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Muscarinic acetylcholine receptors (mAChRs) modulate synaptic function, but whether they influence synaptic structure remains unknown. At neuromuscular junctions (NMJs), mAChRs have been implicated in compensatory sprouting of axon terminals in paralyzed or denervated muscles. Here we used pharmacological and genetic inhibition, and localization studies of mAChR subtypes at mouse NMJs to demonstrate their roles in synaptic stability and growth, but not in compensatory sprouting. M2 mAChRs were present solely in motor neurons, whereas M1, M3, and M5 mAChRs were associated with Schwann cells and/or muscle fibers. Blockade of all five mAChR subtypes with atropine evoked pronounced effects, including terminal sprouting, terminal withdrawal, and muscle fiber atrophy. In contrast, methoctramine, an M2/M4 preferring antagonist, induced terminal sprouting and terminal withdrawal, but no muscle fiber-atrophy. Consistent with this observation, M2-/- but no other mAChR mutant mice exhibited spontaneous sprouting accompanied by extensive loss of parental terminal arbors. Terminal sprouting, however, seemed not to be the causative defect because partial loss of terminal branches was common even in the M2-/- NMJs without sprouting. Moreover, compensatory sprouting after paralysis or partial denervation was normal in mice deficient in M2 or other mAChR subtypes. We also found that many NMJs of M5-/- mice were exceptionally small, and reduced in proportion to the size of parental muscle fibers. These findings show that axon terminals are unstable without M2 and that muscle fiber growth is defective without M5. Subtype-specific muscarinic signaling provides a novel means for coordinating activity-dependant development and maintenance of the tripartite synapse.
Acetylcholine, the neurotransmitter of skeletal neuromuscular junctions (NMJs), activates two structurally and functionally distinct types of receptors- nicotinic (nAChR) and muscarinic (mAChR). nAChRs function as cation channels whereas mAChRs are metabotropic receptors signaling through G-proteins. The five mammalian mAChR subtypes, M1–M5, are comprised of two functionally distinct groups: odd-numbered mAChRs (M1, M3, M5) preferentially activate Gq/G11 type G-proteins; even-numbered mAChRs (M2, M4) activate Gi/Go type G-proteins. Most tissues and cell types express two or more mAChR subtypes that exert diverse physiological actions, depending on the cellular location and identity of receptor subtypes (Wess et al., 2007; Nathanson, 2008). In the nervous system, mAChRs act primarily as modulators of synaptic transmission, regulating cognitive, sensory, motor and autonomic functions, and are implicated in the pathophysiology of illnesses such as Alzheimer's disease, Parkinson's disease, depression and schizophrenia (cf., Wess, 2004; Langmead et al., 2008). It remains unknown, however, whether muscarinic signaling plays a structural role at the synapse.
Paralysis or partial denervation elicits sprouting of motor nerve terminals at adult neuromuscular junctions (NMJs). These terminal sprouts add or restore synaptic contacts on inactive or denervated muscle fibers (Meunier et al., 2002; Rogozhin et al., 2008). Perisynaptic terminal Schwann cells (tSCs) at the NMJ mediate the compensatory process: they become activated in response to paralysis or denervation (Reynolds and Woolf, 1992), and extend fine processes, the hallmark of tSC activation, that induce and guide terminal sprouts (Son and Thompson, 1995a; Ko and Chen, 1996). The molecular events that link synaptic inactivity to tSC activation are unknown but muscarinic signaling may play a role: tSCs monitor transmitter release via unidentified mAChR subtypes (Jahromi et al., 1992; Reist and Smith, 1992) and respond to paralysis or mAChR blockade by reducing internal calcium and upregulating glial fibrillary acidic protein (GFAP; Robitaille et al., 1997; Georgiou et al., 1999; Rochon et al., 2001; Todd et al., 2007). Upregulation of GFAP is a characteristic of activated tSCs (Georgiou et al., 1999; Triolo et al., 2006). Thus, tSC mAChRs are attractive candidates as the long-sought initiators of compensatory plasticity of mature NMJs.
Other evidence suggests that mAChRs, especially M1 and M2, are present in the axon terminals, acting as autoreceptors that inhibit ACh release (Wessler, 1989; Re, 1999; but see Minic et al., 2002; Slutsky et al., 2003). Studies also imply that mAChRs, like nAChRs, are present in muscle fibers, where they are hypothesized to mediate the trophic effects of neurotransmitters (Reyes and Jaimovich, 1996; Furlan and Godinho, 2005).
We sought to test if muscarinic signaling initiates compensatory sprouting or plays other roles at NMJs. Using pharmacological, genetic, electrophysiological and laser-assisted localization studies, we found that compensatory sprouting proceeds normally in the absence of muscarinic signaling. We also discovered that nerve terminal arbors are unstable without presynaptic M2, and that muscle fiber growth is defective without peri- and/or postsynaptically associated M5. These findings are the first to show that mAChRs influence synaptic structure, and that specific subtypes of mAChRs play distinct roles at NMJs.
Mice deficient in each mAChR subtype (i.e., M1 -/-, M2 -/-, M3 -/-, M4 -/-, M5 -/- mice), as well as M1/3 (M1/3-/-) and M2/4 (M2/4-/-) double Knockout (dKO) mice were generated and maintained as described in detail previously (Gomeza et al., 1999; Gomeza et al., 2001; Yamada et al., 2001; Fisahn et al., 2002; Stengel et al., 2002; Wess et al., 2003; Yamada et al., 2003; Gautam et al., 2006). Congenic mouse lines with C57BL/6NTac background were used except M1 -/- and M3 -/- mice that were maintained on a 129SvEv x CF1 background. Mice were amplified at Drexel University. Genotypes were confirmed by PCR analysis of mouse-tail DNA. We used adult female C57Bl/6 mice for pharmacological experiments. Female mice were used because they are easier to handle and more tolerant than male mice of the repeated anesthesia and injections. All animal experiments were performed in accordance with Drexel University College of Medicine's Institutional Animal Care and Use Committee and National Institutes of Health guidelines.
