Advertisement

Functional Coupling between TRPC3 and RyR1 Regulates the Expressions of Key Triadic Proteins*

  • Eun Hui Lee
    Affiliations
    Laboratory of Calcium Communication, Department of Physiology, College of Medicine, The Catholic University of Korea, Seoul 137-701, Korea

    Department of Anesthesiology, Perioperative and Pain Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts 02115
    Search for articles by this author
  • Gennady Cherednichenko
    Affiliations
    Department of Molecular Biosciences and Center for Children's Environmental Health, University of California, Davis, Davis, California 95616
    Search for articles by this author
  • Isaac N. Pessah
    Affiliations
    Department of Molecular Biosciences and Center for Children's Environmental Health, University of California, Davis, Davis, California 95616
    Search for articles by this author
  • P.D. Allen
    Correspondence
    To whom correspondence should be addressed. Tel.: 617-732-6881; Fax: 617-732-6927;
    Affiliations
    Department of Anesthesiology, Perioperative and Pain Medicine, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts 02115
    Search for articles by this author
  • Author Footnotes
    * This work was supported by grants from the National Institutes of Health (P01AR17605) (to P. D. A. and I. N. P.), the Catholic University of Korea (to E. H. L.), and the Medical Research Center, Korea Science and Engineering Foundation, Republic of Korea (R13-2002-005-01002–0). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
    The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. S1–S3.
Open AccessPublished:February 16, 2006DOI:https://doi.org/10.1074/jbc.M600981200
      We have shown that TRPC3 (transient receptor potential channel canonical type 3) is sharply up-regulated during the early part of myotube differentiation and remains elevated in mature myotubes compared with myoblasts. To examine its functional roles in muscle, TRPC3 was “knocked down” in mouse primary skeletal myoblasts using retroviral-delivered small interference RNAs and single cell cloning. TRPC3 knockdown myoblasts (97.6 ± 1.9% reduction in mRNA) were differentiated into myotubes (TRPC3 KD) and subjected to functional and biochemical assays. By measuring rates of Mn2+ influx with Fura-2 and Ca2+ transients with Fluo-4, we found that neither excitation-coupled Ca2+ entry nor thapsigargin-induced store-operated Ca2+ entry was significantly altered in TRPC3 KD, indicating that expression of TRPC3 is not required for engaging either Ca2+ entry mechanism. In Ca2+ imaging experiments, the gain of excitation-contraction coupling and the amplitude of the Ca2+ release seen after direct RyR1 activation with caffeine was significantly reduced in TRPC3 KD. The decreased gain appears to be due to a decrease in RyR1 Ca2+ release channel activity, because sarcoplasmic reticulum (SR) Ca2+ content was not different between TRPC3 KD and wild-type myotubes. Immunoblot analysis demonstrated that TRPC1, calsequestrin, triadin, and junctophilin 1 were up-regulated (1.46 ± 1.91-, 1.42 ± 0.08-, 2.99 ± 0.32-, and 1.91 ± 0.26-fold, respectively) in TRPC3 KD. Based on these data, we conclude that expression of TRPC3 is tightly regulated during muscle cell differentiation and propose that functional interaction between TRPC3 and RyR1 may regulate the gain of SR Ca2+ release independent of SR Ca2+ load.
      TRPCs are one family of transient receptor potential (TRP)
      The abbreviations used are: TRPC3, transient receptor potential channel canonical type 3; TRPC3 KD, TRPC3 knockdown; RyR, ryanodine receptor; DHPR, dihydropyridine receptor; IP3R, inositol 1,4,5-trisphosphate receptor; SOCE, store-operated calcium entry; ECCE, excitation-coupling calcium entry; EC coupling, excitation-contraction coupling; SR, sarcoplasmic reticulum; JP1, junctophilin type 1; CSQ, calsequestrin; HEK, human embryonic kidney.
      2The abbreviations used are: TRPC3, transient receptor potential channel canonical type 3; TRPC3 KD, TRPC3 knockdown; RyR, ryanodine receptor; DHPR, dihydropyridine receptor; IP3R, inositol 1,4,5-trisphosphate receptor; SOCE, store-operated calcium entry; ECCE, excitation-coupling calcium entry; EC coupling, excitation-contraction coupling; SR, sarcoplasmic reticulum; JP1, junctophilin type 1; CSQ, calsequestrin; HEK, human embryonic kidney.
      cation channels. This entire group of channels has been predicted to consist of six transmembrane segment channels that allow the entry of Ca2+ and Na+ into the cell (
      • Ramsey I.S.
      • Delling M.
      • Clapham D.E.
      ,
      • Pedersen S.F.
      • Owsianik G.
      • Nilius B.
      ,
      • Freichel M.
      • Vennekens R.
      • Olausson J.
      • Stolz S.
      • Philipp S.E.
      • Weissgerber P.
      • Flockerzi V.
      ). One of the TRPC subtypes, TRPC3, is highly expressed in the brain, skeletal muscle, and cardiac muscle (
      • Clapham D.E.
      ,
      • Hofmann T.
      • Schaefer M.
      • Schultz G.
      • Gudermann T.
      ,
      • Vandebrouck C.
      • Martin D.
      • Colson-Van Schoor M.
      • Debaix H.
      • Gailly P.
      ). It has been reported that TRPC3-mediated Ca2+ entry can be induced by two different mechanisms: 1) direct activation by binding of exogenous organic molecules or endogenous metabolites such as diacylglycerol analogues (
      • Hofmann T.
      • Obukhov A.G.
      • Schaefer M.
      • Harteneck C.
      • Gudermann T.
      • Schultz G.
      ), and 2) phospholipase C-mediated activation via phospholipase C-coupled receptor. The latter mechanism can be subdivided into two different submechanisms. One submechanism is store-operated Ca2+ entry (SOCE) (
      • Li H.S.
      • Xu X.Z.
      • Montell C.
      ,
      • Vazquez G.
      • Lievremont J.P.
      • St. J. Bird G.
      • Putney Jr., J.W.
      ,
      • Yildirim E.
      • Kawasaki B.T.
      • Birnbaumer L.
