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Originally published In Press as doi:10.1074/jbc.M403316200 on May 15, 2004

J. Biol. Chem., Vol. 279, Issue 30, 31081-31088, July 23, 2004
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ATP Potentiates Agrin-induced AChR Aggregation in Cultured Myotubes

ACTIVATION OF RHOA IN P2Y1 NUCLEOTIDE RECEPTOR SIGNALING AT VERTEBRATE NEUROMUSCULAR JUNCTIONS*

Karen K. Y. Ling, Nina L. Siow{ddagger}, Roy C. Y. Choi§, Annie K. L. Ting, Ling W. Kong, and Karl W. K. Tsim

From the Department of Biology and Molecular Neuroscience Center, Hong Kong University of Science and Technology, Hong Kong, China

Received for publication, March 25, 2004 , and in revised form, April 28, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
At vertebrate neuromuscular junctions, ATP is known to stabilize acetylcholine in the synaptic vesicles and to be co-released with it. We have shown previously that a nucleotide receptor, P2Y1 receptor, is localized at the nmjs, and we propose that this mediates a trophic role for synaptic ATP there. In cultured myotubes, the activation of P2Y1 receptors modulated agrin-induced acetylcholine receptor (AChR) aggregation in a potentiation manner. This potentiation effect in agrin-induced AChR aggregation was reduced by antagonizing the P2Y1 receptors. The guanosine triphosphatase RhoA was shown to be responsible for this P2Y1-potentiated effect. The localization of RhoA in rat and chicken skeletal muscles was restricted at the neuromuscular junctions. Application of P2Y1 agonists in cultured myotubes induced RhoA activation, which showed an additive effect with agrin-induced RhoA activation. Over-expression of dominant-negative mutant of RhoA in cultured myotubes diminished the agrin-induced AChR aggregation, as well as the potentiation effect of P2Y1-specific agonist. Application of UTP in the cultures also triggered similar responses as did 2-methylthioadenosine 5'-diphosphate, suggesting the involvement of other subtypes of P2Y receptors. These results demonstrate that RhoA could serve as a downstream mediator of signaling mediated by P2Y1 receptor and agrin, which therefore synergizes the effects of the two neuron-derived trophic factors in modulating the formation and/or maintenance of post-synaptic apparatus at the neuromuscular junctions.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In developing vertebrate neuromuscular junction (nmj),1 a motor nerve terminal contacts a muscle fiber, acetylcholine receptors (AChRs), acetylcholinesterase (AChE) and certain other proteins become localized and stabilized in a specialized post-synaptic apparatus. The initial stage of AChR aggregation at nmjs is controlled by agrin synthesized by the pre-synaptic motor neuron (1–5). The molecular mechanism of agrin-induced AChR aggregation, however, is poorly understood. The agrin-induced post-synaptic specialization is mediated by muscle-specific receptor tyrosine kinase (MuSK) (6), and indeed, the phenotype of MuSK-deficient mice show similar neuromuscular defects to those in agrin-deficient mice (7). Another key component involved in the downstream signaling of MuSK is rapsyn, a 43-kDa protein closely associated with the intracellular face of AChR (8, 9) at the post-synaptic membrane. Rapsyn knock-out mice lack aggregates of AChRs and other post-synaptic proteins (10), and myotubes derived from rapsyn-deficient mice also fail to respond to agrin in forming AChR aggregates (8). Several downstream effects of agrin and MuSK signaling have been proposed. Agrin has been shown to activate Rho family of guanosine triphosphatase (GTPase) Rac, Cdc-42, and RhoA, and these molecules are required for agrin-induced AChR aggregation in myotubes (11, 12). The mobilization of intracellular Ca2+ in post-synaptic muscle has been demonstrated to regulate the formation and/or the maintenance of agrin-induced AChR aggregation (13–15). In addition, Src-class kinase (16, 17), nitric oxide (18, 19), synaptic MAGI-1c (20), geranylgeranyltransferase (GGT) (21), and Dishevelled (Dvl 1) (22) have also been revealed to associate with agrin and MuSK signaling at the nmjs.

ATP is an additional potential trophic factor at the nmjs (23, 24). In the synaptic vesicles at vertebrate nmjs (25), ATP stabilizes acetylcholine (ACh) and is co-released quantally with it in a ratio of about 1 ATP to 5 ACh. The synaptic ATP, mediated by P2Y1 nucleotide receptors, induces and sustains the expression of AChE and AChR in muscles, and P2Y1 receptor is localized at the nmjs (24, 26). The P2Y1 receptor-mediated gene activations including AChE and different subunits of AChR are acted upon through the mobilization of intracellular inositol triphosphate and Ca2+, and subsequently, the activation of a mitogen-activated protein kinase signaling pathway (27).

Here, we investigate the roles of ATP and its activation of P2Y1 receptors in potentiating the agrin-induced AChR aggregation in cultured myotubes. Several lines of evidence support the notion. There is much evidence that the native P2Y1 receptor in tissues, as well as in muscle, so far examined is linked to the formation of inositol triphosphate and to intracellular Ca2+ mobilization (see references in Refs. 23, 26). ATP potentiates the response to the applied ACh in nerve-muscle co-culture (see references in Ref. 23), which is in line with the activity-dependence of the reshaping of synaptic architecture at the nmjs (4). On the other hand, the application of P2Y1 receptor agonists activated the membrane-bound RhoA and stimulated the actin cytoskeleton organization in cultured vascular myocytes (28). In the present study, we provide evidence that the activation of P2Y1 receptors induced the formation of membrane-bound RhoA, which subsequently potentiated the agrin-induced AChR aggregation on post-synaptic muscle fibers.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials and Purity of Nucleotides—Materials not specified here were obtained as before (26) or from Sigma. Cell culture medium and serum were obtained from Invitrogen. 2-Methylthioadenosine 5'-diphosphate (2-MeSADP) stock solution (100 mM) was pre-incubated with 20 units/ml of yeast hexokinase (Roche Biochemicals, Lewes, UK) in Buffer A (2.5 mM MgCl2, 50 mM HEPES, pH 7.3) containing 25 mM glucose at 37 °C for 30 min to remove all contaminating triphosphates, whereas ATP stock solution (100 mM) was pretreated in Buffer A with 20 units/ml of creatine phosphokinase (Sigma) and 10 mM creatine phosphate (Sigma) at room temperature for 90 min to remove all contaminating diphosphates (26). Anti-RhoA antibody was obtained from Santa Cruz Biotechnology (Santa Cruz, CA); anti-Rac antibody was from Upstate Biotechnology (Lake Placid, NY); others not stated were from Sigma; and peroxidase- or fluorescein-conjugated secondary antibodies were from Cappel (Turnhout, Belgium). Tetramethylrhodamine-conjugated {alpha}-bungarotoxin (TMR-BuTX) was obtained from Molecular Probes (Eugene, OR).