Local in vivo blockade of one or more mAChR subtypes was achieved as previously described (Wright et al., 2007). Briefly, mice were anesthetized with an intraperitoneal injection of ketamine (120 mg/kg; Fort Dodge Laboratories, Inc., Fort Dodge, IA) and xylazine (8 mg/kg; Lloyd Laboratories, Shenandoah, Iowa). Under aseptic conditions, antagonists, dissolved in 50μl sterile physiological saline, were administered by subcutaneous injection over the right Levator auris longus (LAL) muscle. Sources and doses of the antagonists were as follows; atropine (Sigma-Aldrich, St. Louis, MO; for pharmaceutical grade atropine – Voigt Global Distribution, Lawrence, KS), 0.2 mg/kg - 20 mg/kg; Methoctramine (Sigma-Aldrich, St. Louis, MO), 100 - 400μM; 4-DAMP (Sigma-Aldrich, St. Louis, MO), 2.5mg/kg; AFDX-116 and AFDX-384 (Tocris Bioscience, Ellisville, MO); 250μM, 50μM-500μM, respectively; muscarinic toxin-7 (Peptides International, Louisville, KY); 0.1μM - 1μM. Control experiments for 4-DAMP administration were performed with saline containing an equal concentration of DMSO. We injected these reagents twice daily for five or seven days, unless noted in the text.
To assess synaptic transmission in M2-/- NMJs, seven M2-/- and four +/+ mice (2-2.5 months old) were euthanized with carbon dioxide inhalation, and the LAL muscle and its innervation were removed and placed in a chamber continuously perfused with Ringer solution containing (in millimoles per liter) NaCl, 118; KCl, 3.5; CaCl2, 2; MgSO4, 0.7; NaHCO3, 26.2; NaH2PO4, 1.7; glucose, 5.5 (pH 7.3-7.4, 20-22 °C) equilibrated with 95% O2 and 5% CO2. Following pinning, muscle was stained with 10 μM 4-Di-2ASP and imaged with an upright epifluorescence microscope as previously described (Wang et al., 2004; Wang et al., 2005). At this concentration, 4-Di-2ASP staining enables visualization of surface nerve terminals and individual surface muscle fibers. All endplates were imaged and muscles were impaled within 100 μm of the endplates. Muscle fibers were crushed distant from the endplate band and voltage clamped to −45 mV to avoid movement following nerve stimulation (Glavinovic, 1979). A two-electrode voltage clamp was used to measure the amplitude of miniature endplate currents (MEPCs) and endplate currents (EPCs) evoked by nerve stimulation. Quantal content was determined directly by dividing EPC amplitude by the average MEPC amplitude for a given endplate.
To assess reactivity of nerve terminals and tSCs to muscle paralysis, botulinum toxin A (BoTX; 2.0 pg/ 25 g mouse; Sigma-Aldrich, St. Louis MO), dissolved in 50 μl sterile saline, was injected to LAL muscles as described above. Similar dosages of BoTX induce complete paralysis of mouse LAL muscles for up to four weeks (Juzans et al., 1996; Angaut-Petit et al., 1998). To assure prolonged paralysis of LAL muscles, BoTX was administered again on the fourth day and the mice were euthanized on the seventh day after the first injection.
To examine reactive sprouting of nerve terminals and tSCs to partial denervation, the L4 and L5 spinal roots were exposed under aseptic conditions and the right L4 spinal root was transected to denervate partially the extensor digitorum longus (EDL) muscles by ~80% (Albani et al., 1988; Tam et al., 2001). The proximal segment of the transected L4 root was tied with 10.0 suture and the distal segment reflected laterally to prevent reinnervation by L4 axons. Fourteen days after partial denervation, mice were euthanized with an overdose of pentobarbital sodium (300mg/Kg; Abbott Laboratories, North Chicago, IL) and EDL muscles were immunoprocessed.
For NMJ microdissection, sternomastoid muscles of anesthetized mice were exposed and bathed in rhodamine conjugated α-bungarotoxin (Invitrogen, Carlsbad, CA) for 5 min to label postsynaptic nAChRs. Muscles were quickly removed and fresh frozen in isopentane cooled in liquid nitrogen. Muscles were sectioned longitudinally at 10μm thickness and collected on PEN (polyethylene naphthalate) foil slides specially designed for the Leica AS LMD Laser Micro Dissection system (Leica Microsystems, Wetzlar, Germany). Muscle slides were dehydrated with 70% ethanol for 2 min, DEPC-water for 30 sec, 75% ethanol for 30 sec, 95% ethanol for 10 sec, and placed in a 40°C oven to air dry for 7 min. For motor neuron microdissection, essentially the same procedure was followed except that lumbar spinal cord sections prepared in the transverse plane were processed with 1% toluidine blue solution for 30 sec to identify motor neurons. Dehydrated tissues were immediately used for LCM, which was operated as described by the manufacturer.
For each experiment, 350-400 NMJs or motor neurons were microdissected and pooled into PCR tubes for immediate stabilization. Total RNA was isolated using RNase easy Microkit and RNase free DNase kit (Qiagen, Valencia, CA), and stored at -80 °C before testing for expression of specific mRNAs by non-quantitative PCR. The amount of total RNA was determined by UV spectroscopy. Fifty nanograms of cDNA were subjected to a PCR reaction, using Qiagen OneStep RT-PCR kit, for the different mAChR subtypes. The purity of the isolated motor neurons was confirmed by the presence of choline acetyl-transferase (ChAT) and absence of glial fibrillary acidic protein (GFAP) mRNAs. The quality of NMJ isolates was assessed by nAChRε and S100β mRNAs. Isolated brain tissues were used as a positive control for PCR detection of mAChR subtypes. The following primer sequences were used to amplify a region of the third cytoplasmic loop that lacks homology among different subtypes. The specificity of mAChR subtype primers was further verified by sequencing PCR products extracted from gel, using a Gel Extraction Kit (Qiagen, Valencia, CA). M1 forward, TCTGCTCATCAGCTTTGAC CG; M1 reverse, CATCCTCTTCCTCTTCTT CTTT CC; M2 forward,TGTCAGCAATGCCTC C GTTATG; M2 reverse, GCCTTGCCATTCT GGAT CTTG; M3 forward, GGTGTGATGATTG GTCTGGCTTG; M3 reverse, AGAAGCAG AGTTT TCC AGGGAG; M4 forward, TCAAG AG CCCTCTGATGAAGCC; M4 reverse, AGA TTGTC CGA GTCACTTTGCG; M5 forward, GCT GACCTCCAAGGTTCCGATTC; M5 reverse, CCG TC AGCTTTTACCACCAAT; ChAT forward, GCCTGGTATGCCTGGATGGT C; ChAT reverse, TGGAGGGCCACCTGGATGAA G; GFAP forward, CCTCAAGAGGAACA TCGTGG T; GFAP reverse, ACACTGGAGTCATC ACCCTGGA; S100 forward, GCTGAAG AAGTCAG AACTGAAG; S100 reverse, TGATGTG CTAACTTAAAGCAGC; AChRε forward, GGCAG TTTGGAGTGGCCTACGACA; AChRε reverse, GCAGGACGTTGATAGA GACCGTGC.
