Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2013 Jun 15.
Published in final edited form as: Dev Biol. 2012 Apr 20;366(2):255–267. doi: 10.1016/j.ydbio.2012.04.003

β-catenin stabilization in skeletal muscles, but not in motor neurons, leads to aberrant motor innervation of the muscle during neuromuscular development in mice

Yun Liu 1, Yoshie Sugiura 1, Fenfen Wu 2, Wentao Mi 2, Makoto M Taketo 3, Steve Cannon 2, Thomas Carroll 4, Weichun Lin 1,*
PMCID: PMC3358465  NIHMSID: NIHMS371811  PMID: 22537499

Abstract

β-catenin, a key component of the Wnt signaling pathway, has been implicated in the development of the neuromuscular junction (NMJ) in mice, but its precise role in this process remains unclear. Here we use a β-catenin gain-of-function mouse model to stabilize β-catenin selectively in either skeletal muscles or motor neurons. We found that β-catenin stabilization in skeletal muscles resulted in increased motor axon number and excessive intramuscular nerve defasciculation and branching. In contrast, β-catenin stabilization in motor neurons had no adverse effect on motor innervation pattern. Furthermore, stabilization of β-catenin, either in skeletal muscles or in motor neurons, had no adverse effect on the formation and function of the NMJ. Our findings demonstrate that β-catenin levels in developing muscles in mice are crucial for proper muscle innervation, rather than specifically affecting synapse formation at the NMJ, and that the regulation of muscle innervation by β-catenin is mediated by a non-cell autonomous mechanism.

Keywords: neuromuscular junction, synaptic transmission, nerve defasciculation, acetylcholine receptor, mouse genetics

Introduction

The neuromuscular junction (NMJ), the synaptic connection between motor neurons and muscle fibers, is among one of the earliest synapses formed during mammalian development and is vital to animal survival (Ackerman and Greer, 2007; Witzemann, 2006). Normal development of the NMJ requires reciprocal interactions between motor neurons and skeletal muscle fibers. These interactions ultimately lead to a progressive accumulation of synaptic vesicles and proteins at the nerve terminal and juxtaposition of the nerve terminal within areas of the postsynaptic muscle membrane containing a high density of acetylcholine receptors (AChRs) (Burden, 1998; Hall and Sanes, 1993). Studies over the last several decades have led to the emergence of two signaling pathways – the agrin/Lrp4/MuSK and the Wnt signaling pathways – that are critically involved in mediating nerve-muscle interactions during NMJ formation (Speese and Budnik, 2007; Wu et al., 2010).

The agrin/Lrp4/MuSK pathway is triggered by agrin (McMahan, 1990; Nitkin et al., 1987), an extracellular matrix heparan sulfate proteoglycan that is expressed in both motoneurons and skeletal muscles, although motoneuron-derived agrin is much more active than muscle-derived agrin in its biological activity (Ferns et al., 1992; Ruegg et al., 1992). Agrin binds to a membrane complex consisting of a low density lipoprotein (LDL)- receptor related protein 4 (Lrp4) (Kim et al., 2008; Zhang et al., 2008) and a muscle specific receptor kinase (MuSK) and induces MuSK activation and AChR clustering, through an interaction with rapsyn, a cytoplasmic protein in muscle fibers (Apel etal., 1997; Froehner et al., 1981; Gautam et al., 1995). This signaling also requires the presence of Dok-7 (docking protein-7), a protein that contains a phosphotyrosine-binding domain (PTB domain) and lies downstream of MuSK (Inoue et al., 2009; Okada et al., 2006). Mice deficient in Agrin (Burgess et al., 2000; Gautam et al., 1996), Lrp4 (Weatherbee et al., 2006), MuSK (DeChiara et al., 1996), Dok-7 (Okada et al., 2006) or rapsyn (Gautam et al., 1995) all fail to form neuromuscular synapses and die perinatally.

Wnt proteins, like agrin, may also be released from nerve terminals and from muscle, but may induce more complex signaling mechanisms. Mammals have 19 Wnt proteins and 10 Wnt receptors (e.g., Frizzled) (Logan and Nusse, 2004). Interactions of Wnt proteins and their receptors may activate a variety of signaling pathways that are closely integrated and intertwined (van Amerongen and Nusse, 2009). In the canonical Wnt pathway, binding of Wnt to the receptor Frizzled and co-receptor Lrp5/Lrp6 activates the downstream scaffolding protein Dishevelled (Dvl). The activation of Dvl leads to a disassembly of complex consisting of GSK3β, Axin and adenomatous polyposis (APC), thus preventing GSK3β-mediated phosphorylation of β-catenin that normally causes its degradation (Aberle et al., 1997; He et al., 2004; Orford et al., 1997). As a result, β-catenin is stabilized in the cytoplasm, enabling its import into the nucleus, where it regulates gene expression through its association with T cell factor/lymphoid enhancer factor (TCF/LEF) family members (Behrens et al., 1996; Cadigan and Nusse, 1997).

Previous studies suggest that the canonical Wnt pathway plays a role in the development of the NMJ. However, conflicting results have been reported. For instance, although it has been shown that Dvl, a downstream scaffolding protein that regulates β-catenin activation, interacts with MuSK and that disruption of this interaction reduces AChR clustering (Luo et al., 2002), AChR clusters persist (albeit in a broader distribution pattern) in Dvl-1 null muscles (Henriquez et al., 2008). Further, in vitro studies suggested that β-catenin interacts with rapsyn and regulates AChR clustering (Zhang et al., 2007) and that APC binds to the β-subunit of AChR and regulates AChR clustering (Wang et al., 2003). Intriguingly, Wnt1 induced β-catenin accumulation does not seem to affect AChR clustering in C2C12 myotubes (Luo et al., 2002) and AChR cluster sizes are increased in β̃catenin loss-of-function mutation in muscles (Li et al., 2008).

In this study, we sought to clarify the role of β-catenin in NMJ development. We applied a genetic mouse model in which β-catenin can be stabilized (gain-of-function) in tissue-specific manner (Harada et al., 1999). We found that β-catenin stabilization in muscles, but not in motor neurons, results in excessive nerve defasciculation and nerve branching. However, individual neuromuscular synapses developed and functioned normally, despite abnormal innervation pattern. Our findings demonstrate that stabilization of β-catenin, either in motor neurons or in the skeletal muscle, does not affect the formation and function of the NMJ. Thus, β-catenin functions through non-cell autonomous mechanisms in the skeletal muscle to retrogradely regulate the development of motor innervation pattern, rather than specifically affecting neuromuscular synaptogenesis.

Materials and Methods

Animals

β-catenin exon-3 floxed mice (Ctnnb1lox(ex3)/lox(ex3)) (Harada et al., 1999), tissue-specific Cre mice including motor neuron-specific Cre (HB9Cre) (Jackson Laboratories, stock#: 006600) (Arber et al., 1999; Yang et al., 2001) and skeletal muscle specific Cre (Myo-Cre, under myogenin promoter/MEF2c enhancer) (Li et al., 2005), or ROSA26 Cre reporter (Jackson Laboratories, stock # 003474) (Soriano, 1999) have been described previously. To generate mice that express stabilized β-catenin in their skeletal muscle, we bred the Ctnnb1lox(ex3)/lox(ex3) with Myo-Cre+ mice (Li et al., 2005). To generate mice that express stabilized β-catenin specifically in motor neurons, we bred the Ctnnb1lox(ex3)/lox(ex3) mice with HB9Cre mice (Arber et al., 1999; Yang et al., 2001). The littermate mice that did not express Cre (eg., Ctnnb1lox(ex3)/+) were analyzed as controls. The stabilization of β-catenin was conferred by genetically deleting exon-3 from the β-catenin gene to produce a mutant β-catenin that lacks phosphorylation sites (Harada et al., 1999), resulting in β-catenin accumulation and stabilization in Cre-expressing tissues. The day in which a vaginal plug was detected was designated embryonic day 0.5 (E0.5). Embryos were collected by cesarean section from anesthetized pregnant mice. All experimental protocols followed National Institutes of Health Guidelines and were approved by the University of Texas Southwestern Institutional Animal Care and Use Committee.

Western Blot Analysis

The spinal cords and diaphragm muscles of E18.5 mouse embryos were homogenized and separated by SDS-PAGE, transferred to a nitrocellulose membrane, and blocked in 5% milk in Tris-buffered saline. The membrane was incubated with polyclonal antibody against β-catenin (1:3000, Sigma-Aldrich, St. Louis, MO, USA) and monoclonal antibody against α-tubulin (1:1000, Sigma-Aldrich, St. Louis, MO, USA) at 4°C overnight. Membranes then were incubated with goat anti-mouse IRDye 680 and goat anti-rabbit IRDye 800 secondary antibodies (1:10000, Li-Cor Biosciences, Lincoln, NE, USA) at room temperature for 1 hr. Membranes were scanned and analyzed using the Odyssey infrared imaging system (Li-Cor Biosciences). α-tubulin served as a loading control.