Animals—Muscles from adult Sprague-Dawley rats and New Hampshire chickens were collected immediately after the animals were killed. Animals were rapidly frozen in isopentane/liquid nitrogen and stored at –80 °C. All procedures conformed to the Guidelines by Animal Research Panel of Hong Kong University of Science and Technology.

Cell Cultures—Eggs of New Hampshire chickens were purchased from a local farm and hatched in the University Animal Care Facility. Primary chick myotubes were prepared from hindlimb muscles dissected from 11-day-old chick embryos and cultured at 37 °C in a water-saturated 5% CO2 atmosphere, as described previously (26). Myotubes were treated with a mitotic inhibitor (10 µM cytosine arabinoside) at day 3 after plating and used on day 4. Undifferentiated mouse C2C12 myoblasts were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 20% fetal bovine serum, 100 units/ml penicillin and 100 µg/ml streptomycin and were incubated at 37 °Cina water-saturated atmosphere of 95% air/5% CO2. Differentiation of myoblast to myotube was induced by replacing the growth medium with DMEM supplemented with 2% heat-inactivated horse serum, 100 units/ml penicillin, and 100 µg/ml streptomycin, as described previously (29). Skeletal muscle cells in culture release ATP into the medium and also can convert it there to ADP, which, over longer periods, may give some desensitization of P2Y receptors. The cultures were pretreated in all cases with apyrase (2 units/ml; Sigma) for 1 h to eliminate all such free nucleotides, followed by a gentle wash and drug application in apyrase-free medium (26). For longer incubations with an agonist, significant loss thereof due to the ectonucleotidase on muscle cells was prevented (where stated) by maintaining the appropriate enzymatic regeneration system throughout (hexokinase/glucose or creatine phosphokinase/creatine phosphate in Buffer A) as noted above, as well as by three changes of the agonist solution at approximately equal intervals.

cDNA Transfections—For the source of agrin, the sub-confluent HEK293 cells were transfected with chicken agrin cDNA encoding a secretory form of agrin (CBA-1; Ref. 30) by calcium phosphate. The agrin-conditioned medium was collected with the addition of 1 mM phenylmethyl sulfonyl fluoride and 5% glycerol and stored at –80 °C. The unit of agrin was calibrated as reported previously (30). The full-length cDNA encoding chicken P2Y1 receptor (31) in an expression vector pcDNA3 (Invitrogen) was used where stated. RhoA cDNAs encoding the wild-type RhoA (RhoAWT, GenBank accession no. AY026068 [GenBank] ) was generated by RT-PCR from rat muscle cells. The dominant-negative RhoA (RhoAN19) was obtained by site-directed mutagenesis of the RhoAWT, in which amino acid asparagine at position 19 was mutated to threonine (32). All cDNAs were subcloned into pcDNA3, verified therein, and used for transfection.

Myoblasts from 11-day-old chick embryos were cultured at 37 °C for 2 days and transiently transfected with the plasmid constructs (2 µg of plasmid per 35-mm plate or 10 µg per 100-mm plate) with the use of calcium phosphate. Myoblasts were then allowed to fuse to myotubes for the treatments stated, with the methods given in Ref. 26. In C2C12 myoblasts, the transfection was done by using LipofectAMINE Plus (Invitrogen) (29). The transfection efficiency in both cases was determined with enzymatic staining from control cells co-transfected with {beta}-galactosidase cDNA in the same vector; it was consistently ~30%.

Immunohistochemical Staining—Myotube cultures were treated with P2Y1 receptor antagonists or together with agrin-conditioned medium for 16 h to study AChR aggregation. Cell-surface AChR was stained by incubating the treated cultures with 10–8 M TMR-BuTX in DMEM for 1 h, rinsed with phosphate-buffered saline (PBS), and fixed with 2% paraformaldehyde, 5% sucrose in PBS for 10 min, and then dried by increasing amounts of ethanol. The cultures were mounted in Citifluor (Citifluor Ltd., Leicester, UK). The AChR aggregates were counted under a 40x objective on a Zeiss Axiophot equipped with phase-contrast and fluorescence optics as described previously (30, 33). The mean number of AChR aggregates per field in a single myotube was determined by counting 20 different myotubes from different fields. In general, two to three myotubes were counted from a single field. Experiments were repeated at least four times, each with triplicate cultures. For the pre-labeling AChR studies, myotube cultures were incubated with 10–6M TMR-BuTX in DMEM prior to the agrin treatment. AChR aggregation was quantified similarly as above (30, 33).

Muscle sections were prepared from the pectoral and gastrocnemius muscles of adult chicken and rat, respectively, embedded in tissue-freezing medium (Leica Instruments, Nussloch, Germany), and frozen in an isopentane/liquid nitrogen bath. Twenty µm muscle sections generated on cryostat were fixed in 2% paraformaldehyde, 5% sucrose in PBS for 15 min at room temperature, followed by three 5-min washes in PBS and a blocking step in PBS containing 5% bovine serum albumin (BSA) for 30 min. The slides were incubated with ~5 µg/ml anti-RhoA antibody in PBS with 5% BSA for 16 h at 4 °C. RhoA mouse monoclonal antibody recognizes epitope corresponding to amino acids 120–150 of RhoA. For double-staining of AChR, 10–8 M TMR-BuTX was incubated together with anti-mouse secondary antibody for 1 h. Sections were washed 3x with PBS for 10 min each and then dehydrated in ice-cold 100% ethanol; the sections were mounted with Citifluor mounting media and examined with a Zeiss Axiophot microscope equipped with fluorescent and rhodamine optics, using excitation at 555 or 488 nm and emission at 580 or 515 nm for rhodamine or fluorescein, respectively.