Whole mounts of LAL or hindlimb muscles were processed for immunostaining as described previously (Wright and Son, 2007). Briefly, muscles were postfixed in 4% paraformaldehyde for 20 min., rinsed in phosphate-buffered saline (PBS) containing 0.1M glycine, and incubated for 15 minutes with rhodamine-conjugated α-bungarotoxin (Invitrogen, Carlsbad, CA). The muscles were then permeabilized in −20°C methanol for 5 min and blocked for 1 hour in PBS containing 0.2% Triton and 2% BSA. The muscles were subsequently incubated overnight at 4°C in a cocktail of primary antibodies diluted in the blocking solution. Axons and nerve terminals were labeled with mouse monoclonal antibodies to neurofilaments (SMI 312; Sternberger Monoclonals, Baltimore, MD) and to a synaptic vesicle protein, SV2 (Developmental Studies Hybridoma Bank, Iowa City, IA). Schwann cells were labeled with rabbit anti-cow S-100 polyclonal antibody (Dako, Carpentaria, CA). To label activated Schwann cells in partially denervated muscles, we used S100 antibody in combination with an antibody to p75 (Chemicon, Billerica, MA). After incubation with the primary antibodies, muscles were rinsed in PBS and incubated with secondary antibodies in the blocking solution, for 1 hour at room temperature. The secondary antibody for monoclonal antibodies was fluorescein-conjugated goat anti-mouse IgG1 (Roche Molecular Biochemicals, Indianapolis, IN) or Alexa-Fluor 568 conjugated goat anti-mouse IgG1 (Molecular Probes, Eugene, OR). The secondary antibody for polyclonal antibodies was Alexa-Fluor 647 or Alexa-Fluor 568 conjugated goat anti-rabbit (Molecular Probes, Eugene, OR). After incubation with the secondary antibodies, the muscles were rinsed in PBS, mounted in Vectashield (Vector Laboratories, Burlingame, CA) and stored at -20°C. For immunolabeling of mAChR subtypes, methanol permeabilization was omitted. Sources of mAChR antibodies were as follows: anti-M1, -M2, and -M3 rabbit polyclonal antibodies (Alomone Labs, Jerusalem, Israel); anti-M2 monoclonal and anti-M5 rabbit polyclonal antibodies (Abcam Inc. Cambridge, MA); anti-M4 polyclonal antibody (Santa Cruz Biotechnology Inc., Santa Cruz, CA.).
The extent of terminal sprouting was evaluated by the percentage of the junctions with terminal sprouts and by the number and length of the terminal sprouts extended extrasynaptically from each junction. Terminal sprouts were defined as those extended beyond the synaptic boundary delineated by postsynaptic nAChR labels, and the length of secondary and tertiary branches of primary sprouts was included in measuring sprout length. Activated tSCs were defined as those that extended cellular processes extrasynaptically with or without association with terminal sprouts. To evaluate synaptic stability, the spatial alignment of pre- and postsynaptic specializations, fluorescent intensity of postsynaptic nAChR clusters, and measurements of synaptic area (i.e., synapse size) from nAChR labels were determined by a blinded observer with interactive software (Olympus analySIS, Melville, NY).
To evaluate terminal sprouting and reinnervation after partial denervation, each junction was first categorized as: (1) normally innervated by its original axon, (2) completely denervated, (3) denervated but reinnervated by terminal sprout(s), or (4) denervated but reinnervated by nodal sprout(s). At early times after partial denervation (i.e., <2 weeks), several features, such as thinner diameter of preterminal axons or varicosities along the length of preterminal axons, distinguished endplates reinnervated by nodal or terminal sprouts from the endplates innervated by their original axons (i.e., those spared by the partial denervation). Junctions innervated by original axons were further examined both for tSC processes acting as bridges to a denervated end plate and for terminal sprouts growing on these bridges. The frequency of tSC bridges between innervated and denervated endplates was determined by counting the number of junctions linked by a bridge/number of all endplates examined.
The muscles were analyzed using an Olympus BX61 widefield fluorescence microscope equipped with an integrating, cooled CCD camera (ORCA-ER, Hamamatsu, Japan) connected to a PC running image analysis software (Olympus analySIS, Melville, NY). High-resolution confocal images were obtained with a Leica Plan Apo 63× oil objective (1.4NA) on a Leica TCS 4D confocal microscope (Heidelberg, Germany). Z stacks were obtained at 0.3 μm step size for 20-40μm depths and additional optical sections above and below each junction were collected to ensure that the entire synaptic profile was included. Leica TCS-NT acquisition software and Imaris image software (Bitplane AG, Zurich, Switzerland) were used to reconstruct z-series images into maximum intensity projections. Multipanel images presented in the figures were adjusted for brightness and contrast using Adobe Photoshop.
Unpaired Student's t tests were performed to determine significance in experimental versus control groups using StatView software (Abacus Concepts, Inc. Berkley, CA). Differences were considered statistically significant when p <0.05. All data are presented as means ± SEM.
To test the idea that muscarinic signaling regulates sprouting of motor nerve terminals, we blocked mAChRs pharmacologically in vivo and determined whether terminal sprouting was elicited at NMJs. We applied atropine, a non-specific blocker of all five known subtypes of mAChRs (Eglen, 2005; Alexander et al., 2008), to LAL muscles, a thin superficial muscle on the dorsum of the neck, of young adult mice. LAL is uniquely suitable for revealing in vivo pharmacological effects while avoiding the complications of continuous application (Lanuza et al., 2001; Wright and Son, 2007). When we applied atropine subcutaneously twice daily for 7 days (0.2-10mg/kg), nerve terminals at many NMJs developed terminal sprouts that extended beyond the parental synaptic area (e.g., arrows in Fig. 1A, B), and were more frequent with increasing dosage of atropine (Fig. 1C). Terminal sprouts were associated with tSC processes that often led the nerve sprouts (e.g., Fig. 1B; blue arrows) and highly expressed GFAP (Supplemental Fig. S1). These observations therefore supported the intriguing idea that signaling through mAChRs suppresses sprouting in active muscles and may mediate compensatory sprouting in paralyzed muscles.
