with PIP₂ and the pre-S1 shoulder using the cryo-EM structure of TRPC6 (PDB 6UZ8). Within the simulation, the inositol head of PIP₂ was positioned at a surface pocket surrounded by the pre-S1 shoulder and H358/R360 residues. The TRPC6_WT model exhibited the highest docking energy (-6.0 kcal/mol), while R437Q and K442Q pre-S1 shoulder mutants exhibited somewhat lower docking energies (− 5.4 and − 5.6 kcal/mol, respectively). Intriguingly, the docking free energy between PIP₂ and H358/R360 mutant was clearly reduced (− 4.9 kcal/mol). These results reflect the decreased affinity between PIP₂ and the mutant channels. Indeed, we also tested a point mutation at a cysteine residue (C429), because it is located in the pre-S1 domain and is near the center of PIP₂-docking area. Although the docking energy of the C429S mutant was no lower than that of the wild-type channel (− 6.0 kcal/mol), the actual decay was significantly accelerated (121 ± 17 ms, Fig. 4D). These observations further confirm that the pre-S1 shoulder and H358/R360 residues contribute critically to PIP₂ binding.

Functional role of the pre-S1 domain/shoulder

For evaluation the functionality of the PIP₂ binding pocket identified in the docking simulation, TRPC6 channels were co-expressed with muscarinic receptors, which were then stimulated with a muscarinic receptor agonist carbachol (CCh). We found that the maximum current density upon receptor stimulation was significantly suppressed by single or double mutations, H358Y/R360Q, R339Q, K434Q, R437Q, and K442Q (Fig. 5A,B). Moreover, the extents of suppression elicited by these mutations as well as those by different combinations of double or triple mutations K434Q/R437Q, R437Q/K442Q, and K431Q/K434Q/R437Q were nearly identical. This confirms that the surface pocket for the PIP₂ interaction is crucial for receptor-activated channel activity. To then assess the importance of the pre-S1 domain/shoulder for channel localization in the cell membrane, we used confocal microscopy to compare the localizations of TRPC6_WT and R437Q and R437Q/K442Q mutants fused with cyan fluorescence protein (CFP) based on their co-localization with the yellow fluorescence protein (YFP)-fused PH-domain sensor. Co-localization of the channel and PH-domain after transfection into HEK293 cells did not statistically differ between the wild-type and mutant TRPC6 channels (Pearson’s correlation coefficient = 0.68 ± 0.02 for WT and 0.61 ± 0.07 for R437QK442Q, Fig. 5D). This suggests the pre-S1 domain/shoulder mainly contributes to channel functionality, but not to PIP₂-dependent membrane localization.

PIP₂ binding to TRPC channels allows the channels availability, and the less binding is possible to induce channel dysfunctionality, vice versa. It has been shown that impairment of Ca²⁺-dependent inactivation of TRPC6 channels is a cause of FSGS³². We therefore measured the receptor-activated TRPC6 current inactivation of the pre-S1 mutants under low Ca²⁺ buffering conditions. As shown in Fig. 5C, R437Q, R437/K442Q mutants and TRPC6_WT exhibited almost identical residual currents. Thus, binding of PIP₂ to the pre-S1 domain/shoulder is not directly involved in Ca²⁺-dependent inactivation and may not account for the pathogenesis of FSGS.

In addition, here and previously, we demonstrated that reducing PIP₂ inhibits channel activity, even in the presence of DAG⁸. We therefore tested how PIP₂ affects the potency of DAG to activate TRPC6 channel by comparing the concentration-dependent effects of OAG on TRPC6_WT and pre-S1 mutants. Figure 6A shows the concentration dependence of the responses of R437Q, R437Q/K442Q mutant and TRPC6_WT channels to OAG. This dependency was investigated by ratiometric Fura-2 photometry³⁵. The normalized dose–response data were best fit to the Hill equations with EC₅₀ values of 48, 37 and 46 μm and the coefficients (n) of 1.4, 1.5 and 0.8 for R437Q, R437Q/K442Q, and TRPC6_WT, respectively. Thus, mutations in the pre-S1 domain/shoulder cause practically no change in EC₅₀ values. This suggests that PIP₂ binding does not significantly affect DAG binding but instead may affect allosteric activation by DAG.