X-gal staining for LacZ (β-galactosidase)

Wholemount diaphragm muscle or transverse sections (20 µm) of E16.5 embryos were fixed in 2% paraformaldehyde (PFA) in 0.1 M phosphate buffer for 20–30 minutes, washed three times with PBS and then incubated in a buffer containing sodium phosphate (150 mM), MgCl2 (2 mM), sodium deoxycholate (0.01%), NP-40 (0.02%), potassium ferricyanide (5 mM), potassium ferrocyanide (5 mM) and 5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside (1mg/ml, X-Gal). Samples were then washed three times with PBS and mounted in PVA (polyvinyl alcohol) mounting medium. Images were acquired on an Olympus BX51 upright microscope with Nomarski optics.

Analysis of muscle contractile properties

Muscle contractile properties were measured at room temperature according to the procedures described previously (Martin-Caraballo et al., 2000; Wu et al., 2011). First, diaphragm muscles (E18.5) were acutely isolated in oxygenated Ringer’s solution from 3 pairs of Ctnnb1lox(ex3)/+;Myo-Cre– (control) and Ctnnb1lox(ex3)/+; Myo-Cre+ mice. A muscle strip was then removed from the right side of the dorsal portion of the hemi-diaphragm (E18.5) from each animal. The muscle strip was suspended in a 25-ml organ bath (Myobath, World Precision Instruments, Sarasota, FL) filled with oxygenated Ringer’s solution that contains 10 µM tubocurarine to block muscle acetylcholine receptors (Martin-Caraballo et al., 2000). The muscle strip was then mounted vertically in an isometric tissue clamp of a force transducer (FORT25, World Precision Instruments, Sarasota, FL) to detect muscle contraction, which was elicited by field stimulation with parallel wire electrodes. A train of 4–100 pulses, 1-ms duration, 80 mA (A385 Stimulator, World Precision Instruments, Sarasota, FL) at 4, 10, 20, 50 and 100 Hz, delivered 20 s apart was applied to evoke isometric contractions. Data were acquired using a transbridge (4M, World Precision Instruments, Sarasota, FL) and pClamp 9 software (Molecular Devices, Sunnyvale, CA) at a sampling rate of 2K Hz. Finally, the wet weight of the muscle strip was determined immediately after the completion of muscle contraction measurement.

Analysis of neuromuscular synaptic transmission

Electrophysiological analysis of neuromuscular synaptic transmission was carried out as described previously (Liu et al., 2008). Briefly, embryonic phrenic nerve-diaphragm muscles (E18.5) or adult lumbrical muscles (P45) were acutely isolated in oxygenated Ringer’s solution (136.8 mM NaCl, 5 mM KCl, 12 mM NaHCO3, 1 mM NaH2PO4, 1 mM MgCl2, 2 mM CaCl2, and 11 mM d-glucose, pH 7.3) (Liley, 1956). End-plate regions were visually identified under a microscope and impaled with glass micropipettes filled with 2 M potassium citrate containing 10 mM potassium chloride (resistance 20–40 MΩ). Miniature end-plate potentials (mEPPs) and evoked end-plate potentials (EPPs) were acquired using an AxoClamp-2B intracellular amplifier (Molecular Devices, Sunnyvale, CA).

Immunofluorescence staining

Immunofluorescence staining was carried out as previously described (Liu et al., 2008). Briefly, embryonic diaphragm muscles or adult triangularis sterni muscles were fixed in 2% paraformaldehyde (PFA), blocked in antibody dilution buffer (500 mM NaCl, 0.01 M phosphate buffer, 3% BSA and 0.01% thimerosal), and then labelled with Texas-Red conjugated a-bungarotoxin (α-bgt) (2 nM, Molecular Probes) for 30 minutes at room temperature. Wholemounts of muscle samples were then labeled by primary antibodies (rabbit) such as anti-neurofilament (NF150, 1:1000) (Chemicon, Temecula, CA) plus anti- synaptotagmin-2 (1:1000) (gifts from Dr. Thomas Südhof, Stanford University School of Medicine, Palo Alto, CA), anti-synaptophysin (1:100) (Dako, Carpinteria, CA), or anti-S100β (1:1000) (Swant, Bellinzona, Swizerland), followed by secondary antibodies (fluorescein isothiocyanate-conjugated goat anti-rabbit IgG, 1:1000) (Jackson ImmunoResearch Laboratories, Inc. West Grove, PA). Samples were washed in PBS and flat-mounted in Vectashield mounting medium. Images were acquired using a Zeiss LSM 510 confocal microscope.

Acetylcholinesterase (AChE) staining

Detection of AChE was carried out based on the methods previously described (Enomoto et al., 1998). Muscles were fixed with 2% PFA, rinsed in PBS and then incubated in a solution containing 0.2 mM ethopropazine, 4 mM acetylthiocholine iodine, 10 mM glycine, 2 mM cupric sulfate, and 65 mM sodium acetate solution, pH 5.5, for 2–4 h at 37°C. Samples were then developed in sodium sulfide (1.25%, pH 6.0) for 2–5 minutes, followed by extensive wash in water under a fume hood. Muscle samples were then cleared with 50% glycerol in PBS and flat mounted on a glass slide.

Electron Microscopy

Diaphragms muscles with phrenic nerves attached (E18.5) were dissected and fixed in 1% glutaraldehyde (in 0.1 M phosphate buffer, pH 7.4.) for 12 hours at 4°C. The tissue was then rinsed with phosphate buffer, trimmed in to small pieces, and then fixed with 1% osmium tetroxide for 3 hr on ice. The tissue was then dehydrated in a graded series of ethanol and embedded in Epon 812 (Polysciences, Warrington, PA). Ultrathin (70-nm) sections were mounted on Formvar-coated grids and stained with uranyl acetate and lead citrate. Electron micrographs were acquired using a Tecnai (Netherlands) electron microscope at 120 kV.

Statistical Analysis

Data are presented as the mean ± standard error of the mean (SEM). Statistical differences between control and mutant groups are determined by Student’s t test and are considered significant when the P value is less than 0.05.

Results

β-catenin stabilization in skeletal muscle results in neonatal lethality in mice

The exon-3 of β-catenin encodes the highly conserved N-terminal serine and threonine residues that are targets for phosphorylation by glycogen synthase kinase 3β (GSK3β). Phosphorylation of these sites by GSK3β leads to degradation of β-catenin by the ubiquitin-proteasome pathway (Aberle et al., 1997; Orford et al., 1997). Previous studies have shown that Cre mediated deletion of exon-3 in an engineered mouse strain (Ctnnb1lox(ex3)/lox(ex3)) prevents β-catenin from being phosphorylated, resulting in an accumulation and stabilization of β-catenin in Cre-expressing cells (Harada et al., 1999). We applied this model to study the role of β-catenin in the development of the NMJ. Specifically, we sought to determine the effect of stabilizing β-catenin within the muscle cell using Myo-Cre transgenic mice that drive Cre expression in skeletal muscle from embryonic day 9 (E9) onwards (Li et al., 2005). As expected, crossing Myo-Cre mice with ROSA26-LacZ reporter mice (Soriano, 1999) results in specific expression of β-galactosidase in the skeletal muscle (Fig. 1A-C) but not in the spinal cord (Fig. 1D-E) or the cardiac muscle (Fig. 1F-G).

Figure 1. Stabilization of β-catenin in skeletal muscles results in neonatal lethality.

Figure 1

Open in a new tab

A-G: LacZ staining of wholemount diaphragm (A & B) and a cross-section of the eye (C), the spinal cord (D & E) and the heart (F & G) from a ROSA26-LacZ;Myo-Cre+ embryo (E16.5). LacZ expression was detected in the diaphragm muscle (A & B; A, low power; B, high power), extraocular muscle (C), but absent in the spinal cord (D & E; D, low power; E, high power) and the cardiac muscles (F: atrium; G: ventricle). H: Immunoblot analysis of tissue homogenates from two pairs of Ctnnb1lox(ex3)/+;Myo-Cre+ (Cre+) and Ctnnb1lox(ex3)/+; Myo-Cre- (Cre-) embryos (E18.5) probed with anti-β-catenin antibodies and anti-α-tubulin, which served as a loading control. Full-length β-catenin (WT, upper band) was detected in spinal cord and diaphragm muscle in both genotypes, whereas a truncated β-catenin (Mut, lower band) was detected only in the diaphragm muscle in Ctnnb1lox(ex3)/+;Myo-Cre+ mice. I. External phenotype of Ctnnb1lox(ex3)/+;Myo-Cre- (control) and Ctnnb1lox(ex3)/+;Myo-Cre+ (mutant) mice (P0). Mutant pup died within 24 hr after birth. Arrowhead in I points to the white patch of milk in the stomach in the control pup. Mutant pup exhibited a lack of milk in the stomach (J, bottom picture). Scale bars: A: 500 µm; B: 50 µm; C: 100 µm; D: 100 µm; E: 50 µm; F-G: 30 µm.