RhoA and Rac Activation Assay—For the measurement of membrane-bound (activated form) RhoA protein, the treated myotubes were collected in PBS with 10 mM EDTA and centrifuged at 2,300 x g for 7 min. The cell pellets were then resuspended in Buffer B (50 mM Tris-HCl, pH 7.4, 1 mM EGTA, 1 mM EDTA, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 5 mM benzamidine HCl, 10 µg/ml soybean trypsin inhibitor, and 1 mM phenylmethyl sulfonyl fluoride). The cellular suspension was passed through a G-27 gauge needle ten times for lysis of the cells. The cell extracts were separated by low speed centrifugation at 800 x g for 5 min. The supernatant was then collected and further centrifuged at 16,000 x g for 15 min. The cell pellet representing the membrane fraction was resuspended in 50–100 µl of Buffer B and was then ready for Western blot analysis of RhoA.

An affinity precipitation method described in the Rho or Rac activity assay kit (Upstate Biotechnology) was also used. Cultured chick myotubes were treated with agrin or with indicated drugs for indicated times and rinsed with PBS. The cells were then lysed with Buffer C (25 mM HEPES, pH 7.5, 150 mM NaCl, 1% Igepal CA-630, 10 mM MgCl2, 1 mM EDTA, 10% glycerol, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 25 mM sodium fluoride, and 1 mM sodium orthovanadate) on ice for 5 min and centrifuged for 5 min at 16,000 x g at 4 °C. The supernatant was collected as cell lysate, and 2 mg of the lysate were incubated with glutathione-agarose bound to 20 µg of Rho-binding domain (RBD) from the effector protein Rhotekin for RhoA, or 10 µg of Rac-binding domain (p21-binding domain) from the effector protein p21-activated kinase (PAK) for Rac (Upstate Biotechnology) for 45 min at 4 °C. The beads were washed 3x with Buffer C. The bound Rho or Rac proteins were eluted with sample buffer with 50 mM dithiothreitol, separated by SDS-polyacrylamide gel, detected by using an anti-RhoA or anti-Rac antibody in a 1:1000 dilution, followed by peroxidase-conjugated secondary antibody against mouse IgG in a 1:5000 dilution in a Western blot analysis. The Rac antibody recognizes Rac1 and Rac2. The immunocomplexes were visualized by the ECL method (Amersham Biosciences). The intensity of the bands in the control and agonist-stimulated samples, run on the same gel and under strictly standardized ECL conditions, were compared on an image analyzer, in each case using a calibration plot constructed from a parallel gel with serial dilutions of one of those samples.

Other Assays—The concentration of protein was determined by using a Bio-Rad protein assay kit (Hercules, CA); the concentration of sample protein was compared against the standard curve, which was generated from the relative absorbance (595 nm) of the BSA standard (0.1–0.6 µg/ml) added with dye reagent. Statistical tests were made by the PRIMER version 1 software program (47): differences from basal or control values (as shown in the plots in Figs. 1, 2, and 8) were classed as significant, or *, where p < 0.05, and highly significant, or **, where p < 0.001.



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  		FIG. 1.
ATP potentiates the agrin-induced AChR aggregation. Chick (A) and C2C12 (B) myotubes cultured for 5 days were pretreated with apyrase, washed, and treated with the indicated 50 µM agonist (or with control medium), with or without the co-applied recombinant chick agrin (CBA-1; 2 units) for 16 h. In all cases, cultures were fixed by ice-cold 2% paraformaldehyde, 5% sucrose in PBS, stained with TMR-BuTX, and viewed under fluorescence optics. Control cultures did not have drug treatment. *, p < 0.05, significant difference versus control; **, p < 0.001, highly significant difference. Values represent the number of AChR aggregates per field in a single myotube and expressed as mean ± S.E., n = 4, each with triplicate samples. Bar, 20 µm.

 



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  		FIG. 2.
The ATP-potentiation effect is blocked by P2Y1-specific antagonist. A, chick or C2C12 myotubes were treated with agonist 2-MeSADP (50 µM), antagonist A2P5P (100 µM), and agrin (2 units) for 16 h as described in Fig. 1. B, cultured chick myotubes were transiently transfected with chicken P2Y1 cDNA for 4 days before the treatment of apyrase. Drugs (A2P5P at 100 µM; suramin 100 µM) and agrin (2 units) were added to the cultures for 16 h as in Fig. 1. In all cases, 50 µM 2-MeSADP was applied. AChR aggregates were counted and expressed as in Fig. 1. *, p < 0.05, significant difference versus control; **, p < 0.001, highly significant difference.

 



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  		FIG. 8.
P2Y receptor mediates the activation of Rac GTPases. A, 50 µM UTP or ADP was applied onto cultured chick myotubes for 16 h, with or without recombinant chick agrin (2 units). Values represent the number of AChR aggregates per field in a single myotube and are expressed as mean ± S.E., n = 4, each with triplicate samples. Bar, 20 µm. B and C, five-day-old chick myotubes were treated with 50 µM 2-MeSADP or UTP at different time periods, respectively, and 2 mg of cell lysates were assayed for RBD-bound RhoA (B) or PAK-bound Rac (C) as in Fig. 5. Total RhoA or Rac in 20 µg of lysate protein served as internal control. Antibodies against RhoA or Rac were used in both assays. The histograms show the quantitation from the blots by densitometry in arbitrary units; values are expressed as mean ± S.E., n = 4, each with triplicate samples. *, p < 0.05, significant difference versus control; **, p < 0.001, highly significant difference.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Potentiation of Agrin-induced AChR Aggregation by P2Y1 Nucleotide Receptors—Cultured chick and C2C12 myotubes were treated with apyrase to eliminate all free nucleotide in the cultures, which was prerequisite for all pharmacological analyses of nucleotide receptors (26). The application of agrin (2 units) induced an ~4- to 5-fold increase of AChR aggregates in both types of myotube cultures (Fig. 1, A and B). The size of these aggregates was larger than 10 µm, and the smaller aggregates (<10 µm) were not counted in here. In both types of myotubes, the formation of AChR aggregates, induced by agrin, was potentiated by an ~50% increase after the application of 50 µM P2Y1 agonists, ATP (a nonspecific agonist), or 2-Me-SADP (a specific agonist), to the cultured myotubes (Fig. 1). This concentration of P2Y agonists has been shown to be the optimized dosage in inducing the downstream signaling cascade, including the accumulation of inositol triphosphate and the phosphorylation of extracellular signal-regulated kinase (26, 27). In addition, the concentration used for P2Y agonists is within the physiological range of ATP at the synaptic cleft. The sizes of the aggregates remained constant throughout the drug treatments in both chick and C2C12 myotubes. The treatment of P2Y1 agonists slightly increased the basal spontaneous AChR aggregates. Adenosine, an agonist for P1 nucleotide receptor, did not show any potentiation effect upon the agrin-induced AChR aggregation (Fig. 1). Therefore, these results suggest that the potentiation effect of synaptic ATP in regulating the formation of agrin-induced AChR aggregates was mediated by P2Y1 receptor.

