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* This work was supported by grants from the National Institutes of Health and the Walther Cancer Institute (to S. K. and J. E. D).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. ‡ These authors contributed equally to this work. § Supported by a postdoctoral fellowship from the Endocrinology and Metabolism Training Grant, Michigan Diabetes Research Training Center. ¶ Supported by a fellowship from the Norwegian Research Council.
Myotubularin is the archetype of a family of highly conserved protein-tyrosine phosphatase-like enzymes. The myotubularin gene, MTM1, is mutated in the genetic disorder, X-linked myotubular myopathy. We and others have previously shown that myotubularin utilizes the lipid second messenger, phosphatidylinositol 3-phosphate (PI(3)P), as a physiologic substrate. We demonstrate here that the myotubularin-related protein MTMR2, which is mutated in the neurodegenerative disorder, type 4B Charcot-Marie-Tooth disease, is also highly specific for PI(3)P as a substrate. Furthermore, the MTM-related phosphatases MTMR1, MTMR3, and MTMR6 also dephosphorylate PI(3)P, suggesting that activity toward this substrate is common to all myotubularin family enzymes. A direct comparison of the lipid phosphatase activities of recombinant myotubularin and MTMR2 demonstrates that their enzymatic properties are indistinguishable, indicating that the lack of functional redundancy between these proteins is likely to be due to factors other than the utilization of different physiologic substrates. To this end, we have analyzed myotubularin and MTMR2 transcripts during induced differentiation of cultured murine C2C12 myoblasts and find that their expression is divergently regulated. In addition, myotubularin and MTMR2 enhanced green fluorescent protein fusion proteins exhibit overlapping but distinct patterns of subcellular localization. Finally, we provide evidence that myotubularin, but not MTMR2, can modulate the levels of endosomal PI(3)P. From these data, we conclude that the developmental expression and subcellular localization of myotubularin and MTMR2 are differentially regulated, resulting in their utilization of specific cellular pools of PI(3)P.
enhanced green fluorescent protein
Myotubularin (MTM1)1 is a dual specificity protein-tyrosine phosphatase (PTP)-like enzyme that is mutated in X-linked myotubular myopathy, a severe congenital disorder in which muscle cell development is compromised (
). Phylogenetic analysis of MTM family proteins indicates that they can be further divided into at least four distinct subgroups, which include the catalytically active MTM1/MTMR1/MTMR2, MTMR3/MTMR4, and MTMR6/MTMR7/MTMR8 enzymes, as well as the SBF1/LIP-STYX/MTMR10/3-PAP inactive forms (
). Together, these findings suggest that all of the active MTM family members may function to regulate cellular PI(3)P levels. Although it is well established that PI(3)P can serve as a targeting motif for membrane-trafficking and signaling proteins that contain lipid binding modules such as FYVE, pleckstrin homology, and Phox homology domains (
), the precise physiologic roles of MTM family phosphatases as regulators of PI(3)P are unclear.
