Retrograde regulation of motoneuron differentiation by ... - Nature

© 2008 Nature Publishing Group http://www.nature.com/natureneuroscience

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Retrograde regulation of motoneuron differentiation by muscle b-catenin Xiao-Ming Li1–3, Xian-Ping Dong1,3, Shi-Wen Luo1, Bin Zhang1, Dae-Hoon Lee1, Annie K L Ting1, Hannah Neiswender1, Chang-Hoon Kim1, Ezekiel Carpenter-Hyland1, Tian-Ming Gao2, Wen-Cheng Xiong1 & Lin Mei1,2 Synapse formation requires proper interaction between pre- and postsynaptic cells. In anterograde signaling, neurons release factors to guide postsynaptic differentiation. However, less is known about how postsynaptic targets retrogradely regulate presynaptic differentiation or function. We found that muscle-specific conditional knockout of b-catenin (Ctnnb1, also known as b-cat) in mice caused both morphologic and functional defects in motoneuron terminals of neuromuscular junctions (NMJs). In the absence of muscle b-catenin, acetylcholine receptor clusters were increased in size and distributed throughout a wider region. Primary nerve branches were mislocated, whereas secondary or intramuscular nerve branches were elongated and reduced in number. Both spontaneous and evoked neurotransmitter release was reduced at the mutant NMJs. Furthermore, short-term plasticity and calcium sensitivity of neurotransmitter release were compromised in b-catenin–deficient muscle. In contrast, the NMJ was normal in morphology and function in motoneuron-specific b-catenin–deficient mice. Taken together, these observations indicate a role for muscle b-catenin in presynaptic differentiation and function, identifying a previously unknown retrograde signaling in the synapse formation and synaptic plasticity.

Interactions between motoneurons and muscle fibers are essential to NMJ formation1–5. Innervation by motoneurons leads to high concentration of acetylcholine receptors (AChRs) in postjunctional membranes of muscle fibers, a complex process that involves AChR aggregation in subsynaptic areas, dispersion of nonsynaptic AChR-rich sites and local AChR synthesis. The anterograde signals include agrin, a polypeptide used by motoneurons to cluster AChRs6,7 and ACh, which, via activating muscle fibers, suppresses AChR subunit gene expression and disassembles AChR clusters in nonsynaptic areas8–11. Studies of effects of wing bud removal on motoneurons have demonstrated the existence of neurotrophic factors from the muscle that are essential for motoneuron survival12. It has become evident that muscles also generate signals to regulate differentiation and function of presynaptic terminals5. The identity of the retrograde signals remains unknown. Wnt is a family of secreted glycoproteins that have important roles in the development and maturation of the nervous system, including brain patterning, axon guidance and synapse formation13. For example, Wnt7a, secreted by cerebellar granule cells, regulates terminal arborization and presynaptic differentiation of mossy fibers in the mouse cerebellum14. At the glutamatergic NMJ in Drosophila, Wnt is secreted by synaptic boutons. Defects were observed in both presynaptic and postsynaptic differentiations in Wnt and Frizzled mutants15,16. Recently, signaling proteins of the Wnt canonical pathway, including dishevelled (Dvl) and adenomatous polyposis coli, have been

implicated at developing mammalian NMJs17,18. We find it interesting that inhibition of Dvl function in muscle cells not only attenuates AChR clustering, but also causes a reduction in the frequency of spontaneous synaptic currents in neuromuscular synapses in culture17. In this study, we investigate the role of b-catenin in NMJ development and function. b-catenin is a signaling protein downstream of Dvl. After Wnt activation of the canonical pathway, Dvl inhibits b-catenin phosphorylation by GSK3b, resulting in b-catenin accumulation in the cytoplasm and nucleus. Subsequently, b-catenin regulates gene expression by association with T cell factor/LEF1 (ref. 19). Because the b-catenin null mutation causes embryonic lethality20, we generated b-catenin muscle-specific deficient mice by a loxP/cre strategy. Studies of NMJs of b-catenin–deficient mice indicate that muscle b-catenin is essential for NMJ development and function, identifying a previously unknown retrograde signaling downstream of b-catenin. RESULTS Specific deletion of b-catenin in skeletal muscles Mutant mice deficient for b-catenin specifically in skeletal muscles were generated by a loxP/cre strategy. b-catfloxed mice carry an allele of the b-catenin gene where a loxP site was inserted in intron 1 and a loxP-flanked neomycin-TK cassette was inserted downstream in intron 6 (Supplementary Fig. 1 online), which is suitable for Cremediated gene inactivation21. HSA-Cre mice express the Cre gene

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of Developmental Neurobiology, Institute of Molecular Medicine and Genetics, Department of Neurology, Medical College of Georgia, 1120 15th Street, Augusta, Georgia 30912, USA. 2Department of Anatomy and Neurobiology, Southern Medical University, 1023 S Shatai Road, Guangzhou, 510515, China. 3These authors contributed equally to this work. Correspondence should be addressed to L.M. ([email protected]). Received 29 December 2007; accepted 23 January 2008; published online 17 February 2008; doi:10.1038/nn2053