The specificity of the ATP response at the cultured myotubes was further confirmed by using P2Y1-specific antagonist. Application of A2P5P did not alter the formation of agrin-induced AChR aggregates, except for an insignificant reduction of the aggregate observed in C2C12 myotubes (Fig. 2A). The potentiation of agrin-induced AChR aggregation, mediated by 2-Me-SADP, was significantly reduced by the application of A2P5P in the cultures. To enhance the signals, cDNA-encoding chicken P2Y1 receptor was over-expressed in cultured myotubes by DNA transfection. The P2Y1 receptor over-expressed myotubes showed a high background of spontaneous AChR aggregates; nevertheless, the response to agrin was retained (Fig. 2B). Suramin, an antagonist for P2Y receptor, completely blocked the agrin-induced AChR aggregation in the receptor over-expressed myotubes, as well as the background aggregates induced by the over-expression of P2Y1 receptor. In parallel, A2P5P abolished the agrin-induced AChR aggregation.

We have found previously (26, 27) that activation by adenosine triand diphosphates of the P2Y1 receptors present in cultured chick myotubes leads to an increase in the expression of the AChR gene. The increased AChR number after the activation of P2Y1 receptors could explain the increase of the AChR aggregation as revealed here. To eliminate the contribution of newly synthesized AChR after the challenge of 2-MeSADP, the receptors were pre-labeled by TMR-BuTX before the application of agrin and/or the agonist. An increase of over 2-fold pre-labeled AChR aggregate was induced by agrin (Fig. 3). This 2-fold increase of AChR aggregation was much smaller than that of the induction revealed in the above analysis, when all of the AChRs were labeled after agrin treatment. This discrepancy could be due to the rapid turnover of the AChR, because the newly synthesized receptors were not counted in this prelabel study. The potentiation effect of ATP in agrin-induced AChR aggregation, however, was retained. An increase of over 4-fold AChR aggregation was revealed when 2-MeSADP was applied together with agrin (Fig. 3). This result revealed that the P2Y1-potentiated effect upon AChR aggregation was not due to the increased expression of AChR.



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  		FIG. 3.
The P2Y1-potentiated agrin-induced AChR aggregation does not require newly synthesized AChR. Chick myotubes were pre-labeled with TMR-BuTX (1 µM) for 2 h before the treatment with 2-MeSADP (50 µM) or agrin (2 units) for another 16 h. Cultures were fixed with ice-cold 2% paraformaldehyde and 5% sucrose in PBS and viewed under fluorescence optics. Values represent the number of AChR aggregates per field in a single myotube and are expressed as mean ± S.E., n = 4, each with triplicate samples. Bar, 20 µm.

 
Activation of RhoA Is Mediated by ATP and P2Y1 Receptors— Small GTPase is known to play roles in the organization of cytoskeletal elements in cells, as well as in the aggregation of post-synaptic AChRs at vertebrate nmjs. Therefore, the functional role of RhoA was determined here. The localization of RhoA protein in skeletal muscle was determined by using a monoclonal antibody against RhoA. The RhoA immunoreactivity was co-localized with the binding of TMR-BuTX, indicating the restricted localization of RhoA at adult rat nmjs (Fig. 4). That location of RhoA was not only revealed in rat muscle, but it was also co-localized with AChR in adult chicken muscle. A weak staining by anti-RhoA antibody could still be observed in some extra-junctional areas in both species.



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  		FIG. 4.
Synaptic localization of RhoA in skeletal muscles. The localization of RhoA at the nmjs. Rat and chicken muscle sections (20 µm) were used. For each, the same field is shown stained by the anti-RhoA antibody or for AChR by TMR-BuTX. Bar, 20 µm.

 
The activation of RhoA was examined after stimulating the endogenous P2Y1 receptors in cultured myotubes. Active RhoA is known to re-localize from a cytosolic form to a membrane-bound form. Thus, the P2Y1-activated RhoA was determined in the membrane fraction of cultured myotubes. The antibody to RhoA readily detected a band at ~21 kDa in chick myotubes (Fig. 5A, upper panel). In chick myotubes, ATP and 2-MeSADP induced a transient activation of RhoA; the level of membrane-bound RhoA was increased after 40 min of the agonist challenge. Plots of scanned data from four independent experiments show the transient activation peaking at 2- to 3-fold the basal level and peaking at 40 min of exposure to P2Y1-agonist treatment (Fig. 5A, lower panel). The time-course of P2Y1-induced RhoA activation shared a close similarity to that of activation in vascular myocytes (28). The RhoA activation, however, was more robust when it was activated by ATP rather than by 2-MeSADP. The RhoA activation in P2Y1-treated cultures was also determined by using a commercial assay kit in quantifying the amount of RBD-bound RhoA. The application of either ATP or 2-MeSADP increased the amount of RBD-bound RhoA in a time-dependent manner. Again, the RhoA activation was more robust when it was activated by ATP as compared with that of 2-MeSADP (Fig. 5B). The maximum activation, mediated by ATP challenge, reached ~4-fold after 30 min of the treatment (Fig. 5B, lower panel), which was in line with the assay that detected the membrane-bound RhoA; however, the peak was slightly earlier in the RBD-bound RhoA assay. The total amount of RhoA in the drug-treated cultures was not changed, which thus served as an internal control.



