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Previous Article | Next Article 
Molecular and Cellular Biology, May 2005, p. 3630-3638, Vol. 25, No. 9
0270-7306/05/$08.00+0 doi:10.1128/MCB.25.9.3630-3638.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Department of Pharmacology, Skirball Institute of Biomolecular Medicine,1 Department of Pediatrics, Division of Pediatric Cardiology,2 Department of Physiology and Neurosciences, New York University School of Medicine, New York, New York 100163
Received 27 August 2004/ Returned for modification 8 October 2004/ Accepted 31 January 2005
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ABSTRACT |
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INTRODUCTION |
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The genetic finding that loss of MTM function leads to specific phenotypes in mammals and Caenorhabditis elegans indicates that different MTM members, even within the same subgroup, are not functionally redundant with one another. This has led to the suggestion that different MTMs regulate specific subcellular pools of PI(3)P and PI(3,5)P2 whose function is to couple to specific downstream signaling pathways. However, as of yet, the specific downstream events regulated by different MTMs are not known. To gain insight into the specific functions and targets of MTMs, we screened a C. elegans yeast two-hybrid cDNA library for proteins that interacted with the C. elegans MTMR6 (ceMTMR6) CC domain.
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MATERIALS AND METHODS |
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Constructs, cell lines, and cell culture. Flg-tagged Human KCa3.1 (SK4) and Rat KCa2.2 (SK2) were cloned into pcDNA3 and transfected into CHO cells using Lipofectamine (Clontech). Geneticin-resistant colonies were isolated, and homogeneous expression of KCa3.1 (CHO-KCa3.1) and KCa2.2 (CHO-KCa2.2) was confirmed by immunofluorescence using anti-Flg antibodies. Green fluorescent protein (GFP)-tagged human MTMR6, MTMR8, and MTMR1 were generated by amplifying the various cDNAs by PCR and then cloning in frame into the vector pEGFP-C1 to generate an amino-terminal tagged GFP fusion protein. MTMR6 mutant for PT activity [MTMR6-PT(dead)] in which the catalytic cysteine at position 336 was mutated to serine was generated using a Transformer mutagenesis kit (Clontech) and MTMR6 deleted of its CC domain was generated by PCR using a 5' sense primer containing the amino-terminal ATG and a 3' antisense primer 5' to amino acid 514. MTM1/6CC was generated by overlapping PCR to generate a chimeric molecule that consisted of AA 1 to 546 of MTM1 and AA 512 to 560 of MTMR6. Accession numbers for the various constructs are as follows: AAP3609 for human SK4, NP_062187 for rat SK2, CAI39897 for human MTMR6, NP_000243 for human MTM1, and CAI43025 for human MTMR8.
Immunoprecipitation, Western blotting, and immunofluorescence. HEK293 cells were transfected with GFP-tagged MTMR6, MTMR8, or MTMR6 mutants as described above together with Flg-tagged KCa3.1 or Flg-tagged KCa2.2. Cell lysis, immunoprecipitation, and Western blotting were performed as previously described (1).
To determine whether overexpression of MTMR6 affects the amount of KCa3.1 at the plasma membrane (PM), MTMR6 was transfected with KCa3.1 containing an hemagglutinin (HA) epitope tag inserted into the extracellular loops between S3 and S4. Previous studies have demonstrated that KCa3.1 with this extracellular tag functions normally and can be specifically detected at the PM by immunofluorescence on nonpermeabilized cells (38). CHO cells were transfected with HA-KCa3.1 together with the GFP control or the GFP-tagged MTMR6 wild type (GFP-MTMR6-WT), and cell surface expression was performed on nonpermeabilized cells using antibodies to HA as described previously (38).
Patch clamping.