To determine the efficiency of β-catenin stabilization in the muscle, we probed tissue homogenates from two pairs of Ctnnb1lox(ex3)/+;Myo-Cre+ (Cre+) or Ctnnb1lox(ex3)/+;Myo-Cre– (Cre–) mice with anti-β-catenin antibodies. As expected, full-length β-catenin (wildtype β-catenin) was detected in both spinal cord and diaphragm muscle in either Ctnnb1lox(ex3)/+;Myo-Cre+ or Ctnnb1lox(ex3)/+;Myo-Cre– mice, whereas a truncated β-catenin (mutant β-catenin) was detected in the diaphragm muscle in Ctnnb1lox(ex3)/+;Myo-Cre+ mice, but not in Ctnnb1lox(ex3)/+;Myo-Cre– mice (Fig.1H).

Mice expressing stabilized β-catenin in the skeletal muscles were grossly normal throughout embryonic development. Among 97 embryos derived from the breeding of Ctnnb1lox(ex3)/lox(ex3) x Myo-Cre+ mice, 48 were Ctnnb1lox(ex3)/+;Myo-Cre+ (mutant) and 49 Ctnnb1lox(ex3)/+;Myo-Cre– (hereafter referred to as control), and mice were born in the expected Mendelian ratio of 1:1 (Table 1). Furthermore, we examined 29 newborn pups from P0 to P1. At P0, control and mutant pups were indistinguishable; they moved and appeared characteristically pink in color, with no signs of gasping. However, mutant pups died within 1 day after birth. At the time of death, mutant pups appeared grossly normal, except that they lack milk in their stomachs (Figure 1I-J). These results demonstrated that β-catenin stabilization in skeletal muscle resulted in neonatal death in mice.

Table 1.

Survival of the offspring from mating Ctnnblox(ex3)/lox(ex3) with Myo-Cre transgenic mice

Age Total number
of litters		 Total number
of mice		 Ctnnb1lox(ex3)/+
Myo-Cre+ 		Ctnnb1lox(ex3)/+;
Myo-Cre− 		χ2 P
E13.5–15.5 3 32 16 (16) 16(16) 0 1
E18.5 7 65 32 (32.5) 33 (32.5) 0.008 0.930
P0 2 21 10 (10.5) 11 (10.5) 0.023 0.879
P1 2 8 0 (4) 8 (4) 5.333 0.021
P21 4 16 0 (8) 16 (8) 10.667 0.001
Open in a new tab

Mutant mice expressing stabilized β-catenin in the skeletal muscle (Ctnnb1lox(ex3)/+; Myo-Cre+) were born in the expected Mendelian ratio of 1:1 (expected numbers indicated in the parentheses). However, none of these mice survived after P0. Among 118 (E13.5 to P0) progeny from mating of Ctnnb1lox(ex3)/lox(ex3) with Myo-Cre mice, approximately half (n = 58) were identified as Ctnnb1lox(ex3)/+ and Myo-Cre positive (Ctnnb1lox(ex3)/+; Myo-Cre+, mutant) and the remaining half (n = 60) were Ctnnb1lox(ex3)/+ and Myo-Cre negative (Ctnnb1lox(ex3)/+; Myo-Cre−, control). Among 24 mice examined between P1 and P21, all of them were genotyped as Ctnnb1lox(ex3)/+; Myo-Cre negative, indicating none of Ctnnb1lox(ex3)/+; Myo-Cre positive mice survived beyond P1.

β-catenin stabilization in muscles leads to excessive nerve defasciculation and branching, but exerts no adverse effect on the capacity of muscle to form the NMJ

We next investigated whether β-catenin stabilization in muscles affected the formation and function of the NMJ. We began our analyses on embryonic diaphragm muscles, in which the NMJs are among the earliest to be established during development (Bennett and Pettigrew, 1974). We systematically analyzed diaphragm muscles (E13.5-P0) from control and mutant mice expressing stabilized β-catenin (Ctnnb1lox(ex3)/+;Myo-Cre), by wholemount immunostaining using a mixture of antibodies against a neurofilament protein (NF-150) and a synaptic vesicle protein (Syt2) to label pre-synaptic nerves and α-bgt to label post-synaptic AChRs.

We found marked differences in innervation pattern between control and Ctnnb1lox(ex3)/+;Myo-Cre mice. In the control (Fig. 2A-C), the phrenic nerve entered the diaphragm muscle at mid-costal level and extended secondary intramuscular nerve bundles along the central region of the diaphragm muscle. The secondary nerve trunks extended orderly, short tertiary nerve branches near the central end-plate region. In the Ctnnb1lox(ex3)/+;Myo-Cre embryos (Fig. 2D-F), the phrenic nerve reached the diaphragm muscle on schedule. For example, at E13.5, intramuscular nerves fully innervated both the dorsal and the ventral diagram in Ctnnb1lox(ex3)/+;Myo-Cre (Fig. 2D). However, there were increases in tertiary nerve branching and defasciculation compared with the control. For example, the average length of the tertiary nerve branches in the ventral region of the right hemi-diaphragm in Ctnnb1lox(ex3)/+;Myo-Cre mice (E18.5) was 511 ± 138 µm, nearly 10 times longer than that seen in the same region of the diaphragm in litter-mate controls (52 ± 5 µm, P < 0.01). The increase in nerve branching and defasciculation was noticeable at E13.5 (compare Fig. 2D with Fig. 2A), and became more apparent at E14.5 (Fig. 2 E vs. 2B) and persisted throughout the embryonic (Figs. 2, 4–5) and neonatal period (Fig. S1). The increase in nerve branching and defasciculation was more pronounced in the right hemi-diaphragm compared with the left hemi-diaphragm (Fig. S1).

Figure 2. β-catenin stabilization in skeletal muscles leads to excessive nerve defasciculation and branching.

Figure 2

Open in a new tab

A-F: Hemi-diaphragm muscles were labeled by anti-NF150 and anti-Syt2 antibodies to reveal innervation pattern. The primary phrenic nerve (p) enters the diaphragm muscle at mid-costal level and extends secondary (s) and tertiary (t) nerve branches prior to making synaptic contacts with the muscle fibers. Excessive nerve defasciculation and branching are evident in mutant muscles (D-F, Ctnnb1lox(ex3)/+;Myo-Cre) compared with the controls (A-C). G-H: High-magnification views of E18.5 diaphragm muscles double-labeled with antibodies against Syt2 and NF150 and Texas Red conjugated α-bgt. Nerve terminal sprouts (arrowheads) are markedly increased in mutant muscles (H) compared with controls (G). Scale bars: A, D: 200 µm; B, E: 400 µm; C, F, 1000 µm; G-H, 100 µm.

Figure 4. Normal morphology of the NMJ in mice expressing stabilized β-catenin in muscles.

Figure 4

Open in a new tab

A-B: Confocal images of E18.5 diaphragm muscle (A) and triangulris sterni muscle (B), immunostained by a mixture of anti-NF 150 and anti-Syt2 antibodies (green) and α-bgt (red), and the merged images are shown at the bottom row in both A and B. The pre-synaptic nerve terminals (white arrowheads) were juxtaposed with post-synaptic AChR clusters (white arrows) in both controls (left column) and Ctnnb1lox(ex3)/+;Myo-Cre muscles (right column). C-D: Electron micrographs of the NMJ in E18.5 diaphragm muscles from control (C) and Ctnnb1lox(ex3)/+;Myo-Cre (D) muscles. Pre-synaptic nerve terminals (Nt), capped by a terminal Schwann cell, were filled with an abundance of synaptic vesicles in both control and Ctnnb1lox(ex3)/+;Myo-Cre muscles. Black arrow points to the basal lamina between the pre- and post-synaptic membranes, and black arrowhead points to the postsynaptic junctional fold. Scale bars: A-B, 20 µm; C-D, 1 µm.

Figure 5. β-catenin stabilization in muscles leads to formation of the NMJ within the central tendon region of the diaphragm.