      ) (for example, when Ca2+ store depletion through IP3R by phospholipase C pathway triggers TRPC3 activation). The second submechanism is receptor-operated Ca2+ entry (
      • Okada T.
      • Inoue R.
      • Yamazaki K.
      • Maeda A.
      • Kurosaki T.
      • Yamakuni T.
      • Tanaka I.
      • Shimizu S.
      • Ikenaka K.
      • Imoto K.
      • Mori Y.
      ,
      • Zhu X.
      • Jiang M.
      • Birnbaumer L.
      ,
      • Philipp S.
      • Strauss B.
      • Hirnet D.
      • Wissenbach U.
      • Mery L.
      • Flockerzi V.
      • Hoth M.
      ) (for example, in B lymphocytes, when TRPC3 is activated by physical coupling with phospholipase C γ2 and is responsible for the secondary intracellular Ca2+ entry after B-cell receptor activation) (
      • Nishida M.
      • Sugimoto K.
      • Hara Y.
      • Mori E.
      • Morii T.
      • Kurosaki T.
      • Mori Y.
      ).
      Physiological evidence has been demonstrated for the involvement of TRPC3 in many processes. For example, TRPC3 activation via phospholipase C γ-IP3R pathway is essential for brain-derived neurotrophic factor-dependent growth cone guidance in cerebellar granule neurons (
      • Li Y.
      • Jia Y.C.
      • Cui K.
      • Li N.
      • Zheng Z.Y.
      • Wang Y.Z.
      • Yuan X.B.
      ). In arterial smooth muscle cells, TRPC3 is activated by purinergic receptors, and then Ca2+ influx through TRPC3 induces depolarization and vasoconstriction (
      • Reading S.A.
      • Earley S.
      • Waldron B.J.
      • Welsh D.G.
      • Brayden J.E.
      ). Using heterologous expression studies in HEK 293 cells, it has been shown that TRPC3 physically interacts with cytosolic Ca2+ signaling proteins such as IP3R, where it competes with calmodulin for the same binding site on IP3R (
      • Zhang Z.
      • Tang J.
      • Tikunova S.
      • Johnson J.D.
      • Chen Z.
      • Qin N.
      • Dietrich A.
      • Stefani E.
      • Birnbaumer L.
      • Zhu M.X.
      ). In addition, it has been reported that TRPC3 interacts with ryanodine receptors (RyRs) (
      • Kiselyov K.I.
      • Shin D.M.
      • Wang Y.
      • Pessah I.N.
      • Allen P.D.
      • Muallem S.
      ,
      • Lee E.H.
      • Allen P.D.
      ); however, the functional roles of TRPC3 in skeletal muscle have not been well addressed.
      RyRs function as Ca2+ release channels in the sarcoplasmic reticulum (SR) and are essential proteins for excitation-contraction (EC) coupling in striated muscles (
      • Brady A.J.
      ,
      • Sandow A.
      ). RyRs have been shown to have bidirectional communication with a second essential protein in EC coupling, the dihydropyridine receptor (DHPR), which acts as both a sarcolemmal L-type voltage-gated Ca2+ channel (
      • Brady A.J.
      ) and the mechanical trigger for RyR activation (
      • Sandow A.
      ). During EC coupling, as a result of membrane depolarization, Ca2+ influx through DHPR or conformational changes of DHPR via its gating charge movement activates RyR2 in cardiac muscle or RyR1 in skeletal muscle respectively. The Ca2+ stored in SR is then released through RyRs into the cytosol to cause muscle contraction. Recently, functional coupling between RyR and TRPC1 (
      • Sampieri A.
      • Diaz-Munoz M.
      • Antaramian A.
      • Vaca L.
      ) or TRPC3 (
      • Kiselyov K.I.
      • Shin D.M.
      • Wang Y.
      • Pessah I.N.
      • Allen P.D.
      • Muallem S.
      ) has been shown in heterologous expression systems such as Chinese hamster ovary and HEK cells. In the case of TRPC1, it has been proposed that the functional interaction between RyR1 and TRPC1 is a physical interaction with an undefined region in the cytoplasmic foot region of RyR1 (
      • Sampieri A.
      • Diaz-Munoz M.
      • Antaramian A.
      • Vaca L.
      ). Furthermore, involvement of TRPCs has been reported in the abnormal calcium influx observed in muscle disease models such as dystrophic mouse skeletal muscle fibers (
      • Vandebrouck C.
      • Martin D.
      • Colson-Van Schoor M.
      • Debaix H.
      • Gailly P.
      ). Therefore, although in skeletal muscle it is possible for EC coupling to occur in the absence of extracellular Ca2+ (
      • Sandow A.
      ), the possible involvement of extracellular Ca2+ influx and the interactions of RyR1 with Ca2+ ion-conducting channel proteins in sarcolemma (other than the DHPR) during EC coupling still remain to be elucidated. In the current study, we have defined the role of TRPC3 during EC coupling and muscle contraction in primary mouse skeletal muscle myoblasts/myotubes in which TRPC3 mRNA and protein expression was reduced (>97 and >94%, respectively) by using small interference RNA.

      EXPERIMENTAL PROCEDURES

      Materials—Fetal bovine serum, cell culture media, trypsin, l-glutamine, penicillin/streptomycin, and basic fibroblast growth factor were obtained from Invitrogen. Caffeine, KCl, ryanodine, thapsigargin, cyclopiazonic acid, and heat-inactivated horse serum were obtained from Sigma-Aldrich. Monoclonal anti-RyR1 antibody (34C) was provided by Drs. J. Airey and J. Sutko (Developmental Studies Hybridoma Bank, Iowa City, IA). Anti-TRPC3 antibody was obtained from Alomone Labs (Jerusalem, Israel). Anti-DHPR, sarcoplasmic reticulum calcium ATPase (SERCA), triadin, and calsequestrin antibodies were obtained from Affinity BioReagents (Golden, CO). Anti-junctophilin 1 antibody was a kind gift from Dr. Jianjie Ma. Anti-calmodulin, FKBP12, and α-tubulin were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). G-Sepharose beads were obtained from Amersham Biosciences. Collagen was purchased from Upstate Biotechnology (Lake Placid, NY). FuGENE transfection reagent was obtained from Roche Diagnostics.