Interestingly, however, terminal arbors at most NMJs exhibiting sprouting were incomplete, indicating that atropine-elicited sprouting accompanied partial withdrawal of parental terminal branches. Moreover, many other NMJs exhibiting no sprouting displayed additional phenotypic defects, such as postsynaptic loss of nAChRs or complete loss of terminal arbors accompanied by abnormally quiescent tSCs (Fig. 1B). These unanticipated effects virtually destroyed the structural integrity of almost all NMJs in the atropine-treated LAL muscles (n=185 junctions, n=3 LAL muscles treated twice daily with 0.2 mg/kg atropine). Specifically, in approximately 10% of the junctions (Fig. 1C), partially eliminated axon terminals extended sprouts associated with tSC processes but tSCs at the synapse and nAChRs appeared to be intact (e.g., junction 2 in Fig. 1A, B). In 60% of the junctions (Fig. 1D), however, nAChRs were only faintly labeled while their axon terminals and tSCs were largely unaffected (e.g., junction 3 in Fig. 1A, B). In another set of the junctions (Fig. 1E), approximately 6%, postsynaptic nAChRs were unaffected but axon terminals were almost completely retracted and, surprisingly, their tSCs remained quiescent despite absence of nerve terminals (e.g., junction 4 in Fig. 1B). This quiescence of tSCs was surprising because they are normally activated and extend processes when nerve contact is lost (Reynolds and Woolf, 1992; Son and Thompson, 1995b). Additionally, the size of muscle fibers treated with atropine was substantially reduced compared to the saline-treated control muscles (p<0.01; Fig. 1F).
We next examined whether the unexpectedly diverse and severe effects evoked by atropine were due simply to administration of toxic impurities or to inflammation (Verze et al., 1996; Reinert et al., 1998). For this purpose we tested different doses of pharmaceutical grade atropine (Fig. 1), and essentially replicated our observations with industrial-grade atropine (data not shown). We also induced intense inflammation in LAL muscles with subcutaneous Lipopolysaccharide (LPS; 50 μg) but observed none of the structural defects in NMJs that we had found with atropine (Supplemental Fig. S2, n=3 mice). Furthermore, we observed substantial effects even with a very low dose of atropine (0.2 mg/kg twice daily for 7 days; Fig. 1 C-F), which was the same or less than the doses used in most studies of muscarinic signaling (Molinengo et al., 1989; Kociolek et al., 2006). Lastly, when we applied atropine at the lowest dose (0.2mg/kg once-daily for only 5 days; n=6 mice), we found milder defects that were confined to the synaptic area, and both preterminal axons and myelinating Schwann cells appeared completely normal (Supplemental Fig. S3). These observations suggested that atropine acted locally on synapses. We also observed variability in the responses of the junctions; some junctions responded with terminal sprouting and tSC activation (e.g., Supplemental Fig. S3; white and blue arrows, respectively), whereas other junctions responded with selective loss of postsynaptic nAChRs but their nerve terminals and tSCs appeared normal (e.g., Supplemental Fig. S3; arrowhead).
The even- (M2/4) and odd-numbered (M1/3/5) mAChR subtypes differ in their G protein-coupling properties and multiple mAChR subtypes are present at the NMJ (Minic et al., 2002; Garcia et al., 2005). We therefore speculated that specific mAChR subtypes might mediate distinct defects evoked by atropine. To test this idea, we applied subtype-specific mAChR blockers at various doses to LAL muscles of adult mice twice daily for 7 days. Methoctramine, an M2/M4 mAChR-preferring antagonist (Dorje et al., 1991; Caulfield and Birdsall, 1998), evoked terminal sprouting and activation of tSCs in many NMJs (Fig. 2A, white and blue arrowheads, respectively; about 19% junctions of 177 junctions, 4 LAL muscles). No junctions, however, exhibited complete loss of terminal arbors with abnormally quiescent tSCs, postsynaptic loss of nAChRs or fiber atrophy (data not shown). Notably, however, approximately 70% of the junctions exhibiting terminal sprouting also exhibited partial but definite loss of terminal branches (Fig. 2A; e.g., nAChR clusters marked by a white arrow unoccupied by axon terminals). Thus, pharmacological inhibition of M2/4 mAChRs selectively evoked a subset of atropine-elicited defects- sprouting with partial terminal loss.
In contrast to methoctramine, 4-DAMP (4-diphenylacetoxy-N-methylpiperidine) shows low affinity for M2 mAChRs and high affinity for the remaining four mAChR subtypes (Dorje et al., 1991; Caulfield and Birdsall, 1998). We found that 4-DAMP induces complete withdrawal of terminal arbors that strikingly resembles the ‘dying-back’ of nerve terminals observed in degenerative diseases such as amyotrophic lateral sclerosis (Fischer et al., 2004; Schaefer et al., 2005). Following twice-daily application for 7 days, most NMJs partially or completely lost their terminal arbors (Fig. 2B; 86.2 ± 3.5% of 152 junctions, n=3 mice). The withdrawal of axon terminals appeared to be initiated distally as evidenced by 1) no signs of preterminal axon degeneration preceding terminal withdrawal, and 2) retracting axons at the vacant endplates (Fig. 2B; arrowheads) or along the preterminal Schwann cells (white arrows). Postsynaptically, we found no nAChR loss or fiber atrophy. The responses of perisynaptic tSCs were surprising, however, because they remained inactive (i.e., no formation of extrasynaptic processes) despite their loss of axonal contacts. Additional evidence for lack of tSC activation is that 4-DAMP-treated tSCs did not down-regulate S100, although down-regulation is characteristic of activated tSCs and their processes (Fig. 2B, blue arrows indicating tSCs with bright immunofluorescence of S100). Thus, like methoctramine, 4-DAMP induced only a subset of the phenotypes that we observed with atropine. Importantly, however, unlike methoctramine, which elicited sprouting with partial terminal loss, 4-DAMP evoked complete withdrawal of terminal arbors associated with abnormally quiescent tSCs.