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under the control of the human skeletal a-actin (ACTA1, also known as HSA) promoter22. This promoter is active at embryonic day 9.5 (E9.5) in the myotomal region of somites and drives Cre expression specifically in striated muscles22,23. To monitor the specificity of Cre recombination, HSA-Cre mice were crossed with Rosa26lacZ flox transgenic mice, in which the expression of the lacZ gene is activated by Cre recombination24. b-Galactosidase staining was detected in the muscle, but not in the spinal cord or other tissues (Supplementary Fig. 2 online and data not shown). All fibers stained positive for b-galactosidase, suggesting homogenous expression (Supplementary Fig. 2). Thus, HSA-Cre induces Cre recombination specifically in the muscle. Crossing HSA-Cre mice and b-cat floxed mice generated offspring whose b-cat gene is inactivated specifically in the muscle (referred to as HSA–b-cat –/–; Supplementary Fig. 1). Unless otherwise indicated, the control was b-cat loxP/loxP mice of the same litter. Some HSA–b-cat –/– pups died a few hours after birth with signs of cyanosis, probably as a result of a breathing difficulty. Consistent with this notion, the lungs in HSA–b-cat –/– mice were not properly inflated (Supplementary Fig. 1). Western blot analysis revealed specific reduction of b-catenin in HSA–b-cat –/– muscles, but not in the spinal cord, brain, liver or kidney (Supplementary Fig. 1). In agreement with HSA promoter activity, b-catenin was reduced at E13 (Supplementary Fig. 1). The nuclei of muscle fibers in HSA–b-cat –/– mice were located beneath the membrane, not in the central region, suggesting that muscle regeneration was minimal (Supplementary Fig. 1). Muscle fiber size was similar between control and mutant mice (Supplementary Fig. 1). In addition, muscle contractile units appeared to be normal in HSA–b-cat –/– mice (Supplementary Fig. 1). These results

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** Figure 1 Phrenic nerve branch mislocation and 400 ** ** 400 400 40 enlarged central band of AChR clusters in HSA–b200 cat –/– mice. (a) Whole-mount staining of left hemi200 200 20 ** diaphragms of P0 control (left) and HSA–b-cat –/– 0 0 0 0 (right) mice. AChR was labeled with R-BTX (red) and nerves were stained with rabbit antibody to neurofilament/synaptophysin (NF/Syn, green) that were visualized by FITC-conjugated antibodies to rabbit antibody. Asterisks, phrenic nerve entrance points to the diaphragm; arrows, primary nerve branches; arrowheads, secondary nerve branches; D, dorsal; V, ventral; L, lateral; M, medial. (b) Enlarged images of ventral regions of left hemi-diaphragm in a. (c) Enlarged images of AChR clusters around primary branches. The length of the secondary branches was defined as the distance between their roots on primary branches and the point where tertiary branches form, as indicated between two arrowheads. (d) Wider endplate regions in HSA–b-cat –/– muscles. A polygon was drawn to include most peripheral AChR clusters and the myotube length contained in the polygon was measured (h). (e) Increased distance of primary branches from the midline of muscle fibers in HSA–b-cat –/– mice. ** P o 0.01 versus control (n ¼ 6, t-test). (f) Decreased number of secondary/intramuscular nerve branches in HSA–b-cat –/– muscles. ** P o 0.01 versus control (n ¼ 6, t-test). (g) Increased length of secondary branches. ** P o 0.01 versus control (n ¼ 4, t-test). (h) Increased endplate band width. ** P o 0.01 versus control (n ¼ 8, t-test). Primary branch location (µm)


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suggest that b-catenin deficit does not change the overall structure of muscle fibers or cause muscle degeneration in HSA–b-cat –/– mice. Morphological defects in HSA–b-cat –/– NMJs To investigate NMJ defects in HSA–b-cat –/– mice, we whole-mount stained diaphragms for AChR and nerve terminals. In the ventral half of the left hemi-diaphragm of postnatal day 0 (P0) control (b-cat loxP/loxP) littermates, primary phrenic branches traveled in the central region perpendicular to muscle fibers, sending out short secondary or intramuscular branches (Fig. 1)5. AChR clusters, visualized by rhodamineconjugated a-bungarotoxin (R-BTX, red), are localized within 0.5 mm of the primary branches, forming a central band of endplates5. However, in HSA–b-cat –/– mice, primary nerve branches were no longer in the middle of muscle fibers, but were instead mislocated in the tendon region close to the central cavity (Fig. 1a,b, arrows; Fig. 1e). The secondary or intramuscular nerve branches were longer and fewer in HSA–b-cat –/– mice (Fig. 1b,c). The number of secondary branches was decreased from 45.2 ± 5.1 in control to 7.3 ± 2.1 in HSA–b-cat –/– mice (P o 0.01, n ¼ 6), with the length being increased from 24 ± 18 mm to 250 ± 140 mm (P o 0.01, n ¼ 4) (Fig. 1f,g). These results indicate that presynaptic differentiation is defective in the absence of muscle b-catenin. Despite the mislocation of primary phrenic nerve branches, AChR clusters remained in the middle region of muscle fibers in HSA–b-cat –/– mice (Fig. 1a,b). However, the clusters were distributed over a wider area, forming a larger endplate band (Fig. 1d,h). The band width was increased from 182 ± 44 mm in control to 325 ± 80 mm in conditional knockout mice (P o 0.01, n ¼ 8). Nevertheless, the clusters colocalized with rapsyn, a cytoplasmic protein whose mutation abolishes