      Cell Cultures—The method used to derive primary wild-type myoblasts from mouse skeletal muscle was previously described (
      • Felder E.
      • Protasi F.
      • Hirsch R.
      • Franzini-Armstrong C.
      • Allen P.D.
      ,
      • Moore R.A.
      • Nguyen H.
      • Galceran J.
      • Pessah I.N.
      • Allen P.D.
      ,
      • Lee E.H.
      • Lopez J.R.
      • Li J.
      • Protasi F.
      • Pessah I.N.
      • Kim D.H.
      • Allen P.D.
      ). After ∼48 h in growth medium, myoblasts were replated either on 10-cm dishes coated with collagen to prepare cell lysates or on 96-well plates coated with collagen for Ca2+ imaging and Mn2+ quench experiments. When cells reached ∼70% confluence, growth medium was replaced with differentiation medium (containing 5% heat-inactivated horse serum and no growth factors) and placed into an 18% CO2 incubator to induce differentiation. HEK293 cells were grown at 37 °C in a 5% CO2 incubator in high glucose Dulbecco's modified Eagle's medium with 10% fetal bovine serum, 100 units/ml of penicillin, and 100 μg/ml of streptomycin.
      Creation of TRPC3 Knockdown Primary Myoblasts—To knock down the mRNA of TRPC3 in primary myoblasts, vectors expressing short hairpin RNA sequences were used. First, two different sequences were selected using a program from Dharmacon siDESIGN center (Dharmacon, Chicago, IL) based on the cDNA sequence of TRPC3 (GenBank™ accession number NM_019510). BLAST searches confirmed that the selected oligonucleotide sequences did not possess homology to any other genes. Each of two 19-nucleotide sequences (Fig. 1A, Sequences I and II) was inserted into a retroviral vector (pSIREN-RetroQ, Clontech Laboratories, Mountain View, CA). Retroviral particles were packaged by transfecting each short hairpin RNA-expressing vector into HEK293-based packaging cells with FuGENE transfection reagent, and the harvested supernatant was filtered with 0.2 μm non-pyrogenic disc filters (Pall Corporation, Ann Arbor, MI). The filter-through (TRPC3 KD retroviruses I and II) was stored at –70 °C before use. Primary mouse skeletal muscle myoblasts were incubated for 3 h with both TRPC3 KD retroviruses I and II and polybrene (8 μg/ml) on 3 successive days and clonally selected using puromycin (0.3 μg/ml). Each selected clone was expanded and differentiated into myotubes described in “Cell Cultures.”
      Figure thumbnail gr1
      FIGURE 1Deriving a stable cell line (TRPC3 KD) expressing hairpin small interference RNA to target the mRNA of TRPC3. A, to interfere the mRNA of TRPC3, each palindromic double array of 19 base pairs (indicated by dots in Sequences I and II) that were complementary sequences to parts of TRPC3 mRNA was inserted to a type of retroviral vectors using 5′-BamH I and 3′-EcoR I sites. Vector map was adapted from the web site of Clontech Laboratories. PPGK, Pu6, and Psv40, PGK, human U6, and SV40 promoters; Puror or Ampr, puromycin or ampicillin resistance; 5-LTR CMV/MSV, mouse cytomegalovirus type I and sarcoma virus hybrid promoter; 3-LTR, 3′-MoMuLV LTR with poly(A) region; SV40 ori and ColE1 ori, replication initiation sites; Ψ, extended packaging signal. B, real-time PCR analysis of cDNA that was prepared from total mRNA isolated from each myotube clone. Clone numbers 1–5, 1–6, and 1–21 indicated by asterisks showed a >90% reduction in the mRNA level of TRPC3 compared with that of wild-type myotubes. Clone number 2–21 (TRPC3 KD) showed the most dramatic decrease (>97%) and was used for all further studies. The data are the mean ± S.E. of two duplicated independent experiments. C, solubilized cell lysate from TRPC3 KD myotubes was subjected to immunoblot analysis with anti-TRPC3 antibody. The expression of TRPC3 in TRPC3 KD myotubes was reduced >94%. The data are the mean ± S.E. of three independent experiments.
      Total RNA from myoblasts and differentiated myotubes of each TRPC3 KD clone was prepared, subjected to RT-PCR to make cDNA, and analyzed by real-time PCR. The PCR mixture contained 12.5 μl of iTaq supermix with ROX (Bio-Rad, Hercules, CA), 0.4 μg of cDNA, and 2 pm of each primer in a 25-μl final volume. All samples were run in duplicate. The reaction conditions were 50 °C for 2 min, 95 °C for 10 min, and 40 cycles of 95 °C for 15 s and 60 °C for 1 min. Amplification and detection were performed with the ABI PRISM Sequence Detection System (AB Applied Biosystems, Foster City, CA).
      Ca2+ Imaging and Mn2+ Quench Experiments—Ca2+ transients (cytosolic Ca2+ amount) and Mn2+ influx were measured as described previously (
      • Lee E.H.
      • Lopez J.R.
      • Li J.
      • Protasi F.
      • Pessah I.N.
      • Kim D.H.
      • Allen P.D.
      ,
      • Fessenden J.D.
      • Wang Y.
      • Moore R.A.
      • Chen S.R.
      • Allen P.D.
      • Pessah I.N.
      ,
      • Clementi E.
      • Scheer H.
      • Zacchetti D.
      • Fasolato C.
      • Pozzan T.
      • Meldolesi J.
      ). Briefly, differentiated primary wild-type and TRPC3 KD myotubes were loaded with Fluo-4 for Ca2+ imaging and Fura-2 for Mn2+ quench experiments (500 μm MnCl2). Dye-loaded myotubes were imaged with an intensified CCD camera with a ×40 objective. Caffeine, KCl, cyclopiazonic acid, ryanodine, and thapsigargin were dissolved in imaging buffer or Me2SO (<0.1%) and applied to myotubes by a 16-channel perfusion pipette (AutoMate Scientific, Berkley, CA). To maintain osmolarity, in the case of KCl (60 mm) application the concentration of Na+ was lowered and the chloride product was maintained.