The selectivity of most ‘subtype-selective’ muscarinic antagonists is rather limited (Dorje et al., 1991; Caulfield and Birdsall, 1998). We therefore carried out additional experiments using mAChR knockout mice as novel experimental tools. Specifically, we examined NMJs of mutant mice deficient in one or two mAChR subtypes: M1-/-, M2-/-, M3-/-, M4-/-, M5-/-, M1/3 -/- dKO and M2/4-/- dKO. We observed spontaneous sprouting in the NMJs of adult mice lacking M2 mAChRs, but not in the other lines. Many of the nerve terminals in M2 -/- NMJs completely lost the normal, compact branching patterns (Fig. 3A, 34.6 ± 3.1 % of 185 junctions, n=3 LAL muscles) and exhibited unusually long and dispersed terminal branches that resembled terminal sprouts (e.g., short arrows in Fig. 3A, B, Fig. 5). These terminal branches/sprouts also formed varicosities directly apposed to small aggregates of nAChRs (arrowheads), a characteristic feature of terminal sprouts (Yee and Pestronk, 1987; Rogozhin et al., 2008).
M2-/- NMJs with sprouts usually displayed faintly labeled, ‘vacant’ branches of postsynaptic nAChRs that were completely unoccupied by terminal branches (e.g., Fig. 3B, asterisk; see additional examples in Supplemental Fig. S4). This feature resembled the partial terminal loss that we observed in methoctramine-treated NMJs (Fig. 2A), and indicated that nerve terminals in M2-/- NMJs extended sprouts that form new synaptic contacts, while losing synaptic adhesion of parental terminal branches. We were also surprised to find that approximately 23% of M2-/- NMJs without obvious sprouts exhibited ‘vacant’ branches of nAChRs and tSCs that were unoccupied by terminal branches (Fig. 3B, bottom panel; n=185 junctions, 3 M2-/- LAL muscles). This finding shows that withdrawal of terminal branches is likely to precede the loss of tSC processes and postsynaptic nAChRs and, most importantly, the onset of terminal sprouting. The data also suggest that spontaneous sprouting at M2-/- NMJs is an indirect consequence of terminal instability, and that terminal arbors become unstable or unusually mobile in the absence of M2 mAChRs.
To define further the characteristics of the sprouting evoked by the absence of muscarinic signaling, we compared spontaneous sprouting in M2 -/- LAL muscles with sprouting induced in LAL muscles paralyzed for 7 days by botulinum toxin (BoTX, n= 3 muscles). Many of the BoTX-paralyzed nerve terminals extended abundant, lengthy terminal sprouts, while maintaining their original terminal arbors (Fig. 4, asterisk in bottom panel), whereas sprouts (arrows) at M2-/- NMJs partially or completely lost their original terminal arbor and displayed ‘parental’ clusters of nAChRs unoccupied by axons (Fig. 4, asterisk in the upper panel; see additional examples of such M2-/- NMJs in Supplemental Fig. S4). This observation indicates that the sprouting observed in M2-/- NMJs is qualitatively different from compensatory sprouting, and further supports our interpretation of the phenotypic defects at M2-/- NMJs: nerve terminals become unstable or mobile in the absence of M2 mAChRs, causing repeated denervation and reinnervation of synaptic contacts.
We next assessed synaptic transmission at M2-/- NMJs by recording endplate current (EPC) and miniature endplate current (MEPC) in LAL muscles of M2+/+ and -/- mice (n=4 and 7 mice, respectively). Approximately 17 muscle fibers in each LAL muscle were impaled with 2 electrodes and voltage clamped; the nerve was stimulated as described previously (Wang et al., 2004). In 5 of the M2-/- mice, there was no change in EPC amplitude (Fig. 5; 106.8 ± 6.3 nA vs. 103.4 ± 4.9 nA, M2+/+ and -/-, respectively, p = 0.68), MEPC (2.35 ± 0.05 nA vs. 2.28 ± 0.06 nA, M2+/+ and -/-, respectively, p = 0.33) or in quantal content (46.5 ± 2.8 vs. 46.2 ± 1.7, M2+/+ and -/-, respectively, p = 0.94). The EPC time to peak was slightly reduced (0.49 ± 0.01 ms vs. 0.45 ± 0.01 ms, M2+/+ and -/-, respectively, p < 0.05). Notably, however, in 2 of the M2-/- mice, EPCs were smaller than normal (Fig. 5A; n=25 muscle fibers, 2 -/- LAL muscles) and had markedly prolonged time constants of decay (Fig. 5B). The prolongation of EPCs was accompanied by a milder prolongation of MEPC time constant of decay (Fig. 5C; 1.30 and 1.29 ms in the affected M2-/- muscles vs. 0.69 ± 0.02 ms in the other 9 muscles).
Prolongation of EPC and MEPC has been observed at NMJs undergoing early stages of reinnervation after nerve crush (Argentieri et al., 1992), and attributed to re-expression of fetal-type nAChRs which have a longer mean open time than adult-type nAChRs (Wang et al., 2006). It is also known that fetal-type nAChRs are re-expressed in the extra-junctional area of reinnervated or paralyzed adult muscles innervated by terminal sprouts, whereas adult nAChRs continue to be expressed at the original synaptic area (cf., Kues et al., 1995). Subsets of M2-/- NMJs therefore appear to be functionally abnormal due to severe denervation (i.e., marked loss of original terminal arbors enriched with adult-type nAChRs) and reinnervation (i.e., formation of new synaptic contacts enriched with fetal-type nAChRs), consistent with our morphological analysis of M2-/- NMJs.
Unlike M2-/- NMJs, M4-/- NMJs were normal (Supplemental Fig. S5). However, muscles of young M2/4 double KO (dKO) mice included many abnormal NMJs with striking features indicative of terminal instability, similar to M2-/- NMJs (Fig. 6; 65.7 ± 3.2 % of 312 junctions, 3 LAL muscles of 3 week old M2/4 -/-mice). First, portions of nAChR clusters unoccupied by axon terminals were common (Fig. 6A; asterisk and inset), and their axon terminals appeared to have slightly shifted (Fig. 6A; arrows). Second, some terminal arbors appeared to have completely shifted and formed new nAChR clusters next to the previous synaptic site, as indicated by the adjacent, abandoned nAChR clusters whose morphology closely resembled that of ‘shifted’ terminal arbors (e.g., #1 junction in Fig. 6A, B). Third, some terminal arbors appeared to have shifted recently because, although they were not associated with nAChR clusters, abandoned nAChR clusters were usually observed nearby (e.g., #2 junction in Fig. 6A, B).