For electrophysiological studies, CHO-KCa3.1 cells were transfected with GFP-tagged constructs as indicated above and plated on 12-mm-diameter coverslips. After 24 to 48 h, a coverslip was positioned in a recording chamber, and patch clamping was performed under one of two experimental conditions that yielded similar results. One set of solutions consisted of a pipette solution containing K+-aspartate (110 mM), KCl (30 mM), HEPES (5 mM), MgCl2 (1.13 mM), EGTA (1 mM), CaCl2 (0.985 mM; pH 7.2, adjusted with KOH; calculated free Ca2+, 10 µM) and a high-K+ bath solution (KCl [140 mM], CaCl2 [2 mM], MgCl2 [1 mM], HEPES [10 mM], glucose [5 mM; pH 7.4, adjusted with KOH]). The second condition utilized a pipette solution containing: K+-gluconate (100 mM), KCl (30 mM), HEPES (10 mM), MgCl2 (1.15 mM), EGTA (5 mM), CaCl2 (4.27 mM; pH 7.2 with 1 N KOH; calculated free Ca2+, 1 µM), and a bath solution containing NaCl (140 mM), KCl (5 mM), CaCl2 (1 mM), MgCl2 (1 mM), HEPES (10 mM), and glucose (10 mM). Patch clamp pipettes had resistances ranging between 2 and 4 M
. We performed whole-cell patch clamp recordings at room temperature. Current-voltage (IV) relationships were measured using ramp voltage clamp protocols (at 15-s intervals) from a holding potential of –70 mV to –120 mV, followed by ramp depolarization to +60mV (symmetrical ramp rate of 0.18 mV · ms–1). The current-voltage relationship was obtained by plotting the current during the depolarizing ramp phase as a function of the corresponding voltage. In some experiments, a square step voltage clamp protocol was used (200-ms duration) to assess alterations in kinetics. Membrane currents filtered (–3 dB at 1 kHz) and digitized at 10 kHz (pClamp 9.2 with Digidata 1200 ADC interface; Axon Instruments). Cell capacitance and pipette series resistances were compensated (usually >80%) and were obtained using the "membrane test" function of Clampex. Whole-cell current density was expressed as pA/pF.
Currents recorded from stably transfected cells were verified to be Ca2+-activated K+ currents by (i) their sensitivity to cytosolic Ca2+, (ii) the dependence of the reversal potential on the extracellular K+ concentration, (iii) their absence in parental CHO cells, (iv) voltage-independent gating, and (v) pharmacological sensitivities (KCa3.1 was inhibited by Tram 34, but not apamin, whereas KCa2.2 was inhibited by apamin but not by Tram 34).
To determine whether PI(3)-kinase inhibition affects channel activity, stably transfected cells were treated with the PI(3) kinase [PI(3)K] inhibitor LY294002 (10 µM) or wortmannin (100 nM) 15 min prior to patch clamping, and the IV relationship was performed as described above. Phosphatidylinositols (C8) were purchased from Echelon Biosciences and used according to the manufacturer's specifications. Briefly, lipids were resuspended in water and sonicated for 20 min and used at concentrations of 100 nM in the pipette solution.
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RESULTS |
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To determine whether mammalian MTMR6 family members interact with KCa3.1 in cells, we tested whether human MTMR6 (hMTMR6) or a related subfamily member, hMTMR8, coimmunoprecipitate with KCa3.1. HEK-293 cells were transfected with GFP-tagged MTMR6 or GFP-tagged MTMR8 together with Flg-KCa3.1. We found that both GFP-tagged MTMTR6 and GFP-tagged MTMR8 coimmunoprecipitate with Flg antibodies (Fig. 1C, lanes 6 and 9). The interaction between MTMR6 and KCa3.1 did not require phosphatase activity since MTMR6-PT(dead), in which the catalytic cysteine at position 336 was changed to serine, also coimmunoprecipitated to a similar degree as the MTMR6-WT (Fig. 1C, compare lanes 7 and 9). In contrast, mutation of the MTMR6 CC domain by truncating MTMR6 at amino acid 514, which is predicted to delete the CC domain (AA 514 to 546), plus the C-terminal AA 547 to 621, markedly diminished binding (lane 8) despite similar levels of expression (Fig. 1C, compare lanes 3, 4, and 5) and similar amounts of immunoprecipitated KCa3.1 (lower part of lanes 6 to 10). In similar experiments, we found that KCa2.2 (SK2) also coimmunoprecipitated with MTMR6, although to a lesser extent than KCa3.1 (Fig. 1C, lanes 11 and 12).
MTMR6 inhibits KCa3.1 channel activity. To determine whether MTMs regulate KCa3.1 function, a CHO cell line stably expressing Flg-tagged KCa3.1 (CHO-KCa3.1) was generated and the effect of overexpressed GFP-tagged MTMR6 on channel activity was determined. The IV relationships from CHO-KCa3.1 cells transiently transfected with GFP (as a control) are shown in Fig. 2A. Also shown is the IV curve following application of Tram 34. In contrast to these control cells, KCa3.1 currents were markedly reduced when CHO-KCa3.1 cells were cotransfected with GFP-MTMR6 (Fig. 2B), without affecting the reversal potential (Fig. 2B) or time dependence of the currents (not shown). The inhibition of KCa3.1 by MTMR6 required both MTMR6 CC and PT domains, because mutation of either domain abrogated the inhibitory effects of MTMR6 (Fig. 3). In fact, current activity was reproducibly greater in CHO-KCa3.1 cells transfected with MTMR6-PT(dead) than in GFP-transfected cells, suggesting that this mutant may function as dominant negatives to inhibit MTMR6 family members endogenously expressed in these cells. Immunofluorescence studies demonstrated that expression of GFP-MTMR6 or treatment of cells with the PI(3) kinase inhibitor wortmannin (which also inhibits KCa3.1; see Fig. 6) did not affect cell surface expression of KCa3.1, indicating that altered trafficking of KCa3.1 to the plasma membrane does not account for the decrease in current in GFP-MTMR6-transfected cells (Fig. 4).