      Co-immunoprecipitation—Co-immunoprecipitation was done as previously described (
      • Lee E.H.
      • Rho S.H.
      • Kwon S.J.
      • Eom S.H.
      • Allen P.D.
      • Kim D.H.
      ). Briefly, myotubes were homogenized and incubated with a lysis buffer (1% Triton X-100, 10 mm Tris, pH 7.4, 150 mm NaCl, 5 mm EDTA, 1 mm Na3VO4, 10% glycerol, 1 mm phenylmethylsulfonyl fluoride, 20 μg/ml of aprotinin, 12.5 μg/ml of leupeptin, and 20 μg/ml of pepstatin A) to prepare the cell lysate. For triadin, free Ca2+ concentration of the lysis buffer was set at 100 μm by adding 5 mm CaCl2. The cell lysate was then quickly centrifuged to remove insolubilized masses, and the protein concentration of the supernatant was determined. Total protein (500 μg) of each diluted lysate (4×) with Triton X-100-free lysis buffer was incubated with 25 μl of anti-RyR antibody overnight at 4 °C, followed by further incubation with protein G-Sepharose beads for 4 h. Beads were washed three times with the lysis buffer to remove non-specifically bound proteins. The immune complexes were treated with SDS-sample buffer and subjected to SDS-PAGE and immunoblot analysis.
      Statistical Analysis—Results are given as means ± S.E. with the number of experiments mentioned in the figure legends. Significance of the differences was analyzed by the paired or unpaired t-test (GraphPad InStat, v2.04). Differences were considered to be significant when p <0.05. Graphs were prepared using Origin v7.

      RESULTS

      Small Interference RNA Knock Down of TRPC3 Expression—Total mRNA preparations of fourteen different TRPC3 knockdown clones were subjected to RT-PCR and then real-time PCR to examine TRPC3 mRNA levels (Fig. 1B). The most effective clone was clone number 2–21 (with 97.6 ± 1.9% reduction in the mRNA level). Immunoblot analysis with solubilized cell lysate from clone number 2–21 myotubes showed 94.6 ± 0.4% reduction in the protein level (Fig. 1C). This clone was named TRPC3 KD and used for all further functional and biochemical experiments.
      Functional Consequence of TRPC3 Knock Down—We examined cytosolic Ca2+ transients in response to depolarization and direct RyR1 activation in TRPC3 KD myotubes loaded with Fluo-4. Wild-type myotubes were used as a control. To mimic the membrane depolarization of muscle, 60 mm KCl was applied to myotubes for 30 s (Fig. 2A, left). Interestingly, TRPC3 KD myotubes had a 40% decrease in peak amplitude of depolarization-induced Ca2+ release compared with wild-type myotubes (Fig. 2B, normalized peak amplitude of TRPC3 KD myotubes was 0.61 ± 0.05 when the peak amplitude of wild-type myotubes was set to 1, p <0.05), suggesting that the gain of EC coupling was reduced in the absence of TRPC3 expression. Normalized responses compared with those of wild-type myotubes also showed that the transients in TRPC3 KD myotubes had a faster inactivation slope (Fig. 2A, right, dotted lines), which was supported by significantly decreased ratio of residual (at 20 s after KCl application, indicated by a vertical dotted line in panel A, left) versus peak amplitude in TRPC3 KD myotubes (Fig. 2C, 0.35 ± 0.04 (TRPC3 KD versus 0.52 ± 0.05 (wild type)). To rule out the possibility that the main cause of the reduced cytosolic Ca2+ release during EC coupling was due to a reduced amount of SR Ca2+ in TRPC3 KD myotubes, SR Ca2+ content was measured directly by treatment of the myotubes with 10 μm cyclopiazonic acid, an SR Ca2+-ATPase inhibitor; there was no significant difference in the amount of Ca2+ in the SR of TRPC3 KD myotubes compared with that of wild-type myotubes (Fig. 2, D and E).
      Figure thumbnail gr2
      FIGURE 2Decreased cytosolic Ca2+ transients associated with EC coupling in TRPC3 KD myotubes. A, to induce EC coupling, 60 mm KCl was treated to wild-type (black) or TRPC3 KD (gray) myotubes loaded with Fluo-4. TRPC3 KD myotubes showed significantly decreased cytosolic Ca2+ transient during EC coupling (left), suggesting a decreased gain of EC coupling. A representative normalized trace of TRPC3 KD myotubes compared with that of wild-type myotubes is shown on the right. Inactivation slopes were fitted to a linear equation and represented by dotted lines. TRPC3 KD myotubes showed faster inactivation than wild-type myotubes. Histograms are shown for normalized peak amplitude of TRPC3 KD myotubes to that of wild-type myotubes (B) or normalized residual amplitude at 20 s after the peak (vertical dotted line in panel A, left) to its peak amplitude (C). The residual amplitude in TRPC3 KD myotubes was significantly smaller than that of wild-type myotubes. D, SR depletion by treatment with 10 μm cyclopiazonic acid. Depletion traces of wild-type and TRPC3 KD myotubes were colored in black and gray, respectively. E, depletion peak amplitude of TRPC3 KD myotubes was normalized by that of wild-type myotubes. The data are the mean ± S.E. of 115 (Wild type) or 202 (TRPC3 KD) independent experiments. For normalized peak amplitude in panels B and E, peak amplitude of wild-type myotubes was set to 1. For normalized residual amplitude in panel C, each peak amplitude of wild-type or TRPC3 KD myotubes was set to 1. *, significant difference compared with wild type (p <0.05). **, significant difference compared with its peak amplitude (p <0.05).