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NMJ formation5,25 (Fig. 2). The area of individual endplates was increased by 67% (Fig. 2b). When the sizes of AChR clusters (in 5-mm increments) were plotted against the population, it was obvious that the curve was shifted to the right in HSA–b-cat –/– mice (Fig. 2c). Consistent with this increase in the area of individual clusters, their length was increased by 60% (14.7 ± 1.6 mm in control littermates and 23.5 ± 3.1 mm in HSA–b-cat –/– mice, P o 0.05, n ¼ 6; Fig. 2d). The results suggest that b-catenin may be involved in AChR clustering and differentiation of postsynaptic membranes at the NMJ. This notion is consistent with an earlier observation that agrin-induced AChR clustering is impaired when b-catenin is suppressed by siRNA in C2C12 myotubes26. However, agrininduced AChR clusters in cultured HSA–b-cat –/– muscles were similar in size and number to those in control muscles (Supplementary Fig. 3 online). The latter may be caused by in vivo adaptation or redundant compensatory mechanisms in HSA–b-cat –/– mice. To determine whether a loss of b-catenin in muscle regulates initial formation and subsequent consolidation of pre-pattern AChRs, we examined NMJs in HSA–b-cat –/– mice at ages between E14.5, when initial AChR clusters form27,28, and P0, when mutant mice die. Endplate band width and cluster size were larger in HSA–b-cat –/– diaphragms as early as E14.5, indicating that there are defects in the initial formation of AChR clusters or postsynaptic membrane differentiation (Supplementary Fig. 4 online). At age of E12.5, when there are no detectable AChR clusters27,28, growth cones of phrenic nerve axons were in the central region of muscle fibers in HSA–b-cat –/– mice, similar to those in control mice. (Supplementary Fig. 4), suggesting that phrenic nerve terminals were able to reach the central region of diaphragms in mutant mice. As the NMJ developed, primary

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Figure 2 Characterization of postsynaptic differentiation in HSA–b-cat –/– mice. Left hemi-diaphragms of P0 control and HSA–b-cat –/– mice were stained as in Figure 1. (a) Colocalization of AChR and rapsyn. (b) Increase of AChR cluster size in Merge S100 R-BTX HSA–b-cat –/– mice. AChR clusters were traced 30 µm around the perimeters to calculate the area using LSM 5 Image Examiner software (Zeiss). ** P o 0.01 versus control (n ¼ 8, t-test). (c) Rightward shift of the distribution curve of AChR cluster sizes in HSA–b-cat –/– mice. (d) Increase in AChR cluster length in 200 µm 200 µm 10 µm HSA–b-cat –/– mutant mice. ** P o 0.05 versus control (n ¼ 6, t-test). (e) AChRa mRNA was localized in the central region of HSA–b-cat –/– muscles. Diaphragms (E18.5) were subjected to in situ hybridization with digoxygenin-labeled cRNA probe for AChRa. (f) AChE clusters were scattered over a wider region in HSA–b-cat –/– muscles. AChE activity was assayed in situ in left hemi-diaphragms (P0). (g) Diaphragms (P0) were whole-mount stained for S100 with antibody to S100, which was visualized with FITC-conjugated secondary antibodies. HSA–-cat –/–


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branches were dislocated toward the central cavity in HSA–b-cat –/– mice. Note that every AChR remained to be innervated (Fig. 3), despite branch dislocation. The cause of the primary branch mislocation remains unclear and warrants future investigation. In normal muscles, a few nuclei beneath the postsynaptic membrane are active in transcribing genes that encode proteins that are essential for NMJ formation and function5. For example, the mRNAs of AChR subunits are enriched in the synaptic region. To determine whether b-catenin deficiency affects the spatial pattern of gene expression in muscle, we carried out an in situ hybridization analysis using a probe specific to the AChRa subunit gene. Whole-mount in situ hybridization showed that AChRa subunit transcripts were, in general, localized in the middle region of the diaphragms in HSA–b-cat –/– mice (Fig. 2e). Acetylcholinesterase (AChE) is an extracellular protein that localizes with the AChR at the NMJ. AChE, which was visualized by in situ enzymatic assays, was enriched in the central region of muscle fibers (Fig. 2f). However, similar to AChR clusters, the region over which AChRa mRNA and AChE were distributed was wider in HSA–b-cat –/– mice than in control. In addition, S100, a Schwann cell–specific marker, colocalized with AChR clusters (Fig. 2g). These results indicate that synapse-specific transcription, localization of AChE and Schwann cell differentiation are apparently normal. Reduced ACh release in HSA–b-cat –/– NMJs The presynaptic phenotypes in b-catenin HSA–b-cat –/– mice suggest that there may be a role for muscle b-catenin in presynaptic differentiation. To test this hypothesis, we determined whether AChR clusters colocalize with presynatpic terminals by staining for SV2, a