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The CC domain of MTMR6 is sufficient to enable other MTMs to inhibit KCa3.1 channel activity. In contrast to MTMR6, GFP-MTM1 did not inhibit KCa3.1 (Fig. 3). However, a chimeric MTM1 in which the MTM1 CC domain was swapped for the MTMR6 CC (MTM1/6CC)-inhibited KCa3.1 to a similar degree as MTMR6, indicating that the CC domain is sufficient to confer target specificity (Fig. 3).
MTMR6 inhibits KCa3.1 by decreasing intracellular levels of PI(3)P. One function of MTMR6 binding to KCa3.1 may be to localize MTMR6 activity to the plasma membrane adjacent to KCa3.1, thereby enabling MTMR6 to selectively dephosphorylate a local PI(3)P pool in a lipid microdomain adjacent to KCa3.1. The finding that overexpression of MTMR6 turns off channel activity predicts that for this model to be correct, intracellular application of PI(3)P or PI(3,5)P2 should reactivate KCa3.1. Consistent with this hypothesis, dialysis of PI(3)P into CHO-KCa3.1 cells that have been cotransfected with MTMR6 completely restored KCa3.1 channel activity (Fig. 5A to C).
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DISCUSSION |
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An important role for CC domains in regulating specific functions of MTMs has been established. However, unlike the findings reported here, previous studies have focused on the importance of CC domains in mediating specific heterodimerization between a PT-active and PT-inactive MTM family member (24, 33, 34). The importance of this interaction is supported by both biochemical and genetic data. Genetic evidence with humans and C. elegans has shown that mutations in a PT-active and a PT-inactive MTM that heterodimerize give rise to similar phenotypes; mutations in a PT-active (MTMR2) and PT-inactive (MTMR13) MTM give rise to CMT type 4B in humans and mutations in a C. elegans PT-active (CeMTM6), and PT-inactive (CeMTM9) MTM lead to defective fluid phase endocytosis in coelomocytes in C. elegans (3, 8, 37). The finding that heterodimerization increases PT activity has led to the idea that this interaction is critical, at least in part, for optimal PT activity. We now show that MTM CC domains have an additional function, to direct MTMs to specific targets which are regulated by PI(3)P. We propose that at least part of the specificity in MTM function is mediated by the recruitment of different MTM CC domains to specific targets. This model is supported by our finding that while MTM1 does not inhibit KCa3.1, a chimeric MTM1 containing the MTMR6 CC domain inhibits KCa3.1 channel activities. Thus, once MTM1 is targeted to the channel, other domains on MTM1 that are required to inhibit KCa3.1, such as the PT domain, function similarly in MTMR6 and MTM1.
The idea that CC domains specifically target MTMs may also have important implications for understanding the mechanisms whereby loss of MTMs leads to disease. Although altered levels of PI(3)P or PI(3,5)P2 likely account for clinical disease in X-linked myotubular myopathy and CMT, downstream targets or signaling pathways that are altered due to loss of MTM function are not yet known. The identification of binding partners for the CC domains on these MTMs (MTM1, MTMR2, and MTMR13) may thus provide important insights into the mechanisms whereby loss of MTMs leads to disease. One important question that needs to be addressed is whether a CC domain-interacting protein forms a trimolecular complex with a PT-live and a PT-inactive MTM. With regard to MTMR6, genetic and biochemical studies have indicated that MTMR6 family members heterodimerize specifically with MTMR9 (PT inactive) (33, 42). We have found that while C. elegans that has mutations in either MTMR6 and MTMR9 is defective in fluid phase endocytosis, RNA interference that target MTMR6, but not MTMR9, rescues lethality for vps34 in the C. elegans mutant. These data indicate that MTMR6, under some circumstances, can act independently of MTMR9. We have thus far been unable to identify a function of F53A2.8 for C. elegans or whether F53A2.8 functions in a pathway similar to CeMTMR6; C. elegans fed or injected with RNA interference to F53A2.8 to exhibit any obvious phenotype. Future studies will address whether inhibition of a specific target (KCa3.1) by MTMR6 in mammalian cells requires MTMR9.