      We examined several possibilities that could be responsible for the reduced gain of EC coupling. The first hypothesis we tested was that of an inherent impairment of RyR1 function. RyR1 activity was directly measured by challenging the myotubes with a saturating concentration of the channel agonist caffeine (40 mm) and measuring the size of the Ca2+ transient (Fig. 3). We found that the amplitude of the peak cytosolic Ca2+ elicited by caffeine treatment was also significantly lower in TRPC3 KD myotubes compared with wild-type myotubes (Fig. 3A, left, and 3B). However, when a normalized trace of TRPC3 KD myotubes (setting the peak response of wild type to 1) was overlaid with that of wild-type myotubes (Fig. 3A, right), the ratio of residual (at 20 s after caffeine application, indicated by a vertical dotted line in panel A, left) versus peak amplitude was unchanged (Fig. 3C, 0.87 ± 0.07 (TRPC3 KD) versus 0.79 ± 0.08 (Wild type)), suggesting that although the caffeine-induced peak amplitude was significantly decreased, the inactivation kinetics of RyR1 in TRPC3 KD myotubes were similar to those of wild-type myotubes.
      Figure thumbnail gr3
      FIGURE 3Decreased RyR1 activity in TRPC3 KD myotubes. A, wild-type (black) or TRPC3 KD (gray) myotubes loaded with Fluo-4 were exposed to 40 mm caffeine. The peak amplitude of the caffeine response was decreased in TRPC3 KD myotubes (left). A representative normalized trace of TRPC3 KD myotubes compared with that of wild-type myotubes is shown on the right. Inactivation slopes were fitted to linear equation and represented by a dotted line. There was no significant change in inactivation slopes between wild-type and TRPC3 KD myotubes. The histogram for normalized peak amplitudes of TRPC3 KD myotubes by that of wild-type myotubes (B) or normalized residual amplitude at 20 s after the peak (indicated as a vertical dotted line in panel A, left) to its peak amplitude (C) is shown. There was also no significant change in normalized residual amplitudes of wild-type and TRPC3 KD myotubes. The data are the mean ± S.E. of 115 (Wild type) or 202 (TRPC3 KD) independent experiments. For normalized peak amplitude in panel B, peak amplitude of wild-type myotubes was set to 1. For normalized residual amplitude in panel C, each peak amplitude of wild-type or TRPC3 KD myotubes was set to 1. *, significant difference compared with wild-type (p <0.05). **, significant difference compared with its peak amplitude (p <0.05).
      The second hypothesis was that TRPC3 could be the yet to be identified excitation-coupling calcium entry (ECCE) channel (
      • Cherednichenko G.
      • Hurne A.M.
      • Fessenden J.D.
      • Lee E.H.
      • Allen P.D.
      • Beam K.G.
      • Pessah I.N.
      ) and that the decrease in gain and decreased residual ratio after KCl application was secondary to a decrease in depolarization-induced Ca2+ entry. We tested this hypothesis using Mn2+ quench experiments in myotubes loaded with Fura-2 treated with (Fig. 4A) or without (supplemental Fig. S1) 500 μm ryanodine for 30 min to block RyR1 activity. 60 mm KCl was applied to myotubes to mimic membrane depolarization in the presence of extracellular Mn2+, and the rate of Mn2+ quench at the Fura-2 isosbestic point was determined. We found that the initial rate of ECCE in TRPC3 KD myotubes was not significantly different from that of wild-type myotubes (3.30 ± 0.35 (TRPC3 KD) versus 3.44 ± 0.44 (Wild type)), suggesting that TRPC3 is not the ECCE-mediating channel and does not interact with the ECCE-mediating protein(s) or play a role in the three-way communication among RyR1, the DHPR, and the ECCE channel.
      Figure thumbnail gr4
      FIGURE 4Unchanged ECCE and SOCE in TRPC3 KD myotubes. A, to measure Ca2+ influx during excitation (ECCE), Mn2+ influx by the treatment of 60 mm KCl was measured in myotube loaded with Fura-2 with 500 μm ryanodine for 30 min to block RyR1 activity. There was no significant difference in initial rates of influx between wild-type and TRPC3 KD myotubes. The data are the mean ± S.E. of 36 (Wild type) or 14 (TRPC3 KD) independent experiments. B, to measure Ca2+ influx after SR depletion (SOCE), SR of wild-type and TRPC3 KD myotubes loaded with Fluo-4 was depleted of Ca2+ by treatment with 200 nm thapsigargin in the absence of extracellular Ca2+. When the cell would no longer respond to a caffeine or KCl depolarization stimulus, 2 mm Ca2+ was applied to extracellular side (batch), and the rate of Ca2+ entry was determined from the slope of the cytoplasmic Ca2+ transient. There was no significant difference in initial rates of influx between wild-type and TRPC3 KD myotubes; although there was a trend toward a smaller maximal amplitude in TRPC3 KD myotubes, this difference was not significant (p 0.09) (C). The data are the mean ± S.E. of 20 (Wild type) or 17 (TRPC3 KD) independent experiments. Traces from wild-type and TRPC3 KD myotubes are black and gray, respectively. The initial rate of influx was fitted to linear equation and represented by dotted lines.
      Because measurement of SR stores with cyclopiazonic acid (or thapsigargin) is difficult to quantitate, we determined whether the near abolishment of TRPC3 could interfere with SOCE. This was tested in myotubes loaded with Fluo-4 (Fig. 4B) in which SR Ca2+ stores had been previously depleted by the application of 200 nm thapsigargin in the absence of extracellular Ca2+. After total store depletion was confirmed by the lack of any response to KCl or caffeine, 2 mm Ca2+ was applied to the bath and the initial rate of Ca2+ entry was determined. The initial rate of SOCE was not different in TRPC3 KD myotubes compared with that of wild-type myotubes (2.60 ± 0.45 (TRPC3 KD) versus 3.16 ± 0.52 (Wild type)). Although the maximal amplitude of the SOCE transient was reduced in TRPC3 KD myotubes (0.92 ± 0.06 (TRPC3 KD) versus 1.15 ± 0.11 (Wild type)), this difference was not statistically significant (p 0.09) (Fig. 4C).