More mature muscles of M2/4 dKO mice (n=183 junctions, 2 LAL muscles of 3 months old M2/M4 -/-mice) revealed neither abandoned nAChRs nor terminal shifting. Notably, however, approximately 40% of M2/4-/- NMJs were extensively fragmented, similar to aged NMJs (Fig. 6C, D; cf., Prakash and Sieck, 1998). These observations further support the idea that M2 or the even-numbered muscarinic receptors are involved in maintaining the stability of terminal arbors.
We found no obvious defects in the NMJs of the M1-/-, M3-/-, M4-/-, or M1/3 dKO mice (Supplemental Fig. S5). However, M5-/- mice exhibited NMJ defects that differed in several ways from those of M2-/- NMJs. First, although varying among muscles (Fig. 7C; cf., SOL muscles), synaptic size in M5-/- muscles was smaller than that of age and sex matched wildtype mice [p<0.01, e.g., 410.44 ± 10.9 μm2 (n=890, 6 -/- LAL muscles) vs. 657.50 ± 12.3 μm2 (n=931, n= 7 +/+ LAL muscles)], and some NMJs were unusually small (e.g., junctions marked by arrows in Fig. 7A; see inset for a NMJ formed by a single terminal bouton). Second, postsynaptic clusters of nAChRs, at some of the small NMJs (approximately 15% of 345 junctions, n=3 LAL muscles) were faintly labeled and fragmented, indicating postsynaptic disassembly (Fig. 7B; arrowheads). Third, the diameter of muscle fibers was considerably reduced compared to that of wildtype mice (Supplemental Fig. S6; 21.98 ± 0.6 vs. 29.98 ± 1.0 μm, n=3 M5-/- and +/+ LAL muscles, respectively, p < 0.001).
Further analysis of M5-/- muscles showed that synaptic size was highly correlated with the size of parental muscle fibers (Fig. 7C; correlation coefficient, >0.75; n> 200 NMJs, 3 muscles each for LAL, STM, EDL, SOL muscles). Because NMJ size is proportionally regulated by muscle fiber growth (Balice-Gordon et al., 1990; Balice-Gordon and Lichtman, 1990), this finding suggests that NMJs are small in M5-/- muscles because the growth of muscle fibers is limited. In summary, our data indicates that nerve terminals are unstable in the absence of M2 receptors and that muscle fibers are defective in the absence of M5 receptors. These results strengthen the pharmacological evidence that specific subtypes of mAChRs play distinctive roles at NMJs.
To test directly the role of mAChRs in the induction of compensatory sprouting, we next asked whether nerve terminals and tSCs lacking mAChRs react to paralysis. We assessed nerve terminals and tSCs in the LAL muscles of M1/3-/-, M2/4-/-, and M5-/- mice paralyzed by botulinum toxin for 7 days. Terminal sprouting occurred at the NMJs of all these knockout mice (Fig. 8A) and we observed no significant difference from similarly treated wildtype LAL muscles in the number of NMJs exhibiting sprouts (Fig. 8B) or in the number of the primary branches of terminal sprouts extended at the junctions exhibiting sprouting (Fig. 8C). Moreover, as in wildtype LAL muscles, tSCs in the mutant NMJs reacted to paralysis by extending numerous processes that often led terminal sprouts (data not shown). Thus, paralysis can stimulate terminal sprouting and activate tSCs in the absence of M2 or other mAChRs.
Although the number of primary sprouts extended at M5-/- NMJs was not significantly different from M5+/+ NMJs (Fig. 8C), terminal sprouts elongated far more extensively on the M5-/- LAL muscle surface (168.1 ± 13.8 μm vs. 83.0 ± 1.6 μm, n=3 M5 -/- and +/+ muscles, respectively, p < 0.0001). The primary branches of terminal sprouts at M5-/- NMJs often formed secondary and tertiary branches (Fig. 8A), which elongated extensively (Fig. 8D), and the total length of terminal sprouts was much longer in M5-/- muscles than in M5+/+, M1/3-/-, and M2/4-/- muscles (Fig. 8E). Thus, paralysis initiated sprouting normally, but the extension of the sprouts was abnormally enhanced in the paralyzed M5-/- muscles.
When the motor innervation of a muscle is partially denervated, spared motor neurons sprout at nodes of Ranvier and nerve terminals and subsequently reinnervate the denervated muscle fibers (Hoffman, 1950; Brown and Ironton, 1978). Preterminal- and terminal Schwann cells serve as a growth substrate that guides the nodal sprouts to the original endplates of denervated muscle fibers (Son and Thompson, 1995b; Koirala et al., 2000). In addition, tSC processes extending from denervated endplates contact intact nerve terminals and form ‘tSC bridges’ that induce and guide terminal sprouts to denervated endplates (Son and Thompson, 1995a; Love et al., 2003). To test whether muscarinic signaling mediates compensatory sprouting and reinnervation, we transected L4 spinal roots in M1/M3 -/-, M2/M4 -/-, and M5 -/- mice and, 14 days later, analyzed partially denervated EDL muscles (Albani et al., 1988; Tam et al., 2001). As in similarly denervated wildtype EDL muscles, we observed significant sprouting of intact axons and nerve terminals in partially denervated muscles of all mAChR KO mice (Table 1; i.e., L5 axons spared by L4 root transection). In addition, these terminal sprouts frequently reinnervated adjacent denervated endplates, as in wildtype mice, and appeared to be guided by tSC bridges (Table 1, Supplemental Fig. S7). There was no significant difference between the mAChR +/+ and -/- muscles in reinnervation of denervated muscle fibers by terminal or nodal sprouts, or in the ability of tSCs to form ‘tSC bridges’ (Table 1). Taken together, these data show that partial denervation induces both nodal and terminal sprouting independent of mAChR activity, and that reinnervation by the compensatory sprouting proceeds normally with the assistance of tSCs in mAChR -/- mice.