In addition to MTM1 mutations identified in X-linked myotubular myopathy, a second MTM-related gene has been associated with a human genetic disease. Mutations in theMTMR2 gene are causative for the neurodegenerative disorder, type 4B Charcot-Marie-Tooth (CMT) syndrome, which is a hereditary demyelinating peripheral neuropathy that results from improper Schwann cell development (
). The difference in the pathologies of myotubular myopathy and type 4B CMT are particularly intriguing in light of the fact that MTM1 and MTMR2 are both expressed in adult skeletal muscle and neuronal tissues and are highly similar (64% identity, 76% similarity) (
). Although their specific physiologic roles are not known, it is clear from the different pathologies manifested in myotubular myopathy and CMT disease that MTM1 and MTMR2 are not functionally redundant. The lack of functional overlap between these proteins may have several possible explanations. Although they are highly similar at the protein level, MTM1 and MTMR2 may have distinct substrate preferences. Alternatively, although they are expressed in both adult skeletal muscle and neuronal tissues, MTM1 and MTMR2 may play important roles during specific developmental stages and thus be expressed differentially during maturation of specific tissues. Finally, they may both act on PI(3)P but be localized to distinct subcellular compartments, thus regulating different cellular pools of this lipid. Consistent with this notion, a recent study has demonstrated that PI(3)P can be detected at multiple discrete sites within internal vesicular structures, including endosomes and endosomal carrier vesicles as well as the nucleolus (
In the current study, we have undertaken the enzymatic characterization of MTM family phosphatases to determine whether utilization of the inositol lipid, PI(3)P, as a substrate is common among these enzymes. As a first step toward understanding the molecular basis of MTM function in the human genetic disorders, myotubular myopathy and type 4B CMT, we have also conducted a detailed comparison of the enzymatic properties of recombinant MTM1 and MTMR2. Our results demonstrate that like MTM1, MTMR2 can act on PI(3)P, providing a direct link between inositol lipid regulation and type 4B CMT. In addition, we have analyzed the subcellular localization and expression of MTM1 and MTMR2 during myogenic differentiation to determine whether these factors might contribute to their functional regulation. Finally, we present evidence that MTM1 and MTMR2 are likely to act on distinct cellular pools of PI(3)P.
Cell Culture and Transfection
COS-1 and C2C12 cells were maintained at 37 °C with 5% CO2 in Dulbecco's modified Eagle's medium containing 10% fetal calf serum, 50 units/ml penicillin, and 50 μg/ml streptomycin. C2C12 cell differentiation was induced in Dulbecco's modified Eagle's medium containing 5% horse serum, 50 units/ml penicillin, and 50 μg/ml streptomycin for up to 5 days at 37 °C with 5% CO2. COS-1 cells were transiently transfected using Fugene 6 reagent (Roche Molecular Biochemicals) according to the manufacturer's protocol.
Cloning and Expression Constructs
The completeMTM1 open reading frame (1812 nucleotides) used in constructs described herein corresponds to GenBankTMaccession number NM_000252. The complete MTMR1 open reading frame (2022 nucleotides) was amplified by PCR using a mixture of two partial cDNA fragments as a template. The 5′-region ofMTMR1 (nucleotides 1–242) was amplified by PCR using human IMAGE expressed sequence tag clone 2125570 (Research Genetics/Invitrogen, Huntsville, AL) as a template. The 3′-region ofMTMR1 (nucleotides 141–1991 of GenBankTMaccession number AJ224979) was amplified from human skeletal muscle cDNA (CLONTECH, Palo Alto, CA). Equal amounts of gel-purified 5′- and 3′-cDNA fragments were then used as a template to amplify the complete MTMR1 open reading frame. The complete open reading frames of MTMR2 (1932 nucleotides) and MTMR3 (3597 nucleotides) were amplified by PCR from plasmids containing MTMR2 or MTMR3 cDNAs (KIAA1073 and KIAA0371 cDNA clones, respectively) obtained from the Kazusa DNA Research Institute (
). The open reading frame ofMTMR6 (1866 nucleotides) corresponding to GenBankTM accession number AF406619 was amplified by PCR using human skeletal muscle cDNA as a template (CLONTECH).
Vectors for the expression of bacterial recombinant His-tagged MTM1, MTMR1, MTMR2, and MTMR6 were created using the pET21a vector (Stratagene, La Jolla, CA). The pET-MTM1 expression construct has been previously described (
). The pET-MTMR1 vector was created by inserting a DNA fragment containing the MTMR1 open reading frame without a stop codon into the 5′-NheI and 3′-XhoI sites of pET21a in-frame with the six-histidine tag. The pET-MTMR2 construct was created by inserting a DNA fragment containing the MTMR2 open reading frame without a stop codon into the 5′-NheI and 3′-NotI sites of pET21a in-frame with the six-histidine tag. The pET-MTMR6 expression construct was created by inserting a DNA fragment containing the MTMR6open reading frame without a stop codon into the 5′-BamHI and 3′-HindIII sites of pET21a in-frame with the six-histidine tag.