      The Effects of TRPC3 Knock Down on Muscle Protein Expression—To examine expression profiles of the 3 other TRPCs expressed in muscle and 10 triadic proteins involved in EC coupling in TRPC3 KD myotubes, solubilized cell lysates from wild-type and TRPC3 KD myotubes were subjected to immunoblot analysis (Fig. 5). The results of this analysis showed that TRPC1 was up-regulated (1.46 ± 1.91-fold) in response to the knock down of TRPC3, whereas the expression of TRPC4 and TRPC6 were unchanged (Fig. 5A). Of the 10 triadic proteins examined, 7 showed no change in expression (Fig. 5B). However triadin, junctophilin 1 (JP1), and calsequestrin (CSQ) had significant increases in basal expression levels (2.99 ± 0.32-, 1.91 ± 0.26-, and 1.42 ± 0.08-fold, respectively) associated with the 94% reduction of TRPC3 expression. Interestingly, like TRPC3, all three proteins are known to interact with RyR1 and have been shown to regulate RyR1 activity (
      • Brady A.J.
      ,
      • Sandow A.
      ,
      • Collins J.H.
      • Tarcsafalvi A.
      • Ikemoto N.
      ).
      Figure thumbnail gr5
      FIGURE 5Expression level of three other TRPC isoforms and EC-coupling proteins in TRPC3 KD myotubes. A, solubilized cell lysate from TRPC3 KD myotubes was subjected to immunoblot analysis to examine the expression level of three other TRPC isoforms (TRPC1, TRPC4, and TRPC6) expressed in muscle myotubes. There were no changes in TRPC4 and TRPC6. However, expression level of TRPC1 was significantly increased. B, expression levels of 10 EC-coupling proteins were also examined with α-tubulin as a control. Expression levels of triadin, JP1, and CSQ were significantly increased. C, the three proteins showing increased expression in panel A were subjected to co-immunoprecipitation with RyR1 using anti-RyR1 antibody (34C). More triadin and JP1 than CSQ were co-immunoprecipitated with RyR1 in TRPC3 KD myotubes. *, significant difference compared with wild type (p <0.05). The data are the mean ± S.E. of three independent experiments.
      Protein-Protein Interactions Determined by Co-immunoprecipitation—Co-immunoprecipitation experiments were performed with 34C (anti-RyR1 antibody) to examine whether there was any change in the amount of proteins interacting with RyR1 associated with the knock down of TRPC3 (Fig. 5C). As expected, co-immunoprecipitation of TRPC3 with RyR1 was absent in TRPC3 KD myotubes (data not shown). Of the three triadic proteins whose expression was up-regulated, we found that there was an increased complex formation of RyR1 with triadin (3.26 ± 0.59-fold) and JP1 (2.02 ± 0.20-fold), but not CSQ, in the solubilized TRPC3 KD myotube lysates.

      DISCUSSION

      In this study, TRPC3 knock down in skeletal muscle myotubes was accomplished using a retroviral system to stably deliver small interference RNA. One clone (TRPC3 KD, clone number 2–21) had a >97% reduction in mRNA levels and >94% reduction in protein levels (Fig. 1). Myotubes from this clone were subjected to Ca2+ imaging and Mn2+ quench experiments to examine the functional consequences of TRPC3 knock down in muscle cells. These were followed by biochemical assays to reveal underlying mechanisms of the functional changes.
      TRPC3 KD myotubes had a significantly decreased gain of EC coupling, which is reflected as a decreased cytosolic Ca2+ transient during EC coupling (Fig. 2). In accordance with the fact that the major Ca2+ source for muscle contraction is Ca2+ release through RyR1 as a result of a physical interaction between RyR1 and the DHPR during skeletal EC coupling (
      • Sandow A.
      ), RyR1 activity in TRPC3 KD myotubes was significantly reduced (Fig. 3). Because we ruled out store depletion as the cause for this reduction by showing that the SR Ca2+ content is the same in wild-type and TRPC3 KD myotubes (Fig. 2, D and E) and that the initial rate of SOCE is unchanged (Fig. 4B), it is likely that the reduced cytosolic Ca2+ transient during EC coupling in TRPC3 KD myotubes resulted from a direct reduction of RyR1 activity. This possibility is supported by the fact that in addition to a decreased gain of EC coupling, the knock down of TRPC3 is also associated with a decrease in the gain of Ca2+ release in response to direct activation of RyR1 by caffeine (Fig. 3). Two other mechanistic possibilities that link the reduced EC coupling and caffeine-induced release with our protein expression data (Fig. 5) are: 1) by increasing triadin and CSQ, it is very possible that their increased expression affects how tightly Ca2+ is bound to the complex immediately ready for Ca2+ release at the junctional endoplasmic reticulum store or, more likely, 2) the coupling efficiency of RyR1 activation to mobilize the Ca2+ immediately surrounding luminal space is diminished. These interpretations are warranted given that both proteins have been implicated in coupling RyR1 conformation to Ca2+ release from the lumen (
      • Ikemoto N.
      • Ronjat M.
      • Meszaros L.G.
      • Koshita M.
      ,
      • Ikemoto N.
      • Antoniu B.
      • Kang J.J.
      • Meszaros L.G.
      • Ronjat M.
      ,
      • Guo W.
      • Campbell K.P.
      ,
      • Beard N.A.
      • Sakowska M.M.
      • Dulhunty A.F.
      • Laver D.R.
      ).
      One of our original hypotheses was that TRPC3 was the ECCE channel, because TRPC3 could be co-immunoprecipitated with RyR1 (
      • Kiselyov K.I.
      • Shin D.M.
      • Wang Y.
      • Pessah I.N.
      • Allen P.D.
      • Muallem S.
      ) and the newly discovered ECCE current could be blocked by 2-APB, SKF96365, and La3+, all of which are known to block TRPC3 current (
      • Cherednichenko G.
      • Hurne A.M.
      • Fessenden J.D.
      • Lee E.H.
      • Allen P.D.
      • Beam K.G.
      • Pessah I.N.
      ). This hypothesis was unequivocally proven incorrect as the initial rate of ECCE of TRPC3 KD myotubes was not significantly different from that of wild type (Fig. 4A) (1.23 ± 0.15 (TRPC3 KD) versus 1.30 ± 0.08 (wild type)). In addition, these results showed that the decreased gain in EC coupling was not secondary to a decreased ECCE current.