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  		FIG. 5.
P2Y1 receptor agonist stimulates the activation of RhoA GTPase. Five-day-old chick myotubes were treated with 50 µM ATP or 2-MeSADP, respectively, and 2 mg of cell lysates were assayed for RhoA activation. A, in the membrane localization assay, activation of RhoA was measured by the increase of endogenous RhoA that migrated to cellular membranes. 20 µg of membrane protein was used. Ponceau S staining indicates equal loading in each lane. B, in the RBD-bound RhoA assay, total RhoA in 20 µg of lysate protein served as internal control. C, five-day-old chick myotubes were treated with increasing doses of ATP or 2-MeSADP. The membrane-bound RhoA was assayed as in A. D, P2Y1 cDNA was transfected into cultured chick myotubes. After 4 days of transfection, A2P5P (50 or 100 µM) was co-treated with 2-Me-SADP (50 µM) for 30 min. RhoA assay was done as in A. Anti-RhoA antibody was used in these assays. In all cases, the histograms/graphs show the quantitation from the blots of activated RhoA by densitometry in arbitrary units; values are expressed as mean ± S.E., n = 4, each with triplicate samples.

 
The ATP- or 2-MeSADP-induced RhoA activation occurred in a dose-dependent manner; maximum activation was revealed at 50 µM P2Y1 in agonist-treated chick myotubes (Fig. 5C). The over-expression of P2Y1 receptor in cultured myotubes caused a marked increase of the membrane-bound form of activated RhoA, as expected (Fig. 5D). This RhoA activation was blocked by A2P5P in a dose-dependent manner.

RhoA Is a Downstream Signal of Agrin and P2Y1 Nucleotide Receptor—By using both membrane-bound and RBD-bound RhoA activation assays, application of agrin in the cultured chick myotubes activated RhoA by ~4-fold in a time-dependent manner. Because the activation was transient, a decline of activation was observed after the maximum activation at ~10 min (Fig. 6A). This result was in line with a previous report (12), except the elapsed time here is slightly shorter, which could be a result of the difference in myotube species. Weston et al. (12) used C2C12 myotubes, and we used primary chick myotubes. In addition, this time course of RhoA activation was very similar for both types of assays in measuring RhoA activation. Moreover, the time for agrin-induced maximal activation of RhoA was shorter than that of ATP induction, i.e. 10 min instead of 40 min. In addition, the agrin-induced RhoA activation in cultured myotubes occurred in a dose-dependent manner (Fig. 6B); the dosage effect was seen in both types of RhoA assays.



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  		FIG. 6.
Agrin stimulates RhoA activation. A, agrin-induced RhoA activation in chick myotube cultures was measured by the increase of membrane-bound form (left) and RBD-bound form (right) of endogenous RhoA. Five-day-old chick myotubes were treated with agrin (2 units) at different time periods, and lysates were collected for assays. B, agrin at concentrations ranging from 0 to 2 units was applied onto 5-day-old cultured chick myotubes for 10 min. The RhoA activation was measured by both of the assays as in Fig. 5. The histograms show the quantitation from the blots by densitometry in arbitrary units; values are expressed as mean ± S.E., n = 4, each with triplicate samples.

 
The RhoA activation mediated by agrin and P2Y1 receptor showed an additive effect. The agrin-induced RhoA activation could be potentiated by the co-applied 2-MeSADP in cultured myotubes (Fig. 7A), which suggested that RhoA could be the link between these two signaling pathways. The role of RhoA in P2Y1-potentiated agrin effect was further demonstrated in DNA transfection analysis. In RhoA-expressing cultured myotubes, the formation of AChR aggregates induced by agrin was enhanced by >50% (Fig. 7B). In parallel, the potentiation effect of 2-MeSDAP in agrin-induced AChR aggregate formation was also enhanced by ~30%. When transfected into the cultures, the cDNA-encoding dominant-negative RhoA (RhoAN19) completely abolished the formation of AChR aggregation either in agrin-applied or agrin/2-MeSADP-co-applied cultures (Fig. 7B).



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  		FIG. 7.
RhoA activation is a link in the signaling between P2Y1 receptor and agrin. A, five-day-old chick myotubes were treated with 20 µM 2-MeSADP with or without a sub-maximal amount of agrin (1 unit). The amount of RBD-bound RhoA was measured as in Fig. 5. The histograms show the quantitation from the blots by densitometry in arbitrary units, and values are expressed as mean ± S.E., n = 4, each with triplicate samples. B, two-day-old chick cultures on a 35-mm dish were transfected with 2 µg of RhoAWT, RhoAN19, or pcDNA 3 (control). Two days later, 2-MeSADP (50 µM) was applied to the cultures in the presence and absence of agrin (2 units) for 16 h. AChR aggregates were stained and examined as in Fig. 1. Values represent the number of AChR aggregates per field in a single myotube; they are expressed as mean ± S.E., n = 4, each with triplicate samples.

 
The existence of other P2Y receptor subtypes at vertebrate nmjs was reported (24), which could also mediate the ATP-potentiated effect in AChR aggregation at the nmjs. Application of UTP, a specific agonist for P2Y2 and P2Y4 receptors, potentiated the agrin-induced AChR aggregation by ~50% in cultured chick myotubes (Fig. 8A). In contrast, application of ADP, a stronger agonist for P2Y3 and P2Y5 receptors in chicken species, did not show any potentiation effect upon AChR aggregation. Moreover, RhoA activation was not only restricted to P2Y1-specific agonist; the application of UTP increased the amount of RBD-bound RhoA by ~5-fold (Fig. 8B). The UTP-induced RhoA activation occurred in a time-dependent manner; the peak of RhoA activation was less than 15 min, which was faster than that for 2-MeSADP. Another small GTPase Rac was also tested in our culture system. The activated form of Rac (PAK-bound Rac) was increased by 3- to 4-fold in 2-MeSADP-applied myotubes in a time-dependent manner (Fig. 8C). The peak of activation was at ~10 min, which was markedly shorter than the RhoA activation triggered by the same agonist. Additionally, the application of UTP, a specific agonist for P2Y2 and P2Y4 receptors, also showed a similar time-dependent manner in activating the PAK-bound Rac in cultured myotubes (Fig. 8C).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
At vertebrate nmjs, ATP is constantly released by the motor neuron, and the concentration could reach up to 1 mM at the synaptic cleft, which therefore bombards the postsynaptic membrane. This high concentration of ATP at the cleft is maintained whenever the contraction of muscle is triggered. The response of synaptic ATP at the nmjs is strongly supported by the existence of P2Y receptors at the post-synaptic muscle, which in particular has a high level of expression in adult skeletal muscles (23, 24, 26). Together with previous studies, we hypothesize that ATP could have functional roles in affecting the neuromuscular transmission in short- and long-term aspects.