Vectors for the expression of N-terminally FLAG-tagged MTM1 wild type and C375S mutant proteins in mammalian cells have been previously described (
). Mammalian expression vectors for N-terminally FLAG-tagged MTMR1, MTMR2, MTMR3, and MTMR6 proteins were created by inserting cDNA fragments containing the complete open reading frames encoding each of these proteins into the 5′-BamHI/3′-NotI, 5′-NheI/3′-KpnI, 5′-NheI/3′-XbaI, and 5′-BamHI/3′-XbaI, respectively, of pCDNA3.1-NF (
). A cDNA fragment containing the complete open reading frame ofMTMR2 was inserted into the 5′-BglII and 3′-KpnI sites of the pEGFP-C1 vector (CLONTECH) to create pEGFP-MTMR2. A vector for bacterial expression of tandem FYVE domains from the murine hepatocyte growth factor-regulated tyrosine kinase substrate (Hrs) protein fused to GST (GST-2×FYVE; see Ref.
) was the generous gift of Dr. Kathleen Collins (University of Michigan).
Protein Expression and Purification
Bacterial recombinant MTM1 and MTMR2 C-terminally His-tagged fusion proteins used for phosphatase assays were expressed in Escherichia coliBL21(DE3) Codon Plus cells (Stratagene) and purified using Ni2+-agarose affinity resin as previously described forSacIp (
). FLAG-tagged MTM proteins used for phosphatase assays were expressed in COS-1 cells and purified using anti-FLAG M2 affinity resin (Sigma) as previously described for FLAG-tagged MTM1 wild-type and C375S proteins (
). Recombinant GST-2×FYVE protein was expressed in Escherichia coli DH5α cells and purified over glutathione-agarose affinity resin. Briefly, cells harboring the pGEX-2×FYVE plasmid were grown in 2× YT medium containing 100 μg/ml ampicillin to an A600 of 0.7, and then protein expression was induced by the addition of isopropyl β-d-thiogalactopyranoside (0.5 mm) for 2 h at 37 °C. The cells were harvested by centrifugation and resuspended in PBS (pH 7.4) containing 1 mm benzamidine, 1 mm phenylmethylsulfonyl fluoride, and 1 μg/ml (each) aprotinin, leupeptin, and pepstatin (30 ml of lysis buffer per liter of cultured cells). The cells were disrupted by sonication, and Triton X-100 was added to 0.5% (v/v). The crude lysate was centrifuged for 30 min at 18,000 × g to remove unbroken cells and insoluble debris. The soluble fraction was incubated with glutathione-agarose affinity resin (1 ml of 50% slurry in PBS/liter of culture) for 2 h at 4 °C. The resin was washed three times for 5 min each with lysis buffer containing 0.5% Triton X-100, followed by two 5-min washes in lysis buffer without detergent. The GST-2×FYVE fusion protein was eluted from the resin for 2 × 30 min with 0.5 ml of lysis buffer containing 25 mm reduced glutathione (pH 7.4) and filtered through a 0.2-μm syringe filter. Free glutathione was removed by gel filtration chromatography in PBS using a Superdex 200 column and FPLC chromatography system (Amersham Biosciences, Inc.).
Biotinylation of GST-2×FYVE
The GST-2×FYVE fusion protein was biotinylated using the Biotin-Tag microbiotinylation kit as per the manufacturer's protocol (Sigma-Aldrich). The unreacted biotinylation reagent was removed using a PD-10 desalting column (Bio-Rad).