      A possible connection between the reduced RyR1 activity and the absence of TRPC3 is the up-regulation of triadin and subsequent direct inhibition of RyR1 function caused by the increase in the amount of triadin complexed with RyR1 (Fig. 5, B and C). Triadin has been previously suggested to be an direct inhibitory protein of RyR1 activity (
      • Beard N.A.
      • Laver D.R.
      • Dulhunty A.F.
      ), and overexpression of the primary skeletal 95-kDa isoform of triadin, Trisk 95, resulted in the almost complete abolition of skeletal-type depolarization-induced Ca2+ release in the absence of extracellular Ca2+ and a significant decrease in gain in the presence of extracellular Ca2+ (
      • Rezgui S.S.
      • Vassilopoulos S.
      • Brocard J.
      • Platel J.C.
      • Bouron A.
      • Arnoult C.
      • Oddoux S.
      • Garcia L.
      • De Waard M.
      • Marty I.
      ). This is in accordance with our hypothesis. However, arguing against this hypothesis is the fact that in the Trisk 95-overexpressing cells, caffeine-induced Ca2+ release was not different from control. Another interesting protein that shows increased complex formation with RyR1 in TRPC3 KD myotubes is JP1. JP1 has been shown to facilitate junction formation between t-tubule and SR membrane via its N terminus, which has a specific affinity for t-tubule membrane, and its C terminus, which contains an SR membrane-spanning region (
      • Ito K.
      • Komazaki S.
      • Sasamoto K.
      • Yoshida M.
      • Nishi M.
      • Kitamura K.
      • Takeshima H.
      ). However, with the exception of the known affinity of RyR1 for JP1 in co-immunoprecipitation studies, the mechanism by which an increase in JP1 expression would decrease RyR1 function is not known.
      The hypothesis that there is a functional interaction between TRPC3 and RyR1 is supported by the observation that expression of TRPC3 was enhanced by the expression of RyR1 in dyspedic 1B5 (lacking RyR1) myotubes (supplemental Fig. S2, upper left). Regulation of TRPC3 expression by RyR1 was also confirmed with wild-type (RyR1+/+) and RyR1-deficient (RyR–/–) primary myotubes (supplemental Fig. S2, right and histogram), indicating that this regulation is not dependent on the origin of the cells.
      Interestingly, although the expression of two of the three other TRPCs expressed in skeletal muscle, TRPC4 and TRPC6, is unchanged in TRPC3 KD myotubes, there is an increased expression level of TRPC1 (Fig. 5A). Considering recent reports about SOCE through heteromeric TRPC channels composed of endogenous TRPC1, TRPC3, and TRPC7 in HEK293 cells (
      • Zagranichnaya T.K.
      • Wu X.
      • Villereal M.L.
      ) and the proven tandem action of TRPC1 and TRPC3 to mediate SOCE in a rat hippocampal neuronal cell line (
      • Wu X.
      • Zagranichnaya T.K.
      • Gurda G.T.
      • Eves E.M.
      • Villereal M.L.
      ), it is very likely that TRPC1 and TRPC3 are partners in making heteromeric TRPC channels in muscle. Thus it is likely that the increase in the expression of TRPC1 found here is sufficient to compensate for the absence of TRPC3 when the rate of depletion-activated SOCE is measured and prevents the conclusion that TRPC3 plays no role in SOCE in wild-type myotubes (Fig. 4, B and C).
      Studies with neurons have revealed that TRPC3 is highly expressed in the rat hippocampal cell line and mediates the specific SOCE required for its differentiation, but not that associated with proliferation (
      • Wu X.
      • Zagranichnaya T.K.
      • Gurda G.T.
      • Eves E.M.
      • Villereal M.L.
      ). The studies conclude that TRPC3 activity is essential for the guidance of nerve growth cone in rat pontine neurons (
      • Li Y.
      • Jia Y.C.
      • Cui K.
      • Li N.
      • Zheng Z.Y.
      • Wang Y.Z.
      • Yuan X.B.
      ). In the same context, we found that during differentiation of primary wild-type myoblasts to myotubes, expression level of TRPC3 peaked just after starting differentiation (day 1) and then gradually decreased (Ref.
      • Lee E.H.
      • Allen P.D.
      and supplemental Fig. S3). However, the fully differentiated myotubes on day five still expressed more TRPC3 than growing myoblasts (day zero) (2.85 ± 0.21-fold). Thus, in the case of mouse skeletal muscle, regulated TRPC3 expression is associated with muscle cell differentiation, although its role is not yet known.
      In conclusion, in this study we examined roles of TRPC3 in mouse skeletal muscle. We found that there appears to be a direct specific functional interaction between RyR1 and TRPC3 that, when interrupted, lowers the gain of the RyR1 activity without any depletion of the Ca2+ store. It is also possible that, either instead or in addition to this direct interaction, the up-regulation of triadin and JP1 associated with the absence of TRPC3 may play a role in down-regulating RyR1 activity. Our data clearly show that TRPC is not the ECCE-associated protein, and because of an associated up-regulation of the expression of TRPC1, which is known to be a partner in forming heteromeric SOCE channels in other tissues, it was not possible to see any difference in SOCE function associated with the knock down of TRPC3.

      Acknowledgments

      We acknowledge Dr. Jianjie Ma for the kind gift of anti-JP1 antibody.

      Supplementary Material

      References

        • Ramsey I.S.
        • Delling M.
        • Clapham D.E.
        Annu. Rev. Physiol. 2006; 68: 619-647
        • Pedersen S.F.
        • Owsianik G.
        • Nilius B.
        Cell Calcium. 2005; 38: 233-252
        • Freichel M.
        • Vennekens R.
        • Olausson J.
        • Stolz S.
        • Philipp S.E.
        • Weissgerber P.
        • Flockerzi V.
        J. Physiol. 2005; 567: 59-66
        • Clapham D.E.
        Nature. 2003; 426: 517-524
        • Hofmann T.
        • Schaefer M.
        • Schultz G.
        • Gudermann T.
        J. Mol. Med. 2000; 78: 14-25
        • Vandebrouck C.
        • Martin D.
        • Colson-Van Schoor M.
        • Debaix H.
        • Gailly P.