We used our previously described simulation model, to evaluate how PIP₂ binding affects receptor-activated TRPC6 channels¹⁵ (Fig. 6B). By reducing the channel’s binding affinity for PIP₂ while keeping its affinity to DAG, the amplitude of a simulated current was gradually decreased (Fig. 6C). When PIP₂-channel affinity was reduced by one-fifth, the current amplitude was reduced by half (Fig. 6C, black vs. blue traces). This result is consistent with those obtained with the R437Q and K442Q mutants, which exhibited reduced PIP₂ affinities and current densities (Figs. 3I and 5A,B). Taken together, these findings suggest that reducing PIP₂ binding affinity decreased TRPC6 channel activity without altering its membrane localization. Our study therefore uncovers that the pre-S1 domain/shoulder of TRPC6 makes the critical contribution to PIP₂ binding.

K771 mutation in the distal TRP box alters the effect of PIP₂

As shown in Fig. 2, upon the PIP₂ depletion, R758Q and K781Q/K782Q mutations significantly decrease the inhibition ratio (I_post/I_pre) as compared to TRPC6_WT. These residues are located within the distal TRP box, the importance to the thermal sensitivity of which site was shown in earlier studies of PIP₂-mediated regulation of TRPV1 channels^36,37, but lacking such information for TRPC channels. We therefore questioned whether the other basic residues located between R758 and K781/K782 might affect the effects of PIP₂ depletion. Unexpectedly, when K771, an evolutionary highly conserved residue, was mutated to glutamine, the resultant OAG-induced currents were potentiated after activation of DrVSP (I_post/I_pre = 1.24 ± 0.13 n = 7, Fig. 7A–C). This potentiation was observed repeatedly during the recording period (Fig. 7B). By contrast, no potentiation of K771Q-mediated currents was seen when the inactive DrVSP was co-expressed (Fig. 7D, black circles). This eliminates the possibility of a gain-of-function effect on the voltage-dependent activation.

Moreover, using the rapamycin-inducible FKBP12-Inp54p system^40,41, which also depletes PIP₂ by the membrane recruiting of specific inositol 5-phosphatase, the potentiation of TRPC6 currents still occurred (Fig. 7E,G). To explore if this reversed effect of PIP₂ depletion can be generalized to other TRPC channels, we tested the effect of neutralization on the equivalent residue (K716) in TRPC7. As shown in Fig. 7F, the TRPC7 K716Q mutant exhibited a potentiated channel activity upon PIP₂ depletion. The potentiation ratio for TRPC7_K716Q was 1.28 ± 0.06, which is nearly identical to that of TRPC6_K771Q. These results suggest that certain basic residues in the distal C-terminal region (TRP box) act to control the polarity of PIP₂-dependent effects on TRPC6/7.

Discussion

PIP₂ is known to regulate most TRP channels⁴². Earlier studies showed that reductions of PIP₂, either through dephosphorylation by VSP or hydrolysis by PLC, suppress TRPC channel activity^{8,10,12,15,42}. PIP₂ has three phosphate groups, and their negative charges likely form an interactive network through electrostatic interactions or salt bridges with basic residues in target molecules⁴³. Up to now, however, no PIP₂ interaction site for TRPC channel’s activity had yet been determined.

TRPC6 contains over 400 basic residues in the tetrameric complex. In the present study, we introduced neutralizing mutations to various selected basic residues. As a result, 11 of the 29 mutated channels exhibited accelerated deactivation (decay, t_1/2) upon VSP activation, while 12 of the 29 channels showed altered reactivation (recovery, τ). In total, 17 mutated channels exhibited altered deactivation or reactivation kinetics, though only the R437Q and K442Q mutants showed significant effects on both deactivation and reactivation to reduce the effective PIP₂ binding affinity. In previous reports, we have determined decay and recovery constants of TRPC6 and TRPC7 channels⁸. Based on this data, the ratio of recovery (τ)/decay (t_1/2) showed good agreement with the rank order of K_d values for PIP₂-TRPC6 and PIP₂-TRPC7 channels, which are 2 and 5 μM, respectively¹⁵. In current report, the kinetic ratio of TRPC6_WT was 10.01, and R437Q and K442Q were 47.76 and 77.13, respectively (Fig. 3I). Thus, dissociation constants of R437 and K442 to PIP₂ could be decreased to 5–8 folds. However, this estimation can only be effective under a limited condition, where the channel gating kinetics is equal among the respective mutants. Therefore, although it is difficult to obtain the exact dissociation constant, we could still compare relative rank order for PIP₂ affinity, and emphasize the importance of pre-S1 domain for PIP₂ binding. This pre-S1 domain in TRP channels has received particular attention because of the abundance of electrophilic amino acids and its structural proximity to the TRP box^{44,45,46,47,48,49}. The pre-S1 domain contains the pre-S1 elbow and pre-S1 shoulder, the latter of which forms an amphipathic short helix along the inner leaflet of the cell membrane (Fig. 4A). Within this helix, side chains of basic residues, including K431, K434, R437 and K442, extend into the cytoplasm, making them suitable for PIP₂ binding. The docking simulation also supports the importance of basic residues in the pre-S1 shoulder. Nevertheless, an additional pre-S1 shoulder mutation in cysteine at 429 to serine (C429S) did not cause any difference in the docking score as expected, but the VSP-based experimental results indicated the acceleration of PIP₂ dissociation kinetics (Fig. 4C). These inconsistent observations may be due to more direct exposure of the pre-S1 domain to the solvent caused by Cys to Ser mutation, which should greatly affect the recognition of PIP₂⁵⁰.