All phosphoinositide and soluble inositol phosphate substrates used in this work were obtained from Echelon Research Laboratories (Salt Lake City, UT). Phosphatase assays using FLAG-tagged MTM proteins immunoprecipitated from COS-1 cells were carried out on anti-FLAG M2 affinity resin in reaction buffer containing 1.5 μg of di-C6-NBD6-phosphatidylinositol 3-phosphate (Echelon) as described (
). Phosphatase assays with bacterial recombinant MTM1, MTMR1, MTMR2, and MTMR6 using synthetic di-C8-phosphoinositide or soluble inositol phosphate substrates were carried out at 30 °C, and phosphate release was determined using a malachite green-based assay system for inorganic phosphate as described (
). The artificial protein phosphatase substrates myelin basic protein and casein were phosphorylated with [γ-32P]ATP on tyrosyl or seryl/threonyl residues and used as substrates for recombinant MTM1 and MTMR2 as previously described (
Total RNA was isolated from C2C12 cells with TRIzol reagent as recommended by the manufacturer (Invitrogen). RNA samples (15 μg) were then electrophoresed through a 1% agarose-formaldelyde gel and transferred to a nylon membrane. The blots were hybridized with digoxigenin-labeled probes, and mRNAs corresponding to murine MTM1 and MTMR2 were detected using a digoxigenin chemiluminescence detection kit (Roche Molecular Biochemicals). A murine MTM1 cDNA probe corresponding to the 3′-untranslated region (283 bp) was amplified by PCR using expressed sequence tag clone AW911959 (Research Genetics/Invitrogen) as a template. A cDNA probe corresponding to the murine MTMR2 5′-region (nucleotides −5 to +381) was amplified by PCR using expressed sequence tag clone AI561725 (Research Genetics/Invitrogen) as a template.
COS-1 cells were seeded on two-chamber slides (21 × 21 mm) at a density of 5 × 104cells/well in Dulbecco's modified Eagle's medium and cultured as described above. After attachment (∼8 to 12 h), the cells were transiently transfected using Fugene 6 reagent as described above. At 30 h post-transfection, the cells were washed twice with PBS and fixed with 4% paraformaldehyde in PBS for 10 min at room temperature. Coverslips were mounted on the slides using ProLong Antifade mounting medium (Molecular Probes, Eugene, OR). For analyses with biotinylated GST-2×FYVE as a probe for PI(3)P, a modified form of a protocol developed in the laboratory of Dr. Harald Stenmark at the Department of Biochemistry, Institute for Cancer Research, Norwegian Radium Hospital, Oslo, Norway, was used.
Briefly, COS-1 cells were cultured and transfected on chambered microscope slides as described above. At 24 h post-transfection, the cells were washed and fixed overnight at 4 °C in 4% paraformaldehyde in PBS, which permeabilizes most of the cells. The cells were washed once in PBS, then the free aldehyde groups were quenched by incubation with PBS containing 50 mmNH4Cl for 15 min at room temperature. All subsequent steps were carried out at room temperature. The slides were incubated for 1 h with 10% fetal calf serum in PBS as a blocking agent. Next, the slides were incubated with 3% fetal calf serum in PBS containing 50 μg/ml biotinylated GST-2×FYVE for 30 min. The slides were washed for 3 × 5 min with PBS. The slides were then incubated for 30 min with 1 μg/ml Cy3-conjugated streptavidin (Sigma-Aldrich) in PBS containing 3% fetal calf serum. Finally, the slides were washed three times for 5 min each with PBS and mounted using ProLong mounting medium. Fluorescence analyses were performed by conventional fluorescence microscopy or by confocal microscopy (Zeiss LSM 510) as indicated.