        J. Cell Biol. 2002; 158: 1089-1096
        • Hofmann T.
        • Obukhov A.G.
        • Schaefer M.
        • Harteneck C.
        • Gudermann T.
        • Schultz G.
        Nature. 1999; 397: 259-263
        • Li H.S.
        • Xu X.Z.
        • Montell C.
        Neuron. 1999; 24: 261-273
        • Vazquez G.
        • Lievremont J.P.
        • St. J. Bird G.
        • Putney Jr., J.W.
        Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 11777-11782
        • Yildirim E.
        • Kawasaki B.T.
        • Birnbaumer L.
        Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 3307-3311
        • Okada T.
        • Inoue R.
        • Yamazaki K.
        • Maeda A.
        • Kurosaki T.
        • Yamakuni T.
        • Tanaka I.
        • Shimizu S.
        • Ikenaka K.
        • Imoto K.
        • Mori Y.
        J. Biol. Chem. 1999; 274: 27359-27370
        • Zhu X.
        • Jiang M.
        • Birnbaumer L.
        J. Biol. Chem. 1998; 273: 133-142
        • Philipp S.
        • Strauss B.
        • Hirnet D.
        • Wissenbach U.
        • Mery L.
        • Flockerzi V.
        • Hoth M.
        J. Biol. Chem. 2003; 278: 26629-26638
        • Nishida M.
        • Sugimoto K.
        • Hara Y.
        • Mori E.
        • Morii T.
        • Kurosaki T.
        • Mori Y.
        EMBO J. 2003; 22: 4677-4688
        • Li Y.
        • Jia Y.C.
        • Cui K.
        • Li N.
        • Zheng Z.Y.
        • Wang Y.Z.
        • Yuan X.B.
        Nature. 2005; 434: 894-898
        • Reading S.A.
        • Earley S.
        • Waldron B.J.
        • Welsh D.G.
        • Brayden J.E.
        Am. J. Physiol. 2005; 288: H2055-H2061
        • Zhang Z.
        • Tang J.
        • Tikunova S.
        • Johnson J.D.
        • Chen Z.
        • Qin N.
        • Dietrich A.
        • Stefani E.
        • Birnbaumer L.
        • Zhu M.X.
        Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 3168-3173
        • Kiselyov K.I.
        • Shin D.M.
        • Wang Y.
        • Pessah I.N.
        • Allen P.D.
        • Muallem S.
        Mol. Cell. 2000; 6: 421-431
        • Lee E.H.
        • Allen P.D.
        Biophys. J. 2004; 86 (abstr.): 61
        • Brady A.J.
        Annu. Rev. Physiol. 1964; 26: 341-356
        • Sandow A.
        Pharmacol. Rev. 1965; 17: 265-320
        • Sampieri A.
        • Diaz-Munoz M.
        • Antaramian A.
        • Vaca L.
        J. Biol. Chem. 2005; 280: 24804-24815
        • Felder E.
        • Protasi F.
        • Hirsch R.
        • Franzini-Armstrong C.
        • Allen P.D.
        Biophys. J. 2002; 82: 3144-3149
        • Moore R.A.
        • Nguyen H.
        • Galceran J.
        • Pessah I.N.
        • Allen P.D.
        J. Cell Biol. 1998; 140: 843-851
        • Lee E.H.
        • Lopez J.R.
        • Li J.
        • Protasi F.
        • Pessah I.N.
        • Kim D.H.
        • Allen P.D.
        Am. J. Physiol. 2004; 286: C179-C189
        • Fessenden J.D.
        • Wang Y.
        • Moore R.A.
        • Chen S.R.
        • Allen P.D.
        • Pessah I.N.
        Biophys. J. 2000; 79: 2509-2525
        • Clementi E.
        • Scheer H.
        • Zacchetti D.
        • Fasolato C.
        • Pozzan T.
        • Meldolesi J.
        J. Biol. Chem. 1992; 267: 2164-2172
        • Lee E.H.
        • Rho S.H.
        • Kwon S.J.
        • Eom S.H.
        • Allen P.D.
        • Kim D.H.
        J. Biol. Chem. 2004; 279: 26481-26488
        • Cherednichenko G.
        • Hurne A.M.
        • Fessenden J.D.
        • Lee E.H.
        • Allen P.D.
        • Beam K.G.
        • Pessah I.N.
        Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 15793-15798
        • Collins J.H.
        • Tarcsafalvi A.
        • Ikemoto N.
        Biochem. Biophys. Res. Commun. 1990; 167: 189-193
        • Ikemoto N.
        • Ronjat M.
        • Meszaros L.G.
        • Koshita M.
        Biochemistry. 1989; 28: 6764-6771
        • Ikemoto N.
        • Antoniu B.
        • Kang J.J.
        • Meszaros L.G.
        • Ronjat M.
        Biochemistry. 1991; 30: 5230-5237
        • Guo W.
        • Campbell K.P.
        J. Biol. Chem. 1995; 270: 9027-9030
        • Beard N.A.
        • Sakowska M.M.
        • Dulhunty A.F.
        • Laver D.R.
        Biophys. J. 2002; 82: 310-320
        • Beard N.A.
        • Laver D.R.
        • Dulhunty A.F.
        Prog. Biophys. Mol. Biol. 2004; 85: 33-69
        • Rezgui S.S.
        • Vassilopoulos S.
        • Brocard J.
        • Platel J.C.
        • Bouron A.
        • Arnoult C.
        • Oddoux S.
        • Garcia L.
        • De Waard M.
        • Marty I.
        J. Biol. Chem. 2005; 280: 39302-39308
        • Ito K.
        • Komazaki S.
        • Sasamoto K.
        • Yoshida M.
        • Nishi M.
        • Kitamura K.
        • Takeshima H.
        J. Cell Biol. 2001; 154: 1059-1067
        • Zagranichnaya T.K.
        • Wu X.
        • Villereal M.L.
        J. Biol. Chem. 2005; 280: 29559-29569
        • Wu X.
        • Zagranichnaya T.K.
        • Gurda G.T.
        • Eves E.M.
        • Villereal M.L.
        J. Biol. Chem. 2004; 279: 43392-43402