Figure 5

Open in a new tab

A: Low power views of right hemi-diaphragm (E15.5) from control (left panel) and Ctnnb1lox(ex3)/+;Myo-Cre (right panel). B: High power views of the central tendon region bordered by the square, labeled by anti-NF150 and anti-Syt2 antibodies (green) and α-bgt (red). The merged image is shown at the bottom. Arrowheads point to the NMJ. C: A whole diaphragm muscle in from Ctnnb1lox(ex3)/+;Myo-Cre (E18.5) illustrating the aberrant nerves in the central tendon of the diaphragm (arrowhead in C). D: Individual NMJs (arrowheads) in the central tendon regions of the diaphragm in Ctnnb1lox(ex3)/+;Myo-Cre embryos (E18.5), labeled by a mixture of anti-NF150 and anti-Syt2 antibodies (green) and α-bgt (red) (top three panels), or labeled by anti-synaptophysin antibodies (green) and α-bgt (red) (bottom three panels). E: Wholemount AChE staining of E18.5 diaphragm muscle. AChE clusters were detected in extra muscle patches (arrowheads, left panel) within the central tendon of the diaphragm muscle in Ctnnb1lox(ex3)/+;Myo-Cre mice, but not in the control. Scale bars: A, C: 400 µm; B: 10 µm; D: 20 µm; E, 2000 µm.

In addition to increased defasciculation and branching, significantly more nerve terminal sprouts were detected in mutant muscles (arrowheads in Fig. 2H), compared with the control (arrowheads in Fig. 2G). For example, in the ventral sternal region of the right diaphragm muscle (Fig. 2G-H), the number and length of terminal sprouts in mutant muscles (N = 4) were 21 ± 2 and 168 ± 8 µm, respectively, compared with 10 ± 1 and 91 ± 10 µm in control (N = 3), respectively. The terminal nerve sprouts in both control and mutant muscles were accompanied by Schwann cells, as revealed by immunostaining with anti-S100β antibodies, a marker for Schwann cells (Fig. S2)

Consistent with excessive nerve branching, AChR clusters (end-plates) were more broadly distributed, although still confined to the central regions in Ctnnb1lox(ex3)/+;Myo-Cre muscles. The mean width of the end-plate bands in the dorsal region of E14.5 diaphragm muscles was increased by 26%: from 168 ± 12 µm in the controls (N=3) to 212 ± 9µm (N=3), P < 0.05, in the Ctnnb1lox(ex3)/+;Myo-Cre mice (Fig. 3A-B). The end-plate bands remained expanded throughout the embryonic period, as indicated by AChR staining at E18.5 (Fig. 3C-D). However, the size of individual AChR cluster was similar for the Ctnnb1lox(ex3)/+;Myo-Cre mice and their controls (78 ± 3 µm2 vs. 80 ± 4 µm2, respectively; N = 4 mice, each group).

Figure 3. β-catenin stabilization in muscle leads to expansion of the end-plate band.

Figure 3

Open in a new tab

A-D: hole mounts of embryonic diaphragm muscles (A-B: E14.5; C-D: E18.5) labeled with α-bgt for AChRs. In both control (Ctnnb1lox(ex3)/+) and mutant (Ctnnb1lox(ex3)/+;Myo-Cre), AChR clusters (white arrowheads) are aligned along the central region of the muscle, forming a centrally located end-plate band. However, the end-plate band occupied a broader region in the mutant muscle (B, D: Ctnnb1lox(ex3)/+;Myo-Cre) than in the control muscle (A, C: Ctnnb1lox(ex3)/+). Insets in C and D show high power views of individual AChR clusters. Scale bars: A-B, 200µm; C-D, 200µm; insets in C and D, 20 µm.

Furthermore, individual neuromuscular synapses developed normally in Ctnnb1lox(ex3)/+;Myo-Cre mice. At light microscopic level, we found that presynaptic nerve terminals in Ctnnb1lox(ex3)/+;Myo-Cre mice were intensely labeled by anti-Syt2 antibodies and formed in juxtaposition with postsynaptic AChR clusters in the diaphragm, triangularis sterni or hindlimb muscles, similar to those in the control (Fig. 4A-B, Fig. S3). At the electron microscopic level, we found that the ultrastructure of the NMJ was indistinguishable between control and mutant muscles (Fig. 4C-D). In both control (n = 76 nerve terminals, N = 4 mice) and mutant (n = 53 nerve terminals, N = 4 mice), pre-synaptic nerve terminals were capped by a terminal Schwann cell and were juxtaposed with postsynaptic membranes. Basal lamina was well defined at the synaptic cleft between the pre- and post-synaptic membrane (black arrow in Fig. 4C-D). Thus, stabilization of β-catenin in muscles exerted no adverse effect on the capacity of the muscle to establish the NMJ.

Intriguingly, additional NMJs were also detected within the central tendon region in Ctnnb1lox(ex3)/+;Myo-Cre mice (Fig. 5). These NMJs were innervated by aberrant nerve branches (arrowhead in Fig. 2D-F; Fig. 5A, C) extended from the intramuscular nerve trunk. The nerve terminals of these aberrant nerves were intensely labeled by synaptic vesicle proteins (Fig. 5B, D) and juxtaposed to AChR clusters (Fig. 5B, D). Wholemount AChE staining of the diaphragm muscle further revealed that small muscle patches containing AChE clusters were present within the central tendon of the diaphragm in Ctnnb1lox(ex3)/+;Myo-Cre mice (arrowheads in Fig. 5E) but not in the control.

β-catenin stabilization in muscles leads to an increase in motor axon number

From wholemount staining of the phrenic nerve/diaphragm preparation, we noticed that the phrenic nerve bundles were thicker in the stabilized β-catenin mutant embryos compared with their littermate controls (Fig. 2), suggesting a possible increase of axon number and/or axon size in the phrenic nerve in the mutant embryos. We therefore examined cross sections of the phrenic nerves by electron microscopy. We found that the motor axon numbers were moderately, but significantly (P < 0.01) increased in the phrenic nerve of the mutant embryos (Fig. 6B) compared with the control (Fig. 6A). For example, the average number of motor axons was 261 ± 7 (N = 6) in the mutant embryos versus 228 ± 9 (N = 7) in the controls. The size of individual axons, however, was similar between control and mutant embryos and all motor axons were wrapped by Schwann cell processes (arrows in Fig. 6C and D) that distributed throughout the entire span of the nerve. These results indicate that β-catenin stabilization in muscles leads to an increase in motor axon number.

Figure 6. β-catenin stabilization in muscles leads to an increase in motor axon numbers in the phrenic nerve.

Figure 6

Open in a new tab

Cross-sections of the phrenic nerve (E18.5) were examined under an electron microscope: A-B: Low power views of the entire nerve cross-section. The number of axons was significantly increased (t-test, P < 0.01) in the mutants (261 ± 7 per nerve, n = 6 nerves) compared with the controls (228 ± 9 per nerve, n = 7). C-D: High power views illustrating individual axons (Ax) wrapped by Schwann cell (SC) processes (arrow). Scale bars: A-B: 10 µm; C-D: 1 µm.

β-catenin stabilization in muscles leads to a reduction in muscle fiber size

The expression of β̃catenin is developmentally regulated during myogenesis (Cifuentes-Diaz et al., 1998; Goichberg et al., 2001) and increased level of membrane-bound β̃catenin is associated with abnormal myofibril organization in skeletal muscles (Nastasi et al., 2004). We therefore sought to determine if stabilization of β̃catenin in muscle affects muscle development. Cross-sections of the embryonic diaphragm muscle (E18.5) showed that muscle nuclei in Ctnnb1lox(ex3)/+;Myo-Cre mice were peripherally localized, suggesting no muscle fiber necrosis or muscle fiber regeneration occurred in these mice (Fig. 7A-B). However, the size of individual muscle fibers was markedly reduced in Ctnnb1lox(ex3)/+;Myo-Cre mice; the average size of muscle fibers was 139.6 ± 4.1 µm2 (n = 407 fibers) in Ctnnb1lox(ex3)/+;Myo-Cre mice, vs. 167.9 ± 3.9 µm2 (n = 408 fibers, P < 0.001) in the control mice (Fig. 7C). We further examined longitudinal sections of the diaphragm muscle under EM; we found that the ulstrastructure of the Z-lines and sarcomeres were similar between Ctnnb1lox(ex3)/+;Myo-Cre and control mice (Fig. 7D-E). Thus, β-catenin stabilization in muscles resulted in a reduction in muscle fiber size but did not appear to affect the assembly of sarcomeres.

Figure 7. β-catenin stabilization in muscles leads to a reduction in muscle fiber caliber.