An important physiological role for KCa3.1 channels is to regulate membrane potential and calcium signaling in a variety of cell types. In response to a rise in cytosolic calcium, KCa3.1 channels are activated via calmodulin bound to the C terminus of KCa3.1, leading to cross-linking of the channel, pore opening, and subsequent K+ efflux (11, 36). Efflux of K+ maintains the membrane potential at fairly negative values, providing an electrical gradient that facilitates Ca2+ influx. Beyond the role of Ca2+, little is known about the regulation of KCa3.1 channel activity. Kinases, such as protein kinase A, have been proposed to play a role in KCa3.1 activation (13, 14), although direct phosphorylation of KCa3.1 has not been shown. In addition, arachidonic acid has been shown to inhibit KCa3.1 by binding the S5-pore-S6 region (16). We now provide evidence that KCa3.1 is activated by PI(3)P and that MTMs negatively regulate KCa3.1, possibly by decreasing the concentration of PI(3)P in the plasma membrane. The finding that KCa2.2 is not regulated by PI(3)P provides further evidence that while KCa2.2 and KCa3.1 share conserved amino acid sequences and structure and both require Ca2+ for activation, they also have distinct mechanisms for regulation. This is consistent with previous studies that demonstrated that KCa3.1, but not KCa2.2, is regulated by kinases and arachidonic acid (14, 16).
While a number of channels have been shown to be regulated by phosphoinositides, we believe that this is the first demonstration of specific regulation of a channel by PI(3)P. It is not clear whether PI(3)P activates KCa3.1 directly or whether PI(3)P activates KCa3.1 indirectly by binding and recruiting a different molecule to a membrane (such as the PM), which then activates KCa3.1. Direct regulation of channels, such as the Ca2+ transporter NCX1 and voltage-gated K channels, by PIP2 involves the interaction of PIP2 with cationically charged inhibitory domains in the cytoplasmic face of the channel, leading to conformational change and channel activation (17, 18). The ability of PIP2 to activate channels under these circumstances has been attributed to its high physiologic concentrations (>1% of anionic phospholipids), which are far in excess of the concentration of PI(3)P in membranes. Thus, it is unlikely that PI(3)P regulates KCa3.1 by a similar mechanism. Rather, we favor the hypothesis that PI(3)P regulates KCa3.1 by specifically binding an as-yet-unidentified PH or FYVE domain-containing protein, which in turn regulates KCa3.1. In support of this model, we have preliminary evidence that PI(3)P does not activate KCa3.1 in isolated membrane patches (data not shown).
The finding that MTMR6 family members downregulate KCa3.1 suggests that this family of PI(3)P PTs plays unexpected roles in downregulating biological responses that are mediated by KCa3.1. KCa3.1 is required for optimal Ca2+ influx which, in turn, leads to proliferation by a variety of cells, including T lymphocytes, keratinocytes, vascular smooth muscle cells, and cancer cells (2, 5, 12, 15, 21, 25, 27, 40). In response to stimulation, KCa3.1 is upregulated in naïve and central memory T cells and naïve and memory B cells (12, 15, 23, 40, 41). Moreover, KCa3.1 is required for proliferation of preactivated naïve T cells and naive B cells, as demonstrated by the finding that the specific KCa3.1 inhibitor Tram 34 suppresses proliferation of these T and B cell subsets (15, 41). KCa3.1 is also upregulated in proliferative vascular smooth muscle cells, and the inhibition of KCa3.1 inhibits angioplasty-induced restenosis in animal models (27). KCa3.1 also contributes to the proliferation of a number of different cancer cell lines. Thus, MTM6 family members may function physiologically to negatively regulate immune cells and to inhibit mitogenic responses in conditions associated with pathological proliferation, such as cancer and restenosis after angioplasty. Finally, KCa3.1 also permits the passive efflux of Cl–, driving water and Na+ secretion from epithelia and red blood cells (9, 20, 32). Agonists of KCa3.1 have been proposed as therapy for cystic fibrosis, in which enhanced K+ secretion via KCa3.1 can be a means to promote Cl– secretion. However, only poor KCa3.1 agonists exist, and they are ineffective because they result in downregulation of KCa3.1 (26, 35). Thus, the inhibition of MTM6 family members may provide a novel target strategy for upregulating KCa3.1 activity.
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ACKNOWLEDGMENTS |
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This work is supported by NIH grants GM58573 and DK49207 to E.Y.S.
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FOOTNOTES |
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CC) (lanes 4 and 9), or GFP alone (lanes 2 and 10). Lysates were immunoprecipitated with 
-Flg antibodies, separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (10%) and then Western blotted with 