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to respond to ACh or defects in vesicle release from presynaptic terminals. First, we determined whether muscle fibers in HSA–b-cat –/– 40 mice failed to form NMJs. Individual muscle ** R-BTX SV2 Merge 20 fibers were isolated and stained with R-BTX. The number of AChR clusters per muscle fiber 0 was the same in control littermates and in HSA–b-cat –/– mice (Fig. 4a,b). Considering 10 µm that every AChR cluster co-stained with SV2 2 mV 100 ms (Fig. 3a), these results indicate that each muscle fiber forms at least one neuromuscular loxP/loxP d e f g -cat synapse, arguing against the first hypothesis. –/– HSA–-cat 0.10 Next, we determined whether AChR 100 1.0 1.0 responsiveness is altered in the absence of 0.8 ** 0.05 0.6 muscle b-catenin. Muscle fibers were stimu50 0.5 0.4 lated by carbachol (CCh), a specific and non0.2 0.0 0.00 0 hydrolyzable cholinergic agonist, which was 0.0 0.5 1.0 1.5 2.0 delivered by a microelectrode positioned at mEPP the central region of the diaphragm. CCh amplitude (mV) stimulation, but not vehicle stimulation, elicited voltage changes in control NMJs Figure 3 Defective spontaneous ACh release in HSA–b-cat –/– NMJs. (a) Reduced SV2 staining at (Fig. 4c). Notably, CCh was able to elicit HSA–b-cat –/– NMJs. Left hemi-diaphragm was whole-mount stained for SV2 with antibody to SV2 voltage changes in HSA–b-cat –/– muscle and visualized with FITC-conjugated secondary antibody. (b) Quantitative analysis of images in a. fibers. There was no difference between the ** P o 0.01 versus control (n ¼ 6, t-test). (c) Representative traces of mEPPs in control and number of CCh-responding muscle fibers in HSA–b-cat –/– NMJs. (d,e) Reduced frequencies, but no change in amplitude, of successful mEPPs. ** P o 0.01 versus control (n ¼ 31, t-test). (f) Cumulative mEPPs amplitude distribution in control control and HSA–b-cat –/– mice (Fig. 4d), and HSA–b-cat –/– mice. P 4 0.05 versus control (n ¼ 37 in 6 mice, Kolmogorov-Smirnov test). indicating that muscle fibers that failed to (g) Percentage of muscle fibers to produce mEPPs in a period of 3 min in control (n ¼ 49 in 6 mice) generate mEPPs were able to respond to and HSA–b-cat –/– mice. P o 0.01 versus control (n ¼ 46 of 6 mice, w 2-test). cholinergic stimulation. The rise-time and decay-time of CCh-evoked voltage changes, vesicle protein. In control littermates, R-BTX staining showed almost however, appeared to be increased in HSA–b-cat –/– NMJs in comparcomplete registration to SV2. In contrast, however, the area stained by ison with control (Fig. 4c). The mechanism of this phenomenon was both R-BTX and SV2 was reduced at HSA–b-cat –/– NMJs, although unclear. It may be caused by increased times for CCh, delivered by every AChR cluster was positive for SV2 (Fig. 3a,b), suggesting that pipettes positioned in the middle of muscle fibers, to diffuse to AChR there are potential defects in nerve terminals. To determine whether clusters in HSA–b-cat –/– mice. Note that AChR clusters were distribmuscle b-catenin depletion alters neurotransmission, we measured uted over a wider region in HSA–b-cat –/– diaphragms (Fig. 1d,h). A miniature endplate potentials (mEPPs), an indicator of spontaneous similar phenomenon was observed in Cdk5 mutant mice where AChR vesicle release. Muscle b-catenin deficiency did not appear to clusters are distributed over a wider central region29. Together, these affect mEPP amplitudes, suggesting that AChRs are functional in observations provide evidence that impaired neurotransmission may HSA–b-cat –/– mice (Fig. 3c,d), in general agreement with morpholo- be caused by defects in vesicle release. The above characterization of mEPPs indicates impaired spontagical characterization (Fig. 2). mEPP frequency, however, was markedly reduced in HSA–b-cat –/– mice (P o 0.01, n ¼ 6; Fig. 3e), suggesting neous release of ACh in HSA–b-cat –/– mice, suggesting that muscle that there are potential defects in spontaneous ACh release from b-catenin may be necessary for presynaptic differentiation and/or presynaptic terminals. Although a decrease in mEPP frequency usually function. To further investigate presynaptic defects, we analyzed reflects a reduction in presynaptic transmitter release, it can also be a result of a decrease in postsynaptic AChR responsiveness because some a -cat loxP/loxP b c -cat loxP/loxP d small-amplitude events are reduced to a level below the detection limits 1.6 of recording. In the latter case, there should be a preferential decrease in 100 1.2 small-amplitude events. However, muscle b-catenin deficiency did 0.8 50 HSA–-cat –/– Saline CCh not appear to increase the number of small-amplitude mEPPs in 0.4 HSA– -cat –/– the cumulative mEPPs amplitude distribution (Fig. 3f). Thus, the 0.0 0 50 µm reduction in mEPP frequency may not be caused by compromised AChR responsiveness; rather, a presynaptic mechanism may be 2 mV 5s involved. The notion was further supported by pair-pulse analysis of CCh neuromuscular transmission (see below). It is worth pointing out that the failure rate of mEPP recording was Figure 4 Responsiveness of muscle fibers to exogenous cholinergic agonist increased in HSA–b-cat –/– muscle fibers. We were unable to detect CCh. (a) Single fibers were stained for AChR with R-BTX. (b) Quantitative analysis of data in a. P 4 0.05 versus control (n ¼ 12, t-test). mEPPs in 70% of HSA–b-cat –/– muscle fibers in the 3-min recording (c) Representative traces of synaptic response to vehicle or CCh in control period (Fig. 3g, P o 0.01, n ¼ 49, 46 clusters of 6 mice). The failure to and HSA–b-cat –/– NMJs. Arrows indicate puff delivery. (d) Percentage of detect mEPPs in HSA–b-cat –/– muscle fibers could result from three muscle fibers with synaptic currents in response to CCh in control (n ¼ 17 possibilities: lack of synapses in muscle fibers, inability of AChR clusters in 3 mice) and HSA–b-cat –/– (n ¼ 23 of 3 mice) mice. P 4 0.05, w 2-test. mEPP amplitude (mV)