Figure 7

Open in a new tab

A-B: Cross-section (semi-thin, 1 µm) of the mid-costal regions of E18.5 diaphragm muscle stained with toluidine blue. Muscle nuclei (arrows) are peripherally localized in both Ctnnb1lox(ex3)/+;Myo-Cre and control mice. C: Quantification of the average area of muscle fiber size (left bar graph) and size distribution (right bar graph) in the control and Ctnnb1lox(ex3)/+;Myo-Cre mice. The average size of muscle fibers was 139.6 ± 4.1 µm2 (n = 407 fibers) in Ctnnb1lox(ex3)/+;Myo-Cre mice, vs. 167.9 ± 3.9 µm2 (n = 408 fibers, P < 0.001) in the control mice. D-E: Longitudinal sections of the diaphragm muscle under EM. The organization of the Z-lines (arrowheads) and sarcomeres (unit between Z-lines) is similar between the control (D) and Ctnnb1lox(ex3)/+;Myo-Cre mice (E). Scale bars: A-B: 30 µm; D-E: 1 µm.

Normal synaptic transmission and muscle contraction properties in mice expressing stabilized β-catenin in muscles

To determine whether β-catenin stabilization in muscle affects NMJ function or muscle contractile properties, we carried out electrophysiological analyses. First, we measured mEPPs (Fig. 8A) and EPPs (Fig. 8C) in acutely isolated phrenic nerve/diaphragm preparations (E18.5). We found that mEPP frequencies (events/min−1) were similar between Ctnnb1lox(ex3)/+;Myo-Cre (0.90 ± 0.15, N = 4 mice, n = 25 fibers) and control (0.93 ± 0.15, N = 3 mice, n = 18 fibers). These low-frequencies of mEPPs are consistent with previous reports on wildtype mammalian embryonic muscles (Dennis et al., 1981; Diamond and Miledi, 1962). Similarly, mEPP amplitude, mEPP rise time (10–90%), mEPP half width (Fig. 8B) and EPP amplitude (Fig. 8D) were also similar between the two genotypes, and action potentials were evoked by electrical stimulation of the phrenic nerves in both control and Ctnnb1lox(ex3)/+;Myo-Cre muscles (Fig. 8E).

Figure 8. Normal synaptic transmission at the NMJ in mice expressing stabilized β-catenin in muscles.

Figure 8

Open in a new tab

A: Sample mEPP traces (superimposed 120 X 1-second traces) from E18.5 diaphragm muscle in control and Ctnnb1lox(ex3)/+;Myo-Cre mice. B: Quantification of mEPP frequencies, amplitudes, 10–90% rise time and half width – mEPP frequencies (event/minute): 0.90 ± 0.15 (Ctnnb1lox(ex3)/+;Myo-Cre) vs. 0.93 ± 0.15 (control); mEPP amplitudes 2.60 ± 0.25 mV (Ctnnb1lox(ex3)/+;Myo-Cre) vs. 2.12 ± 0.26 mV (control); mEPP rise time: 3.87 ± 0.24 ms (Ctnnb1lox(ex3)/+;Myo-Cre) vs. 3.84 ± 0.56 ms (control); mEPP half width: 12.82 ± 0.24 ms (Ctnnb1lox(ex3)/+;Myo-Cre) vs. 11.74 ± 0.74 ms (control). No statistical differences were found between the Ctnnb1lox(ex3)/+;Myo-Cre and the controls. C-D: Sample traces and quantification of EPP amplitudes. EPP amplitudes were indistinguishable between controls (10.67 ± 0.96 mV) and Ctnnb1lox(ex3)/+;Myo-Cre (10.39 ± 1.3 mV). Number of sample analyzed: Ctnnb1lox(ex3)/+;Myo-Cre (n = 25 cells, N = 4 mice); control (n = 18 cells, N = 3 mice). E: Action potentials were evoked by electrical stimulation of the phrenic nerves in both control and Ctnnb1lox(ex3)/+;Myo-Cre diaphragm muscles.

Next, we measured muscle contractile properties in E18.5 diaphragm muscles. We found that muscle twitch responses and twitch force were similar between control and Ctnnb1lox(ex3)/+;Myo-Cre muscles (Fig. 9). In both control and Ctnnb1lox(ex3)/+;Myo-Cre diaphragm muscles, muscle twitch force increased in response to stimulation frequency increases, and reached the maximal twitch force (normalized by muscle weight) at 0.78 ± 0.06 g/mg (control, N = 3) and 0.70 ± 0.07 g/mg (Ctnnb1lox(ex3)/+;Myo-Cre, N = 3). Together, these results demonstrated that both neuromuscular synaptic transmission and muscle contractile properties were normal in mice expressing stabilized β-catenin in muscles.

Figure 9. Normal muscle contractile properties in mice expressing stabilized β-catenin in muscles.

Figure 9

Open in a new tab

A: Muscle twitch responses at 4, 10, 20 and 50 Hz in control and Ctnnb1lox(ex3)/+;Myo-Cre diaphragm muscles (E18.5). B: Plot of twitch force (expressed as % of maximal force) against stimulation frequency. In both control and Ctnnb1lox(ex3)/+;Myo-Cre diaphragm muscles, muscle twitch force increases as stimulation frequency increases. C: Normalized maximal twitch force by muscle weight. No significant difference was detected between control (0.78 ± 0.06 g/mg, N = 3 embryos) and Ctnnb1lox(ex3)/+;Myo-Cre (0.70 ± 0.07 g/mg, N = 3 embryos).

Normal formation and function of the NMJ in mice with β-catenin stabilization specifically in motor neurons

Previous studies have demonstrated that Wnt signaling in pre-synaptic cells promotes nerve terminal differentiation in cerebellar synapses (Hall et al., 2000; Lucas and Salinas, 1997), in Drosophila NMJs (Ataman et al., 2008; Packard et al., 2002), and regulates synaptic specificity in C. elegans NMJs (Klassen and Shen, 2007; Maro et al., 2009). In hippocampal neurons, β̃catenin regulates the localization of the synaptic vesicle pool (Bamji et al., 2003), synaptic size and synaptic strength (Murase et al., 2002; Okuda et al., 2007) and dendritic morphogenesis (Yu and Malenka, 2003). We therefore sought to determine the effect of β-catenin stabilization in motor neurons. We generated and analyzed mice that express stabilized β-catenin specifically in motor neurons by breeding the Ctnnb1lox(ex3)/lox(ex3) mice with HB9Cre mice, in which Cre recombinase has been shown to be expressed specifically in motor neurons from E9 onwards (Arber et al., 1999; Yang et al., 2001), a stage similar to the onset of Myo-Cre expression described above.

In contrast to Ctnnb1lox(ex3)/+;Myo-Cre mice, which died shortly after birth, mice expressing stabilized β-catenin in motor neurons (Ctnnb1lox(ex3)/+;HB9Cre) survived postnatally. Furthermore, in contrast to the aberrant innervation pattern detected in Ctnnb1lox(ex3)/+;Myo-Cre mice (Figs 2–5), the pattern of innervation in Ctnnb1lox(ex3)/+;HB9Cre/+ mice was normal (Fig. 10A-B). High-power examination of the pre-synaptic nerve terminals and postsynaptic AChRs at E18.5 (Fig. 10C-H) and P45 (Fig. 10I-N) showed normal formation and maturation of the NMJ in Ctnnb1lox(ex3)/+;HB9Cre/+mice. Electrophysiological analysis further demonstrated that the neuromuscular synapses functioned normally in Ctnnb1lox(ex3)/+;HB9Cre/+ mice: mEPP frequencies, amplitude, rise time and half-width were similar between Ctnnb1lox(ex3)/+;HB9Cre/+ and their littermate controls (Ctnnb1lox(ex3)/+) (Fig. 11A-B). EPP amplitudes, rise time and half width were also similar between Ctnnb1lox(ex3)/+;HB9Cre/+ and their littermate controls (Ctnnb1lox(ex3)/+) (Fig. 11C-D), as was the rate of EPP run-down, a measure of short-term synaptic plasticity of the NMJ (Fig. 11E-F). Together, these findings indicate that activation of the Wnt/β-catenin signaling in motor neurons does not affect the patterning, formation, or function of the NMJ.

Figure 10. Neuromuscular synapses develop normally in mice with β-catenin stabilization in motor neurons.

Figure 10

Open in a new tab

Hemi-diaphragm muscles (E18.5) were double-labeled with anti-NF150 and anti-Syt2 antibodies and α-bgt. Similar innervation patterns were seen in the control (A) and the mice expressing stabilized β-catenin specifically in motor neurons (B, Ctnnb1lox(ex3)/+;HB9cre). High-power magnification of the diaphragm muscle in A and B, demonstrating individual NMJs were developed in control (C-E) and Ctnnb1lox(ex3)/+;HB9cre mice (F-H). I-N: NMJs in triangularis sterni muscles (P45) of control (I-K) and Ctnnb1lox(ex3)/+;HB9cre mice (L-N). Scale bars: A-B, 400 µm; C-H, 20µm; I-N, 20 µm.