RESULTS AND DISCUSSION
The association of MTM1 and MTMR2 with the respective genetic disorders myotubular myopathy and type 4B Charcot-Marie-Tooth syndrome is of significant interest because although they are highly similar proteins and exhibit overlapping expression patterns in human tissues, they do not appear to be functionally redundant (
). As a first step toward understanding the functional differences between MTM1 and MTMR2, we analyzed their enzymatic properties to determine whether they possessed similar activity and/or substrate specificity. Recombinant MTM1 and MTMR2 fusion proteins were expressed and purified as described under “Experimental Procedures,” and their phosphatase activity was tested toward a panel of substrates including phosphoinositides, soluble inositol phosphates, and radiolabeled artificial protein substrates. As shown in Table I, the activity of MTMR2 was indistinguishable from that of MTM1 over the panel of substrates tested. Both enzymes dephosphorylated PI(3)P >50-fold more efficiently than any other substrate, demonstrating that they are highly specific for this lipid (Table I). Although MTM1 and MTMR2 were able to dephosphorylate the PI(3)P soluble headgroup analog, inositol 1,3-bisphosphate, their activity with this substrate was 10-fold lower than with PI(3)P, indicating a strong preference for the lipid as a substrate. Neither enzyme exhibited significant activity toward artificial protein substrates phosphorylated on tyrosyl or seryl/threonyl residues, confirming that they are extremely poor protein phosphatases. Most importantly, because the truncatingMTMR2 mutations first associated with type 4B CMT syndrome would cause loss of MTMR2 phosphatase activity, our findings suggest that this disorder, like myotubular myopathy, results from the failure to regulate cellular PI(3)P levels (
Limit of detection in these assays was 10 pmol of phosphate released.
Phosphatase assays using purified recombinant MTM1 and MTMR2 proteins were carried out at 30 °C with the indicated amount of substrate as described under “Experimental Procedures.” The specific activities shown are expressed as mol of phosphate released/min/mol of enzyme and represent the mean of triplicate determinations. S.E. of each determination was less than 5%.
The identification of MTM1 and MTMR2 as PI(3)P-specific phosphatases led us to question whether activity toward this inositol lipid is a conserved property among other active myotubularin family enzymes. We therefore tested the ability of the myotubularin family phosphatases MTM1, MTMR1, MTMR2, MTMR3, and MTMR6, which represent each of the three active MTM subgroups (see Ref.
), to function as PI(3)P-specific inositol lipid phosphatases. FLAG epitope-tagged MTM proteins were transiently overexpressed in COS-1 cells and immunoprecipitated with anti-FLAG affinity resin, and their ability to dephosphorylate PI(3)P was determined using a qualitative inositol lipid phosphatase assay that employs a water-soluble fluorescent substrate (di-C6-NBD6-phosphatidylinositol 3-phosphate) combined with thin layer chromatography as previously described (
). As shown in Fig. 1A, human MTM1, MTMR1, MTMR2, MTMR3, and MTMR6 immunoprecipitates each converted PI(3)P to phosphatidylinositol (Fig. 1A, lanes 3–7, respectively). As expected, immunoprecipitates from cells transfected with vector alone (Fig. 1A, lane 1) or a vector encoding the catalytically inactive MTM1 C375S mutant (Fig. 1A,lane 2) showed no activity toward PI(3)P. The expression of MTM proteins in the lysates of transfected cells used for anti-FLAG immunoprecipitation and lipid phosphatase assays was confirmed by immunoblotting with anti-FLAG antibody (Fig. 1B). Because both MTM1 and MTMR2 are highly specific for PI(3)P as a substrate (Table I), we wanted to determine how the substrate preferences of MTMR1 and MTMR6 might compare with those of MTM1 and MTMR2. To accomplish this, we performed lipid phosphatase assays using bacterial recombinant MTMR1 and MTMR6 proteins with the complete panel of phosphoinositide substrates. As shown in Fig.2, both MTMR1 and MTMR6 were also highly specific for PI(3)P as a substrate. It should be noted that the specific activities of recombinant MTMR1 and MTMR6 toward PI(3)P are ∼30- and 100-fold lower than those of MTM1 and MTMR2 (Fig. 2, TableI) but are similar to those observed for recombinant MTMR3 and MTMR4 (
). However, we have been unable to detect activity toward PI(3,5)P2 utilizing FLAG-tagged MTMR3 immunoprecipitated from mammalian cells (not shown). It is unclear whether this discrepancy may be the result of differences in the preparations of MTMR3 used or in lipid phosphatase assay conditions. Regardless, the results presented here not only identify MTMR1, MTMR2, and MTMR6 as PI(3)P-specific inositol lipid phosphatases but further support the hypothesis that PI(3)P is likely to be a common substrate among all of the active MTM proteins. Furthermore, the finding that MTMR2 is specific for PI(3)P as a substrate suggests a possible link between type 4B CMT and phosphoinositide signaling.