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whether vesicle release evoked by depolarization was affected in HSA–b-cat –/– mice. Muscles were stimulated by bath application of 40 mM KCl, which depolarizes nerve terminals and thus enhances vesicle release29. The stimulation caused robust and synchronous ACh release that often resulted in large depolarizing potentials in control muscles (Fig. 5a). However, the frequency of 40 mM KCl-evoked mEPPs was suppressed in HSA–b-cat –/– muscles (Fig. 5b), indicating that there were defects in vesicle releases by depolarization. Short-term plasticity in HSA–b-cat –/– NMJs To further determine whether this defect is presynaptic or postsynaptic, we characterized the short-term synaptic plasticity via paired-pulse ratios (PPRs) of EPPs elicited by two consecutive stimulations. The PPRs increased from 1.24 ± 0.11 in control to 1.8 ± 0.13 in HSA–bcat –/– NMJs (n ¼ 35 for control and 31 for HSA–b-cat –/–, P o 0.01; Fig. 6a) at physiological concentrations of Ca2+ and Mg2+ (2 mM and 1 mM, respectively). This result provides further evidence that there are presynaptic defects in neurotransmission. We found it interesting that an increase in extracellular Ca2+ concentration reduced PPRs, suggesting that the release machinery is less sensitive to Ca2+ in mutant mice (Fig. 6a). To test this hypothesis, we characterized EPP Ca2+ sensitivity in HSA–b-cat –/– NMJs. The EPP amplitude increase was smaller in mutant NMJs than in controls, although both showed a Ca2+ concentration–dependent increase (Fig. 6b). The EC50 values for HSA–b-cat –/– mice (3.41 ± 0.41 mM, n ¼ 48) were significantly larger than those of control littermates (1.9 ± 0.17 mM, n ¼ 42, P o 0.01). The K values were reduced from 0.71 ± 0.13 (n ¼ 48) in controls to 0.53 ± 0.13 (n ¼ 42) in HSA–b-cat –/– mice (P o 0.01). These results are consistent with the notion that calcium sensitivity was compromised at mutant NMJs. Note that we observed B40% decrease in maximal EPP amplitudes of mutant NMJs (Emax being 14.7 ± 1.8 mV and 26.4 ± 3.7 mV for HSA–b-cat –/– and control mice, respectively) at 8 mM Ca2+. This suggests that there are additional deficits that cannot be rescued by increasing extracellular calcium concentration. It is possible that the readily releasable pool is reduced in HSA–b-cat –/– NMJs. Normal NMJ morphology and function in HB9–b-cat –/– NMJs b-catenin can bind to cadherin to regulate the interaction between b-catenin and the cytoskeleton. The process is central to homophilic cadherin-mediated cell-cell adhesion30. It is possible that b-catenin–dependent adhesion between motoneurons and muscle fibers may be involved in the initial stages of NMJ formation, although