Figure 11. Neuromuscular synapses function normally in mice with β-catenin stabilization in motor neurons.

Figure 11

Open in a new tab

A: Typical mEPP traces (superimposed 20 X 0.5-second traces) from the lumbrical muscles at P45 in control and Ctnnb1lox(ex3)/+;HB9Cre mice. B: Quantification of mEPP frequencies, amplitudes, 10–90% rise time and half width – mEPP frequencies (Hz) were 0.62 ± 0.08 in control and 0.62 ± 0.12 in Ctnnb1lox(ex3)/+;HB9Cre; mEPP amplitudes: 1.63 ± 0.13 mV (control), 1.68 ± 0.16 mV (Ctnnb1lox(ex3)/+;HB9Cre); mEPP rise time: 2.20 ± 0.24 ms (control), 2.26 ± 0.26 ms (Ctnnb1lox(ex3)/+;HB9Cre); mEPP half width: 4.12 ± 0.24 ms (control), 4.33 ± 0.29 ms (Ctnnb1lox(ex3)/+;HB9Cre). No statistical differences were found between control and Ctnnb1lox(ex3)/+;HB9Cre mice. C-D: Sample traces (C) and quantification of EPP (D). EPP amplitudes (control: 23.96 ± 1.44 mV; Ctnnb1lox(ex3)/+;HB9Cre: 22.67 ± 1.41 mV), EPP rise time (control: 1.24 ± 0.04 ms; Ctnnb1lox(ex3)/+;HB9Cre: 1.30 ± 0.05 ms) and EPP half width (control: 3.92 ± 0.09 ms; Ctnnb1lox(ex3)/+;HB9Cre: 4.09 ± 0.14 ms) were indistinguishable between control and Ctnnb1lox(ex3)/+;HB9Cre mice. E-F: Sample EPP traces evoked by electrical stimulation of the nerve at 70 Hz (E) and quantification of EPP rundown ratio. The rate of EPP rundown is similar between controls and Ctnnb1lox(ex3)/+;HB9cre mice. Number of sample analyzed: control (n = 13); Ctnnb1lox(ex3)/+;HB9Cre (n = 11).

Discussion

Previous studies have indicated that β-catenin is involved in the development of the NMJ. However, the precise role for β-catenin in NMJ formation remains unclear. To gain insight into the role of β-catenin in the development of the NMJ, we expressed a stabilized form of β-catenin in either motor neurons or skeletal muscles. Surprisingly, β-catenin stabilization, either in motor neurons or in muscles, does not adversely affect NMJ formation or function. However, β-catenin stabilization in skeletal muscle leads to abnormal motor innervation of the muscle, including increased defasciculation and nerve branching. These abnormal patterns displayed in mutant mice expressing stabilized β-catenin in muscles resemble that reported in a previous study in mutant mice lacking β-catenin in skeletal muscles (Li et al., 2008). Thus, β-catenin levels in the skeletal muscle appear to be crucial for the establishment of normal motor innervation pattern. The fact that these defects are detected only when β-catenin is altered in muscles, but not in motor neurons, indicates that β-catenin regulates motor innervation through a non-cell autonomous mechanism.

During the development of zebrafish NMJ, Wnt-11r binds to unplugged/MuSK ectodomain and regulates axon guidance and AChR prepatterning (Jing et al., 2010; Jing et al., 2009). As interactions of Wnt proteins and their receptors may activate a variety of signaling pathways (van Amerongen and Nusse, 2009), it remains to be determined whether β-catenin signaling is involved in AChR prepatterning in zebrafish NMJ. We found that β-catenin stabilization in mouse skeletal muscles results in a broadening of the end-plate band in muscles, but it does not appear to disrupt muscle pre-patterning, as the end-plate band, though broadened, still lies in the central region of the muscle.

Our data have demonstrated that β-catenin stabilization in muscles, but not in motor neurons, leads to neonatal lethality – mutant mice died within 1 day of birth. The direct cause of the lethality remains to be determined, as neonatal lethality in mutant mice can result from impairments of multiple physiological systems (Turgeon and Meloche, 2009). At the time of death, mutant pups lacked milk in their stomachs, indicating that they were unable to feed, which may have contributed to the lethality. Furthermore, aberrant innervation pattern and the presence of muscle patches in the central tendon of the diaphragm may also contribute, at least in part, to the lethality. The diaphragm muscle is unique to mammals and is the major muscle of respiration. The central tendon of the diaphragm normally does not contain muscles (Allan and Greer, 1997; Greer et al., 1999) and diaphragmatic defects often lead to perinatal lethality (Ackerman and Greer, 2007). Consistent with this hypothesis, the presence of muscle patches in the central tendon of the diaphragm has also been reported in human birth defects (Blair, 1923; Colacino and Pettersen, 1978).

The increases in nerve defasciculation and branching in the intramuscular nerve bundles on the phrenic nerve in mice expressing stabilized β-catenin can be detected as early as E13.5, at a stage after the completion of naturally occurring motor neuron cell death in the cervical spinal segments (Yamamoto and Henderson, 1999) but prior to the establishment of functional NMJs between phrenic motor neurons and their target diaphragm muscles (Witzemann, 2006). Therefore, the increase in nerve branching may be attributed to an increase in motor neuron numbers but is likely independent of the level on neuromuscular activity. Indeed, neither neuromuscular synaptic activity nor muscle contraction is affected by β-catenin stabilization in muscles.

The precise mechanism for how β-catenin regulates motor innervation remains to be elucidated. Remarkably, innervation defects displayed in the muscle expressing stabilized β-catenin resemble the defects seen in Slit2 or Robo1/Robo2 mutant mice (Jaworski and Tessier-Lavigne, 2012). This raises an intriguing possibility that β-catenin stabilization in muscles may down-regulate Slit2 and/or Robo1/Robo2, or down-regulate the signaling components downstream of Slit2/Robo1/Robo2. This further suggests a potential functional interaction between Wnt/β-catenin signaling and Slit2/Robo1/Robo2 signaling during the development of motor innervation in mouse embryonic muscles.