In the short-term effect, ATP potentiates the muscle responses to ACh as well as the synaptic current (34, 35). The long-term effect of synaptic ATP is to affect the formation and maintenance of post-synaptic specializations at the nmjs. First, the degradation rate of AChR in rat muscle tissue culture was modulated by exogenously applied ATP. Although a high dose of ATP was used in the study, the induction was possibly mediated by post-synaptic P2Y receptors, because the effect could be mimicked by the application of phospholipase A2 activator (36). Second, the transcriptional activity of genes encoding AChE and different subunits of AChR were induced by the application of ATP, or P2Y1-specific agonist in cultured myotubes (26); the effective concentration of the applied agonist was in the µM range. Application of P2Y1-specific agonist induced the accumulation of intracellular inositol triphosphate and Ca2+ in cultured myotubes. This activation subsequently triggered a downstream mitogen-activated protein kinase signaling pathway involving Raf-1, extracellular signal-regulated kinase, and Elk-1; this process has been described as responsible for the post-synaptic gene activation (27). Here, we further extend the long-term functional role of synaptic ATP and its P2Y receptors in potentiating the formation of AChR aggregates mediated by agrin. Synaptic ATP activates P2Y receptors at the nmjs and subsequently activates the intracellular small GTPase, e.g. RhoA and Rac; this activation potentiates the role of agrin in directing the formation and maintenance of the post-synaptic specializations. This long-term effect of synaptic ATP at the nmjs is mainly to strengthen the post-synaptic specialization, i.e. increase the expression and aggregation of AChR/AChE at the junction. Indeed, this proposed function of the synaptic ATP is strongly supported by the increase of TMR-BuTX-stained areas at the nmjs after high-intensity of exercise (37) that resulted in a high-frequency release of the synaptic ATP.

Different types of P2Y receptors could mediate the postsynaptic response of ATP; in particular, these P2Y receptors share very similar downstream signaling mechanisms. The recognition of these P2Y receptors is dependent upon their pharmacological properties. By using P2Y1-specific agonist (2-MeSADP) and antagonist (A2P5P), we clearly show that P2Y1 is one of the P2Y receptors that can account for the potentiation effect. On the other hand, UTP, a strong agonist for P2Y2 receptor, also activated the RhoA, which suggests the possible role of post-synaptic P2Y2 receptor. Indeed, P2Y2 receptor was detected in skeletal muscles and co-localized with AChR at the adult rat and chicken nmjs.2 The pharmacological properties of P2Y4 receptor show a close similarity with that of P2Y2 receptor; UTP is known to be the strongest agonist for these two receptors. Although the expression of P2Y4 transcript was reported by using RT-PCR in rat embryonic muscles (38), the expression of transcript-encoding P2Y4 receptor in rat muscle and C2C12 myotube was below the detection level in our Northern blot analysis.2 Lastly, the insensitivity of ADP in the cultured myotubes excluded the possible involvement of P2Y3 and P2Y5 receptors. In line with this observation, the expression of P2Y3 and P2Y5 receptors were not detected in skeletal muscles (39).

The signaling mechanism of agrin-induced Rac and RhoA activation in cultured myotubes has been described previously (11, 12). The agrin-induced Rac activation is rapid and transient, and it has been proposed to be a prerequisite for the activation of RhoA. Moreover, Rac is proposed to play a role in the formation of agrin-induced small AChR aggregates; RhoA is required, subsequently, to condense those small aggregates as large agrin-induced AChR aggregates. The small GTPases are known to induce the re-arrangement of intracellular cytoskeleton (40), which, therefore, could explain the clustering of AChRs in the post-synaptic muscle (12, 41). Here, we demonstrate further the potentiation role of P2Y1 receptor in agrin-induced AChR aggregation could also involve the small GTPases. In cultured vascular myocytes, the activation of P2Y receptors including P2Y1, P2Y2, P2Y4, and P2Y6 are known to induce RhoA and RhoA kinase activation, formation of actin stress fiber and an increase in F- to G-actin ratio (28). Similar to the action of agrin, P2Y1 receptor is also involved in the transient activation of RhoA and Rac in cultured myotubes. Regardless of the activation of the small GTPases by ATP application in cultured myotubes, the P2Y-mediated GTPase activation is not sufficient to cause the aggregation of AChR, i.e. ATP application by itself has no significant effect upon AChR aggregation, at least below our detection level. This suggests that other signals, besides small GTPase, could also be involved in agrin-induced AChR aggregation, and ATP is playing a synergistic role with agrin. Moreover, the time course of P2Y1-mediated Rac activation (maximum at ~10 min) preceded the RhoA activation (maximum at ~40 min). Although we have not determined the role of P2Y1 receptor in the formation of agrin-induced small AChR aggregates, the activation of Rac by 2-MeSADP suggests the possible role of P2Y1 receptor in directing the formation of these small aggregates.

The signaling cascade of small GTPase in the agrin/MuSK-induced AChR aggregation in muscle has been determined. Downstream of the small GTPase may be PAK, a cytoplasmic kinase involved in cytoskeleton regulation (22), whereas the activation of GGT could occur upstream of GTPase. In C2C12 myotubes, GGT was rapidly tyrosine phosphorylated with an increase of activity after agrin/MuSK stimulation (21). The activation of GGT results in prenylation of GTPase (e.g. RhoA and Rac), and which subsequently causes the migration of GTPase from cytosol to membrane in activated form. Whether the activation of P2Y receptors in muscle could result in GGT activation, however, has not been determined.