The distinct NMJ phenotypes observed in pharmacological and genetic studies suggest that specific mAChR subtypes are selectively associated with different types of cells at NMJs. To test this possibility and to identify mAChR subtypes expressed at NMJs, we screened numerous commercially available ‘subtype-specific’ mAChR antibodies and observed brightly labeled cells at NMJs with many of the antibodies (e.g., Fig. 9A). We failed, however, to confirm their specificity in subtype-specific mAChR KO mice (data not shown). Most of the antibodies seemed to detect more than one mAChR subtype (cf., Jositsch et al., 2008; Pradidarcheep et al., 2008). One of these antibodies, which brightly labels nerve terminals, tSCs, and postsynaptic membranes in wildtype muscles, did not label terminal branches in M2-/- NMJs (arrow in Fig. 9A, see inset for selective disappearance of terminal branch- associated labels in M2-/-). Consistent with this observation, our analysis of mRNA expression in the cell bodies of spinal motor neurons and NMJs prepared with laser-assisted microdissection (Fig. 9B), demonstrated that motor neurons express M2 receptors and no other subtypes of mAChRs (Fig. 9C). In contrast, we observed only odd-numbered receptors (M1, M3, M5) in the microdissected NMJ preparations. These data show that motor neurons express M2 mAChRs, whereas Schwann cells and/or muscle fibers express multiple odd-numbered mAChRs. These findings also agree with the results of our studies with mAChR blockers and mAChR KO mice.
In the present study, we have analyzed the structural consequences of pharmacological inhibition and genetic deletion of all or a subset of mAChRs at mouse NMJs. These studies lead to the conclusion that muscarinic signaling does not mediate compensatory sprouting but that motor nerve terminals are unstable without M2 receptors, and that muscle fiber growth is defective in the absence of M5 receptors. These findings are the first to show that mAChRs not only modulate synaptic function but also influence synaptic structure, and that specific mAChR subtypes play distinct roles at NMJs.
The idea that mAChRs on tSCs mediate paralysis- or denervation-induced terminal sprouting was attractive because tSCs monitor synaptic activity via mAChRs (Jahromi et al., 1992; Reist and Smith, 1992) and because paralysis or mAChR blockade upregulates GFAP in tSCs (Georgiou et al., 1999). Consistent with this view, we observed tSC activation and terminal sprouting following non-selective (i.e., atropine) and selective (methoctramine and M2-/- KO) inhibition of mAChRs. We also found, however, that spontaneous sprouting at M2-/- NMJs was accompanied by the loss of terminal arbors, that M2-/- NMJs lost terminal branches even without sprouting and that reactive sprouting in response to paralysis and denervation proceeded normally without M2. Our finding that M2 mAChRs are located exclusively in motor neurons, and not in tSCs, further supports our conclusion that M2 mAChRs do not initiate compensatory sprouting.
Compensatory sprouting was also inducible in M1/3-/- dKO and M5-/- NMJs, making it unlikely that mAChRs other than M2 are involved. We observed that M5-/- nerve terminals extend much longer sprouts in response to paralysis than those in wildtype or other KO mice but that the number of primary terminal sprouts was not significantly different. The lack of M5 receptors might therefore promote sprout ‘elongation’ rather than ‘initiation’ by altering the muscle surface to promote growth, after sprouting has been initiated by non-muscarinic mechanisms. Our finding that the primary defects associated with the lack of M5 mAChRs were in muscle fibers rather than in nerve terminals or tSCs further supports this interpretation.
Surprisingly, atropine not only evoked terminal sprouting, as anticipated, but even in small doses destroyed the structural integrity of many NMJs. These dramatic effects were unexpected because atropine is commonly used in the clinic (Das, 1989; Kociolek et al., 2006). The effects are unlikely to be indirect or due to overdosage. First, we observed alterations of NMJ structure even at our smallest dose (0.2 mg/kg, once-daily for 5days), but not with considerably higher doses of other mAChR blockers, including muscarinic toxin-7 and AFDX-384. Second, pharmacological inhibition and genetic deletion of specific mAChRs elicited only subsets of the effects evoked by atropine. Third, we found that at least four mAChR subtypes are expressed by the cells at the NMJ. Thus, the dramatic changes caused by atropine are most likely due to the complete lack of muscarinic signaling at NMJs. These atropine-induced changes in NMJ structure have not been appreciated before and raise the worrisome possibility that atropine could have similar effects in patients when applied locally and repeatedly.
M1-M5 mAChR knockout mice are viable and fertile, but display distinct phenotypic changes (reviewed in Wess, 2004; Wess et al., 2007). For example, M1-/- mice exhibited a profound increase in locomotor activity, and M2-/- mice displayed impaired regulation of heart rate and deficits in working memory. M3-/- mice were hypophagic and lean, associated with a reduction in food intake. M4-/- mice showed enhanced central dopaminergic transmission, and M5-/- mice displayed reduced cerebral blood flow, associated with deficits in hippocampal LTP. The present study represents the first systematic analysis of muscarinic signaling at neuromuscular synapses using M1-M5 mAChR knockout mice. We found that M2 mAChRs are solely expressed in motor neurons and that motor neurons express no other mAChR receptor subtypes. Pharmacological and genetic inhibition of M2 mAChRs caused exclusively presynaptic defects: terminal sprouting and partial terminal withdrawal. Terminal sprouting did not seem to be the causative defect because partial loss of terminal branches was common even in the M2-/- NMJs that exhibited no sprouting. In addition, NMJs exhibiting sprouting and terminal loss displayed abandoned nAChR clusters in both the original and adjacent synapse areas, indicating that nerve terminals lacking M2 receptors are abnormally mobile and repeatedly form transient synapses. Furthermore, NMJs of adult M2/4 -/- dKO mice were markedly fragmented, and terminal arbors of developing M2/4-/- NMJs appeared to have shifted and left portions of nAChR clusters unoccupied. Whereas terminal fragmentation and shifting indicated the lack of stable adhesion between nerve terminal and muscle membrane, we observed no terminal sprouting in M2/4-/- dKO NMJs, in contrast to M2-/- NMJs. This observation therefore further supports our interpretation of M2-/- phenotypes: the lack of stable adhesion of nerve terminals rather than terminal sprouting is the primary defect in M2-/- NMJs.