Supplementary Material

01

Acknowledgments

We thank Drs. Eric Olson and Rhonda Bassel-Duby (UT Southwestern Medical Center) for providing Myo-Cre mice. This work was supported by grants (to W. Lin) from NIH/NINDS (NS055028, NS055028-03S1), the Edward Mallinckrodt, Jr. Scholar Program and the Cain Foundation in Medical Research.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  1. Aberle H, Bauer A, Stappert J, Kispert A, Kemler R. beta-catenin is a target for the ubiquitin-proteasome pathway. Embo J. 1997;16:3797–3804. doi: 10.1093/emboj/16.13.3797. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Ackerman KG, Greer JJ. Development of the diaphragm and genetic mouse models of diaphragmatic defects. Am J Med Genet C Semin Med Genet. 2007;145C:109–116. doi: 10.1002/ajmg.c.30128. [DOI] [PubMed] [Google Scholar]
  3. Allan DW, Greer JJ. Embryogenesis of the phrenic nerve and diaphragm in the fetal rat. J Comp Neurol. 1997;382:459–468. doi: 10.1002/(sici)1096-9861(19970616)382:4<459::aid-cne3>3.0.co;2-1. [DOI] [PubMed] [Google Scholar]
  4. Apel ED, Glass DJ, Moscoso LM, Yancopoulos GD, Sanes JR. Rapsyn is required for MuSK signaling and recruits synaptic components to a MuSK-containing scaffold. Neuron. 1997;18:623–635. doi: 10.1016/s0896-6273(00)80303-7. [DOI] [PubMed] [Google Scholar]
  5. Arber S, Han B, Mendelsohn M, Smith M, Jessell TM, Sockanathan S. Requirement for the homeobox gene Hb9 in the consolidation of motor neuron identity. Neuron. 1999;23:659–674. doi: 10.1016/s0896-6273(01)80026-x. [DOI] [PubMed] [Google Scholar]
  6. Ataman B, Ashley J, Gorczyca M, Ramachandran P, Fouquet W, Sigrist SJ, Budnik V. Rapid activity-dependent modifications in synaptic structure and function require bidirectional Wnt signaling. Neuron. 2008;57:705–718. doi: 10.1016/j.neuron.2008.01.026. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Bamji SX, Shimazu K, Kimes N, Huelsken J, Birchmeier W, Lu B, Reichardt LF. Role of beta-catenin in synaptic vesicle localization and presynaptic assembly. Neuron. 2003;40:719–731. doi: 10.1016/s0896-6273(03)00718-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Behrens J, von Kries JP, Kuhl M, Bruhn L, Wedlich D, Grosschedl R, Birchmeier W. Functional interaction of beta-catenin with the transcription factor LEF-1. Nature. 1996;382:638–642. doi: 10.1038/382638a0. [DOI] [PubMed] [Google Scholar]
  9. Bennett MR, Pettigrew AG. The formation of synapses in striated muscle during development. J Physiol. 1974;241:515–545. doi: 10.1113/jphysiol.1974.sp010670. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Blair DM. A Study of the Central Tendon of the Diaphragm. J Anat. 1923;57:203–215. [PMC free article] [PubMed] [Google Scholar]
  11. Burden SJ. The formation of neuromuscular synapses. Genes Dev. 1998:133–148. doi: 10.1101/gad.12.2.133. [DOI] [PubMed] [Google Scholar]
  12. Burgess RW, Skarnes WC, Sanes JR. Agrin isoforms with distinct amino termini: differential expression, localization, and function. J Cell Biol. 2000;151:41–52. doi: 10.1083/jcb.151.1.41. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Cadigan KM, Nusse R. Wnt signaling: a common theme in animal development. Genes Dev. 1997;11:3286–3305. doi: 10.1101/gad.11.24.3286. [DOI] [PubMed] [Google Scholar]
  14. Cifuentes-Diaz C, Goudou D, Mege RM, Velasco E, Nicolet M, Herrenknecht K, Rubin L, Rieger F. Distinct location and prevalence of alpha-, beta-catenins and gamma-catenin/plakoglobin in developing and denervated skeletal muscle. Cell Adhes Commun. 1998;5:161–176. doi: 10.3109/15419069809040289. [DOI] [PubMed] [Google Scholar]
  15. Colacino SC, Pettersen JC. Analysis of the gross anatomical variations found in four cases of trisomy 13. Am J Med Genet. 1978;2:31–50. doi: 10.1002/ajmg.1320020106. [DOI] [PubMed] [Google Scholar]
  16. DeChiara TM, Bowen DC, Valenzuela DM, Simmons MV, Poueymirou WT, Thomas S, Kinetz E, Compton DL, Rojas E, Park JS, Smith C, DiStefano PS, Glass DJ, Burden SJ, Yancopoulos GD. The receptor tyrosine kinase MuSK is required for neuromuscular junction formation in vivo. Cell. 1996;85:501–512. doi: 10.1016/s0092-8674(00)81251-9. [DOI] [PubMed] [Google Scholar]
  17. Dennis MJ, Ziskind-Conhaim L, Harris AJ. Development of neuromuscular junctions in rat embryos. Dev Biol. 1981;81:266–279. doi: 10.1016/0012-1606(81)90290-6. [DOI] [PubMed] [Google Scholar]
  18. Diamond J, Miledi R. A study of foetal and new-born rat muscle fibres. J Physiol. 1962;162:393–408. doi: 10.1113/jphysiol.1962.sp006941. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Enomoto H, Araki T, Jackman A, Heuckeroth RO, Snider WD, Johnson EM, Jr., Milbrandt J. GFR alpha1-deficient mice have deficits in the enteric nervous system and kidneys. Neuron. 1998;21:317–324. doi: 10.1016/s0896-6273(00)80541-3. [DOI] [PubMed] [Google Scholar]
  20. Ferns M, Hoch W, Campanelli JT, Rupp F, Hall ZW, Scheller RH. RNA splicing regulates agrin-mediated acetylcholine receptor clustering activity on cultured myotubes. Neuron. 1992;8:1079–1086. doi: 10.1016/0896-6273(92)90129-2. [DOI] [PubMed] [Google Scholar]
  21. Froehner SC, Gulbrandsen V, Hyman C, Jeng AY, Neubig RR, Cohen JB. Immunofluorescence localization at the mammalian neuromuscular junction of the Mr 43,000 protein of Torpedo postsynaptic membranes. Proc Natl Acad Sci U S A. 1981;78:5230–5234. doi: 10.1073/pnas.78.8.5230. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Gautam M, Noakes PG, Moscoso L, Rupp F, Scheller RH, Merlie JP, Sanes JR. Defective neuromuscular synaptogenesis in agrin-deficient mutant mice. Cell. 1996;85:525–535. doi: 10.1016/s0092-8674(00)81253-2. [DOI] [PubMed] [Google Scholar]
  23. Gautam M, Noakes PG, Mudd J, Nichol M, Chu GC, Sanes JR, Merlie JP. Failure of postsynaptic specialization to develop at neuromuscular junctions of rapsyn-deficient mice. Nature. 1995;377:232–236. doi: 10.1038/377232a0. [DOI] [PubMed] [Google Scholar]
  24. Goichberg P, Shtutman M, Ben-Ze'ev A, Geiger B. Recruitment of beta-catenin to cadherin-mediated intercellular adhesions is involved in myogenic induction. J Cell Sci. 2001;114:1309–1319. doi: 10.1242/jcs.114.7.1309. [DOI] [PubMed] [Google Scholar]
  25. Greer JJ, Allan DW, Martin-Caraballo M, Lemke RP. An overview of phrenic nerve and diaphragm muscle development in the perinatal rat. J Appl Physiol. 1999;86:779–786. doi: 10.1152/jappl.1999.86.3.779. [DOI] [PubMed] [Google Scholar]
  26. Hall AC, Lucas FR, Salinas PC. Axonal remodeling and synaptic differentiation in the cerebellum is regulated by WNT-7a signaling. Cell. 2000;100:525–535. doi: 10.1016/s0092-8674(00)80689-3. [DOI] [PubMed] [Google Scholar]
  27. Hall ZW, Sanes JR. Synaptic structure and development: the neuromuscular junction. Cell. 1993;72(Suppl):99–121. doi: 10.1016/s0092-8674(05)80031-5. [DOI] [PubMed] [Google Scholar]
  28. Harada N, Tamai Y, Ishikawa T, Sauer B, Takaku K, Oshima M, Taketo MM. Intestinal polyposis in mice with a dominant stable mutation of the beta-catenin gene. Embo J. 1999;18:5931–5942. doi: 10.1093/emboj/18.21.5931. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. He X, Semenov M, Tamai K, Zeng X. LDL receptor-related proteins 5 and 6 in Wnt/beta-catenin signaling: arrows point the way. Development. 2004;131:1663–1677. doi: 10.1242/dev.01117. [DOI] [PubMed] [Google Scholar]
  30. Henriquez JP, Webb A, Bence M, Bildsoe H, Sahores M, Hughes SM, Salinas PC. Wnt signaling promotes AChR aggregation at the neuromuscular synapse in collaboration with agrin. Proc Natl Acad Sci U S A. 2008;105:18812–18817. doi: 10.1073/pnas.0806300105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Inoue A, Setoguchi K, Matsubara Y, Okada K, Sato N, Iwakura Y, Higuchi O, Yamanashi Y. Dok-7 activates the muscle receptor kinase MuSK and shapes synapse formation. Sci Signal. 2009;2:ra7. doi: 10.1126/scisignal.2000113. [DOI] [PubMed] [Google Scholar]
  32. Jaworski A, Tessier-Lavigne M. Autocrine/juxtaparacrine regulation of axon fasciculation by Slit-Robo signaling. Nat Neurosci. 2012;15:367–369. doi: 10.1038/nn.3037. [DOI] [PubMed] [Google Scholar]