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such a mechanism may not occur at adult NMJs because of a large synaptic cleft5. Earlier studies have shown that b-catenin interacts with cadherin, and this interaction has been shown to influence the size and strength of CNS synapses31. Deletion of b-catenin in hippocampal pyramidal neurons in null mutant mice caused a reduction in the number of reserved pool vesicles and an impaired response to prolonged repetitive stimulation32. To investigate whether the impaired neurotransmission in HSA–bcat –/– NMJs is the result of disruption of homophilic cadherinmediated cell adhesion and to determine whether motoneuron b-catenin is necessary for NMJ development and function, we crossed the b-cat floxed mice with Mnx1 (HB9)-Cre mice to generate HB9–b-cat –/– mice (Supplementary Fig. 5 online). HB9 is a homeodomain transcription factor that is expressed selectively by motoneruons in the developing spinal cord (E9.5)33 and is essential for differentiation of postmitotic motoneurons33,34. HB9-Cre mice express the Cre gene specifically in motoneurons (Supplementary Fig. 2) and have previously been used to manipulate gene expression in motoneurons28. In situ hybridization indicated that b-catenin mRNA was diminished in motoneurons in HB9–b-cat –/– mice (Supplementary Figs. 5 and 6 online). Unlike HSA–b-cat –/– mice, which died soon after birth, HB9–b-cat –/– mice were vital and showed no apparent behavioral defects within several months of experiments. Notably, there was no observable abnormality in phrenic nerve branch location and the number and length of secondary branches (Supplementary Fig. 5). AChR clusters were distributed in a narrow central region in the middle of diaphragms similar to that seen in the b-cateninloxP/loxP control littermates (Supplementary Fig. 5). No difference was observed in AChR cluster size between the control and HB9–b-cat –/– mice (Supplementary Fig. 5). Neurotransmission at HB9–b-cat –/– NMJs was normal in terms of mEPP amplitude and frequency and EPP amplitude (Supplementary Fig. 5). The data could indicate that b-catenin–dependent homophilic adhesion may be dispensable for differentiation or function of motoneurons. In agreement with this, cadherin and a-catenin distribution in muscle and NMJs showed no difference between HSA–b-cat –/– and control mice (Supplementary Fig. 7 online). These results suggest that b-catenin in muscle fibers, but not motoneurons, is essential for NMJ development and function and that the presynaptic deficits may be caused by impaired b-catenin–dependent transcription in the muscle.

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Figure 6 Compromised short-term plasticity and calcium sensitivity in HSA– b-cat –/– NMJs. (a) PPRs from control (n ¼ 35) and HSA–b-cat –/– (n ¼ 31) NMJs at different [Ca2+]o. Inset, representative traces induced by paired stimuli separated by 25 ms at 2 mM [Ca2+]o. ** P o 0.01, t-test. (b) Reduced calcium sensitivity in HSA–b-cat –/– NMJs. EPPs were elicited at indicated [Ca2+]o. Data were fitted by the dose-response equation Emax E¼ , where Emax is the maximal EPP amplitude (evoked by 8 mM EC ½Ca2+ o 1+e

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Ca2+) and E is EPP amplitude at the indicated [Ca2+]o. EC50 is the [Ca2+]o at which E is 50% of Emax, and K is the Hill coefficient, indicating the slope of the curve. n ¼ 42 and 48 for control and HSA–b-cat –/– NMJs, respectively.

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ARTICLES DISCUSSION This study demonstrates that b-catenin in the muscle is essential for NMJ development. In HSA–b-cat –/– mice, the primary branches of phrenic nerves were no longer located in the central region of diaphragm muscle fibers. Secondary branches were extended to innervate larger AChR clusters that were distributed in a wider area in the central region of muscle fibers. Both spontaneous and evoked neurotransmitter release was reduced at HSA–b-cat –/– NMJs, and was accompanied by reduced SV2 staining and increased PPR. Furthermore, calcium sensitivity of neurotransmitter release was compromised when b-catenin was deficient in the muscle. There was no evidence of regeneration, as the nuclei of each fiber were located beneath the membrane and muscle contractile units appeared to be normal in HSA–b-cat –/– mice. These observations indicate that muscle b-catenin is involved in the post- and presynaptic development of the NMJs and identify a b-catenin–dependent retrograde signaling for synapse development. This notion was supported by the normal NMJ morphology and function in HB9–b-cat –/– mice. Hypoplasia in the spinal cord is observed after removal of a limb bud, and is now known to be caused by apoptosis of motoneurons12. This finding led to the eventual discovery of the neurotrophic factors of the NGF family. However, these factors have a limited role in motoneuron survival35. Muscle-derived factors are thought to be essential for differentiation and function of presynaptic terminals in addition to motoneuron survival5,36. The current study demonstrates an important role for muscle b-catenin in presynaptic differentiation and function. b-catenin is a key regulator of gene expression. In the canonical pathway of Wnt signaling, activated Dvl prevents b-catenin degradation by the destruction complex, which includes GSK3b, axin and adenomatous polyposis coli protein14,19. Stabilized b-catenin translocates to the nucleus, provides transcriptional activator functions to T cell factor and regulates gene expression. In addition to gene regulation, b-catenin acts as a component of the cadherin-based cell-adhesion system. It bridges cadherins to a-catenin and thus regulates the cytoskeleton30. Cadherins and associated catenins appear to be important for synapse formation and synaptic plasticity. In developing synapses, cadherins have been shown to mediate a homophilic, attractive interaction between growth cones and their targets37. N-cadherin and b-catenin are diffusely distributed along the length of free dendritic filopodia that are not in contact with axons, but rapidly accumulate at filopodia-axon contact sites following target recognition38,39 and are required to promote synapse stabilization and structural and functional maturation40. In Drosophila N-cadherin missense mutants, an overabundance of synaptic vesicles are observed at photoreceptor-interneuron synapses41, where inhibition of N-cadherin function leads to the suppression of synaptic vesicle accumulation and recycling at presynaptic sites39,40. After synapse formation, the N-cadherin/b-catenin complex may regulate structural plasticity of synapses by directly interacting with synaptic proteins including AMPA receptors31,42. b-catenin in hippocampal neurons has been implicated in localizing the reserved pool of vesicles at presynaptic sites32. The localization of cadherin and a-catenin in HSA–b-cat –/– muscle was similar to that observed in control mice. Furthermore, motoneuron-specific b-catenin depletion seemed to have little effect on NMJ development or function. Both NMJ morphology and function (in terms of mEPPs and EPPs) were normal in HB9–b-cat –/– mice. These results indicate that b-catenin–dependent homophilic adhesion may be dispensable for differentiation or function of motoneurons. They also demonstrate that b-catenin in muscle, and not motoneurons, is important for NMJ formation. In light of the importance of b-catenin in gene regulation, it is plausible to speculate that presynaptic