Our interpretation that M2 receptor signaling contributes to stable adhesion of terminal arbors to the postsynaptic muscle membrane is consistent with the role attributed to muscarinic signaling in integrin-mediated cell-cell adhesion and migration of non-neuronal cell types (Quigley et al., 1998; Shafer et al., 1999; Nguyen et al., 2003). The integrins are major receptors for the laminins and collagens important for stable synaptic adhesion between the nerve terminal and muscle membrane at NMJs (Martin et al., 1996; Patton et al., 1998; Son et al., 2000). M2 mAChRs may also influence synaptic stability via cAMP-regulated protein kinases implicated in terminal withdrawal (Renger et al., 2000; Lanuza et al., 2001). More recent data have also shown that nAChR clustering can be induced by laminin acting via Rho GTPases (Weston et al., 2007), a downstream pathway for muscarinic signaling (Linseman et al., 2000). It is therefore tempting to speculate that signaling through the mAChR-laminin-Rho GTPase-nAChR pathway may be responsible for nAChR declustering where terminal loss was evident in M2-/- NMJs.
The notion that M2 receptors may inhibit evoked ACh release was based on earlier pharmacological studies reporting reduced evoked ACh release following application of muscarine (a non-subtype-selective agonist) and increased evoked ACh release with methoctramine (an M2/M4 receptor-preferring antagonist; reviewed in Re, 1999). However, atropine (a non-subtype-selective antagonist) had no effect on ACh release (Minic et al., 2002). A recent study showed that the inhibitory effect of muscarine and the stimulatory effect of methoctramine on ACh release are abolished in M2-/- mice, but that the amount of ACh released at M2-/- NMJs is virtually normal at physiological calcium levels (Slutsky et al., 2003). This observation is therefore consistent with our electrophysiological analysis of M2-/- NMJs, which revealed normal transmission at most M2-/- NMJs (Fig. 5). However, we also observed subsets of M2-/- NMJs with marked prolongation of EPCs, a characteristic of NMJs undergoing denervation and reinnervation (Argentieri et al., 1992).
Prolongation of EPCs is known to be caused by fetal-type nAChRs, which have a longer opening time than adult-type nAChRs (Wang et al., 2006). It is unlikely, however, that M2 receptors regulate fetal- or adult-type nAChR expression, because many M2-/- NMJs were functionally normal and displayed no prolongation of EPCs (Fig. 5). Instead, our interpretation is that the M2-/- NMJs with marked prolongation of EPCs are those in which adhesion of axon terminals to the postsynaptic membrane was particularly unstable, which lost their original terminal branches and parental nAChRs (i.e., adult-type), and relied on synaptic transmission mostly through the new fetal-type nAChRs that were clustered by spontaneous terminal sprouts. This interpretation receives support from the observation that regenerating axons and terminal sprouts induce clusters of fetal-type, rather than adult-type, nAChRs, in the extrajunctional area of paralyzed or reinnervated muscle fibers (cf., Kues et al., 1995). Our morphological analysis also showed a differing extent of terminal loss or instability among M2-/-NMJs.
We found that M5-/- NMJs display phenotypic defects distinct from those of M2-/- NMJs: small NMJs and small muscle fibers. The size of M5-/- NMJs was highly correlated with the size of parental muscle fibers. Because synaptic size is proportionally regulated by muscle fiber size (Balice-Gordon et al., 1990; Balice-Gordon and Lichtman, 1990), this result indicates that the primary function of M5 receptor signaling is to regulate muscle fiber growth. In agreement with this interpretation, several lines of evidence suggest that skeletal muscle fibers express both mAChRs and nAChRs (Reyes and Jaimovich, 1996; Welsh and Segal, 1997; Liu et al., 2002; Jordan et al., 2003; Furlan and Godinho, 2005), and that mAChRs mediate protective effects of cholinergic drugs on denervated muscles (Urazaev et al., 1997, 2000). Moreover, downstream effectors of mAChRs such as MAPKs (mitogen activated protein kinases), ubiquitin-proteasome pathways, GEF (guanine nucleotide exchange factor), and Rho GTPases have been strongly implicated in muscle growth and atrophy (Tawa et al., 1997; Pawson et al., 2008; Wang et al., 2007; Lamon et al., 2009; Shi et al., 2009). Therefore an exciting implication of the present results, in combination with those of earlier studies, is that mAChRs, predominantly M5 receptors, present in muscle fibers may mediate activity-dependent growth and atrophy of NMJs and muscle fibers.
Previous studies have reported expression of some or all five mAChR receptors at NMJs (Minic et al., 2002; Garcia et al., 2005). The lack of mAChR subtype specificity of the reagents used in the pharmacological and histological studies, however, has limited their implications (Jositsch et al., 2008; Pradidarcheep et al., 2008). We found that, in contrast with prior reports, motor neurons express only M2 receptors, whereas tSCs and/or muscle fibers express odd-numbered mAChRs (i.e., M1, 3, 5). Because most tissues and cell types typically express two or more mAChR subtypes (Caulfield and Birdsall, 1998; Abrams et al., 2006), and because pure cultures of Schwann cells and muscle cells express all three odd-numbered receptors (Wright et al., unpublished observation), both tSCs and muscle fibers are likely to express multiple odd-numbered mAChRs. Future studies using tissue-specific double or triple KO mice lacking odd-numbered mAChRs will address this issue and determine whether simultaneous inhibition of multiple odd-numbered receptors evokes more striking defects at NMJs than we observed after selective inhibition of M1, M3 or M5 receptors.
Like synapses elsewhere in the nervous system the organization of the NMJ is ‘tripartite’: they are composed of presynaptic nerve terminals, postsynaptic muscle cells and perisynaptic Schwann cells. Each of the three cell types detects neural activity and changes dramatically in response to synaptic dysfunction (Pun et al., 2002; Midrio, 2006; Burns et al., 2007; Feng and Ko, 2007; Santafe et al., 2007; Kong et al., 2009). Little is known, however, about how each cell type coordinates synaptic activity with growth and maintenance of functional synapses, or how the structure of the synapse disintegrates under non-functional or pathological conditions. Synaptic activity with concomitant ACh release can signal through metabotropic mAChRs, which are broadly expressed among the three cell types at the NMJ. It is tempting to speculate that mAChRs provide a fundamental pathway that orchestrates the activity-dependant growth or maintenance of the tripartite synapse. Definitive cellular localization of mAChR subtypes and generation of tissue- or cell-specific mutant mice lacking different combinations of receptor subtypes will be necessary to fully define their roles at synapses.
We thank John Houle and Benjamin Keeler for help with laser microdissection and PCR analysis. We also thank Srishti Bhagat for technical assistance, and Tim Himes and the members of the Son lab for their comments on the manuscript. This work was supported by NS045091, NS062320 (Y-J.S.), NS057228 (M.R.), VA (A.T.), and NIH intramural funds (J.W.).