  33. Jing L, Gordon LR, Shtibin E, Granato M. Temporal and spatial requirements of unplugged/MuSK function during zebrafish neuromuscular development. PLoS One. 2010;5:e8843. doi: 10.1371/journal.pone.0008843. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Jing L, Lefebvre JL, Gordon LR, Granato M. Wnt signals organize synaptic prepattern and axon guidance through the zebrafish unplugged/MuSK receptor. Neuron. 2009;61:721–733. doi: 10.1016/j.neuron.2008.12.025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Kim N, Stiegler AL, Cameron TO, Hallock PT, Gomez AM, Huang JH, Hubbard SR, Dustin ML, Burden SJ. Lrp4 is a receptor for Agrin and forms a complex with MuSK. Cell. 2008;135:334–342. doi: 10.1016/j.cell.2008.10.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Klassen MP, Shen K. Wnt signaling positions neuromuscular connectivity by inhibiting synapse formation in C. elegans. Cell. 2007;130:704–716. doi: 10.1016/j.cell.2007.06.046. [DOI] [PubMed] [Google Scholar]
  37. Li S, Czubryt MP, McAnally J, Bassel-Duby R, Richardson JA, Wiebel FF, Nordheim A, Olson EN. Requirement for serum response factor for skeletal muscle growth and maturation revealed by tissue-specific gene deletion in mice. Proc Natl Acad Sci U S A. 2005;102:1082–1087. doi: 10.1073/pnas.0409103102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Li XM, Dong XP, Luo SW, Zhang B, Lee DH, Ting AK, Neiswender H, Kim CH, Carpenter-Hyland E, Gao TM, Xiong WC, Mei L. Retrograde regulation of motoneuron differentiation by muscle beta-catenin. Nat Neurosci. 2008;11:262–268. doi: 10.1038/nn2053. [DOI] [PubMed] [Google Scholar]
  39. Liley AW. An investigation of spontaneous activity at the neuromuscular junction of the rat. J Physiol. 1956;132:650–666. doi: 10.1113/jphysiol.1956.sp005555. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Liu Y, Padgett D, Takahashi M, Li H, Sayeed A, Teichert RW, Olivera BM, McArdle JJ, Green WN, Lin W. Essential roles of the acetylcholine receptor {gamma}-subunit in neuromuscular synaptic patterning. Development. 2008;135:1957–1967. doi: 10.1242/dev.018119. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Logan CY, Nusse R. The Wnt signaling pathway in development and disease. Annu Rev Cell Dev Biol. 2004;20:781–810. doi: 10.1146/annurev.cellbio.20.010403.113126. [DOI] [PubMed] [Google Scholar]
  42. Lucas FR, Salinas PC. WNT-7a induces axonal remodeling and increases synapsin I levels in cerebellar neurons. Dev Biol. 1997;192:31–44. doi: 10.1006/dbio.1997.8734. [DOI] [PubMed] [Google Scholar]
  43. Luo ZG, Wang Q, Zhou JZ, Wang J, Luo Z, Liu M, He X, Wynshaw-Boris A, Xiong WC, Lu B, Mei L. Regulation of AChR clustering by Dishevelled interacting with MuSK and PAK1. Neuron. 2002;35:489–505. doi: 10.1016/s0896-6273(02)00783-3. [DOI] [PubMed] [Google Scholar]
  44. Maro GS, Klassen MP, Shen K. A beta-catenin-dependent Wnt pathway mediates anteroposterior axon guidance in C. elegans motor neurons. PLoS One. 2009;4:e4690. doi: 10.1371/journal.pone.0004690. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Martin-Caraballo M, Campagnaro PA, Gao Y, Greer JJ. Contractile and fatigue properties of the rat diaphragm musculature during the perinatal period. J Appl Physiol. 2000;88:573–580. doi: 10.1152/jappl.2000.88.2.573. [DOI] [PubMed] [Google Scholar]
  46. McMahan UJ. The agrin hypothesis. Cold Spring Harb Symp Quant Biol. 1990;55:407–418. doi: 10.1101/sqb.1990.055.01.041. [DOI] [PubMed] [Google Scholar]
  47. Murase S, Mosser E, Schuman EM. Depolarization drives beta-Catenin into neuronal spines promoting changes in synaptic structure and function. Neuron. 2002;35:91–105. doi: 10.1016/s0896-6273(02)00764-x. [DOI] [PubMed] [Google Scholar]
  48. Nastasi T, Bongiovanni A, Campos Y, Mann L, Toy JN, Bostrom J, Rottier R, Hahn C, Conaway JW, Harris AJ, D'Azzo A. Ozz-E3, a muscle-specific ubiquitin ligase, regulates beta-catenin degradation during myogenesis. Dev Cell. 2004;6:269–282. doi: 10.1016/s1534-5807(04)00020-6. [DOI] [PubMed] [Google Scholar]
  49. Nitkin RM, Smith MA, Magill C, Fallon JR, Yao YM, Wallace BG, McMahan UJ. Identification of agrin, a synaptic organizing protein from Torpedo electric organ. J Cell Biol. 1987;105:2471–2478. doi: 10.1083/jcb.105.6.2471. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Okada K, Inoue A, Okada M, Murata Y, Kakuta S, Jigami T, Kubo S, Shiraishi H, Eguchi K, Motomura M, Akiyama T, Iwakura Y, Higuchi O, Yamanashi Y. The muscle protein Dok-7 is essential for neuromuscular synaptogenesis. Science. 2006;312:1802–1805. doi: 10.1126/science.1127142. [DOI] [PubMed] [Google Scholar]
  51. Okuda T, Yu LM, Cingolani LA, Kemler R, Goda Y. beta-Catenin regulates excitatory postsynaptic strength at hippocampal synapses. Proc Natl Acad Sci U S A. 2007;104:13479–13484. doi: 10.1073/pnas.0702334104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Orford K, Crockett C, Jensen JP, Weissman AM, Byers SW. Serine phosphorylation-regulated ubiquitination and degradation of beta-catenin. J Biol Chem. 1997;272:24735–24738. doi: 10.1074/jbc.272.40.24735. [DOI] [PubMed] [Google Scholar]
  53. Packard M, Koo ES, Gorczyca M, Sharpe J, Cumberledge S, Budnik V. The Drosophila Wnt, wingless, provides an essential signal for pre- and postsynaptic differentiation. Cell. 2002;111:319–330. doi: 10.1016/s0092-8674(02)01047-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Ruegg MA, Tsim KW, Horton SE, Kroger S, Escher G, Gensch EM, McMahan UJ. The agrin gene codes for a family of basal lamina proteins that differ in function and distribution. Neuron. 1992;8:691–699. doi: 10.1016/0896-6273(92)90090-z. [DOI] [PubMed] [Google Scholar]
  55. Soriano P. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat Genet. 1999;21:70–71. doi: 10.1038/5007. [DOI] [PubMed] [Google Scholar]
  56. Speese SD, Budnik V. Wnts: up-and-coming at the synapse. Trends Neurosci. 2007;30:268–275. doi: 10.1016/j.tins.2007.04.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Turgeon B, Meloche S. Interpreting neonatal lethal phenotypes in mouse mutants: insights into gene function and human diseases. Physiol Rev. 2009;89:1–26. doi: 10.1152/physrev.00040.2007. [DOI] [PubMed] [Google Scholar]
  58. van Amerongen R, Nusse R. Towards an integrated view of Wnt signaling in development. Development. 2009;136:3205–3214. doi: 10.1242/dev.033910. [DOI] [PubMed] [Google Scholar]
  59. Wang J, Jing Z, Zhang L, Zhou G, Braun J, Yao Y, Wang ZZ. Regulation of acetylcholine receptor clustering by the tumor suppressor APC. Nat Neurosci. 2003;6:1017–1018. doi: 10.1038/nn1128. [DOI] [PubMed] [Google Scholar]
  60. Weatherbee SD, Anderson KV, Niswander LA. LDL-receptor-related protein 4 is crucial for formation of the neuromuscular junction. Development. 2006;133:4993–5000. doi: 10.1242/dev.02696. [DOI] [PubMed] [Google Scholar]
  61. Witzemann V. Development of the neuromuscular junction. Cell Tissue Res. 2006 doi: 10.1007/s00441-006-0237-x. [DOI] [PubMed] [Google Scholar]
  62. Wu F, Mi W, Burns DK, Fu Y, Gray HF, Struyk AF, Cannon SC. A sodium channel knockin mutant (NaV1.4-R669H) mouse model of hypokalemic periodic paralysis. J Clin Invest. 2011;121:4082–4094. doi: 10.1172/JCI57398. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Wu H, Xiong WC, Mei L. To build a synapse: signaling pathways in neuromuscular junction assembly. Development. 2010;137:1017–1033. doi: 10.1242/dev.038711. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Yamamoto Y, Henderson CE. Patterns of programmed cell death in populations of developing spinal motoneurons in chicken, mouse, and rat. Dev Biol. 1999;214:60–71. doi: 10.1006/dbio.1999.9413. [DOI] [PubMed] [Google Scholar]
  65. Yang X, Arber S, William C, Li L, Tanabe Y, Jessell TM, Birchmeier C, Burden SJ. Patterning of muscle acetylcholine receptor gene expression in the absence of motor innervation. Neuron. 2001;30:399–410. doi: 10.1016/s0896-6273(01)00287-2. [DOI] [PubMed] [Google Scholar]
  66. Yu X, Malenka RC. Beta-catenin is critical for dendritic morphogenesis. Nat Neurosci. 2003;6:1169–1177. doi: 10.1038/nn1132. [DOI] [PubMed] [Google Scholar]
  67. Zhang B, Luo S, Dong XP, Zhang X, Liu C, Luo Z, Xiong WC, Mei L. Beta-catenin regulates acetylcholine receptor clustering in muscle cells through interaction with rapsyn. J Neurosci. 2007;27:3968–3973. doi: 10.1523/JNEUROSCI.4691-06.2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Zhang B, Luo S, Wang Q, Suzuki T, Xiong WC, Mei L. LRP4 serves as a coreceptor of agrin. Neuron. 2008;60:285–297. doi: 10.1016/j.neuron.2008.10.006. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

01
NIHMS371811-supplement-01.pdf (2MB, pdf)

RESOURCES