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differentiation and function at the NMJ requires a b-catenin–dependent retrograde factor. Candidates for this factor include BDNF, NT-3, GDNF, Wnt7a, FGF22, S-laminin, SynCAMs and neuroligins36,43–45. However, expression of BDNF, NT-3, GDNF and FGF22 was not different in control and mutant mice in a preliminary analysis (data not shown), suggesting that the factor(s) may be previously unidentified. Impaired postsynaptic activity at neuromuscular synapses has been shown to delay the withdrawal of presynaptic terminals, a mechanism that has been implicated in synapse elimination46. In vitro studies of agrin-induced AChR clustering have indicated that b-catenin serves as a potential linker between AChRs and the a-catenin–associated cytoskeleton26. In the absence of muscle b-catenin in vivo, AChR clusters are larger in size and distributed over a wider central region in the diaphragms. These results suggest that b-catenin may be involved in AChR clustering and cluster location. Similar postsynaptic phenotypes were observed in mutant mice where muscle activity was compromised, including Chat –/–, Cdk5 –/– and Chrng e/e (AChRg) mice9,10,29,47,48. Whether such AChR cluster defects are responsible for abnormal presynaptic development or function in HSA–b-cat –/– NMJs warrants further investigation. It is worth pointing out that AChR clusters appeared to be functional in HSA–b-cat –/– NMJs. There was no difference between mEPP amplitudes in control and HSA–b-cat –/– conditional knockout muscles. Of the muscle fibers that were unable to produce mEPPs, CCh was able to elicit comparable synaptic currents. Moreover, analysis of the number of AChR clusters in dissociated single muscle fibers failed to show a difference between HSA–b-cat –/– and control mice, arguing against a differential synapse formation or elimination. In summary, we demonstrate that muscle b-catenin is involved in the development of both pre- and postsynaptic membranes. It is plausible that b-catenin–dependent transcription is necessary for the expression of a necessary retrograde signal protein. Further studies along this line will contribute to a better understanding of muscular dystrophies and motoneuron disorders including amyotrophic lateral sclerosis. METHODS Generation of b-catenin mutant mice. We generated and studied two b-catenin conditional knockout transgenic lines: b-catenin muscle-specific– deficient and b-catenin motoneuron-specific–deficient mice by a loxP/cre strategy (Supplementary Figs. 1, 5 and Supplementary Methods online). Immunohistochemistry. Whole-mount staining was carried out as described previously49. Unless otherwise indicated, staining was performed on the ventral region of the left hemi-diaphragms. Briefly, dissected muscles were fixed in 4% paraformaldehyde (w/v) at 4 1C overnight, rinsed with PBS (pH 7.3) at 22 1C, incubated with 0.1 M glycine in PBS for 1 h and rinsed with 0.5% Triton X-100/ PBS. Muscles were blocked in the blocking buffer (3% BSA, 5% goat serum and 0.5% Triton X-100 in PBS) for 2–4 h at 22 1C or overnight at 4 1C. They were then incubated with primary antibodies (neurofilament, 1:1,000, AB1983, Chemicon; synaptophysin, 1:2,000, A0010, Dako; SV2, 1:1,000, Developmental Studies Hybridoma Bank; S100, 1:1,000, Z0311, Dako; or rapsyn, 1:1,000) in the blocking buffer overnight at 4 1C. After washing three times for 1 h each with 0.5% Triton X-100 in PBS, the muscles were incubated with fluorescenceconjugated antibody to rabbit or mouse IgG (1:500, Molecular Probes) and R-BTX (1:2,500, Molecular Probes) for 2–4 h at 22 1C or overnight at 4 1C. Samples were washed three times for 1 h each with 0.5% Triton X-100 in PBS, rinsed once with PBS and flat-mounted in Vectashield mounting medium (H-1000, Vector laboratories). Z serial images were collected with a Zeiss confocal laser scanning microscope (LSM 510 META 3.2) and collapsed into a single image. The area and length (the longest axis) of AChR clusters and the percentage of endplates covered by SV2 were determined with LSM 5 Image Examiner (Carl Zeiss). To label single muscle fibers, whole-mount diaphragms were fixed in 4% paraformaldehyde at 4 1C overnight, rinsed with PBS (pH 7.3) at 22 1C and

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ARTICLES incubated with PBS containing Alexa Fluor 633–conjugated phalloidin (1:2,000, Molecular Probes) and R-BTX (1:2,500, Molecular Probes) for 2–4 h at 22 1C. Samples were washed three times for 0.5 h each in PBS. Single fibers were teased and flat-mounted in Vectashield mounting medium (H-1000, Vector laboratories). Z serial images were collected with a Zeiss confocal laser scanning microscope (LSM 510 META 3.2) and collapsed into a single image.