The explanation for the ATP potentiation effect in agrin signaling could be not only the activation of small GTPase; other downstream signals of P2Y receptor may also play roles in mediating this post-synaptic specialization. In fibroblasts or astrocytes, the activation of RhoA could result in rapid tyrosine phosphorylation of cytoskeleton-associated proteins (42), which subsequently may generate binding sites for Src and phosphatidylinositol 3-kinase (43). Indeed, Src kinases are known to be involved in agrin/MuSK signaling in muscle (16, 17, 20). Moreover, application of P2Y agonist triggered the intracellular Ca2+ mobilization and inositol triphosphate accumulation in muscle; thus, intracellular signal changes might also be required for the action of agrin in directing AChR aggregation (13–15).

We also note that the P2Y1 receptor is, exceptionally for P2Y receptors, widely expressed on brain neurons (44). ATP is known to be generally co-released at central and peripheral cholinergic and bioaminergic (and even some GABA-ergic; Ref. 45) neuronal synapses. Further, it has been reported that an ionotropic ATP receptor, P2X7, is widespread at brain excitatory pre-synaptic terminals (46), suggesting ATP co-transmission there. Hence, further investigation is indicated as to whether the post-synaptic actions of the P2Y1 receptor now being uncovered at the nmj have a wider relevance at such central synapses.


   FOOTNOTES
 
* This work was supported by Grants HKUST 6098/02 M and 6283/03 M from the Research Grants Council of Hong Kong (to K. W. K. T.). 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. Back

{ddagger} Recipient of a Croucher Foundation Scholarship. Back

§ Supported by the post-doctoral matching fund from Hong Kong University of Science and Technology. Back

To whom correspondence should be addressed: Dept. of Biology, Hong Kong University of Science and Technology, Clear Water Bay Road, Kowloon, Hong Kong SAR, China. Tel.: 852-2358-7332; Fax: 852-2358-1559; E-mail: botsim{at}ust.hk.

1 The abbreviations used are: nmj, neuromuscular junction; AChR, acetylcholine receptor; AChE, acetylcholinesterase; MuSK, muscle-specific receptor tyrosine kinase; GTPase, guanosine triphosphatase; ACh, acetylcholine; TMR-BuTX, tetramethylrhodamine-conjugated {alpha}-bungarotoxin; PBS, phosphate-buffered saline; RBD, Rho-binding domain; PAK, p21-activated kinase; 2-MeSADP, 2-methylthioadenosine 5'-diphosphate. Back