Having established that MTM1 and MTMR2 are both specific and efficient catalysts toward PI(3)P, we next asked whether other factors such as subcellular localization, developmentally regulated expression, or utilization of distinct substrate pools might be responsible for the differences in their function(s). To analyze their subcellular localization, MTM1 and MTMR2 were expressed as EGFP fusion proteins in COS-1 cells and visualized by fluorescence microscopy. As expected, cells expressing EGFP alone exhibited staining in both the nucleus and cytoplasm (Fig. 3A). EGFP-MTM1 (Fig. 3B) and EGFP-MTMR2 (Fig. 3C) displayed a cytosolic staining pattern, with the highest staining intensity in the perinuclear region. In contrast to EGFP-MTMR2, EGFP-MTM1 staining was also detected in large membrane projections and at the cell periphery (Fig. 3B). This staining pattern is similar to that observed previously for an MTM1 putative substrate-trapping mutant protein, which localized to plasma membrane extensions in HeLa cells (
). The dramatic difference in the morphology of COS-1 cells transfected with either EGFP-MTM1 or EGFP-MTMR2 fusion protein expression constructs also provides evidence of their distinct properties. EGFP-MTM1 overexpression resulted in the formation of membrane projections (Fig.3B), whereas EGFP-MTMR2 did not (Fig. 3C). Although the precise molecular basis of this phenomena is unclear, it highlights a difference between the effects of MTM1 and MTMR2 overexpression. Collectively, these findings suggest the possibility that MTM1 and MTMR2 may perform different functions by targeting to different subcellular environments.
We have also examined the expression of MTM1 and MTMR2 during myogenic differentiation in murine C2C12 myoblasts, which can be induced to differentiate into myotubes by growth in low mitogen medium. To accomplish this, proliferating C2C12 cells were cultured to confluence in high mitogen medium (Dulbecco's modified Eagle's medium containing 10% fetal calf serum), and differentiation was then induced for 5 days by switching the cells to low mitogen medium (Dulbecco's modified Eagle's medium containing 5% horse serum). Differentiation of C2C12 cells was monitored visually by the formation of multinucleated myotubes and by analyzing the expression of a myogenic differentiation marker, myosin heavy chain, which commenced ∼40 h after the cells were switched to low mitogen media (Fig.4A). To probe the expression of MTM1 and MTMR2 during myogenic differentiation, total RNA was isolated from day 0 to day 5 following the change to low mitogen medium, and the levels of MTM1 and MTMR2 transcripts were analyzed by Northern hybridization using transcript-specific probes. As shown in Fig. 4B, MTM1 expression was up-regulated during the differentiation of C2C12 cells, whereas MTMR2 expression steadily declined over the same period. Identical results were obtained using quantitative RT-PCR (not shown). These data are consistent with a role for MTM1 at a late step of myogenic differentiation/maturation and suggest that the expression of MTM1 and MTMR2 is differentially regulated during myogenic development.