We also observed that recovery kinetics was slowed in the H358Y/R360Q mutant (Fig. 2). Both H358 and R360 are located within the linkage domain between the ARD and the pre-S1, and are facing basic residues in the pre-S1 shoulder (Fig. 4B). These residues likely facilitate the interaction between PIP₂ and the pre-S1 shoulder, as supported by the result of docking simulation. In the receptor activation experiments, the current density was lower with the H358Y/R360Q mutant than TRPC6_WT (Fig. 5B).This highlights the idea that the positively charged H358 and R360 cooperatively contribute to PIP₂ binding with pre-S1 domain/shoulder. This may be consistent with a recent structural study showing that in TRPM8 channels, the head group of PIP₂ is situated within a pocket coordinated by the pre-S1, TRP box, the linker region between the pre-S1 and the TRPM homology region (MHR) domain⁴⁹. This TRPM8-PIP₂ binding may be similar to TRPP channels where pre-S1 and TRP box-like domain coupling supports the negative effect of PIP₂⁴⁸. However, in TRPM8 and TRPV5, the S4-S5 segment also forms part of the PIP₂ binding site^47,49. This does not seem to be the case in TRPC6, as the mutation (R629Q) in this segment had faster inhibition, but also showed faster recovery from DrVSP inhibition (Fig. 2). This residue, and the other untested positive charge in this region seems to be not close enough to the pre-S1 PIP₂ interacting residues. Consequently, the contribution of the pre-S1 domain/shoulder may be greater than in TRPM8 and TRPV5. Thus, we assume that in TRPML and PI(3,5)P₂ structures, the location of the PIP₂ near pre-S1 (though missing in the structure) is close to TRPC6 channel⁴⁷. To summarize this mutational study in the structural aspect, we have mapped residues either only for decay (t_1/2) or recovery (τ) and both are altered in the TRPC6 channel structure (Supplemental Fig. 1). This mapping reveals that basic residues locate near the inner leaflet contribute to decay (t_1/2). Contrary to these, residues contributing for recovery (τ) locate in the intracellular domain. These observations indicate that basic residues near the transmembrane would retain PIP₂ binding, and those in the intracellular region of the channel would attract PIP_2.

To investigate the pathophysiological importance of PIP₂ binding to the pre-S1 domain/shoulder, we assessed its contribution to Ca²⁺-dependent inactivation of the TRPC channel. It has been argued that the dissociation of PIP₂ could be involved in the desensitization or inactivation of TRP channels⁵¹. The inactivation kinetics seen with the R437Q and R437/K442Q mutant is not very different from the wild-type (Fig. 5C). Thus, we guess that inactivation kinetics is more directly controlled by feedback of intracellular Ca²⁺ or direct modulation via Ca²⁺ bound calmodulin to this channel but not via PIP₂ interaction with the pre-S1domain/shoulder.

Another important issue is the binding mechanism of PIP₂ and DAG in TRPC3/6/7 channels. These lipids essentially share the same acyl-chain structure, though neither the proximity nor the cross-reactivity of PIP₂ and DAG binding to these TRPC channel has been determined. Our study suggests that PIP₂ and DAG bind at independent sites, which is consistent with the dose–response curves obtained with the pre-S1 mutants in the present study. In fact, a unique DAG recognition site has recently been identified in the pore domain of TRPC3 channel⁵². These observations indicate that DAG-activated TRPC3/C6/C7 channels are accessible to both PIP₂ and DAG within a single channel complex.