AChE staining and in situ hybridization. See the Supplementary Methods. Electrophysiology. Electrophysiological recording was carried out as we previously described49. Briefly, diaphragms with ribs and intact phrenic nerve were dissected from P0 mice, pinned on Sylgard gel in a dish and perfused with an oxygenated solution containing 145 mM NaCl, 5.4 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 10 mM Hepes-NaOH, pH 7.3, and 13 mM glucose at 22 1C for at least 1 h. Microelectrodes, 20–50 MO when filled with 3 M KCl, were placed in the central region of muscle fibers where AChR clusters were enriched. Positions were adjusted for maximal response with a mEPPs rise-time o3 ms. We recorded induced massive activity after treatment with 40 mM KCl. To record nerve-evoked EPPs, we sucked phrenic nerves into an electrode and stimulated at o1 mA, 0.5 Hz for a duration of 0.2 ms. A series of 50 to 300 EPPs were recorded for each endplate. For exogenous agonist-evoked synaptic potentials, 100 mM CCh or saline (control) was loaded into glass microelectrodes that were then positioned at the central region of the diaphragm. CCh delivery was executed by the Toobey Pressure System IIe at 800 ms and 15 psi pressure. We used 2.5 mM m-conotoxin GIIIB in perfusing solution to block muscle contraction. Membrane potentials in muscle fibers were maintained at about –45 mV by injecting currents. EPP amplitudes larger than 5 mV were corrected for by nonlinear summation using a previously described method50. Data were collected and analyzed using pClamp 9.2 (Axon Instruments). Statistical Analysis. Data was presented as mean ± s.e.m. We used two-tailed Student’s tests, Kolmogorov-Smirnov tests or w2-tests for statistical analysis. Changes were identified as significant if the P value was less than 0.05. Note: Supplementary information is available on the Nature Neuroscience website.

ACKNOWLEDGMENTS We are grateful to J. Melki and S. Arber for valuable mouse lines. This work was supported in part by grants from the US National Institutes of Health and the Muscular Dystrophy Association to L.M. and W.-C.X., National Natural Science Foundation of China (U0632007) and Program of Changjian Scholars and Innovative Research Team in University (IRT0731) to T.-M.G. T.-M.G. and L.M. are Chang Jiang Scholars. Published online at http://www.nature.com/natureneuroscience Reprints and permissions information is available online at http://npg.nature.com/ reprintsandpermissions 1. Kalinovsky, A. & Scheiffele, P. Transcriptional control of synaptic differentiation by retrograde signals. Curr. Opin. Neurobiol. 14, 272–279 (2004). 2. Markus, A., Patel, T.D. & Snider, W.D. Neurotrophic factors and axonal growth. Curr. Opin. Neurobiol. 12, 523–531 (2002). 3. Sudhof, T.C. The synaptic vesicle cycle. Annu. Rev. Neurosci. 27, 509–547 (2004). 4. Waites, C.L., Craig, A.M. & Garner, C.C. Mechanisms of vertebrate synaptogenesis. Annu. Rev. Neurosci. 28, 251–274 (2005). 5. Sanes, J.R. & Lichtman, J.W. Development of the vertebrate neuromuscular junction. Annu. Rev. Neurosci. 22, 389–442 (1999). 6. McMahan, U.J. et al. Agrin isoforms and their role in synaptogenesis. Curr. Opin. Cell Biol. 4, 869–874 (1992). 7. Gautam, M. et al. Defective neuromuscular synaptogenesis in agrin-deficient mutant mice. Cell 85, 525–535 (1996). 8. Xiong, W.C. & Mei, L. An unconventional role of neurotransmission in synapse formation. Neuron 46, 521–523 (2005). 9. Brandon, E.P. et al. Aberrant patterning of neuromuscular synapses in choline acetyltransferase–deficient mice. J. Neurosci. 23, 539–549 (2003). 10. Misgeld, T. et al. Roles of neurotransmitter in synapse formation: development of neuromuscular junctions lacking choline acetyltransferase. Neuron 36, 635–648 (2002). 11. Schaeffer, L., de Kerchove d’Exaerde, A. & Changeux, J.P. Targeting transcription to the neuromuscular synapse. Neuron 31, 15–22 (2001). 12. Hamburger, V. Trophic interactions in neurogenesis: a personal historical account. Annu. Rev. Neurosci. 3, 269–278 (1980). 13. Ciani, L. & Salinas, P.C. WNTs in the vertebrate nervous system: from patterning to neuronal connectivity. Nat. Rev. Neurosci. 6, 351–362 (2005).

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