In addition to their differential expression, another possible explanation for the lack of functional redundancy between MTM1 and MTMR2 is that they may regulate different subcellular pools of PI(3)P. To test this hypothesis, we used a biotinylated bacterial recombinant GST-2×FYVE fusion protein as a probe to specifically label intracellular PI(3)P. FYVE domains are modified zinc finger-like domains that specifically recognize and bind PI(3)P, thus serving to target proteins to distinct sites within cells (
). Treatment of COS-1 cells with the phosphatidylinositol 3-kinase inhibitor, wortmannin, completely abolished the punctate staining pattern (Fig.5B), demonstrating that endosomal targeting of the GST-2×FYVE probe is dependent on D3-phosphorylated phosphoinositides as previously described (
). To test the effects of MTM1 and MTMR2 overexpression on specific intracellular pools of PI(3)P, COS-1 cells were transfected with either pEGFP-MTM1 or pEGFP-MTMR2. At 24 h post-transfection, the cells were fixed, stained using the biotinylated GST-2×FYVE probe, and analyzed for red and green fluorescence by confocal microscopy. As seen in Fig. 5, D and E, overexpression of EGFP-MTM1 disrupted the punctate endosomal staining pattern, suggesting that MTM1 was able to dephosphorylate endosomal PI(3)P, thus abrogating lipid-dependent targeting of the GST-2×FYVE probe. In contrast, cells in which EGFP-MTMR2 was overexpressed displayed a punctate staining pattern identical to that observed for untransfected cells, suggesting that the EGFP-MTMR2 fusion protein was unable to dephosphorylate endosomal PI(3)P (Fig. 5,G and H). We have analyzed the lipid phosphatase activity of EGFP-MTM1 and EGFP-MTMR2 fusion proteins immunoprecipitated from COS-1 cells using fluorescent di-C6-NBD6-PI(3)P substrate as previously described (
). Both of the fusion proteins were expressed at comparable levels and possessed comparable phosphatidylinositol 3-phosphatase activity under these conditions, indicating that the differences in GST-2×FYVE staining of PI(3)P observed in COS-1 cells overexpressing EGFP-MTM1 or EGFP-MTMR2 were not due to differences in their lipid phosphatase activities. Although we cannot completely rule out the notion of an MTMR2-specific cellular inhibitor, the mild conditions used during immunoprecipitation of EGFP-MTM1 and EGFP-MTMR2 fusion proteins for activity analysis would be likely to preserve such interactions. Furthermore, expression of catalytically inactive (Cys to Ser) MTM1 and MTMR2 EGFP fusion proteins had no effect on PI(3)P staining by the GST-2×FYVE probe (not shown). It should also be noted that endosomal staining of PI(3)P by GST-2×FYVE was lowered in a small number of cells exhibiting the highest amount of EGFP-MTMR2 as assessed by green fluorescence (not shown), whereas GST-2×FYVE staining was affected in all cells expressing detectable levels of EGFP-MTM1. EGFP-MTMR2 staining in those cells was clearly not restricted to the perinuclear region, being more diffusely cytoplasmic. Together, these results suggest that when overexpressed to a sufficiently high level, EGFP-MTMR2 localization becomes unrestricted, rendering it capable of depleting endosomal PI(3)P. We therefore conclude that MTM1 and MTMR2 are likely to regulate different subcellular pools of PI(3)P by virtue of their subcellular localization and differential expression during development.
We have provided the first evidence that MTMR2, a phosphatase mutated in type 4B Charcot-Marie-Tooth syndrome, utilizes the lipid second messenger, PI(3)P, as its physiologic substrate. Because mutations in the MTMR2 gene associated with type 4B CMT would disrupt the phosphatase activity of MTMR2, it is probable that this disease results from improper regulation of PI(3)P. Although the exact mechanisms by which failure to dephosphorylate PI(3)P contributes to myotubular myopathy and type 4B CMT are unclear, the identification of this lipid as a specific target for MTM1, MTMR2, and other MTM family phosphatases represents a first step toward understanding their roles in human neuromuscular diseases. The similarity of MTM1 and MTMR2 protein sequences and enzymatic properties has also led us to question what factors might underlie the apparent lack of functional redundancy between these phosphatases. Collectively, our findings suggest that MTM1 and MTMR2 can be regulated not only by tissue-specific expression during development, but also by their subcellular localization and use of specific cellular pools of PI(3)P. It will now be important to identify downstream effectors of PI(3)P whose function is regulated by MTM1 or MTMR2 in order to pinpoint the signaling pathways that are affected by mutations associated with myotubular myopathy and type 4B CMT.
We thank M. J. Wishart and C. D. Worby for helpful suggestions and critical evaluation of the manuscript.