We also found that K771Q mutation completely reverses the inhibitory effect of PIP₂ depletion without significantly affecting PIP₂ binding, as indicated by the absence of a significant change in the value of decay (t_1/2) and recovery (τ) from the VSP experiments (Fig. 7C). This observation implies the existence of a polarity switch for the PIP₂ binding effect. Similar to the K771Q mutation, the R758Q mutation and K781Q/K782Q double mutation also altered the effect of PIP₂ depletion (Fig. 2, bottom). Using Swiss-model (https://swissmodel.expasy.org), we have created a model of human TRPC6 based on TRPC4 (PDBID: 7B1G), in which residues equivalent to K771, K781 and K782 are resolved. According to this modeled structure, the three residues are located very close to the pre-S1 PIP₂ interacting residues, most likely forming a part of the binding site (Supplemental Fig. 2). However, for K771, because this residue is positioned at a flexible outer skirt region, it is not likely to be a sole determinant of PIP₂ binding or switching polarity yet. These findings shed light on functionality of PIP₂, apparently localized between R758 and K782 in TRPC6/7.

This switching mechanism for PIP₂ may be physiological relevant in vascular native cells, where the depletion of PIP₂ effect enhances TRPC6-like currents^3,53. As previously revealed through R721A mutation of TRPV1, a basic residue in the distal TRP box modifies their selectivity from PIP₂ to PIP for maintaining capsaicin-activated currents²⁷. This suggests that a similar effector mechanism underlies the actions of PIP_s in both TRPV1 and TRPC6/7 channels. Understanding how the pre-S1 domain/shoulder and the distal TRP domain orchestrate PIP₂ regulation is an intriguing mechanism for future study.

Our present findings summarized above provide new insights into the molecular interactions between PIP₂ and TRPC6 channel. Specifically, the pre-S1 domain/shoulder is a critical segment for PIP₂ interaction that probably acts in concert with the TRP box and linker segment between the ARD and pre-S1. The results also reveal the switching of the mode of PIP₂ binding action that can be made by single point mutation in the basic amino acid residues in the distal TRP box. These provide inevitable structural and functional insights into the biochemical and physiological mechanisms governing TRP channel activity.

Materials and methods

Plasmids and cells

A pcDNA3 expression vector encoding human TRPC6 (accession number: NM_004621) was provided by Dr. Thomas Hofmann (Institut für Pharmakologie und Toxikologie) and was transferred into a kanamycin resistant pIRES2 vector (excluded eGFP region). cDNA encoding hTRPC7 was cloned from the Human Brain Library (Invitrogen) and inserted into the pIRES2 vector. Single amino acid mutations in TRPC6 and TRPC7 were generated using a QuikChange Site-Directed Mutagenesis kit (Stratagene) according to the manufacturer’s instructions. Danio rerio VSP (DrVSP) and its inactive mutant in pIRES-eGFP vector (Invitrogen) were identical to those described previously^24,25. To detect PIP₂, we used PIP₂ sensor molecules consisting of monomeric superenhanced CFP or YFP fused with a PH domain (CFPmse-PHd and YFPmse-PHd, respectively)¹⁵. Muscarinic type-I (M₁R) receptor was provided by Dr. Tatsuya Haga. All PCR products were sequenced entirely.

HEK293 cells (ATCC) were maintained in Dulbecco’s modified Eagle’s medium (DMEM; Invitrogen) supplemented with 10% FBS (Gibco) and antibiotics (penicillin and streptomycin; Gibco) at 37 °C (5% CO₂). For transfection, the cells were seeded onto poly-L-lysine-coated glass coverslips (Matsunami) in 35-mm culture dishes and transfected with a mixture of plasmid vector-incorporated DNAs using SuperFect transfection reagent (Qiagen). To screen for PIP₂ binding sites, plasmids encoding TRPC6 and DrVSP were mixed at a 2:1 molar ratio.