2 E. K. K. Tung, unpublished results. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Tina Dong and H. Y. Choi from our laboratory for their expert technical assistance.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Gautam, M., Noakes, P. G., Moscoso, L., Rupp, F., Scheller, R. H., Merlie, J. P., and Sanes, J. R. (1996) Cell 85, 525–535[CrossRef][Medline] [Order article via Infotrieve]
  2. Pun, S., and Tsim, K. W. K. (1997) Mol. Cell. Neurosci. 10, 87–99[CrossRef][Medline] [Order article via Infotrieve]
  3. Burgess, R. W., Nguyen, Q. T., Son, Y. J., Lichtman, J. W., and Sanes, J. R. (1999) Neuron 23, 33–44[CrossRef][Medline] [Order article via Infotrieve]
  4. Sanes, J. R., and Lichtman, J. W. (1999) Annu. Rev. Neurosci. 22, 389–442[CrossRef][Medline] [Order article via Infotrieve]
  5. Lin, W., Burgess, R. W., Dominguez, B., Pfaff, S. L., Sanes, J. R., and Lee, K. F. (2001) Nature 410, 1057–1064[CrossRef][Medline] [Order article via Infotrieve]
  6. DeChiara, T. M., Bowen, D. C., Valenzuela, D. M., Simmons, M. V., Poueymirou, W. T., Thomas, S., Kinetz, E., Compton, D. L., Rojas, E., Park, J. S., Smith, C., DiStefano, P. S., Glass, D. J., Burden, S. J., and Yancopoulos, G. D. (1996) Cell 85, 501–512[CrossRef][Medline] [Order article via Infotrieve]
  7. Glass, D. J., Bowen, D. C., Stitt, T. N., Radziejewski, C., Bruno, J., Ryan, T. E., Gies, D. R., Shah, S., Mattsson, K., Burden, S. J., DiStefano, P. S., Valenzuela, D. M., DeChiara, T. M., and Yancopoulos, G. D. (1996) Cell 85, 513–523[CrossRef][Medline] [Order article via Infotrieve]
  8. Fuhrer, C., Gautam, M., Sugiyama, J. E., and Hall, Z. W. (1999) J. Neurosci. 19, 6405–6416[Abstract/Free Full Text]
  9. Watty, A., Neubauer, G., Dreger, M., Zimmer, M., Wilm, M., and Burden, S. J. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 4585–4590[Abstract/Free Full Text]
  10. Gautam, M., Noakes, P. G., Mudd, J., Nichol, M., Chu, G. C., Sanes, J. R., and Merlie, J. P. (1995) Nature 377, 232–236[CrossRef][Medline] [Order article via Infotrieve]
  11. Weston, C., Yee, B., Hod, E., and Prives, J. (2000) J. Cell Biol. 150, 205–212[Abstract/Free Full Text]
  12. Weston, C., Gordon, C., Teressa, G., Hod, E., Ren, X. D., and Prives, J. (2003) J. Biol. Chem. 278, 6450–6455[Abstract/Free Full Text]
  13. Megeath, L. J., and Fallon, J. R. (1998) J. Neurosci. 18, 672–678[Abstract/Free Full Text]
  14. Borges, L. S., Lee, Y., and Ferns, M. (2002) J. Neurobiol. 50, 69–79[CrossRef][Medline] [Order article via Infotrieve]
  15. Megeath, L. J., Kirber, M. T., Hopf, C., Hoch, W., and Fallon, J. R. (2003) Neuroscience 122, 659–668[CrossRef][Medline] [Order article via Infotrieve]
  16. Smith, C. L., Mittaud, P., Prescott, E. D., Fuhrer, C., and Burden, S. J. (2001) J. Neurosci. 21, 3151–3160[Abstract/Free Full Text]
  17. Mohamed, A. S., Rivas-Plata, K. A., Kraas, J. R., Saleh, S. M., and Swope, S. L. (2001) J. Neurosci. 21, 3806–3818[Abstract/Free Full Text]
  18. Jones, M. A., and Werle, M. J. (2000) Mol. Cell. Neurosci. 16, 649–660[CrossRef][Medline] [Order article via Infotrieve]
  19. Jones, M. A., and Werle, M. J. (2004) Mol. Cell. Neurosci. 25, 195–204[CrossRef][Medline] [Order article via Infotrieve]
  20. Strochlic, L., Cartaud, A., Labas, V., Hoch, W., Rossier, J., and Cartaud, J. (2001) J. Cell Biol. 153, 1127–1132[Abstract/Free Full Text]
  21. Luo, Z. G., Je, H. S., Wang, Q., Yang, F., Dobbins, G. C., Yang, Z. H., Xiong, W. C., Lu, B., and Mei, L. (2003) Neuron 40, 703–717[CrossRef][Medline] [Order article via Infotrieve]
  22. Luo, Z. G., Wang, Q., Zhou, J. Z., Wang, J., Luo, Z., Liu, M., He, X., Wynshaw-Boris, A., Xiong, W. C., Lu, B., and Mei, L. (2002) Neuron 35, 489–505[CrossRef][Medline] [Order article via Infotrieve]
  23. Tsim, K. W. K., and Barnard, E. A. (2002) Neurosignals 11, 58–64[CrossRef][Medline] [Order article via Infotrieve]
  24. Tsim, K. W. K., Choi, R. C. Y., Siow, N. L., Ling, K. K. Y., Jiang, J. X. S., Tung, E. K. K., Lee, H. C., Xie, Q. H., Simon, J., and Barnard, E. A. (2003) J. Neurocytol. 32, 603–617[CrossRef][Medline] [Order article via Infotrieve]
  25. Silinsky, E. M., and Redman, R. S. (1996) J. Physiol. 492, 815–822[Abstract/Free Full Text]
  26. Choi, R. C. Y., Man, M. L. S., Ling, K. K. Y., Ip, N. Y., Simon, J., Barnard, E. A., and Tsim, K. W. K. (2001) J. Neurosci. 21, 9224–9234[Abstract/Free Full Text]
  27. Choi, R. C., Siow, N. L., Cheng, A. W., Ling, K. K., Tung, E. K., Simon, J., Barnard, E. A., and Tsim, K. W. (2003) J. Neurosci. 23, 4445–4456[Abstract/Free Full Text]
  28. Sauzeau, V., Le Jeune, H., Cario-Toumaniantz, C., Vaillant, N., Gadeau, A., Desgranges, C., Scalbert, E., Chardin, P., Pacaud, P., and Lairand, G. (2000) Am. J. Physiol. 278, H1751–H1761
  29. Siow, N. L., Choi, R. C. Y., Cheng, A. W. M., Jiang, J. X. S., Wan, D. C. C., Zhu, S. Q., and Tsim, K. W. K. (2002) J. Biol. Chem. 277, 36129–36136[Abstract/Free Full Text]
  30. Tsim, K. W. K., Ruegg, M. A., Escher, G., Kröger, S., and McMahan, U. J. (1992) Neuron 8, 677–689[CrossRef][Medline] [Order article via Infotrieve]
  31. Webb, T. E., Simon, J., Krishek, B. J., Bateson, A. N., Smart, T. G., King, B. F., Burnstock, G., and Barnard, E. A. (1993) FEBS Lett. 324, 219–225[CrossRef][Medline] [Order article via Infotrieve]
  32. Feig, L. A. (1999) Nat. Cell Biol. 1, 25–27
  33. Wallace, B. G. (1986) J. Cell Biol. 102, 783–794[Abstract/Free Full Text]
  34. Lu, Z., and Smith, D. O. (1991) J. Physiol. 436, 45–56[Abstract/Free Full Text]
  35. Fu, W. M., Chan, Y. H., Lee, K. F., and Liou, J. C. (1997) Eur. J. Neurosci. 9, 676–685[CrossRef][Medline] [Order article via Infotrieve]
  36. O'Malley, J. P., Moore, C. T., and Salpeter, M. M. (1997) J. Cell Biol. 138, 159–165[Abstract/Free Full Text]
  37. Deschenes, M. R., Maresh, C. M., Crivello, J. F., Armstrong, L. E., Kraemer, W. J., and Covault, J. (1993) J. Neurocytol. 22, 603–615[CrossRef][Medline] [Order article via Infotrieve]
  38. Cheung, K. K., Ryten, M., and Burnstock, G. (2003) Dev. Dyn. 228, 254–266[CrossRef][Medline] [Order article via Infotrieve]
  39. Webb, T. E., Henderson D., King, B. F., Wang, S., Simon, J., Bateson, A. N., Burnstock, G., and Barnard, E. A. (1996) Mol. Pharmacol. 50, 258–265[Abstract]
  40. Hall, A. (1998) Science 280, 2074–2075[Free Full Text]
  41. Moransard, M., Borges, L. S., Willmann, R., Marangi, P. A., Brenner, H. R., Ferns, M. J., and Fuhrer, C. (2003) J. Biol. Chem. 278, 7350–7359[Abstract/Free Full Text]
  42. Ramakers, G. J., and Moolenaar, W. H. (1998) Exp. Cell Res. 245, 252–262[CrossRef][Medline] [Order article via Infotrieve]
  43. Chen, H. C., Appeddu, P. A., Isoda, H., and Guan, J. L. (1996) J. Biol. Chem. 271, 26329–26334[Abstract/Free Full Text]
  44. Moore, D., Chambers, J., Waldvogel, H., Faull, R., and Emson, P. (2000) J. Comp. Neurol. 421, 374–384[CrossRef][Medline] [Order article via Infotrieve]
  45. Jo, Y. H., and Schlichter, R. (1999) Nat. Neurosci. 2, 241–245[CrossRef][Medline] [Order article via Infotrieve]
  46. Deuchars, S. A., Atkinson, L., Brooke, R. E., Musa, H., Milligan, C. J., Batten, T. F., Buckley, N. J., Parson, S. H., and Deuchars, J. (2001) J. Neurosci. 21, 7143–7152[Abstract/Free Full Text]
  47. Glantz, S. A. (1988) Primer of Biostatistics, McGraw-Hill, New York

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