Electrophysiology

The pipette solution for whole-cell recording contained (mM): 120 CsOH, 120 aspartate, 20 CsCl, 2 MgCl₂, 5 EGTA, 1.5 CaCl₂, 10 HEPES, 2 ATP-Na₂, 0.1 GTP, 10 glucose (pH 7.2, adjusted with Tris base; 285–290 mOsm, adjusted with glucose). For measurement of Ca²⁺-dependent inactivation of mutant channels in Fig. 5C, 5 EGTA and 1.5 CaCl₂ in the pipette solution were replaced to 1 EGTA and 0.3 CaCl₂ to reduce Ca²⁺ buffering capacity. The standard external solution contained (mM): 140 NaCl, 5 KCl, 1 CaCl₂, 1.2 MgCl₂, 10 HEPES, 10 glucose (pH 7.4, adjusted with Tris base; 300 mOsm, adjusted with glucose). 100 μM DIDS (4,4-diisothiocyanostilbene-2,2-disulphonic acid, 2Na, purchased from Calbiochem) was applied in the external solutions when needed. Stock solutions for 1-o-oleoyl-2-acetyl-sn-glycerol (OAG, Cayman chemical) and RHC80267 (Calbiochem) were dissolved in dimethyl sulfoxide (DMSO, Wako) at concentrations more than 1000-fold higher than used in the experiments. Carbachol (Sigma-Aldrich) was diluted in the standard external solution from its stock (100 mM) in H₂O. The cells were continuously gravity perfused with the external solution at a flow rate of 1 mL/min. Perfusion was turned on and off using electromagnetic solenoid microvalves (Takasago Electric). The currents were recorded at a holding potential of − 50 mV. After applying 50 μM OAG in bath solution, TRPC6 currents were gradually increased and reached the peak amplitudes around 30–120 s. During this recording, strong depolarizing pulse (+ 100 mV, 700 ms) was applied every 15–25 s. For measurements of effect of PIP₂ depletion and replenishment, several pulses (2–4) near the peak amplitudes were averaged as for the data of the individual cells.

FRET detection

Fluorescence from voltage-clamped cells was detected using a Nikon TE300 Eclipse microscope (60×, 0.9 N.A. objective) equipped with a beam-splitter (Dual-View2, Photometrics) and an EMCCD camera (Evolve512, Photometrics). Excitation light filtered at 427/10 nm and 504/12 nm was alternately introduced via an optical fiber from a lamp-house equipped with a high-speed excitation wavelength selector (75 W xenon lamp, OSP-EXA; Olympus). Optical filters were obtained from Semrock except for the splitter (Chroma). The duration of camera exposures was 100 ms occurring within 150-ms periods of illumination at each excitation wavelength. Images were captured with an EM gain of 300 and then digitized as 512 × 512 pixels in 16-bit arrays by the microscope software (Micro-manager v.1.4). Averaged intensities from the whole-cell region (typically 20 × 20 to 40 × 40 square pixels) were analyzed to calculate FRET using a custom-written MATLAB program. Finally, the FRET ratio (FR) was calculated as described previously⁵⁴.

Docking simulation

The TRPC6 (PDBID: 6UZ8)-PIP₂ docking process was simulated using AutoDock 4.2⁵⁵. A 15 × 15 × 15 Å (x, y, and z) grid box was centered on the binding pocket with 0.375 nm spacing for each dimension. AutoGrid 4.2 was used to prepare grid maps. Docking parameters were set as follows: exhaustiveness = 8, num_modes = 100, and energy_range = 4. Other parameters were set at their default value.

Confocal microscopy

HEK293 cells were transfected with wild-type TRPC6 (TRPC6_WT) fused with CFP on the N-terminal side (CFP-TRPC6_WT) and YFP-PHd or CFP-TRPC6_mutant/YFP-PHd. After 48 h, the cells were seeded into a glass-bottomed culture dish (MatTek) and incubated for more than 3 h prior to imaging. Fluorescence and optical images were obtained using an inverted confocal microscope (LSM830, Carl Zeiss) equipped with a 63× oil objective lens (1.25 N.A.) at 1024 × 1024 resolution with a pixel dwell time of 6.4–25.4 µs. During confocal microscopy, the cells were bathed in the standard external solution identical to that used for the electrophysiological experiments.

Barium influx for dose–response to OAG

Ba²⁺ influx was measured as described in previous report⁵⁶. Briefly, transfected HEK293 cells were seeded onto glass coverslips and allowed to attach for 3 h. The coverslips were then incubated with 1 μM Fura-2/AM for 30 min and transferred to a custom-made glass bottom chamber apparatus on an inverted microscope stage (IX-83, Olympus). The solution flow rate was set to 1 ml/min using gravity-fed system. The bath solution had an identical composition used in the electrophysiological experiments only by the replacement of external calcium with barium. Cells exhibiting YFP fluorescence were selected for measurement. Fluorescence images of the cells were captured with a CCD camera (EXi Blue, QImaging) and recorded with a software (cellSens, Olympus). The 340/380 nm Fura-2 ratios from images were obtained on a pixel-by-pixel basis every 5 s for a total of 400 s. OAG (0.01–100 μM) was applied at 120 s of after the recording up to 420 s. Ratiometric F₃₄₀/F₃₈₀ values during the imaging were calculated by averaging the ROI intensity and analyzed by a custom written program in MATLAB (Mathworks). Delta Fura-2 ratio (ΔF₃₄₀/F₃₈₀) was calculated from the ratio of the maximal response minus the basal level.

Data analysis

Electrophysiological data were analyzed using MATLAB and Excel (Microsoft). The recovery (τ) of the TRPC6 inward current (I) was determined with least-squares fits of the exponential as follows: I = I₀ + A·exp(− t/τ), where I₀ is the current size after repolarization, A is a scale factor, and τ (recovery) is the reactivation kinetics of the PIP₂ binding. The value of t_1/2 of the TRPC6 outward current decay was determined by fitting with a logarithmic equation as follows: I = I_min + I_d/(1 + exp[(t − t_1/2)/f_s]), where I_min is the minimum current induced by the inhibition, I_d is the inhibited current amplitude, f_s is a slope factor, and t_1/2 is a measure of the deactivation kinetics of the PIP₂ dissociation. For confocal imaging, data were analyzed using the colocalization function in Fiji (National Institutes of Health).

Data are presented as means ± SEM. Statistical significance were evaluated using analysis of variance (ANOVA) with Tukey’s and Dunnett’s post-hoc tests for single and multiple comparisons, respectively.

Data availability

The datasets generated during and/or analysed during the current study are available from the corresponding author on reasonable request.

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Acknowledgements

We thank Dr. Thomas Hofmann for providing TRPC6 plasmid, Dr. Tatsuya Haga for providing M₁R plasmid, and Dr. Kees Jalink for fluorophore-tagged PH domain plasmids. We appreciate technical support of the Research Support Center at the UOEH. This work was supported by KAKENHI grant 26460296 (M.X.Mori), Institute for Molecular Science IMS program (21-IMS-C220), and MEXT Promotion of Distinctive Joint Research Center Program #JPMXP 0621467946.

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Authors and Affiliations

Laboratory of Biomaterials and Chemistry, School of Medicine, University of Occupational and Environmental Health, 1-1, Iseigaoka, Yahatanishi-ku, Kitakyushu, Fukuoka, 807-8555, Japan
Masayuki X. Mori, Ryo Okada & Reiko Sakaguchi
Human Information and Life Sciences, School of Health Sciences, University of Occupational and Environmental Health, Kitakyushu, Fukuoka, Japan
Ryo Okada
Laboratory of Molecular Biology, Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan
Masayuki X. Mori, Hideharu Hase, Onur K. Polat & Yasuo Mori
Department of Physiology, School of Medicine, Fukuoka University, Fukuoka, Japan
Yuko Imai & Ryuji Inoue
Exploratory Research Center on Life and Living Systems/Institute for Molecular Science, National Institutes of Natural Sciences, Okazaki, Aichi, Japan
Satoru G. Itoh & Hisashi Okumura
Department of Structural Molecular Science, SOKENDAI (The Graduate University for Advanced Studies), Okazaki, Aichi, Japan
Satoru G. Itoh & Hisashi Okumura
Laboratory of Integrative Physiology, Department of Physiology, Graduate School of Medicine, Osaka University, Suita, Osaka, Japan
Yasushi Okamura

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Contributions

R.O., R.S., H.H., O.K.P., Y.I. and M.X.M. performed experiment and analyzed the data. M.X.M. designed the experiments. R.O., R.S., R.I., and M.X.M. wrote the paper. S.G.I., H.O., Y.O., and Y.M. provided technical advice.

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Correspondence to Masayuki X. Mori.

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Mori, M.X., Okada, R., Sakaguchi, R. et al. Critical contributions of pre-S1 shoulder and distal TRP box in DAG-activated TRPC6 channel by PIP₂ regulation. Sci Rep 12, 10766 (2022). https://doi.org/10.1038/s41598-022-14766-x

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Received: 14 April 2022
Accepted: 13 June 2022
Published: 24 June 2022
DOI: https://doi.org/10.1038/s41598-022-14766-x

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